We thank the NCFPB Integrated Core Facility for Functional Genomics (Taipei, Taiwan) and the NRPB Pharmacogenomics Lab (Taipei, Taiwan) for their technical support. This work was supported by research grants from the Taiwan Health Foundation (L-I.L.) and the Ministry of Science and Technology, Taipei, Taiwan, ROC [MOST 105-2320-B-002-047] for Woei-horng Fang, [MOST 105-2628-B-002 -051-MY3] for Kang-Yi Su, and [MOST-105-2320-B-002-051-MY3] for Liang-In Lin.

1. Primer/template Preparation
<ol>
	<li>Design primers/templates with a balanced G+C content between 40% and 60% as in a sequencing or PCR primer design. Use primers of 18 to 21 nt for an appropriate annealing and better MS signals.</li>
	<li>Design the template by setting 50 &#176;C as the minimum melting temperature for the duplex region with at least 7 nt of 5&#39;-overhang to separate the signals between the primer and the template. 
	NOTE: For example, for the substrate P21/T28 in Table 1, the 21-nt primer is paired with a 28-nt template. Option: the use of alternative nucleic acids such as nuclease-resistant phosphorothioate bonds to replace the last 4 phosphodiester bonds at the 3&#39; end of the template can prevent possible interference by a non-specific 3&#39; end degradation of the templates (e.g., T28S4 in Table 1), although this will add some cost for the template preparation.</li>
	<li>Using standard desalted oligonucleotides without any further purification is satisfactory for this study. Use oligonucleotide primers/templates at a concentration of 100 pmol/&#181;L in H2O as a stock and store it at -20 &#176;C. Check the quality and purity of the primers and templates by running the oligonucleotides on MALDI-TOF MS (steps 4 - 7) to ensure unique peak signals and a good signal to noise ratio.</li>
	<li>Determine the relative MALDI-TOF MS signal intensities (step 7) of the equal molar primers of the relevant sequences in the reaction for calibration and concentration normalization (Figure 2).</li>
</ol>
2. Proofreading Reactions
Note: The same proofreading reaction condition and protocol applies to a DNA repair of A-I, G-I, C-I, and T-I.
<ol>
	<li>Using a T-G mismatch of substrate P21/T28 for the proofreading in Table 1 as an example, dilute primer P21 and template T28 to 12.5 pmol/&#181;L with H2O; then transfer 12 &#181;L of the diluted primer and 12 &#181;L of the diluted template to a 1.5-mL sterilized microcentrifuge tube. 
	NOTE: The amount of template/primer mix is sufficient for 2 reactions; proportionately increase the volume of the reagents accordingly for multiple reactions.</li>
	<li>Close the tube tightly. Incubate it for 30 min in a covered 65-&#176;C water bath, followed by 30 min in a 37-&#176;C water bath, and finally on ice to ensure a proper annealing.</li>
	<li>To a 1.5-mL sterilized microcentrifuge tube, add 8 &#181;L of primer and template mix (50 pmol), 2 &#181;L of 10x proofreading reaction buffer, 4 &#181;L of 4 ddNTPs mix (2 mM of each 4 dNTPs), and H2O up to 18 &#181;L. Flick the tube to mix. 
	NOTE: 1x proofreading reaction buffer contains 50 mM NaCl, 10 mM MgCl2, 1 mM dithiothreitol, and 10 mM Tris-HCl (pH 7.9 at 25 &#176;C). See the Table of Materials for the commercial source of 10x proofreading reaction buffer. The final concentration of each ddNTP in the reaction is 0.1 mM.</li>
	<li>Dilute DNA polymerase in ice-cold 1x proofreading reaction buffer to the desired concentration [e.g., to a 1.5-mL microcentrifuge tube, add 4 &#181;L of 1x proofreading reaction buffer and 1 &#181;L of 5-U Klenow polymerase from a commercial source (Table of Materials)]. Store the diluted enzyme on ice at all times. 
	NOTE: The amount of diluted DNA polymerase is sufficient for 2 reactions; proportionately increase the volume of the enzymes accordingly for multiple reactions. A volume of 2.0 &#181;L, containing 2.0 units of Klenow polymerase, can proofread more than half of 50 pmol of T-G terminal mismatched P21/T28 in 5 min.</li>
	<li>Prewarm the microcentrifuge tube with the substrate mix from step 2.3 to the desired reaction temperature (e.g., typically 37 &#176;C for Klenow polymerase and 25 &#176;C for T4 DNA polymerase). Then transfer 2.0 &#181;L of the diluted DNA polymerase from step 2.4 to the prewarmed substrate mixture and flick the tube to mix its contents.</li>
	<li>Centrifuge the tubes with enzyme and substrate for a few seconds at 3,200 x g at ambient temperature to spin down the components of the reactions. Immediately transfer the reaction to a heating block or a water bath at the desired incubation temperature as in step 2.5.</li>
	<li>Terminate the reaction
	<ol>
		<li>Add an equal volume of buffered phenol, mix it by vortexing for a few seconds, and centrifuge it in a microcentrifuge at 3,200 x g at ambient temperature for 5 min.
		<ol>
			<li>Transfer the aqueous phase to a clean microcentrifuge tube and add an equal volume of chloroform, mix it by vortexing for a few seconds and centrifuge it in a microfuge at 3,200 x g at ambient temperature for 3 min. Transfer the aqueous phase to a clean microfuge tube and incubate it at 95 &#176;C for 5 min; then place it on ice. 
			CAUTION: Phenol and chloroform are hazardous chemicals. Handling and waste disposal should follow institutional regulations.</li>
		</ol>
		</li>
		<li>Alternatively, use an acid quenching protocol. For this, add 2 &#181;L of 1 M HCl to stop the reaction on ice for 6 min, then add 0.64 &#181;L of 1 M diethanolamine (DEA) to neutralize the pH of the reaction mixture and incubate it at 95 &#176;C for 5 min; then place it on ice.</li>
	</ol>
	</li>
</ol>
3. Resin Addition to Eliminate Salt Contamination
<ol>
	<li>Using the resin scoop from the commercial desalting kit (see the Table of Materials), take sufficient resin to cover the dimples of the 384-dimple, 6-mg resin dimple plate.</li>
	<li>Distribute the resin across all the dimples evenly (6 mg/dimple) by scraping across the top of the plate. Recover any excess resin and replace it in the bottle.</li>
	<li>Transfer the final reaction products from step 2.7 from the microfuge tubes to a 384-well plate. Dispense 16 &#181;L of H2O to dilute the final reaction products in each well and then seal the plate and centrifuge it to remove any bubbles.</li>
	<li>Remove the seal, turn the 384-well plate with the reaction products upside down to cover the resin-filled dimple plate.</li>
	<li>Turn over the well-to-dimple plates sandwich and carefully let the resin drop into the 384-well plate (be sure the 384-well plate is flush against the metal peg on the resin-filled dimple plate).</li>
	<li>Remove the dimple plate on top, seal the 384-well plate containing the mixtures of the reaction products and resin, briefly centrifuge it at 3,200 x g to remove any bubbles and place it on a rotator for at least 15 min at room temperature for desalting its contents.</li>
	<li>After the resin cleanup, centrifuge the 384-well plate at 3,200 x g for 5 min to compact the resin to the bottom of the wells.</li>
</ol>
4. Transfer Reaction Products to a Matrix Chip
<ol>
	<li>Set the sample plates in the nanoliter dispenser (nanodispenser, see the Table of Materials).</li>
	<li>Put the matrix chip (see the Table of Materials) and the 384-well plate from section 3.7 in the corresponding tray.</li>
	<li>Operate the nanodispenser to spot the reaction products to the chip; push the RUN button on the touch screen of the nanodispenser to start.</li>
	<li>Check the automated captured image of the sample spots containing saturation information on the screen to ensure the overall spotted volume on the chip is around 5 - 10 nL. Repeat the sample spotting if the volume is insufficient.</li>
</ol>
5. Setup the Assay Information on the Mass Spectrometry
<ol>
	<li>Prepare a file in .xls format containing the anticipated signal information for importing. For example, use the setting of &#34;P21_T28.xls&#34; (Table 2) for the proofreading reaction of P21/T28 in steps 2, 3, and 4.</li>
	<li>Create and define a new assay in the application program (see the Table of Materials) by right-clicking Import Assay Group in Designer Format and select the .xls file (e.g., &#34;P21_T28.xls&#34; from step 5.1).</li>
	<li>Establish the target assay plate via the Customer:Project:Plate option tree on the left side of the screen by right-clicking the top of the tree and giving it a file name (e.g., &#34;CTT20171201&#34; represents the assay code and date).</li>
	<li>Select the 384-well plate type and press OK; a blank plate will appear.</li>
	<li>Select the appropriate assay (e.g., P21_T28) on the left side of the screen.</li>
	<li>Assign the selected assay (e.g., P21_T28) for each sample spotted sample position on the chip by highlighting the well and right-clicking to select Add Plex.</li>
	<li>Prepare a working list of all the tests on the chip in .xlsx format with no header (e.g., 1201.xlsx of Table 3). Import the working list via the Add New Sample Project option.</li>
	<li>Assign the test in the working list (e.g., from 1201.xlsx) to each position of the P21_T28 assay and choose by right-clicking.</li>
</ol>
6. MALDI-TOF MS Operation
<ol>
	<li>Link the mass spectrometer to the chip using the application program (see the Table of Materials).</li>
	<li>Select the default setting at the right side of the screen.</li>
	<li>Fill in the assay name from step 5.3 (e.g., CTT20171201), chip ID at the corner of the chip, and save the settings.</li>
	<li>Start the MS control program (see the Table of Materials).</li>
	<li>Press the In/Out button and take out the scout plate. Place the spotted chip from step 4.4 onto the scout plate and press the In/Out button for the spotted chip to enter the mass spectrometer.</li>
	<li>Click the Acquire button of the application program (see the Table of Materials) to start the mass spectrometry and acquire data.</li>
</ol>
7. Data Analysis
<ol>
	<li>Open the program for the data analysis (see Table of Materials).</li>
	<li>Browse the tree of the database at the upper left corner and select the chip ID from step 6.3. Open the data file on the right side of the screen.</li>
	<li>Click the spectrum icon to display the mass spectrum. To crop a specific range of m/z, right-click to select Customization Dialog and in the new window click on X-Axis and set the desired upper and lower limit and press OK to show the specified range of the spectrum.</li>
	<li>Export the spectrum for record keeping by right-clicking Export. 
	NOTE: Using a scale of 1,600 width/1,200 unit is a reasonable size for data inspection on a computer screen.
	<ol>
		<li>Select the JPEG file type, click on Destination, select Browse Disc (e.g., flash disc E:), type the file name (e.g., 1201-1.jpg), and then click on Export.</li>
	</ol>
	</li>
	<li>For data quantitation, use the cursor and click on the peak to show the peak height at the upper left-hand corner.
	<ol>
		<li>Alternatively, print out an exported JPEG file and measure the peak height with a ruler manually.</li>
	</ol>
	</li>
</ol>

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The authors have nothing to disclose.

This study described a step-by-step proofreading activity assay analyzed by the chosen commercial instrument (see the Table of Materials) using MALDI-TOF MS. The major advantages include that the primer and template are label free and easy to perform, allowing for greater flexibility in designing experiments. A stream-lime complete processing of 30 proofreading tests would take 4 h, including 3 h for manually performing the proofreading reactions and their cleanup, while the MALDI-TOF MS analyses using the aforementioned protocol takes 1 h to complete. The primary drawback of this method is the amount of substrate required for the assay. To obtain strong signals and consistent results, we used 50 pmol of DNA substrate in a 20-&#181;L reaction; thus, the amount of DNA is much higher than with radiolabeled DNA experiments8,9,10. Reducing the volumes and the sample size and fine-tuning the MS operation can lower the amount of substrate required.
Synthetic oligonucleotides obtained from commercial sources can be used for the proofreading study directly without any further purification or treatment. However, it is advisable to check the purity and quality of the DNA by MALDI-TOF MS. The phosphorothioate bonds were introduced to replace the last 4 phosphodiester bonds at the 3&#39; end of the template (Table 1) to reduce the non-specific degradation of the template strand (Figure 3G), albeit at a higher cost per sample. Interestingly, the signals from the template and primer DNA can be well separated by MS by virtue of their differences in size; thus, this phosphorothioate modification is not required. In Figure 3E, for example, all the possible products of the primer P21 extension and the proofreading excision and resynthesis were less than 22 nt (m/z &#60; 7,000), while all the degradation fragments from template T28 were greater than 24 nt (m/z &#62; 7,000). The results in Figure 4A were from the same P21/T28 duplex set as those in Figure 3E. An enlarged window of m/z 6,000-7,000 shows only signals from the primer DNA without any confounding signals from the template.
The strategy of using primers with a ddNMP extension proved to be very stable, and, thus, ddNMP containing products are good indicators for the proofreading assay. The addition of ddNMP to excision products could &#39;freeze&#39; the proofreading products in the state for analysis immediately post-excision.&#160;Alternatively, using a single correct dNTP (Figure 5B) instead of 4 ddNTPs to suppress the non-specific hydrolytic activity of the proofreading exonuclease is also feasible.
Stopping the reactions with a standard phenol/chloroform extraction procedure proved to be the best means for the reaction termination since, in one step, it enables the reactions to be stopped while also removing any protein interference. Alternatively, acid quenching, which is amenable to automation, then neutralized by non-metal alkaline without a phase separation, also demonstrated reliable results. Sodium ion adducts (Na+ in Figure 4, 5, and 7) are of concern since they can increase background and complicate the signal discrimination. A proper use of the clean resin can reduce such interference; yet, even when such adducts are present, they normally have much smaller peaks relative to their parent-signal, and this property can be used to distinguish between them (Figure&#160;4, 5, and 7). Generally, the peak height of sodium adducts was proportional to the major peaks; the inclusion or exclusion of sodium adducts in the calculation did not significantly affect the quantification results. Therefore, for the quantification, we used the major peak height only for the calculation.
The proofreading experiments described here took advantage of internal mismatches to demonstrate the importance of the exonuclease domain partition assay of KF26 and the detailed DNA polymerase I capability to efficiently proofread the mismatch at up to 4 nt upstream from the primer 3&#39;-end (Figure 5, MM1 - 4) possibly due to the base-pairing stability at the primer-template junctions27. In addition, this study also used primers containing penultimate deoxyinosine lesions to study a pol I repair of endo V-nicked repair intermediates. The results showed that T-I was repaired more efficiently than A-I and G-I; however, C-I was refractory to the pol I repair (Table 1 and Figure 7). These results confirm that a MALDI-TOF MS analysis can be very useful for proofreading and various short-patch DNA repair assays28 due to its high resolution.

The proofreading functions of DNA polymerases during DNA replication are essential to ensure the high fidelity of genetic information that needs to be transferred to progeny1,2,3,4,5,6,7. Being able to assess the contributions of polymerase proofreading exonucleases would clarify the mechanisms safeguarding genetic stability.
Radioisotope labeling and gel-based assays in combination with densitometric analyses of the autoradiograms or phosphor imaging8,9,10 have traditionally been used to detect proofreading activity of DNA polymerases. While functional, these assays are laborious, expensive, and not amenable to high-throughput formats. In addition, radioisotopes suffer safety issues including waste disposal. Alternatively, proofreading activities have been analyzed by fluorometric techniques. For example, 2-aminopurine (2-AP) can be incorporated into extension products during in vitro polymerase proofreading assays to produce a fluorescent signal11,12. Unfortunately, these approaches suffer from a low specificity, since 2-AP can pair with both thymine and cytosine. More recent approaches include a sensitive G-quadruplex-based luminescent switch-on probe for a polymerase 3&#39;-5&#39; exonuclease assay13 as well as a singly-labeled fluorescent probe for a polymerase proofreading assay that overcomes some of the aforementioned drawbacks14. Enthusiasm for these fluorometric methods is diminished due to the need for the specific labeling of DNA substrates.
In contrast, a MALDI-TOF MS for DNA analysis has been employed in the PinPoint assay, where the primer extension reactions with unlabeled 4 ddNTPs can be used to identify polymorphisms at a given locus15,16,17 and has been widely adopted in clinical applications for mutation detections and cancer diagnoses18. Using these basic principles, we have created a label-free assay for the in vitro determination of DNA polymerase proofreading activity exploiting the high resolution, high specificity, and high-throughput potential of MALDI-TOF MS. Using the E. coli DNA polymerase I Klenow fragment as a model enzyme, dideoxyribonucleotide triphosphates (ddNTPs) as substrates can take a &#34;snapshot&#34; of proofreading products after a single nucleotide extension via MALDI-TOF MS (Figure 1).
Likewise, this method was also developed for a DNA repair assay where primers containing 3&#39; penultimate dI lesions are subjected to a pol I repair assay which mimics endo V nicked repair intermediates. While not fully understood, the endo V repair pathway is the only repair system known to employ the pol I proofreading exonuclease activity for lesion excision19,20. Using MALDI-TOF MS, we show a clearly defined repair patch where dI can be excised by pol I when occurring in the last 2 nt of the primer before adding the correct complemented nucleotide.
For the study of proofreading and DNA repair, this method is faster and less laborious than previous methods and provides additional information towards mechanism and function.

<table><tbody><tr><td>Oligonucleotides</td><td>Mission Biotech (Taiwan)</td><td></td><td></td></tr><tr><td>phosphorothioate modified oligonucleotides</td><td>Integrated DNA Technologies (Taiwan)</td><td></td><td></td></tr><tr><td>DNA polymerase I, Large (Klenow) Fragment</td><td>New England Biolabs, MA</td><td>M0210L</td><td></td></tr><tr><td>Klenow fragment (3&#039;&amp;rarr;5&#039; exo-)</td><td>New England Biolabs, MA</td><td>M0212L</td><td></td></tr><tr><td>NEBuffer 2.1 (10X)</td><td>New England Biolabs, MA</td><td>B7202S</td><td></td></tr><tr><td>2&#039;, 3&#039; ddATP</td><td>Trilink Biotechnologies, CA</td><td>N4001</td><td></td></tr><tr><td>2&#039;, 3&#039; ddGTP</td><td>Trilink Biotechnologies, CA</td><td>N4002</td><td></td></tr><tr><td>2&#039;, 3&#039; ddTTP</td><td>Trilink Biotechnologies, CA</td><td>N4004</td><td></td></tr><tr><td>2&#039;, 3&#039; ddCTP</td><td>Trilink Biotechnologies, CA</td><td>N4005</td><td></td></tr><tr><td>dATP, dGTP, dCTP, dTTP set</td><td>Clubio, Taiwan</td><td>CB-R0315</td><td></td></tr><tr><td>SpectroCHIP array</td><td>Agena Bioscience, CA</td><td>#01509</td><td></td></tr><tr><td>MassARRAY</td><td>Agena Bioscience, CA</td><td></td><td></td></tr><tr><td>Typer 4.0 software</td><td>Agena Bioscience, CA</td><td>#10145</td><td></td></tr><tr><td>Clean Resin Tool Kit</td><td>Agena Bioscience, CA</td><td>#08040</td><td></td></tr></tbody></table>

proofreading and dna repair assay using single nucleotide extension and maldi-tof mass spectrometry analysis

The maintenance of the genome and its faithful replication is paramount for conserving genetic information. To assess high fidelity replication, we have developed a simple non-labeled and non-radio-isotopic method using a matrix-assisted laser desorption ionization with time-of-flight (MALDI-TOF) mass spectrometry (MS) analysis for a proofreading study. Here, a DNA polymerase [e.g., the Klenow fragment (KF) of Escherichia coli DNA polymerase I (pol I) in this study] in the presence of all four dideoxyribonucleotide triphosphates is used to process a mismatched primer-template duplex. The mismatched primer is then proofread/extended and subjected to MALDI-TOF MS. The products are distinguished by the mass change of the primer down to single nucleotide variations. Importantly, a proofreading can also be determined for internal single mismatches, albeit at different efficiencies. Mismatches located at 2-4-nucleotides (nt) from the 3' end were efficiently proofread by pol I, and a mismatch at 5 nt from the primer terminus showed only a partial correction. No proofreading occurred for internal mismatches located at 6 - 9 nt from the primer 3' end. This method can also be applied to DNA repair assays (e.g., assessing a base-lesion repair of substrates for the endo V repair pathway). Primers containing 3' penultimate deoxyinosine (dI) lesions could be corrected by pol I. Indeed, penultimate T-I, G-I, and A-I substrates had their last 2 dI-containing nucleotides excised by pol I before adding a correct ddN 5'-monophosphate (ddNMP) while penultimate C-I mismatches were tolerated by pol I, allowing the primer to be extended without repair, demonstrating the sensitivity and resolution of the MS assay to measure DNA repair.

Watch this Scientific Journal Video about Proofreading and DNA Repair Assay Using Single Nucleotide Extension and MALDI-TOF Mass Spectrometry Analysis at JoVE.com

Proofreading and DNA Repair Assay Using Single Nucleotide Extension and MALDI-TOF Mass Spectrometry Analysis

A non-labeled, non-radio-isotopic method for DNA polymerase proofreading and a DNA repair assay was developed by using high-resolution MALDI-TOF mass spectrometry and a single nucleotide extension strategy. The assay proved to be very specific, simple, rapid, and easy to perform for proofreading and repair patches shorter than 9-nucleotides.

The maintenance of the genome and its faithful replication is paramount for conserving genetic information. To assess high fidelity replication, we have ...

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Unit 1

DNA Replication

DNA Replication and Repair

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9.6K Views.  National Taiwan University. Een niet-label niet-radio-isotopische methode voor DNA-polymerase proeflezen en een DNA repair assay werd ontwikkeld met behulp van hoge-resolutie MALDI-TOF massa spectrometrie en een strategie voor de uitbreiding van één nucleotide. De test bleek te zijn van de zeer specifieke, eenvoudig, snel en gemakkelijk uit te voeren voor het proeflezen en reparatie patches korter dan 9-nucleotiden.

Video: Proofreading and DNA Repair Assay Using Single Nucleotide Extension and MALDI-TOF Mass Spectrometry Analysis

Atomic Force Microscopy Investigations of DNA Lesion Recognition in Nucleotide Excision Repair

AFM imaging is a powerful technique for the study of protein-DNA interactions. This single molecule method allows the simultaneous resolution of different molecules and molecular assemblies in a heterogeneous sample. In the particular context of DNA interacting protein systems, different protein complex forms and their corresponding binding positions on target sites containing DNA fragments can thus be distinguished. Here, an application of AFM to the study of DNA lesion recognition in the prokaryotic and eukaryotic nucleotide excision DNA repair (NER) systems is presented. The procedures of DNA and protein sample preparations are described and experimental as well as analytical details of the experiments are provided. The data allow important conclusions on the strategies by which target site verification may be achieved by the NER proteins. Interestingly, they indicate different approaches of lesion recognition and identification for the eukaryotic NER system, depending on the type of lesion. Furthermore, distinct structural properties of the two different helicases involved in prokaryotic and eukaryotic NER result in and explain the different strategies observed for these two systems. Importantly, these experimental and analytical approaches can be applied not only to the study of DNA repair but also very similarly to other DNA interacting protein systems such as those involved in replication or transcription processes.

Here, the study of different DNA lesion recognition approaches via single molecule AFM imaging is demonstrated with the nucleotide excision repair system as an example. The procedures of DNA and protein sample preparations and experimental as well as analytical details for the AFM experiments are described.

AFM imaging is a powerful technique for the study of protein-DNA interactions. This single molecule method allows the simultaneous resolution of different ...

Genetics

Detection and Visualization of DNA Damage-induced Protein Complexes in Suspension Cell Cultures Using the Proximity Ligation Assay

The DNA damage response orchestrates the repair of DNA lesions that occur spontaneously, are caused by genotoxic stress, or appear in the context of programmed DNA breaks in lymphocytes. The Ataxia-Telangiectasia Mutated kinase (ATM), ATM- and Rad3-Related kinase (ATR) and the catalytic subunit of DNA-dependent Protein Kinase (DNA-PKcs) are among the first that are activated upon induction of DNA damage, and are central regulators of a network that controls DNA repair, apoptosis and cell survival. As part of a tumor-suppressive pathway, ATM and ATR activate p53 through phosphorylation, thereby regulating the transcriptional activity of p53. DNA damage also results in the formation of so-called ionizing radiation-induced foci (IRIF) that represent complexes of DNA damage sensor and repair proteins that accumulate at the sites of DNA damage, which are visualized by fluorescence microscopy. Co-localization of proteins in IRIFs, however, does not necessarily imply direct protein-protein interactions, as the resolution of fluorescence microscopy is limited. 
In situ Proximity Ligation Assay (PLA) is a novel technique that allows the direct visualization of protein-protein interactions in cells and tissues with unprecedented specificity and sensitivity. This technique is based on the spatial proximity of specific antibodies binding to the proteins of interest. When the interrogated proteins are within ~40 nm an amplification reaction is triggered by oligonucleotides that are conjugated to the antibodies, and the amplification product is visualized by fluorescent labeling, yielding a signal that corresponds to the subcellular location of the interacting proteins. Using the established functional interaction between ATM and p53 as an example, it is demonstrated here how PLA can be used in suspension cell cultures to study the direct interactions between proteins that are integral parts of the DNA damage response.

Here, it is demonstrated how the in situ Proximity Ligation Assay (PLA) can be used to detect and visualize the direct protein-protein interactions between ATM and p53 in suspension cell cultures exposed to genotoxic stress.

The DNA damage response orchestrates the repair of DNA lesions that occur spontaneously, are caused by genotoxic stress, or appear in the context of ...

A Simple, Rapid, and Quantitative Assay to Measure Repair of DNA-protein Crosslinks on Plasmids Transfected into Mammalian Cells

The purpose of this method is to provide a flexible, rapid, and quantitative technique to examine the kinetics of DNA-protein crosslink (DPC) repair in mammalian cell lines. Rather than globally assaying removal of xenobiotic-induced or spontaneous chromosomal DPC removal, this assay examines the repair of a homogeneous, chemically defined lesion specifically introduced at one site within a plasmid DNA substrate. Importantly, this approach avoids the use of radioactive materials and is not dependent on expensive or highly-specialized technology. Instead, it relies on standard recombinant DNA procedures and widely available real-time, quantitative polymerase chain reaction (qPCR) instrumentation. Given the inherent flexibility of the strategy utilized, the size of the crosslinked protein, as well as the nature of the chemical linkage and the precise DNA sequence context of the attachment site can be varied to address the respective contributions of these parameters to the overall efficiency of DPC repair. Using this method, plasmids containing a site-specific DPC were transfected into cells and low molecular weight DNA recovered at various times post-transfection. Recovered DNA is then subjected to strand-specific primer extension (SSPE) using a primer complementary to the damaged strand of the plasmid. Since the DPC lesion blocks Taq DNA polymerase, the ratio of repaired to un-repaired DNA can be quantitatively assessed using qPCR. Cycle threshold (CT) values are used to calculate percent repair at various time points in the respective cell lines. This SSPE-qPCR method can also be used to quantitatively assess the repair kinetics of any DNA adduct that blocks Taq polymerase.

The goal of this protocol is to quantify the repair of defined DNA-protein crosslinks on plasmid DNA. Lesioned plasmids are transfected into recipient mammalian cell lines and low-molecular weight harvested at multiple time points post-transfection. DNA repair kinetics are quantified using strand-specific primer extension followed by qPCR.

The purpose of this method is to provide a flexible, rapid, and quantitative technique to examine the kinetics of DNA-protein crosslink (DPC) repair in ...

Rare Event Detection Using Error-corrected DNA and RNA Sequencing

Conventional next-generation sequencing techniques (NGS) have allowed for immense genomic characterization for over a decade. Specifically, NGS has been used to analyze the spectrum of clonal mutations in malignancy. Though far more efficient than traditional Sanger methods, NGS struggles with identifying rare clonal and subclonal mutations due to its high error rate of ~0.5&#8211;2.0%. Thus, standard NGS has a limit of detection for mutations that are &#62;0.02 variant allele fraction (VAF). While the clinical significance for mutations this rare in patients without known disease remains unclear, patients treated for leukemia have significantly improved outcomes when residual disease is &#60;0.0001 by flow cytometry. In order to mitigate this artefactual background of NGS, numerous methods have been developed. Here we describe a method for Error-corrected DNA and RNA Sequencing (ECS), which involves tagging individual molecules with both a 16 bp random index for error-correction and an 8 bp patient-specific index for multiplexing. Our method can detect and track clonal mutations at variant allele fractions (VAFs) two orders of magnitude lower than the detection limit of NGS and as rare as 0.0001 VAF.

Next-generation sequencing (NGS) is a powerful tool for genomic characterization that is limited by the high error rate of the platform (~0.5&#8211;2.0%). We describe our methods of error-corrected sequencing that allow us to obviate the NGS error rate and detect mutations at variant allele fractions as rare as 0.0001.

Conventional next-generation sequencing techniques (NGS) have allowed for immense genomic characterization for over a decade. Specifically, NGS has been ...

Using Next Generation Sequencing to Identify Mutations Associated with Repair of a CAS9-induced Double Strand Break Near the CD4 Promoter

Double strand breaks (DSBs) in DNA are the most cytotoxic type of DNA damage. Because a myriad of insults can result in these lesions (e.g., replication stress, ionizing radiation, unrepaired UV damage), DSBs occur in most cells each day. In addition to cell death, unrepaired DSBs reduce genome integrity and the resulting mutations can drive tumorigenesis. These risks and the prevalence of DSBs motivate investigations into the mechanisms by which cells repair these lesions. Next generation sequencing can be paired with the induction of DSBs by ionizing radiation to provide a powerful tool to precisely define the mutations associated with DSB repair defects. However, this approach requires computationally challenging and cost prohibitive whole genome sequencing to detect the repair of the randomly occurring DSBs associated with ionizing radiation. Rare cutting endonucleases, such as I-Sce1, provide the ability to generate a single DSB, but their recognition sites must be inserted into the genome of interest. As a result, the site of repair is inherently artificial. Recent advances allow guide RNA (sgRNA) to direct a Cas9 endonuclease to any genome locus of interest. This could be applied to the study of DSB repair making next generation sequencing more cost effective by allowing it to be focused on the DNA flanking the Cas9-induced DSB. The goal of the manuscript is to demonstrate the feasibility of this approach by presenting a protocol that can define mutations that stem from the repair of a DSB upstream of the CD4 gene. The protocol can be adapted to determine changes in the mutagenic potential of DSB associated with exogenous factors, such as repair inhibitors, viral protein expression, mutations, and environmental exposures with relatively limited computation requirements. Once an organism's genome has been sequenced, this method can be theoretically employed at any genomic locus and in any cell culture model of that organism that can be transfected. Similar adaptations of the approach could allow comparisons of repair fidelity between different loci in the same genetic background.

Presented here is sgRNA/CAS9 endonuclease and next-generation sequencing protocol that can be used to identify the mutations associated with double strand break repair near the CD4 promoter.

Double strand breaks (DSBs) in DNA are the most cytotoxic type of DNA damage. Because a myriad of insults can result in these lesions (e.g., replication ...

Identification of Rare Bacterial Pathogens by 16S rRNA Gene Sequencing and MALDI-TOF MS

There are a number of rare and, therefore, insufficiently described bacterial pathogens which are reported to cause severe infections especially in immunocompromised patients. In most cases only few data, mostly published as case reports, are available which investigate the role of such pathogens as an infectious agent. Therefore, in order to clarify the pathogenic character of such microorganisms, it is necessary to conduct epidemiologic studies which include large numbers of these bacteria. The methods used in such a surveillance study have to meet the following criteria: the identification of the strains has to be accurate according to the valid nomenclature, they should be easy to handle (robustness), economical in routine diagnostics and they have to generate comparable results among different laboratories. Generally, there are three strategies for identifying bacterial strains in a routine setting: 1) phenotypic identification characterizing the biochemical and metabolic properties of the bacteria, 2) molecular techniques such as 16S rRNA gene sequencing and 3) mass spectrometry as a novel proteome based approach. Since mass spectrometry and molecular approaches are the most promising tools for identifying a large variety of bacterial species, these two methods are described. Advances, limitations and potential problems when using these techniques are discussed.

Matrix-assisted laser desorption/ionization time of flight mass spectrometry (MALDI-TOF MS) and molecular techniques (16S rRNA gene sequencing) permit the identification of rare bacterial pathogens in routine diagnostics. The goal of this protocol lies in the combination of both techniques which leads to more accurate and reliable data.

There are a number of rare and, therefore, insufficiently described bacterial pathogens which are reported to cause severe infections especially in ...

Immunology and Infection

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 ...

Identification of RNA Fragments Resulting from Enzymatic Degradation using MALDI-TOF Mass Spectrometry

RNA is a biopolymer present in all domains of life, and its interactions with other molecules and/or reactive species, e.g., DNA, proteins, ions, drugs, and free radicals, are ubiquitous. As a result, RNA undergoes various reactions that include its cleavage, degradation, or modification, leading to biologically relevant species with distinct functions and implications. One example is the oxidation of guanine to 7,8-dihydro-8-oxoguanine (8-oxoG), which may occur in the presence of reactive oxygen species (ROS). Overall, procedures that characterize such products and transformations are largely valuable to the scientific community. To this end, matrix-assisted laser desorption ionization time-of-flight (MALDI-TOF) mass spectrometry is a widely used method. The present protocol describes how to characterize RNA fragments formed after enzymatic treatment. The chosen model uses a reaction between RNA and the exoribonuclease Xrn-1, where enzymatic digestion is halted at oxidized sites. Two 20-nucleotide long RNA sequences [5&#39;-CAU GAA ACA A(8-oxoG)G CUA AAA GU] and [5&#39;-CAU GAA ACA A(8-oxoG)(8-oxoG) CUA AAA GU] were obtained via solid-phase synthesis, quantified by UV-vis spectroscopy, and characterized via MALDI-TOF. The obtained strands were then (1) 5&#39;-phosphorylated and characterized via MALDI-TOF; (2) treated with Xrn-1; (3) filtered and desalted; (4) analyzed via MALDI-TOF. This experimental setup led to the unequivocal identification of the fragments associated with the stalling of Xrn-1: [5&#39;-H2PO4-(8-oxoG)G CUA AAA GU], [5&#39;-H2PO4-(8-oxoG)(8-oxoG) CUA AAA GU], and [5&#39;-H2PO4-(8-oxoG) CUA AAA GU]. The described experiments were carried out with 200 picomols of RNA (20 pmol used for MALDI analyses); however, lower amounts may result in detectable peaks with spectrometers using laser sources with more power than the one used in this work. Importantly, the described methodology can be generalized and potentially extended to product identification for other processes involving RNA and DNA, and may aid in the characterization/elucidation of other biochemical pathways.

MALDI-TOF was used to characterize fragments obtained from the reactivity between oxidized RNA and the exoribonuclease Xrn-1. The present protocol describes a methodology that can be applied to other processes involving RNA and/or DNA.

RNA is a biopolymer present in all domains of life, and its interactions with other molecules and/or reactive species, e.g., DNA, proteins, ions, drugs, ...

Quantifying DNA Double Strand Break Repair with Nanoluciferase Reporter Assays

Assessment of DNA Double Strand Break Repair Activity Using High-throughput and Quantitative Luminescence-Based Reporter Assays

The repair of DNA double strand breaks (DSBs) is crucial for the maintenance of genome stability and cell viability. DSB repair (DSBR) in cells is mediated through several mechanisms: homologous recombination (HR), non-homologous end joining (NHEJ), microhomology-mediated end joining (MMEJ), and single strand annealing (SSA). Cellular assays are essential to measure the proficiency and modulation of these pathways in response to various stimuli.
Here, we present a suite of extrachromosomal reporter assays that each measure the reconstitution of a nanoluciferase reporter gene by one of the four major DSBR pathways in cells. Upon transient transfection into cells of interest, repair of pathway-specific reporter substrates can be measured in under 24 h by the detection of Nanoluciferase (NanoLuc) luminescence.
These robust assays are quantitative, sensitive, titratable, and amenable to a high-throughput screening format. These properties provide broad applications in DNA repair research and drug discovery, complementing the currently available toolkit of cellular DSBR assays.

Author Spotlight: Developing Novel Anticancer Therapeutics Targeting the DNA Damage Response

We present protocols for the assessment of double strand break repair pathway proficiency in cells using a suite of luminescence-based extrachromosomal reporter substrates.

The repair of DNA double strand breaks (DSBs) is crucial for the maintenance of genome stability and cell viability. DSB repair (DSBR) in cells is ...

Cancer Research

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 ...

Proofreading and DNA Repair Assay Using Single Nucleotide Extension and MALDI-TOF Mass Spectrometry Analysis

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