This work was funded by the National Institutes of Health (ES023350). Lisa Chesner is supported by Training Grant 5T32HL007741. We thank Natalia Tretyakova (University of Minnesota) and Ashis Basu (University of Connecticut) and their lab members for support and technical advice during the early and intermediate stages of this work.

1. Generation of DPC-containing Plasmid
<ol>
	<li>Combine 80 pmol of oligonucleotide containing an 8-oxoguanine residue (20 &#181;L) with 10 units of T4 polynucleotide kinase (1 &#181;L) in 10x ligase buffer (5 &#181;L). Add water to reach a total volume of 50 &#181;L and incubate at 37 &#176;C for 30 min in a heated water bath.
	<ol>
		<li>Combine the phosphorylated oligonucleotide (50 &#181;L), 80 pmol of single-stranded DNA (80 &#956;L), 100 units Taq polymerase (20 &#181;L) in 10x Taq reaction buffer (25 &#181;L), 100 mM ATP (20 &#181;L), 10 mM dNTPs (20 &#181;L), NEB buffer 2 (25 &#181;L), and 8 &#181;g BSA (20 &#181;L) to total 260 &#181;L. Incubate the sample in a thermocycler set at 75 &#176;C for 15 min followed by 37 &#176;C for 5 min. Next, add 60 U of T4 polymerase (20 &#181;L), 8,000 U T4 ligase (20 &#181;L), 100 mM ATP (20 &#181;L), 10 mM dNTPs (20 &#181;L), NEB buffer 2 (15 &#181;L), and 8 &#181;g BSA (20 &#181;L) to total 375 &#181;L. Incubate the reaction overnight at 37 &#176;C (Figure 1A).</li>
	</ol>
	</li>
	<li>Create a 50:50 buffer-saturated phenol:chloroform mixture. Add equal volumes of each component, mix, and spin in a table-top centrifuge at 21,130 x g for 5 min. Chloroform drives water out of the buffer-saturated phenol to form two layers: an upper, aqueous layer and a bottom, organic layer. Add 375 &#181;L of the lower, organic layer to the primer extension reaction, mix, and spin in a table-top centrifuge at 21,130&#160;x g for 5 min. 
	CAUTION: Phenol and chloroform are hazardous substances and precautions should be taken to avoid contact with the eyes or skin.
	<ol>
		<li>Following centrifugation, carefully extract the top layer and mix with ammonium acetate added to a final concentration of 0.3 M followed by 2 volumes of 100% ethanol. Store the solution at -20 &#176;C for a minimum of 30 min or overnight.</li>
		<li>Spin the sample in a table-top centrifuge at 15,000 x rpm&#160;at 4 &#176;C for 10 min. Remove the supernatant and wash the pellet in 1 mL of 70% ethanol. Spin the sample in a table-top centrifuge at 15,000 x rpm&#160;at 4 &#176;C for 5 min. Remove the supernatant and resuspend the pellet in 100 &#181;L of water.</li>
	</ol>
	</li>
	<li>Combine 50 &#181;L of DNA from the previous step with 16 &#181;L 6x gel-loading dye, and 34 &#181;L water and subject to electrophoresis on a 0.8% low-melt agarose gel containing 0.5 &#181;g/mL ethidium bromide. Run the gel at 2 V/cm on a 10-cm gel for 6 h in 1x TAE buffer. Excise the supercoiled band using a razor blade and weigh the gel slice (Figure 1A). Run a positive control to serve as a marker for where the covalently closed, supercoiled DNA migrates. 
	CAUTION: Ethidium bromide is a mutagen and precautions should be taken to avoid contact with the skin. Also, it is important to minimize the time that the plasmid sample is exposed to UV in order to reduce the formation of UV photoproducts.
	<ol>
		<li>Digest the gel slice by adding 10% &#946;-agarase reaction buffer for every mg of gel weight. Incubate at 65 &#176;C for 10 min and cool to 42 &#176;C. Add 10 U of &#946;-agarase and incubate at 42 &#176;C for 1 h.</li>
		<li>Following incubation, measure the volume and add ammonium acetate to a final concentration of 0.3 M and chill on ice for 15 min. Centrifuge at 15,000 x g for 15 min at room temperature, collect the supernatant, and add 2 volumes of isopropanol. Chill at -20 &#176;C overnight.</li>
		<li>Centrifuge the purified, supercoiled DNA for 10 min at 15,000 x rpm&#160;in a table-top centrifuge at 4 &#176;C and resuspend the pellet in 40 &#181;L of water.</li>
	</ol>
	</li>
	<li>Crosslink 12 pmol (15 &#181;L) of DNA to 36 pmol of oxoguanine glycosylase (1 &#181;L) in buffer containing 100 mM NaCl (3 &#181;L), 1 mM MgCl2 (3 &#181;L), 20 mM Tris-HCl pH 7.0 (6 &#181;L), and 10 mM sodium cyanoborohydride (2 &#181;L) to reach a final volume of 30 &#181;L at 37 &#176;C for 30 min26. Remove 1 &#181;L of crosslinked sample and dilute in 500 &#181;L of water to serve as a 0 h data point and analyze on PCR in step 3.</li>
</ol>
2. KCl/SDS Precipitation
<ol>
	<li>To visualize crosslinking efficiency, restriction digest 1 &#181;g of the crosslinked plasmid with 1 &#956;L BspDI in 10x buffer at 37 &#176;C for 1 h to generate two different sized DNA fragments. 
	NOTE: One fragment will be crosslinked to protein (4.4 kb) and the other will not (2.8 kB) (Figure 1B).
	<ol>
		<li>Divide the samples in half, add SDS to both to a final concentration of 0.5%, and incubate at 65 &#176;C for 10 min.</li>
		<li>Add KCl (final concentration 100 mM) to one of the samples and incubate on ice for 5 min.</li>
		<li>Centrifuge both samples at 12,000 x g for 5 min at 4 &#176;C and run the supernatants on an agarose gel to estimate the percent conjugation of protein to DNA12. 
		NOTE: KCl/SDS precipitation is used for quality control, not substrate preparation.</li>
	</ol>
	</li>
</ol>
3. Transfection into Mammalian Cells
<ol>
	<li>1 day prior to transfection, plate 0.5 x 106 cells/well in a 6-well plate. The next day, mix 1.5 &#181;g (30 &#181;L) of the DPC-containing plasmid (from step 1.6) with 300 &#181;L of serum-free culture media. In another tube, mix 12 &#181;L of transfection reagent and 300 &#181;L of serum-free culture media. Combine 300 &#181;L of diluted DNA with 300 &#181;L of diluted transfection reagent and incubate for 5 min at room temperature. Add 250 &#181;L of the complexes to each of two wells and incubate for a minimum of 1 h at 37 &#176;C.</li>
	<li>Following incubation, remove media, add 1 mL of 0.6% SDS/0.01 M EDTA, and incubate at room temperature for 10 -&#160;15 min. Scrape the cells using a rubber policeman and transfer to a 1.5 mL microfuge tube. Add 200 &#181;L of 5 M NaCl (final concentration of 1 M), invert gently 5 times, and incubate at 4 &#176;C overnight27.</li>
	<li>Centrifuge the samples at 21,130&#160;x g in a table-top centrifuge at 4 &#176;C for 30 min, collect the supernatant, ethanol precipitate as described above, and resuspend in 50 &#181;L of water.</li>
</ol>
4. SSPE-qPCR
<ol>
	<li>Mix together 1 &#181;L of recovered DNA from each time point (including 0 h), 2x SYBR green PCR master mix&#160;(30 &#181;L), 27 &#181;L of water, and 1 &#181;L (100 pMol) of primer complementary to the damage strand of the plasmid (Primer R, Figure 2). Perform PCR using the following conditions: Initial pre-melt for 10 min at 90 &#176;C, followed by 8 cycles of: 90 &#176;C, 15 s, 65 &#176;C, 1 min. At the completion of cycle 8 add 1 &#181;L (100 pMol) of the second primer to total 60 &#181;L (Primer L, Figure 2)28 (see Table of Materials for reagent details).</li>
	<li>Mix 1 &#181;L of unamplified recovered DNA from each time point (including 0 h), 2x master mix (30 &#181;L), 27 &#181;L of water, and 1 &#181;L of each primer (100 pMol) to total 60 &#181;L.</li>
	<li>Load a 96-well PCR plate with the samples from steps 4.1 and 4.2 (20 &#181;L/well, in triplicate) and perform qPCR for 30 cycles using the conditions described above.</li>
	<li>Average the CT values from each set of triplicate samples. Subtract the CT values generated in step 4.1 from those in step 4.2 to obtain the delta CT value for each sample. Subtract the 0 h time point from the delta CT value to remove any background (See Representative Results section for a detailed description).</li>
	<li>Convert the delta CT value into percent repair using the formula: 
	<img alt="Equation" src="/files/ftp_upload/57413/57413eq1v2.jpg" /></li>
</ol>

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M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Transfection+by+Electroporation.">Transfection by Electroporation.</a> Current protocols in molecular biology. , (2003).</li></ol>

The authors have nothing to disclose.

The SSPE-qPCR method offers numerous advantages over other approaches by examining the repair of a homogenous population containing a single, defined DPC lesion. It is noteworthy that in addition to controlling the identity of the protein and the type of chemical crosslink used to connect the protein to the DNA, it is possible to easily manipulate the sequence context into which the DPC lesion is introduced. We have explored the influence on DPC repair of introducing the lesion on either the template or coding strand of a plasmid downstream of an active promotor locus. Similarly, we are in the process of investigating the influence on DPC repair of replication using a M13 plasmid containing an SV40 origin of replication transfected into HEK293T cells. The assay described herein directly measures DPC repair, as opposed to other strategies such as host cell reactivation that indirectly estimates repair activity24,25. In addition, the system is robust, sensitive, and quantitative. Unlike other systems, which measure DPC removal, this assay only detects complete repair events, i.e., it requires not only that the DPC lesion be removed but that the integrity of the duplex DNA be fully restored17. This is because abasic sites and nicks or breaks in the phosphodiester backbone block the assay as effectively as the original DPC lesion29.
While this report focuses on a particular type of DPC, which was created by borohydride trapping of an enzyme reaction intermediate, we are currently developing approaches to study repair of DPCs involving other proteins and lesions in which the protein linkage to the DNA occurs through the nucleoside base, rather than the ribose position. Using reductive amination, we have created protein and peptide crosslinks attached to the guanine or cytosine base of a DNA primer30. These oligonucleotides were purified to homogeneity and used to generate supercoiled plasmids containing DNA-protein and DNA-peptide crosslinks. While the efficiency of these reactions is somewhat reduced relative to that of the oxoguanine glycosylase crosslink approach described in detail above, they were nevertheless successful and permit the examination of the repair of these substrates in wild-type and nucleotide excision repair-deficient mammalian cell lines. We have also used the SSPE-qPCR assay to study the repair of oxoguanine lesions and a synthetic ribose-cholesterol conjugate, which was previously shown to be repaired via the cellular nucleotide excision repair machinery31. As Figure 2 graphically depicts, repair of any lesion that blocks primer extension by Taq polymerase can, in principle, be measured using the SSPE-qPCR assay.
Current models of DPC repair suggest that larger DPCs (&#62;10 kDa) are subjected to proteolytic processing to smaller peptide lesion prior to removal32,33,34. Most likely candidates responsible for this proteolysis are the proteasome or a specific protease in human cells named Spartan20,21,35,36,37,38. Further investigations into the roles of these proteases can be conducted using the SSPE-qPCR assay. Proteasome inhibitors could be used to pretreat cells prior to transfection with DPC-containing substrates. Alternatively, Spartan knockdown cell lines could be transfected with damaged plasmids to elucidate its role in proteolysis of larger DPCs.
Irrespective of the method used to create the crosslink or of the nature of the DNA lesion, it is worth stressing that the SSPE-qPCR methodology is critically dependent on the ability to generate substantial amounts of homogeneous DPC-plasmid substrate. Steps essential for this analysis include purification of covalently, closed-circular plasmid following primer extension to eliminate any nicked or linear plasmid molecules. These contaminants must be eliminated to ensure that any subsequent DPC repair observed is not due to nick-directed or double-strand break-directed repair processes. Failure to obtain sufficient quantities of supercoiled DNA following primer extension reactions may be overcome by varying the ratio of oligonucleotide to single-stranded DNA. Another important step for the SSPE-qPCR method, is the crosslinking efficiency of the protein to the plasmid. In this study, an efficient, enzymatic reaction was used to crosslink the oxoguanine glycosylase protein to an 8-oxo-guanine lesion. Sub-optimum crosslinking can be improved by varying the amount of protein added to the reaction.
While the results described in this report relied on lipofection to introduce DPC repair substrates into recipient cells, there is no reason, a priori, other transfections methods could not be employed. We have performed preliminary studies and observed that electroporation39 can also be used. However, it is worth noting that in our experience, electroporation transfection efficiency is reduced compared to lipofection, and we found it necessary to use carrier DNA (up to 5 &#181;g) in addition to the 1.5 &#181;g of DPC substrate to obtain repair data. Overall, the SSPE-qPCR method described above provides an innovative way to exclusively examine DPC repair on plasmid DNA and generate new insight into the DNA damage response.

Described herein is a PCR-based assay termed SSPE-qPCR. The purpose of this method is to quantify DPC repair on plasmid DNA transfected into repair deficient and proficient mammalian cells. This assay is rapid, quantitative, extremely flexible, and directly measures repair activity. While this report focuses on the use of this methodology to study repair of DPCs, results presented below illustrate that repair of any lesion that blocks Taq polymerase can be studied using this methodology.
The rationale behind the development of this method is to gain insight into the mechanisms through which mammalian cells repair DPCs. Unlike other types of DNA damage, DPCs are massively diverse1,2. Studies have demonstrated that hundreds of cellular proteins can become crosslinked to DNA and that for each protein there are, in principle, numerous amino acid side chains that could become covalently attached to cellular DNA3. In addition, there are numerous chemical attachment points for proteins onto the DNA backbone, including several positions on the nucleotide bases as well as on the ribose sugar4,5. This chemical diversity raises the prospect that distinct biochemical pathways may be relied upon to repair different types of DPCs. It was with this concern in mind that the SSPE-qPCR assay was developed.
Several techniques have been developed to gain insight into the molecular biology of cellular DPC repair. The following provides an overview of the major approaches that have been developed, with a summary of the major strengths and weaknesses each possess. It is worth stressing that while this summary focuses on studies of DPC repair in mammalian cell culture systems, significant contributions to the current model of DPC repair have been made using microbial and cell-free systems that are not discussed in this manuscript.
Perhaps the easiest strategy that can be taken to gain insight into the genetics of DPC repair is to assess the respective sensitivity to cell death observed in wild-type and mutant cells exposed to agents that induce DPCs6,7. This strategy is relatively fast, inexpensive, and does not require specialized expertise beyond the ability to perform basic cell culture techniques. Counterbalancing these advantages are numerous limitations to this approach including the following. First, the assay does not directly measure DNA repair. The working assumption underlying this strategy is that inactivating mutations in genes encoding relevant DNA repair proteins results in an accumulation of DNA damage that triggers programmed cell death. However, mutations in genes encoding non-DNA-repair proteins could, in principal, enhance (or reduce) cellular sensitivity to xenobiotic-induced cell death. Second, agents that create DPCs invariably induce other types of DNA damage (one exception is 5-aza-2&#39;-deoxycytadine, but this agent also depletes cellular methyltransferase levels8). Consequently, it is conceivable that enhanced cellular hypersensitivity to the agent in question may reflect defects in repair of interstrand crosslinks or other lesions. Third, as was mentioned above, DPCs represent a vastly heterogeneous class comprised of different types of chemical crosslinks, involving different protein partners. It is possible that while repair of one or more sub-types of these lesions may be altered in a particular genetic background, this difference may not be sufficient to significantly alter cellular hypersensitivity to death induced by this agent. In summary, while this strategy represents an attractive starting point, the limitations outlined above highlight the importance of pursuing other, more direct methods to study the kinetics of DPC repair.
A number of related approaches have been developed to achieve this objective. For instance, investigators have developed methods to distinguish between &#39;free&#39; DNA and &#39;protein-bound&#39; DNA9,10,11. Using these approaches, it is possible to compare steady-state levels of DPCs or following exposure to a DPC-producing agent in different genetic backgrounds. The two strategies that have been most widely used involve separating DPC-containing DNA from free DNA using either a nitrocellulose membrane binding strategy or KCl/SDS precipitation12,13. In the former approach cells are lysed and passed through a nitrocellulose filter. Because nitrocellulose binds protein, the filter retains protein-linked DNA, permitting free DNA to pass through. In the latter strategy protein-bound DNA is separated from free DNA based on the fact that SDS binds to protein but not DNA, and can be precipitated by the addition of KCl. Consequently, protein-linked DNA becomes insoluble while unbound DNA remains in solution. DPC-containing DNA can then be quantitated using radiolabeled thymidine (if cells were initially metabolically labeled) or by a DNA-selective fluorescent dye like Hoechst 33258. These methods are reproducible and require a small number of steps. However, they do not provide information regarding the nature of the chemical crosslink through which protein is attached to DNA. Furthermore, it is important to note, these assays may over-estimate DPC repair by falsely scoring incomplete repair, i.e., proteolytic processing to smaller DNA-peptide crosslinks that may not be as easily trapped or precipitated, as bona fide DNA repair.
Comet assays can be used to visualize DPC formation in cells14. In these experiments, the presence of DPCs decrease DNA migration which can then be reversed by pretreating with proteinase K. Therefore, the length of the tail can be used to estimate DPC formation. However, as mentioned above, DPC-forming drugs create other types of DNA damage which could alter tail length. This protocol is also highly technical and requires expertise and training in confocal imaging.
Mass spectrometry can be used to study DPC repair kinetics following treatment with crosslinking agents15,16,17. These experiments treat cells with DPC-forming agents and isolate DPCs via biotin capture or phenol:chloroform (1:1) extraction. Mass spectrometry can then be used to identify the crosslinked proteins or quantitate the amount of DPCs formed over time. The major advantage of this approach is the nature of the data produced. It is possible to precisely catalog the types of proteins that become crosslinked following exposure to a xenobiotic, however, this protocol is expensive, time consuming, and is limited by the type of crosslink that can be detected.
Maizels et al. developed a sensitive &#39;RADAR&#39; (rapid approach to DNA adduct recovery) assay to quantitate immunodetection of DNA-protein adducts as well as an ELISA-based RADAR assay18,19. These assays are especially useful for trapping DNA-protein intermediates that transiently form in cells and generate samples suitable for mass spectroscopy to identify new protein adducts. This immunodetection assay relies on the availability of antibodies to capture the DNA-protein crosslink and, therefore, may not be capable of detecting degraded DNA-peptide adducts that form during repair. Recently, a specific DPC repair pathway linked to DNA replication and a DNA-dependent metalloprotease Spartan was discovered in which DPCs are proteolyzed to smaller peptides during repair20,21. Inherited mutations in this gene are associated with Ruijs-Aalfs syndrome in humans, a disease characterized by genomic instability, premature ageing, and liver cancer22. Mice with genetically engineered Spartan gene defects display similar phenotypes23.
Host-cell reactivation of transcriptional activity has been used to study the repair of defined lesions present on transfected plasmid DNA substrates24,25. In these experiments, plasmid containing DPCs (or other types of DNA lesions) that block the transcription of a reporter, such as luciferase, are transfected into cells. Luminescence measurements taken 24 - 72 h later are then correlated with DPC repair. However, these indirect repair assays are incapable of detecting repair events earlier than 24 h post-transfection and cannot distinguish between RNA polymerase bypass of partially repaired substrates and complete repair.
Each of the methods described above has advantages and has contributed to the current model of DPC repair. However, the SSPE-qPCR assay circumvents several of the limitations associated with these other approaches and consequently can provide more specific insight into DPC repair mechanisms. For example, the SSPE-qPCR assay can directly measure repair of site-specific DPCs on DNA in intact mammalian cells. This method is versatile and has been used to obtain repair results following transfection in hamster and human cell lines. Transfection of the plasmid can be performed using lipofection or electroporation in cultured mammalian cell lines. It also ensures that only repair of defined DPCs is measured, and not other types of DNA damage induced by most DPC-forming agents. The SSPE-qPCR is easy to perform, inexpensive, and rapid. Results obtained using this assay have detected repair events as early as 2 h post-transfection. Using this method, variables that may influence DPC repair outcomes can be studied in a manner that is sensitive and efficient. For example, the role of transcription in DPC repair has yet to be rigorously evaluated. Due to the flexibility of the SSPE-qPCR assay, the crosslinking site of the DPC can be manipulated to address this question. In addition, introduction of an origin of replication into the DPC-bearing plasmid can be used to address the influence of replication on DPC repair. Additionally, multiple crosslinks can be created on the plasmid to examine differences in repair of a single DPC versus multiple crosslinks. These are questions that would be difficult to answer using chromosomal DNA but can easily be addressed using the SSPE-qPCR assay. Overall, the SSPE-qPCR assay requires purified, plasmid DNA in microgram quantities containing a lesion of a known location. Adducts besides DPC can be used in this assay, however, the lesion must be capable of blocking extension by Taq polymerase.

<table><tbody><tr><td>8-oxoguanine modified oligo</td><td>Midland Certified Reagent Company</td><td>NA</td><td>Sequence: 5&amp;rsquo;AGGGTTTTCCA (8-oxo-dG)TCACGACGTT 3&amp;rsquo;</td></tr><tr><td>T4 PNK</td><td>NEB</td><td>M0201S</td><td></td></tr><tr><td>Single-stranded M13</td><td>NEB</td><td>N4040S</td><td></td></tr><tr><td>Taq polymerase</td><td>NEB</td><td>M0267L</td><td></td></tr><tr><td>ATP, 100mM</td><td>Thermo Scientific</td><td>R0441</td><td></td></tr><tr><td>dNTP Mix, 10mM each</td><td>Thermo Scientific</td><td>R0192</td><td></td></tr><tr><td>BSA</td><td>NEB</td><td>B9001S</td><td></td></tr><tr><td>T4 polymerase</td><td>NEB</td><td>M0203L</td><td></td></tr><tr><td>T4 ligase</td><td>NEB</td><td>M0202L</td><td></td></tr><tr><td>&amp;beta;-agarase</td><td>NEB</td><td>M0392S</td><td></td></tr><tr><td>Oxoguanine glycosylase 1</td><td>NEB</td><td>M0241S</td><td></td></tr><tr><td>SYBR Green PCR Master Mix</td><td>Applied Biosystems</td><td>4309155</td><td>green PCR master mix</td></tr><tr><td>Primer R</td><td>UMN Genomic Center</td><td>NA</td><td>5&amp;rsquo;CGGCTCGTATGTTGTGTG 3&amp;rsquo;</td></tr><tr><td>Primer L</td><td>UMN Genomic Center</td><td>NA</td><td>5&amp;rsquo; GCTGCAAGGCGATTAAGT 3&amp;rsquo;</td></tr><tr><td>Lipofectamine Reagent</td><td>Invitrogen</td><td>18324-012</td><td>transfection reagent&amp;nbsp;</td></tr><tr><td>BspD1</td><td>NEB</td><td>R0557S</td><td></td></tr><tr><td>NEB buffer 2</td><td>NEB</td><td>B7002S</td><td></td></tr><tr><td>StepOnePlus Real Time PCR System</td><td>Applied Biosystems</td><td>4376600</td><td></td></tr><tr><td>UltraPure Buffer-Saturated Phenol</td><td>Invitrogen</td><td>15513-047</td><td></td></tr><tr><td>Chloroform, reagent ACS, spectro grade</td><td>ACROS</td><td>40463-5000</td><td></td></tr></tbody></table>

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.

In order to utilize the SSPE-qPCR assay to assess cellular DPC repair, a sufficient quantity (&#181;g amounts) of high-quality DPC repair substrate must be prepared. To obtain this product, an oligonucleotide containing an 8-oxoguanine residue was annealed to complementary single-stranded DNA, primer extended, and gel purified to ensure that only covalently, closed circular product was used for transfections (Figure 1A). This report focuses on substrates in which borohydride-trapping is used to create a covalent crosslink between recombinant human oxoguanine glycosylase and a ribose unit on a double-stranded circular plasmid molecule, although analogous approaches can be used to obtain chemically diverse DPC substrates. The oxoguanine glycosylase/borohydride trapping strategy is especially attractive given the extremely high efficiency of the crosslinking reaction. As shown in Figure 1B, KCl/SDS precipitation performed on a DPC substrate that had been digested into two fragments selectively and nearly quantitatively depleted the DNA fragment harboring the DPC. These results support the conclusion that essentially 100% of the plasmid substrate molecules contain a protein crosslink.
The SSPE-qPCR assay described in the Protocol section utilizes CT values generated by qPCR to calculate the percent of DPC repair following transfection in mammalian cells. As Figure 2 illustrates, the protein crosslink blocks Taq polymerase from extending the &#39;R&#39; primer annealed to the DPC-containing strand. A second, critical feature of this assay is the incorporation of eight rounds of SSPE reactions (using the R primer) prior to performing the qPCR assay. While undamaged (or repaired) DNA will be extended during these SSPE reactions, creating a binding site for the downstream &#39;L&#39; primer, damaged DNA (or incompletely repaired DNA) will not be extended. Consequently, SSPE increases the abundance of each undamaged (or repaired) strand by eight-fold. This means that repaired samples in which the SSPE step has been performed prior to qPCR will display a CT value that is three units lower than that obtained for an identical sample in which SSPE was not performed prior to qPCR (Figure 2). The three-unit difference in CT values observed for the two treatments, referred to as &#916;CT, reflects the fact that 8 = 23. This delta CT value can be used to calculate cellular DNA repair activity by using the formula: percent DNA repair = (2&#916;CT/23) x 100. Control experiments confirmed that a delta CT value of approximately 3 was consistently observed when undamaged substrate plasmids were subjected to SSPE-qPCR (data not shown).
Table 1 depicts example CT values that were generated from DPC-containing plasmid substrates that were recovered from Chinese hamster lung fibroblasts 3 h and 8 h post-transfection (Table 1A), as well as from time zero samples, i.e., samples that were prepared as described in the Protocol section, but not transfected into cells (Table1B). The table also provides the &#916;CT, and percent repair values (calculated using the formula 2&#916;CT/23 x 100) obtained from these samples. As illustrated in Table 1A, the percent repair calculated from samples harvested 3 h and 8 h post-transfection was 66% and 93%, respectively. These values are then subtracted from that obtained from analysis of the 0 h sample to determine the percent repair. Table 1B depicts CT values for two different 0 h samples in which efficient crosslinking was or was not achieved prior to transfection. Efficiently crosslinked sample resulted in a delta CT value of 0.8 whereas a poorly crosslinked sample resulted in a delta CT value of 2.5. To date, it has not been possible to precisely determine the source of the 0.8 background signal in the efficiently crosslinked 0 h sample. It is unlikely that this background reflects a low level of either spontaneous DPC removal, or is due to a low level of Taq polymerase &#39;bypass&#39; synthesis through the DPC lesion: extensive experimentation performed on highly purified single-stranded M13 substrate consistently yielded a delta CT value of approximately 0.8, i.e., essentially identical to this &#39;background&#39; extension seen in DPC-containing substrates. Consequently, it is likely that there is a small degree of mis-priming of the &#39;R&#39; primer during SSPE that results in the generation of a small amount of product that can subsequently be amplified when the &#39;L&#39; primer is added, and qPCR performed. Efforts to eliminate this background by manipulating PCR conditions have, to date, failed to remedy this defect. However, the subtraction strategy described above permits accurate estimation of DPC repair activity. It is worth noting that inefficient crosslinking of the protein onto the plasmid will result in an elevated background value. This is depicted in the sample data provided in Table 1B where inefficient protein-DNA crosslinking resulted in a higher delta CT value for time 0. Therefore, control experiments are invariably performed prior to initiating transfections and substrates with delta CT values elevated above the 0.8 threshold level are discarded. As mentioned in the qPCR protocol, each sample is divided into 3 wells to ensure pipetting consistency. These replicates are then averaged to obtain the CT value for that sample. Replicates that deviate more than &#177; 0.1 are eliminated from the data set. If all three replicates deviate more than &#177; 0.1 from each other, the SSPE-qPCR assay is redone until consistent values are obtained. Experience shows that at least 3 independent transfections must be performed to obtain reliable average values that can then be subjected to statistical analysis to determine the influence of various parameters on DPC repair efficiency.
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/57413/57413fig1.jpg" /> 
Figure 1. Generation of a DPC-containing plasmid. (A) An oligonucleotide containing an 8-oxo-guanine residue (red) is annealed to single-stranded DNA (bold black circle) and extended to create double-stranded plasmid (primer extension product indicated by dashed line). Following electrophoresis in ethidium bromide, the band corresponding to supercoiled DNA (red box) is excised from a low-melt agarose gel and digested with &#946;-agarase. Lanes: (1) Molecular weight marker, (2) single-stranded DNA, (3) double-stranded DNA, (4) primer extended sample. (B) Oxoguanine glycosylase (abbreviated hOGG1, depicted as an orange circle) is crosslinked to the 8-oxo-guanine residue via sodium cyanoborohydride and the resulting DPC substrate digested to generate a 2,800 base-pair, free DNA fragment and a 4,400 base-pair fragment attached to the protein. SDS is added to the sample which is then divided into two portions, one of which is treated with KCl and centrifuged to sediment DPC-containing DNA (depicted as an orange pellet). The supernatant from this latter sample (depicted as blue fluid in centrifuge tube) and the sample not exposed to KCl are subjected to gel electrophoresis. Lanes: (1) Molecular weight marker, (2) -KCl, (3) +KCl. <a href="https://cloudflare.jove.com/files/ftp_upload/57413/57413fig1large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 2" class="xfigimg" src="/files/ftp_upload/57413/57413fig2.jpg" /> 
Figure 2: Quantification of damaged plasmid via SSPE-qPCR. Plasmid DNA is denatured and a primer complementary to the damaged strand (R) is annealed and extended. This process is repeated for a total of 8 cycles. With a damaged substrate (top), Taq polymerase is blocked by the DPC lesion and will not produce full length product strands. In contrast, with a repaired substrate (bottom), Taq polymerase will produce full-length product strands containing the binding site for primer L. After 8 cycles, primer L is added, and cycle threshold values are determined using qPCR. Example amplification results for damaged and undamaged substrates are illustrated on the right with the red line representing DNA that underwent primer extension prior to qPCR and blue representing DNA that was not primer extended prior to qPCR. <a href="https://cloudflare.jove.com/files/ftp_upload/57413/57413fig2large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td colspan="2">Table 1A</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td rowspan="2">Time</td>
			<td rowspan="2">- Primer extension</td>
			<td rowspan="2">+ Primer extension</td>
			<td rowspan="2">&#916; CT</td>
			<td>Percent Repair</td>
		</tr>
		<tr>
			<td>(2&#916;CT/23x 100)&#160;</td>
		</tr>
		<tr>
			<td>3 h</td>
			<td>23.5</td>
			<td>21.1</td>
			<td>2.4</td>
			<td>66</td>
		</tr>
		<tr>
			<td>8 h</td>
			<td>29.4</td>
			<td>26.5</td>
			<td>2.9</td>
			<td>93</td>
		</tr>
		<tr>
			<td colspan="2">Table 1B</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td rowspan="2">Time</td>
			<td rowspan="2">- Primer extension</td>
			<td rowspan="2">+ Primer extension</td>
			<td rowspan="2">&#916; CT</td>
			<td>Percent Background</td>
		</tr>
		<tr>
			<td>(2&#916;CT/23x 100)&#160;</td>
		</tr>
		<tr>
			<td>0 h</td>
			<td>15.4</td>
			<td>14.6</td>
			<td>0.8</td>
			<td>22</td>
		</tr>
		<tr>
			<td>0 h</td>
			<td>15.4</td>
			<td>12.9</td>
			<td>2.5</td>
			<td>71</td>
		</tr>
	</tbody>
</table>
Table 1: Calculating percent repair using CT values generated from SSPE-qPCR. (A) Repair of a DPC-containing substrate transfected in V79 Chinese hamster lung fibroblasts 3 h and 8 h post-transfection. (B) SSPE-qPCR of 0 h samples not transfected into cells. In one sample the efficiency of the crosslinking reaction between DNA and oxoguanine glycosylase was high (this sample yielded a low delta CT value) while in the other sample the efficiency of protein crosslinking was low (this sample yielded a relatively high delta CT value). See text of Representative Results for details.

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

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 ...

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8.2K Views.  University of Minnesota. 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.

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

Tools to Study the Role of Architectural Protein HMGB1 in the Processing of Helix Distorting, Site-specific DNA Interstrand Crosslinks

High mobility group box 1 (HMGB1) protein is a non-histone architectural protein that is involved in regulating many important functions in the genome, such as transcription, DNA replication, and DNA repair. HMGB1 binds to structurally distorted DNA with higher affinity than to canonical B-DNA. For example, we found that HMGB1 binds to DNA interstrand crosslinks (ICLs), which covalently link the two strands of the DNA, cause distortion of the helix, and if left unrepaired can cause cell death. Due to their cytotoxic potential, several ICL-inducing agents are currently used as chemotherapeutic agents in the clinic. While ICL-forming agents show preferences for certain base sequences (e.g., 5&#39;-TA-3&#39; is the preferred crosslinking site for psoralen), they largely induce DNA damage in an indiscriminate fashion. However, by covalently coupling the ICL-inducing agent to a triplex-forming oligonucleotide (TFO), which binds to DNA in a sequence-specific manner, targeted DNA damage can be achieved. Here, we use a TFO covalently conjugated on the 5&#39; end to a 4&#39;-hydroxymethyl-4,5&#39;,8-trimethylpsoralen (HMT) psoralen to generate a site-specific ICL on a mutation-reporter plasmid to use as a tool to study the architectural modification, processing, and repair of complex DNA lesions by HMGB1 in human cells. We describe experimental techniques to prepare TFO-directed ICLs on reporter plasmids, and to interrogate the association of HMGB1 with the TFO-directed ICLs in a cellular context using chromatin immunoprecipitation assays. In addition, we describe DNA supercoiling assays to assess specific architectural modification of the damaged DNA by measuring the amount of superhelical turns introduced on the psoralen-crosslinked plasmid by HMGB1. These techniques can be used to study the roles of other proteins involved in the processing and repair of TFO-directed ICLs or other targeted DNA damage in any cell line of interest.

Targeted DNA damage can be achieved by tethering a DNA damaging agent to a triplex-forming oligonucleotide (TFO). Using modified TFOs, DNA damage-specific protein association, and DNA topology modification can be studied in human cells by the utilization of modified chromatin immunoprecipitation assays and DNA supercoiling assays described herein.

High mobility group box 1 (HMGB1) protein is a non-histone architectural protein that is involved in regulating many important functions in the genome, ...

Application of Laser Micro-irradiation for Examination of Single and Double Strand Break Repair in Mammalian Cells

Highly coordinated DNA repair pathways exist to detect, excise and replace damaged DNA bases, and coordinate repair of DNA strand breaks. While molecular biology techniques have clarified structure, enzymatic functions, and kinetics of repair proteins, there is still a need to understand how repair is coordinated within the nucleus. Laser micro-irradiation offers a powerful tool for inducing DNA damage and monitoring the recruitment of repair proteins. Induction of DNA damage by laser micro-irradiation can occur with a range of wavelengths, and users can reliably induce single strand breaks, base lesions and double strand breaks with a range of doses. Here, laser micro-irradiation is used to examine repair of single and double strand breaks induced by two common confocal laser wavelengths, 355 nm and 405 nm. Further, proper characterization of the applied laser dose for inducing specific damage mixtures is described, so users can reproducibly perform laser micro-irradiation data acquisition and analysis.

Confocal fluorescence microscopy and laser micro-irradiation offer tools for inducing DNA damage and monitoring the response of DNA repair proteins in selected sub-nuclear areas. This technique has significantly advanced our knowledge of damage detection, signaling, and recruitment. This manuscript demonstrates these technologies to examine single and double strand break repair.

Highly coordinated DNA repair pathways exist to detect, excise and replace damaged DNA bases, and coordinate repair of DNA strand breaks. While molecular ...

Cancer Research

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 ...

Biochemistry

Detection of Post-Replicative Gaps Accumulation and Repair in Human Cells Using the DNA Fiber Assay

The DNA fiber assay is a simple and robust method for the analysis of replication fork dynamics, based on the immunodetection of nucleotide analogs that are incorporated during DNA synthesis in human cells. However, this technique has a limited resolution of a few thousand kilobases. Consequently, post-replicative single-stranded DNA (ssDNA) gaps as small as a few hundred bases are not detectable by the standard assay. Here, we describe a modified version of the DNA fiber assay that utilizes the S1 nuclease, an enzyme that specifically cleaves ssDNA. In the presence of post-replicative ssDNA gaps, the S1 nuclease will target and cleave the gaps, generating shorter tracts that can be used as a read-out for ssDNA gaps on ongoing forks. These post-replicative ssDNA gaps are formed when damaged DNA is replicated discontinuously. They can be repaired via mechanisms uncoupled from genome replication, in a process known as gap-filling or post-replicative repair. Because gap-filling mechanisms involve DNA synthesis independent of the S phase, alterations in the DNA fiber labeling scheme can also be employed to monitor gap-filling events. Altogether, these modifications of the DNA fiber assay are powerful strategies to understand how post-replicative gaps are formed and filled in the genome of human cells.

Here we describe two modifications of the DNA fiber assay to investigate single-stranded DNA gaps in replicating DNA after lesion induction. The S1 fiber assay enables the detection of post-replicative gaps using the ssDNA-specific S1 endonuclease, while the gap-filling assay allows visualization and quantification of gap repair.

The DNA fiber assay is a simple and robust method for the analysis of replication fork dynamics, based on the immunodetection of nucleotide analogs that ...

Genetic Studies of Human DNA Repair Proteins Using Yeast as a Model System 

Understanding the roles of human DNA repair proteins in genetic pathways is a formidable challenge to many researchers. Genetic studies in mammalian systems have been limited due to the lack of readily available tools including defined mutant genetic cell lines, regulatory expression systems, and appropriate selectable markers. To circumvent these difficulties, model genetic systems in lower eukaryotes have become an attractive choice for the study of functionally conserved DNA repair proteins and pathways. We have developed a model yeast system to study the poorly defined genetic functions of the Werner syndrome helicase-nuclease (WRN) in nucleic acid metabolism. Cellular phenotypes associated with defined genetic mutant backgrounds can be investigated to clarify the cellular and molecular functions of WRN through its catalytic activities and protein interactions. The human WRN gene and associated variants, cloned into DNA plasmids for expression in yeast, can be placed under the control of a regulatory plasmid element. The expression construct can then be transformed into the appropriate yeast mutant background, and genetic function assayed by a variety of methodologies. Using this approach, we determined that WRN, like its related RecQ family members BLM and Sgs1, operates in a Top3-dependent pathway that is likely to be important for genomic stability. This is described in our recent publication [1] at <a href="http://www.impactaging.com">www.impactaging.com</a>. Detailed methods of specific assays for genetic complementation studies in yeast are provided in this paper.

Genetic Studies of Human DNA Repair Proteins Using Yeast as a Model System

Genetic studies in yeast can be employed to investigate the molecular and cellular functions of human genes in cellular DNA metabolism. Methods are described for the genetic characterization of the human WRN gene product defective in the premature aging disorder Werner syndrome in functionally conserved pathways using yeast as a tractable model system.

Understanding the roles of human DNA repair proteins in genetic pathways is a formidable challenge to many researchers.  Genetic studies in mammalian ...

Biology

Analysis of DNA Double-strand Break (DSB) Repair in Mammalian Cells

DNA double-strand breaks are the most dangerous DNA lesions that may lead to massive loss of genetic information and cell death. Cells repair DSBs using two major pathways: nonhomologous end joining (NHEJ) and homologous recombination (HR). Perturbations of NHEJ and HR are often associated with premature aging and tumorigenesis, hence it is important to have a quantitative way of measuring each DSB repair pathway. Our laboratory has developed fluorescent reporter constructs that allow sensitive and quantitative measurement of NHEJ and HR. The constructs are based on an engineered GFP gene containing recognition sites for a rare-cutting I-SceI endonuclease for induction of DSBs. The starting constructs are GFP negative as the GFP gene is inactivated by an additional exon, or by mutations. Successful repair of the I-SceI-induced breaks by NHEJ or HR restores the functional GFP gene. The number of GFP positive cells counted by flow cytometry provides quantitative measure of NHEJ or HR efficiency.

Analysis of DNA Double-strand Break DSB Repair in Mammalian Cells

This article describes GFP-based fluorescence in vivo assays that separately quantify homologous recombination and nonhomologous end joining in mammalian cells.

DNA double-strand breaks are the most dangerous DNA lesions that may lead to massive loss of genetic information and cell death.  Cells repair DSBs using ...

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 ...

Measuring Ubiquitylated, SUMOylated, and ADP-Ribosylated DNA-Protein Crosslinks

Quantitative Detection of DNA-Protein Crosslinks and Their Post-Translational Modifications

DNA-protein crosslinks (DPCs) are frequent, ubiquitous, and deleterious DNA lesions, which arise from endogenous DNA damage, enzyme (topoisomerases, methyltransferases, etc.) malfunctioning, or exogenous agents such as chemotherapeutics and crosslinking agents. Once DPCs are induced, several types of post-translational modifications (PTMs) are promptly conjugated to them as early response mechanisms. It has been shown that DPCs can be modified by ubiquitin, small ubiquitin-like modifier (SUMO), and poly-ADP-ribose, which prime the substrates to signal their respective designated repair enzymes and, in some cases, coordinate the repair in sequential manners. As PTMs transpire quickly and are highly reversible, it has been challenging to isolate and detect PTM-conjugated DPCs that usually remain at low levels. Presented here is an immunoassay to purify and quantitatively detect ubiquitylated, SUMOylated, and ADP-ribosylated DPCs (drug-induced topoisomerase DPCs and aldehyde-induced non-specific DPCs) in vivo. This assay is derived from the RADAR (rapid approach to DNA adduct recovery) assay that is used for the isolation of genomic DNA containing DPCs by ethanol precipitation. Following normalization and nuclease digestion, PTMs of DPCs, including ubiquitylation, SUMOylation, and ADP-ribosylation, are detected by immunoblotting using their corresponding antibodies. This robust assay can be utilized to identify and characterize novel molecular mechanisms that repair enzymatic and non-enzymatic DPCs&#160;and has the potential to discover small molecule inhibitors targeting specific factors that regulate PTMs to repair DPCs.

Author Spotlight: Quantitative Detection of DNA Protein Crosslinks and Their Post-Translational Modifications  

The present protocol highlights a modified method to detect and quantify DNA-protein crosslinks (DPCs) and their post-translational modifications (PTMs), including ubiquitylation, SUMOylation, and ADP-ribosylation induced by topoisomerase inhibitors and by formaldehyde, thereby allowing the study of the formation and repair of DPCs and their PTMs.

DNA-protein crosslinks (DPCs) are frequent, ubiquitous, and deleterious DNA lesions, which arise from endogenous DNA damage, enzyme (topoisomerases, ...

High-throughput Measurement of Plasma Membrane Resealing Efficiency in Mammalian Cells

In their physiological environment, mammalian cells are often subjected to mechanical and biochemical stresses that result in plasma membrane damage. In response to these damages, complex molecular machineries rapidly reseal the plasma membrane to restore its barrier function and maintain cell survival. Despite 60 years of research in this field, we still lack a thorough understanding of the cell resealing machinery. With the goal of identifying cellular components that control plasma membrane resealing or drugs that can improve resealing, we have developed a fluorescence-based high-throughput assay that measures the plasma membrane resealing efficiency in mammalian cells cultured in microplates. As a model system for plasma membrane damage, cells are exposed to the bacterial pore-forming toxin listeriolysin O (LLO), which forms large 30-50 nm diameter proteinaceous pores in cholesterol-containing membranes. The use of a temperature-controlled multi-mode microplate reader allows for rapid and sensitive spectrofluorometric measurements in combination with brightfield and fluorescence microscopy imaging of living cells. Kinetic analysis of the fluorescence intensity emitted by a membrane impermeant nucleic acid-binding fluorochrome reflects the extent of membrane wounding and resealing at the cell population level, allowing for the calculation of the cell resealing efficiency. Fluorescence microscopy imaging allows for the enumeration of cells, which constitutively express a fluorescent chimera of the nuclear protein histone 2B, in each well of the microplate to account for potential variations in their number and allows for eventual identification of distinct cell populations. This high-throughput assay is a powerful tool expected to expand our understanding of membrane repair mechanisms via screening for host genes or exogenously added compounds that control plasma membrane resealing.

Here we describe a high-throughput fluorescence-based assay that measures the plasma membrane resealing efficiency through fluorometric and imaging analyses in living cells. This assay can be used for screening drugs or target genes that regulate plasma membrane resealing in mammalian cells.

In their physiological environment, mammalian cells are often subjected to mechanical and biochemical stresses that result in plasma membrane damage. In ...

Immunology and Infection

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 ...

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

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Chapters in this video

Tools to Study the Role of Architectural Protein HMGB1 in the Processing of Helix Distorting, Site-specific DNA Interstrand Crosslinks

Application of Laser Micro-irradiation for Examination of Single and Double Strand Break Repair in Mammalian Cells

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

Detection of Post-Replicative Gaps Accumulation and Repair in Human Cells Using the DNA Fiber Assay

Genetic Studies of Human DNA Repair Proteins Using Yeast as a Model System

Analysis of DNA Double-strand Break (DSB) Repair in Mammalian Cells

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

Quantitative Detection of DNA-Protein Crosslinks and Their Post-Translational Modifications

High-throughput Measurement of Plasma Membrane Resealing Efficiency in Mammalian Cells

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