We are grateful to Michela de Martino, Wilfred R. Hagen, and Kourosh Honarmand Ebrahimi for useful discussions. This work was supported by start-up funding from the Delft University of Technology.

1. Dps Expression and Purification
Obtaining protein of high purity is an essential first step for the DNA-protection assay. Purification of Dps protein can be performed in 4 to 5 days.
<ol>
	<li>Transform a protease-deficient strain of E. coli (such as BL21(DE3) pLysS) with a pET vector (such as pET17) into which the Dps protein-encoding sequence has been cloned.</li>
	<li>Streak the transformed cells out onto Luria Broth (LB) agar plates containing appropriate antibiotics (such as ampicillin and chloramphenicol for the examples provided in 1.1). Incubate the plates overnight at 37 &deg;C.</li>
	<li>Inoculate a single colony from the plate into 30 ml of LB medium containing appropriate antibiotics. Incubate overnight at 37 &deg;C while shaking at 200-250 rpm.</li>
	<li>Inoculate 2 x 1 L LB medium containing appropriate antibiotics with 10 ml each of the overnight culture. Grow at 37 &deg;C, while shaking, to O.D.600 of approximately 0.6. (Growing cells to higher O.D. increases protein yield, without noticeably compromising the purity of the final product.) Induce expression of Dps by adding IPTG to a concentration of 0.3 mM, and incubate at 37 &deg;C for 3-4 hr while shaking.</li>
	<li>Harvest cells by centrifugation at 6,000 x g for 15 min. Resuspend the cell pellets in 7.5 ml of DEAE buffer A (50 mM HEPES-KOH, pH 7.5, 100 mM NaCl, 0.1 mM EDTA) per L of induced cell culture. Samples may be frozen in liquid nitrogen and stored at -80 &deg;C, if desired. The procedure may then be continued by thawing the samples in a water bath at 4 &deg;C.</li>
	<li>Add a mixture of protease inhibitors (such as Calbiochem set III protease inhibitors, at 0.167 &mu;l per ml of cell suspension) to the cell suspension to prevent degradation of overexpressed Dps.</li>
	<li>Prepare cell-free extract using a French press. Prime the French press with 20 ml DEAE Buffer A (if using a continuous-flow model), then disrupt the sample twice at 20 kpsi. Centrifuge the lysate at 30,000 x g for 35 min at 4 &deg;C to clarify insoluble particles, and save the supernatant. The supernatant may then be frozen using liquid nitrogen and stored at -80 &deg;C, if desired.</li>
	<li>Equilibrate a 30 ml DEAE Sepharose CL-6B column with DEAE Buffer A, using an FPLC. Run buffer through the column at 1-2 ml/min with a maximum pressure of 0.2 MPa over background. Load the cell-free extract onto the column, and begin to collect the flow-through once the O.D.280 signal begins to increase above baseline. Wash the column with DEAE Buffer A until the O.D.280 value returns to baseline, while continuously collecting the flow-through. The flow-through should contain the majority of the Dps protein, free from bound DNA. Wash the column with 100% Buffer B (50 mM HEPES-KOH, pH 7.5, 1 M NaCl, 0.1 mM EDTA). This eluate contains Dps-DNA complexes as well as other proteins and can be discarded.</li>
	<li>Remove additional contaminating proteins through ammonium sulfate precipitation. Determine the volume of pooled flow-through. Slowly (over a period of 10-20 min) add dry small-grain ammonium sulfate to 62% saturation (390 mg per ml of solution) at 4 &deg;C while stirring well. Stir the mixture for an additional 20-30 min after adding the last portion of ammonium sulfate to ensure complete equilibration. Dps will remain soluble, while contaminating proteins will precipitate out of solution.</li>
	<li>Remove precipitated proteins by centrifugation at 20,000 x g for 30 min at 4 &deg;C and save the supernatant.</li>
	<li>Slowly add an additional 227 mg ammonium sulfate per ml of supernatant to reach 94% saturation and stir for 20-30 min to ensure complete equilibration. Dps precipitates at this high salt concentration and can be collected by centrifugation at 20,000 x g for 30 min at 4 &deg;C. Dps will be in the pellet; the supernatant can be discarded. The pellet can be stored at -80 &deg;C if desired.</li>
	<li>Resuspend the Dps-containing pellet in Resuspension Buffer (50 mM HEPES-KOH, pH 7.5, 150 mM NaCl, 0.1 mM EDTA). Adjust the sample volume to 2.5 ml.</li>
	<li>Buffer exchange to remove the ammonium sulfate by using a PD-10 gel filtration column. Equilibrate the gel bed with 25 ml Resuspension Buffer. Apply the 2.5 ml of Dps sample to the top of the PD-10 column and allow it to soak in. Discard the flow-through. Collect Dps by eluting with 3.5 ml Resuspension Buffer.</li>
	<li>Centrifuge the buffer-exchanged sample at 16,000 x g for 10 min at 4 &deg;C to remove any insoluble components, and dilute it with 6-10 ml Dilution Buffer (50 mM HEPES-KOH, pH 7.5, 0.1 mM EDTA) to ensure that the salt concentration is low enough for Dps to bind to the SP Sepharose column. Particularly concentrated samples of Dps may become less soluble upon dilution into a lower-salt buffer, as evidenced by cloudiness of the liquid, but can be completely resolubilized by the addition of a small amount of 5 M NaCl solution (an additional 10-20 mM).</li>
	<li>Equilibrate a 30 ml SP Sepharose Fast Flow column with SP Buffer A (50 mM HEPES-KOH, pH 7.5, 50 mM NaCl, 0.1 mM EDTA). Run buffer through the column at 1.5-2 ml/min with a pressure limit of 0.3 MPa over background. Load the sample onto the column. Wash with SP Buffer A until the O.D.280 trace returns to baseline, and discard the flow-through. Run a 150 ml linear gradient from 0 to 100% Buffer B, while collecting 2 ml fractions. Dps will elute in a sharp peak at around 50% B, though variation by 10% is possible.</li>
	<li>Run the elution fractions on a 15% SDS-PAGE gel, and check for DNA contamination by measuring the O.D.260/ O.D.280 ratio using a spectrophotometer. The gel should only show one band around 19 kDa, and the O.D.260/ O.D.280 is typically around 0.7. Pool the purest Dps fractions. Use a centrifugal filter unit (such as the Amicon Ultra Filtration Unit) with a 10K molecular weight cut-off to concentrate the Dps and exchange it into a storage buffer (50 mM HEPES-KOH, pH 7.5, 50 mM NaCl). Aliquot the purified Dps, and freeze in liquid nitrogen for storage at -80 &deg;C.</li>
</ol>
2. DNA Protection Assay
The assay involves multiple time-sensitive steps and should be timed to the second to obtain reproducible results. Many ingredients and pipetting steps are involved, so a pipetting table (see Table 1) is advisable to keep track of steps and volumes. The total time required for the assay is no more than 3 hr.
<ol>
	<li>Measure 30 ml water into an airtight vial; flush the water with N2 for 10 min (using two syringe needles, one in the fluid, one above it) to remove oxygen from solution. Add 0.0168 g FeSO4&bull;7H2O to the water to obtain a 2 mM stock solution. Seal the vial and flush with nitrogen for 5 min more, then mix. The solution must be clear; if any hint of yellow is detectable, prepare a new iron solution.</li>
	<li>Make a 100 mM H2O2 solution (10.87 &mu;l of 9.2M H2O2 into 1 ml water); keep on ice and shielded from light by wrapping in aluminum foil. Use only for about 1-2 hr after preparation due to spontaneous decay. The hydrogen peroxide stock is similarly subject to breakdown, which may be partially ameliorated by proper storage (in the dark, at 4 &deg;C). Regular checks of the stock concentration by measuring the O.D.240 are recommended.</li>
	<li>Thaw an aliquot of purified Dps rapidly in a room-temperature water bath, and keep on ice. Centrifuge for 10 sec at 4,000 x g in a table top picocentrifuge to precipitate aggregates. Measure concentration after centrifuging using a spectrophotometer (O.D.280=1.547 indicates a concentration of 100 &mu;M monomer Dps).</li>
	<li>Dilute linear DNA from high-concentration stock using 12x reaction buffer (1 M MOPS-KOH pH 7.0, 1 M NaCl) to a concentration of 100 ng/&mu;l. Linear DNA is used to make quantification easier, but plasmid DNA may be used as well.</li>
	<li>Pipette 1 &mu;l of the DNA-buffer mixture into a PCR tube. Add enough water into the reaction so that the final reaction volume (minus SDS) will be 12 &mu;l. This amount should be calculated in advance using the pipetting table. Add Dps to a final dodecamer concentration of 3 &mu;M. Incubate for 15 min at room temperature to allow for Dps to bind to DNA.</li>
	<li>Add enough 2 mM FeSO4 solution to reach the desired concentration. A typical experiment uses a range of concentrations from 0 to 1 mM FeSO4. Higher concentrations will cause more extensive DNA degradation but may also lead to formation of ferric precipitates in the reaction mixture. Quickly add 1 &mu;l of the 100 mM H2O2 solution (to 10 mM final concentration), and incubate at room temperature for 5 min to allow the Fenton-mediated degradation reaction to take place.</li>
	<li>Add 0.8 &mu;l of 20% SDS (sodium dodecyl sulfate). Mix and incubate at 85 &deg;C for 5 min to disrupt the Dps-DNA complexes. The SDS destabilizes the Dps-DNA complex and prevents unwanted gel shifts for the DNA, which improves the ease of quantification. An optional step is to stop the reaction by catalytically degrading the hydrogen peroxide into non-toxic products. For the conditions described in this protocol, this step is not needed to obtain a broad range of DNA degradation.</li>
	<li>Incubate on ice for 1 min, add loading dye, and load on an agarose gel. Run the gel, and stain for DNA using ethidium bromide. The gel should be stained post-electrophoresis rather than prior to electrophoresis, to prevent the SDS from interfering with the ethidium bromide distribution in the gel.</li>
</ol>

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We have nothing to disclose.

The purification process of Dps as described in this article is very robust. Purity has consistently been high (&gt; 99%); no other proteins appear on SDS-PAGE gels as visible bands. Despite this, some batches of purified Dps appear to have nuclease activity, as evidenced by partial DNA degradation when incubated with very high concentrations of Dps. This might indicate the presence of highly active DNases at low concentration that we were unable to remove through purification. However, this DNA degradation is only obser...

Aerobic organisms must constantly contend with reactive oxygen species that can damage their DNA as well as other crucial biological macromolecules. One potent tool to counteract the toxic effects of oxidative damage is the DNA-binding protein from starved cells (Dps). Since its discovery in 1992 from starved E. coli culture 1, Dps has been identified in more than 300 species of bacteria and archaebacteria 2. Massive upregulation of Dps during stationary phase makes it the most highly expressed nucleoid-associated protein of E. coli under starvation conditions 3, 4. Additionally, Dps has been shown to preserve both bacterial viability and DNA integrity during many various stresses, including starvation, high iron concentration, UV light exposure, heat shock, and oxidative stress 5, 6.
Structurally, Dps self-associates into a stable homo-oligomeric complex of 12 monomers, which assemble into a spherical hollow shell. The ~4.5 nm-wide internal cavity is accessible to the exterior solvent via pores that allow the passage of small molecules 7, and it can sequester mineralized metals such as iron 8. The protective effect of Dps derives from its several biochemical activities, which include non-specific DNA binding 1, ferroxidase activity, and iron storage 8.
Detailed study of the beneficial biochemical activities of Dps first requires its purification. Dps purification is an elaborate procedure, as Dps must be separated not only from other proteins, but also from any bound DNA 7. Our optimized purification process uses many common techniques, consisting of two ion-exchange columns and an ammonium sulfate precipitation step. Several buffer exchanges are needed, as highly concentrated Dps can precipitate out of solution in low salt conditions. Once Dps protein has been purified, it may be applied to assays that directly measure its ferroxidase activity 8, DNA-binding stoichiometry 9, and mechanisms of iron binding 10. Purified Dps also has other potential applications. The stable hollow spherical structure of Dps has been used as scaffolding for storing hydrophobic particles inside the protein cavity 11 and even as a reaction chamber to synthesize novel magnetic nanoparticles 12.
The protective ability of Dps to mediate damage due to reactive oxygen species can be clearly and directly demonstrated using the DNA protection assay 13, 14. In this in vitro procedure, radical species are produced when iron catalyzes H2O2 degradation through Fenton chemistry. These radicals directly damage DNA present in the reaction and can completely degrade it at high concentrations. Two key Dps activities may both directly counteract the effects of Fenton-mediated radical production. Dps lowers the concentration of catalytic iron through mineralization, consuming the available hydrogen peroxide in the process. Additionally, Dps binding to DNA may potentially shield it physically from radical damage and condenses it into a smaller volume with less reactive surface area. The combination of these two properties makes the DNA protection assay well suited for the purpose of measuring protective Dps activity.
The DNA protection assay is quite versatile and can be used for a variety of applications beyond Dps characterization. Radical damage is a common form of stress in cells, and many different proteins and chemicals are used to counteract it. The general principle of the assay, using DNA integrity as a marker for radical damage, can be used in combination with almost any radical-producing reaction or counteracting agent. Among others, the assay has been successfully used to determine anti-oxidative properties of K. paniculata extract for use in food industry 15, to characterize the effects of uric acid on hydroxyl damage mediation 16, and to gain new insights into the function of Fur transcriptional regulator proteins 17.
Despite the numerous uses of the assay in published papers, we found that many optimization and troubleshooting steps were required, which makes setting up the assay for the first time an unnecessarily laborious process for many researchers. The protocol we present in this article aims to remove this barrier for entry.

<table><tbody><tr><td>BL21(DE3)pLysS competent cells</td><td>Promega</td><td>L1195</td><td></td></tr><tr><td>pET-17b DNA</td><td>EMD-Millipore</td><td>69663-3</td><td></td></tr><tr><td>LB broth powder</td><td>Sigma-Aldrich</td><td>L3022</td><td></td></tr><tr><td>Ampicillin sodium salt</td><td>Sigma-Aldrich</td><td>A0166</td><td></td></tr><tr><td>Chloramphenicol</td><td>Sigma-Aldrich</td><td>C0378</td><td></td></tr><tr><td>IPTG</td><td>Sigma-Aldrich</td><td>I6758</td><td></td></tr><tr><td>HEPES</td><td>BDH</td><td>441476L</td><td></td></tr><tr><td>Potassium hydroxide</td><td>Merck</td><td>105033</td><td></td></tr><tr><td>Sodium chloride</td><td>VWR</td><td>443824T</td><td></td></tr><tr><td>EDTA</td><td>Sigma-Aldrich</td><td>E9884</td><td></td></tr><tr><td>Protease Inhibitor Cocktail Set III</td><td>EMD-Millipore</td><td>539134</td><td></td></tr><tr><td>DEAE-Sepharose</td><td>Sigma-Aldrich</td><td>DFF100</td><td></td></tr><tr><td>Ammonium sulphate</td><td>Sigma-Aldrich</td><td>A4418</td><td></td></tr><tr><td>PD-10 Desalting Columns</td><td>GE Healthcare LS</td><td>17-0851-01</td><td></td></tr><tr><td>SP-Sepharose</td><td>Sigma-Aldrich</td><td>S1799</td><td></td></tr><tr><td>Amicon Ultra Centr. Filter (10K MWCO)</td><td>Millipore</td><td>UFC901024</td><td></td></tr><tr><td>Ferrous Sulphate heptahydrate</td><td>Sigma-Aldrich</td><td>F8048</td><td></td></tr><tr><td>Hydrogen peroxide solution</td><td>Sigma-Aldrich</td><td>216763</td><td></td></tr><tr><td>MOPS</td><td>Calbiochem</td><td>475898</td><td></td></tr><tr><td>SDS solution</td><td>Bio-Rad</td><td>161-0418</td><td></td></tr><tr><td>Ethidium bromide</td><td>Sigma-Aldrich</td><td>E1510</td><td></td></tr><tr><td>Static incubator</td><td>Hettich</td><td>INE500</td><td></td></tr><tr><td>Shaking Incubator </td><td>New Brunswick Sc.</td><td>Inova 44</td><td></td></tr><tr><td>Cooled centrifuge</td><td>Beckman Coulter</td><td>Avanti J-E</td><td></td></tr><tr><td>Table-top centrifuge</td><td>Eppendorf</td><td>5424</td><td></td></tr><tr><td>Cell disrupter</td><td>Constant Systems Ltd.</td><td>TS2/40/AA/AA</td><td></td></tr><tr><td>FPLC Purifier</td><td>General Electric</td><td>AKTA</td><td></td></tr><tr><td>Airtight vials</td><td>Cole-Parmer</td><td>EW-08918-85</td><td></td></tr><tr><td>Syringe needles</td><td>BD</td><td>305128</td><td></td></tr><tr><td>Pipettes</td><td>Eppendorf</td><td>Z683779-1EA, Z683795-1EA</td><td></td></tr></tbody></table>

application of an in vitro dna protection assay to visualize stress mediation properties of the dps protein

Oxidative stress is an unavoidable byproduct of aerobic life. Molecular oxygen is essential for terrestrial metabolism, but it also takes part in many damaging reactions within living organisms. The combination of aerobic metabolism and iron, which is another vital compound for life, is enough to produce radicals through Fenton chemistry and degrade cellular components. DNA degradation is arguably the most damaging process involving intracellular radicals, as DNA repair is far from trivial. The assay presented in this article offers a quantitative technique to measure and visualize the effect of molecules and enzymes on radical-mediated DNA damage.
The DNA protection assay is a simple, quick, and robust tool for the in vitro characterization of the protective properties of proteins or chemicals. It involves exposing DNA to a damaging oxidative reaction and adding varying concentrations of the compound of interest. The reduction or increase of DNA damage as a function of compound concentration is then visualized using gel electrophoresis. In this article we demonstrate the technique of the DNA protection assay by measuring the protective properties of the DNA-binding protein from starved cells (Dps). Dps is a mini-ferritin that is utilized by more than 300 bacterial species to powerfully combat environmental stressors. Here we present the Dps purification protocol and the optimized assay conditions for evaluating DNA protection by Dps.

The purification process for Dps described here is very reproducible. Dps purification according to the described protocol, using 2 L of E. coli culture as a starting point, will typically yield 2.5 ml of protein containing Dps12 at concentrations between 5 and 12 &mu;M. Longer induction times (4 hr) seem to reduce this variability. Protein purity is consistently above 99%, as evidenced by SDS-PAGE gels (Figure 1). The level of DNA contamination is consistently negligible, as evidence...

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Application of an In vitro DNA Protection Assay to Visualize Stress Mediation Properties of the Dps Protein

The DNA-binding protein from starved cells (Dps) plays a crucial role in combating bacterial stress. This article discusses the purification of E. coli Dps and the protocol for an in vitro assay demonstrating Dps-mediated protection of DNA from degradation by reactive oxygen species.

Application of an In vitro DNA Protection Assay to Visualize Stress Mediation Properties of the Dps Protein

Oxidative stress is an unavoidable byproduct of  aerobic life.  Molecular oxygen is  essential for terrestrial metabolism, but it also takes part in many ...

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Research

JoVE Journal

Biology

11.1K Views.  Delft University of Technology. De DNA-bindend eiwit van uitgehongerde cellen (DPS) speelt een cruciale rol in de strijd tegen bacteriële stress. Dit artikel bespreekt de zuivering van E. coli Dps en het protocol voor een In vitro Assay tonen Dps-gemedieerde bescherming van DNA tegen afbraak door reactieve zuurstof species.

Video: Application of an In vitro DNA Protection Assay to Visualize Stress Mediation Properties of the Dps Protein

Single Molecule Analysis of Laser Localized Psoralen Adducts

The DNA Damage Response (DDR) has been extensively characterized in studies of double strand breaks (DSBs) induced by laser micro beam irradiation in live cells. The DDR to helix distorting covalent DNA modifications, including interstrand DNA crosslinks (ICLs), is not as well defined. We have studied the DDR stimulated by ICLs, localized by laser photoactivation of immunotagged psoralens, in the nuclei of live cells. In order to address fundamental questions about adduct distribution and replication fork encounters, we combined laser localization with two other technologies. DNA fibers are often used to display the progress of replication forks by immunofluorescence of nucleoside analogues incorporated during short pulses. Immunoquantum dots have been widely employed for single molecule imaging. In the new approach, DNA fibers from cells carrying laser localized ICLs are spread onto microscope slides. The tagged ICLs are displayed with immunoquantum dots and the inter-lesion distances determined. Replication fork collisions with ICLs can be visualized and different encounter patterns identified and quantitated.

Lasers are frequently used in studies of the cellular response to DNA damage. However, they generate lesions whose spacing, frequency, and collisions with replication forks are rarely characterized. Here, we describe an approach that enables the determination of these parameters with laser localized interstrand crosslinks.

The DNA Damage Response (DDR) has been extensively characterized in studies of double strand breaks (DSBs) induced by laser micro beam irradiation in live ...

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 ...

Biochemistry

Investigation of Protein Recruitment to DNA Lesions Using 405 Nm Laser Micro-irradiation

The DNA Damage Response (DDR) uses a plethora of proteins to detect, signal, and repair DNA lesions. Delineating this response is critical to understand genome maintenance mechanisms. Since recruitment and exchange of proteins at lesions are highly dynamic, their study requires the ability to generate DNA damage in a rapid and spatially-delimited manner. Here, we describe procedures to locally induce DNA damage in human cells using a commonly available laser-scanning confocal microscope equipped with a 405 nm laser line. Accumulation of genome maintenance factors at laser stripes can be assessed by immunofluorescence (IF) or in real-time using proteins tagged with fluorescent reporters. Using phosphorylated histone H2A.X (&#947;-H2A.X) and Replication Protein A (RPA) as markers, the method provides sufficient resolution to discriminate locally-recruited factors from those that spread on adjacent chromatin. We further provide ImageJ-based scripts to efficiently monitor the kinetics of protein relocalization at DNA damage sites. These refinements greatly simplify the study of the DDR dynamics.

Studying DNA damage repair kinetics requires a system to induce lesions at defined sub-nuclear regions. We describe a method to create localized double-stranded breaks using a laser-scanning confocal microscope equipped with a 405 nm laser and provide automated procedures to quantify the dynamics of repair factors at these lesions.

The DNA Damage Response (DDR) uses a plethora of proteins to detect, signal, and repair DNA lesions. Delineating this response is critical to understand ...

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 ...

Isolation of Labile Multi-protein Complexes by in vivo Controlled Cellular Cross-Linking and Immuno-magnetic Affinity Chromatography

The dynamic nature of cellular machineries is frequently built on transient and/or weak protein associations. These low affinity interactions preclude stringent methods for the isolation and identification of protein networks around a protein of interest. The use of chemical crosslinkers allows the selective stabilization of labile interactions, thus bypassing biochemical limitations for purification. Here we present a protocol amenable for cells in culture that uses a homobifunctional crosslinker with a spacer arm of 12 &Aring;, dithiobis-(succinimidyl proprionate) (DSP). DSP is cleaved by reduction of a disulphide bond present in the molecule. Cross-linking combined with immunoaffinity chromatography of proteins of interest with magnetic beads allows the isolation of protein complexes that otherwise would not withstand purification. This protocol is compatible with regular western blot techniques and it can be scaled up for protein identification by mass spectrometry1.
Stephanie A. Zlatic and Pearl V. Ryder contributed equally to this work.

Isolation of Labile Multi-protein Complexes by in vivo Controlled Cellular Cross-Linking and Immuno-magnetic Affinity Chromatography

The cell permeable crosslinker DSP [dithiobis-(succinimidyl propionate)] stabilizes transient and labile interactions in vivo, which allows their isolation using stringent protein complex purification techniques. Here we present a technique for crosslinking cells grown in culture followed by isolation of protein complexes by immunoprecipitation.

The dynamic nature of cellular machineries is frequently built on transient and/or weak protein associations. These low affinity interactions preclude ...

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, ...

Two- and Three-Dimensional Live Cell Imaging of DNA Damage Response Proteins

Double-strand breaks (DSBs) are the most deleterious DNA lesions a cell can encounter. If left unrepaired, DSBs harbor great potential to generate mutations and chromosomal aberrations1. To prevent this trauma from catalyzing genomic instability, it is crucial for cells to detect DSBs, activate the DNA damage response (DDR), and repair the DNA. When stimulated, the DDR works to preserve genomic integrity by triggering cell cycle arrest to allow for repair to take place or force the cell to undergo apoptosis. The predominant mechanisms of DSB repair occur through nonhomologous end-joining (NHEJ) and homologous recombination repair (HRR) (reviewed in2). There are many proteins whose activities must be precisely orchestrated for the DDR to function properly. Herein, we describe a method for 2- and 3-dimensional (D) visualization of one of these proteins, 53BP1.
The p53-binding protein 1 (53BP1) localizes to areas of DSBs by binding to modified histones3,4, forming foci within 5-15 minutes5. The histone modifications and recruitment of 53BP1 and other DDR proteins to DSB sites are believed to facilitate the structural rearrangement of chromatin around areas of damage and contribute to DNA repair6. Beyond direct participation in repair, additional roles have been described for 53BP1 in the DDR, such as regulating an intra-S checkpoint, a G2/M checkpoint, and activating downstream DDR proteins7-9. Recently, it was discovered that 53BP1 does not form foci in response to DNA damage induced during mitosis, instead waiting for cells to enter G1 before localizing to the vicinity of DSBs6. DDR proteins such as 53BP1 have been found to associate with mitotic structures (such as kinetochores) during the progression through mitosis10.
In this protocol we describe the use of 2- and 3-D live cell imaging to visualize the formation of 53BP1 foci in response to the DNA damaging agent camptothecin (CPT), as well as 53BP1's behavior during mitosis. Camptothecin is a topoisomerase I inhibitor that primarily causes DSBs during DNA replication. To accomplish this, we used a previously described 53BP1-mCherry fluorescent fusion protein construct consisting of a 53BP1 protein domain able to bind DSBs11. In addition, we used a histone H2B-GFP fluorescent fusion protein construct able to monitor chromatin dynamics throughout the cell cycle but in particular during mitosis12. Live cell imaging in multiple dimensions is an excellent tool to deepen our understanding of the function of DDR proteins in eukaryotic cells.

This protocol describes a method for visualizing a DNA double-strand break signaling protein activated in response to DNA damage as well as its localization during mitosis.

Double-strand breaks (DSBs) are the most deleterious DNA lesions a cell can encounter. If left unrepaired, DSBs harbor great potential to generate ...

Genome-wide Protein-protein Interaction Screening by Protein-fragment Complementation Assay (PCA) in Living Cells

Proteins are the building blocks, effectors and signal mediators of cellular processes. A protein&#8217;s function, regulation and localization often depend on its interactions with other proteins. Here, we describe a protocol for the yeast protein-fragment complementation assay (PCA), a powerful method to detect direct and proximal associations between proteins in living cells. The interaction between two proteins, each fused to a dihydrofolate reductase (DHFR) protein fragment, translates into growth of yeast strains in presence of the drug methotrexate (MTX). Differential fitness, resulting from different amounts of reconstituted DHFR enzyme, can be quantified on high-density colony arrays, allowing to differentiate interacting from non-interacting bait-prey pairs. The high-throughput protocol presented here is performed using a robotic platform that parallelizes mating of bait and prey strains carrying complementary DHFR-fragment fusion proteins and the survival assay on MTX. This protocol allows to systematically test for thousands of protein-protein interactions (PPIs) involving bait proteins of interest and offers several advantages over other PPI detection assays, including the study of proteins expressed from their endogenous promoters without the need for modifying protein localization and for the assembly of complex reporter constructs.

Genome-wide Protein-protein Interaction Screening by Protein-fragment Complementation Assay PCA in Living Cells

Proteins interact with each other and these interactions determine in a large part their functions. Protein interaction partners can be identified at high-throughput in vivo using a yeast fitness assay based on the dihydrofolate reductase protein-fragment complementation assay (DHFR-PCA).

Proteins are the building blocks, effectors and signal mediators of cellular processes. A protein&#8217;s function, regulation and localization often ...

Screening for Functional Non-coding Genetic Variants Using Electrophoretic Mobility Shift Assay (EMSA) and DNA-affinity Precipitation Assay (DAPA)

Population and family-based genetic studies typically result in the identification of genetic variants that are statistically associated with a clinical disease or phenotype. For many diseases and traits, most variants are non-coding, and are thus likely to act by impacting subtle, comparatively hard to predict mechanisms controlling gene expression. Here, we describe a general strategic approach to prioritize non-coding variants, and screen them for their function. This approach involves computational prioritization using functional genomic databases followed by experimental analysis of differential binding of transcription factors (TFs) to risk and non-risk alleles. For both electrophoretic mobility shift assay (EMSA) and DNA affinity precipitation assay (DAPA) analysis of genetic variants, a synthetic DNA oligonucleotide (oligo) is used to identify factors in the nuclear lysate of disease or phenotype-relevant cells. For EMSA, the oligonucleotides with or without bound nuclear factors (often TFs) are analyzed by non-denaturing electrophoresis on a tris-borate-EDTA (TBE) polyacrylamide gel. For DAPA, the oligonucleotides are bound to a magnetic column and the nuclear factors that specifically bind the DNA sequence are eluted and analyzed through mass spectrometry or with a reducing sodium dodecyl sulfate polyacrylamide gel electrophoresis (SDS-PAGE) followed by Western blot analysis. This general approach can be widely used to study the function of non-coding genetic variants associated with any disease, trait, or phenotype.

Screening for Functional Non-coding Genetic Variants Using Electrophoretic Mobility Shift Assay EMSA and DNA-affinity Precipitation Assay DAPA

We present a strategic plan and protocol for identifying non-coding genetic variants affecting transcription factor (TF) DNA binding. A detailed experimental protocol is provided for electrophoretic mobility shift assay (EMSA) and DNA affinity precipitation assay (DAPA) analysis of genotype-dependent TF DNA binding.

Population and family-based genetic studies typically result in the identification of genetic variants that are statistically associated with a clinical ...

Studying the Protein Quality Control System of D. discoideum Using Temperature-controlled Live Cell Imaging

The complex lifestyle of the social amoebae Dictyostelium discoideum makes it a valuable model for the study of various biological processes. Recently, we showed that D. discoideum is remarkably resilient to protein aggregation and can be used to gain insights into the cellular protein quality control system. However, the use of D. discoideum as a model system poses several challenges to microscopy-based experimental approaches, such as the high motility of the cells and their susceptibility to photo-toxicity. The latter proves to be especially challenging when studying protein homeostasis, as the phototoxic effects can induce a cellular stress response and thus alter to behavior of the protein quality control system. 
Temperature increase is a commonly used way to induce cellular stress. Here, we describe a temperature-controllable imaging protocol, which allows observing temperature-induced perturbations in D. discoideum. Moreover, when applied at normal growth temperature, this imaging protocol can also noticeably reduce photo-toxicity, thus allowing imaging with higher intensities. This can be particularly useful when imaging proteins with very low expression levels. Moreover, the high mobility of the cells often requires the acquisition of multiple fields of view to follow individual cells, and the number of fields needs to be balanced against the desired time interval and exposure time.

Studying the Protein Quality Control System of D. discoideum Using Temperature-controlled Live Cell Imaging

The social amoebae Dictyostelium discoideum has recently been established as a system to study protein misfolding and proteostasis. Here, we describe a new imaging-based methodology to study temperature-induced protein aggregation and the cellular stress response in D. discoideum.

The complex lifestyle of the social amoebae Dictyostelium discoideum makes it a valuable model for the study of various biological processes. Recently, we ...

Application of an In vitro DNA Protection Assay to Visualize Stress Mediation Properties of the Dps Protein

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