Name: visualization of dna replication in the vertebrate model system dt40 using the dna fiber technique
Uploaded: 2011-10-27T12:00:00
Duration: 7 min 18 s
Description: Watch this Scientific Journal Video about Visualization of DNA Replication in the Vertebrate Model System DT40 using the DNA Fiber Technique at JoVE.com

We thank Dr T. Helleday and Dr E. Petermann for help with the fiber technique as well as members of the lab for helpful discussion. WN is supported by a Senior International Research Fellowship from the Association for International Cancer Research and by the Polish State Committee for Scientific Research grant (N301 165935).

The method described here was used in the following publication: Schwab et al., EMBO J., 29(4):806-185.
1. DT40 cell culture
Materials: Fetal bovine serum (FBS), chicken serum, penicillin/streptomycin, 2-mercaptoethanol, RPMI
<ol>
<li>Prepare cell culture medium: add 7% FBS, 3% chicken serum, 1x penicillin/streptomycin and 10 &mu;M 2-mercaptoethanol to RPMI medium.</li>
<li>DT40 cells are grown in sterile cell culture flasks at 38 &deg;C and 5% CO2 in a humidified incubator. Split cells every day to a density of 5 x 105 cells/ml6.</li>
</ol>
2. In vivo labeling
Materials: IdU, CldU
<ol>
<li>Prepare stock solutions of nucleotide analogues: dissolve IdU to 5 mM and CldU to 2.5 mM in medium. Heat both solutions briefly to 60 &deg;C and vortex until the nucleotide analogues are dissolved completely.</li>
<li>Add IdU to a final concentration of 25 &mu;M to exponentially growing DT40 cells and mix the cell suspension well. Incubate cells for 20 minutes at 38 &deg;C and 5% CO2.</li>
<li>After incubation with the first label, add CldU to a final concentration of 250 &mu;M and treat cells as described in the previous step.</li>
<li>Wash cells with ice cold PBS and resuspend them at a concentration of 7.5 x 105 cells/ml in cold PBS. Keep the labeled cells on ice.</li>
</ol>
The labeling procedure described is merely a suggestion and can be modified to address specific questions. Please refer to the discussion section for more information on experimental design.
3. Cell lysis and DNA spreading
Materials: Glass slides, fiber lysis solution
<ol>
<li>Prepare the fiber lysis solution: 50 mM EDTA and 0.5% SDS in 200 mM Tris-HCl, pH 7.5</li>
<li>Spot 2 &mu;l of the cell suspension to one end of the glass slide. Air-dry for 5 minutes or until the volume of the drop is greatly reduced but not dry.</li>
<li>Pipet 7 &mu;l lysis solution on top of the cell suspension and gently stir with the pipette tip to mix the solutions. Incubate for 2 minutes for cell lysis to proceed.</li>
<li>Tilt slides to 15&deg; to allow the fibers to spread along the slide.</li>
<li>Once the fiber solution has reached the bottom of the slide, place it horizontally to air-dry. After drying, a thin, opaque line should be visible along the slide. At this point, the beginning of the stretched fibers should be marked with a pencil as after staining the line will not be visible any longer. This mark will later help to locate the fibers under the microscope.</li>
</ol>
4. Immunofluorescence staining
Materials: Methanol, acetic acid, HCl, 5% BSA in PBS, staining jar, anti-BrdU (mouse) antibody, anti-BrdU (rat) antibody, sheep anti-mouse Cy3 antibody, goat anti-rat Alexa Fluor 488 antibody, Vectashield mounting medium, coverslips, nail polish
<ol>
<li>Immerse slides in methanol/acetic acid (3:1) in a staining jar and incubate for 10 minutes.</li>
<li>Wash slides in distilled H2O, then immerse in 2.5 M HCl for 80 minutes.</li>
<li>Wash slides three times in PBS for 5 minutes.</li>
<li>Remove slides from the staining jar and collect excess PBS with a paper towel. Place the slides horizontally and pipet 5% BSA on top of each slide. Cover the slides gently with a coverslip to spread the BSA evenly over the slide and incubate for 20 minutes.</li>
<li>Dilute the primary antibodies in 5% BSA at the following concentrations: 1:25 anti-BrdU (mouse) and 1:400 anti-BrdU (rat).</li>
<li>Move the coverslip gently down the glass slide to remove it. Do not apply force if the cover slip sticks to the slide. The slide can be rehydrated in PBS until the coverslip becomes loose and can be removed with ease. Collect excess BSA with a paper towel and place the slides horizontally. Pipet 50 &mu;l of the primary antibody solution onto each slide. Cover again with a coverslip to spread the antibody solution evenly over the slide and incubate in a humidified chamber for 2 hours.</li>
<li>After removing the coverslips, wash the slides three times in PBS for 5 minutes</li>
<li>Dilute the secondary antibodies in 5% BSA at the following concentrations: 1:500 sheep anti-mouse Cy3 and 1:400 goat anti-rat Alexa Fluor 488.</li>
<li>Apply 50 &mu;l of the secondary antibody solution as described for the primary antibodies. Protect the slides from light and incubate for 1 hour.</li>
<li>After removing the coverslips, wash the slides three times in PBS for 5 minutes.</li>
<li>Add a drop of Vectashield mounting medium onto each slide and apply a coverslip. Gently press the coverslips and remove excess fluid around it with a paper towel. Seal coverslips with transparent nail polish and let them dry. Store the slides at -20 &deg;C.</li>
</ol>
5. Image acquisition
Materials: fluorescence microscope, camera
Place a drop of immersion oil onto a slide close to the pencil mark and start locating the fibers. Typically, there is a main fiber bundle, but these fibers are too entangled and cannot be analyzed later. Move away from the main bundle to find areas where fibers are clearly separated from each other (Fig. 2). We would select pictures only using one color channel in order to avoid bias. Then, we would take approximately 10 pictures of each time point, concentration or cell line. However, the number of pictures depends on how many fibers can be counted on each picture. Move along the slide to take the different pictures, as one area of a slide may not provide representative fiber lengths or replication structures.
6. Data analysis
Materials: Picture analysis program, e.g. ImageJ (<a href="http://rsbweb.nih.gov/ij/" target="_blank">http://rsbweb.nih.gov/ij/</a>) 
Import pictures into a picture analysis program. Measure the lengths of the fiber tracts and/or count the different replication structures (Fig 3). Only count fibers which are clearly visible and which do not extend over the edge of the picture. We typically measure the length of about 100 fibers and count 150-200 replication structures and we would repeat individual experiments at least three times.
7. Representative Results: 
Newly replicated DNA can be visualized as lines of antibody-labeled nucleotide analogues. In our experiments an ongoing fork is represented as adjacent red and green signals (Fig 3). The double labeling protocol also allows us to define four major classes of replication structures: 1) new initiation events can be divided into origins that have fired while the cells were incubated with the first label and origins that have fired during the incubation with the second label. The former consist of neighboring green-red-green signals and the latter of a green line only. 2) termination events manifest as adjoining red-green-red signals. 3) interspersed origins are sites in the genome with closely spaced origins. Such sites consist of consecutive origins and termination signals. 4) depending on the experimental design, stalled/collapsed forks can either be defined as a red only signal7 or a red line followed by a short green tract8,9.
Using the conditions described here, wild type DT40 cells have an average fork speed of 0.4 &mu;m/min. We can detect approximately 63% ongoing forks, 10% origins, 16% stalled forks (red only tracts), 8% terminations and 3% interspersed fibers. 
<img src="/files/ftp_upload/3255/3255fig1.jpg" alt="Figure 1" /> 
Figure 1. (A) The left hand side of the cartoon depicts the first step of the DNA labeling procedure: adding IdU to exponentially growing DT40 cells leads the nucleotide analogue to be readily incorporated into the newly synthesized leading and lagging DNA strands. This can be visualized by using a specific antibody, and will appear as a red line when viewed under the microscope. Subsequently, the same procedure is repeated with CldU as shown on the right side of the figure. After incubation with the second nucleotide analogue, an active fork will be visible as an adjoining red-green tract. The average fiber length is proportional to the incubation time of the nucleotide analogues. (B) DNA from lysed DT40 cells is stretched by gravity onto a microscope slide and the incorporated nucleotide analogues are visualized by use of specific antibodies and fluorescence microscopy. 
<img src="/files/ftp_upload/3255/3255fig2.jpg" alt="Figure 2" /> 
Figure 2. A representative fluorescence image shows fibers from wild type DT40 cells subsequently labeled with IdU and CldU for 20 minutes each. Most of the fibers are well separated from each other and can thus be easily analyzed. The white bar represents 10 &mu;m. 
<img src="/files/ftp_upload/3255/3255fig3.jpg" alt="Figure 3" /> 
Figure 3. The double-labeling fiber technique allows one to distinguish between different replication structures; 1 - actively replicating forks, 2 - new sites of DNA replication (firing of new origins), 3 - fork terminations (two converging forks), 4 - interspersed fibers (sites of closely spaced origins) and 5 - stalled forks (red only signal).

<ol>
	<li>Caldwell, R. B., Fiedler, P., Schoetz, U., Buerstedde, J. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Gene+function+analysis+using+the+chicken+B-cell+line+DT40.">Gene function analysis using the chicken B-cell line DT40.</a> Methods. Mol. Biol. 408, 193-210 (2007).</li><li>Petes, T. D., Williamson, D. H. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Fiber+autoradiography+of+replicating+yeast+DNA.">Fiber autoradiography of replicating yeast DNA.</a> Exp. Cell. Res. 95, 103-110 (1975).</li><li>Takeuchi, F. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Altered+frequency+of+initiation+sites+of+DNA+replication+in+Werner's+syndrome+cells.">Altered frequency of initiation sites of DNA replication in Werner's syndrome cells.</a> Hum. Genet. 60, 365-368 (1982).</li><li>Jackson, D. A., Pombo, A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Replicon+clusters+are+stable+units+of+chromosome+structure:+evidence+that+nuclear+organization+contributes+to+the+efficient+activation+and+propagation+of+S+phase+in+human+cells.">Replicon clusters are stable units of chromosome structure: evidence that nuclear organization contributes to the efficient activation and propagation of S phase in human cells.</a> J. Cell. Biol. 140, 1285-1295 (1998).</li><li>Schwab, R. A., Blackford, A. N., Niedzwiedz, W. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=ATR+activation+and+replication+fork+restart+are+defective+in+FANCM-deficient+cells.">ATR activation and replication fork restart are defective in FANCM-deficient cells.</a> EMBO. J. 29, 806-818 (2010).</li><li>Saribasak, H., Arakawa, H. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Basic+cell+culture+conditions.">Basic cell culture conditions.</a> Subcell. Biochem. 40, 345-346 (2006).</li><li>Conti, C., Seiler, J. A., Pommier, Y. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+mammalian+DNA+replication+elongation+checkpoint:+implication+of+Chk1+and+relationship+with+origin+firing+as+determined+by+single+DNA+molecule+and+single+cell+analyses.">The mammalian DNA replication elongation checkpoint: implication of Chk1 and relationship with origin firing as determined by single DNA molecule and single cell analyses.</a> Cell. Cycle. 6, 2760-2767 (2007).</li><li>Petermann, E., Orta, M. L., Issaeva, N., Schultz, N., Helleday, T. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Hydroxyurea-stalled+replication+forks+become+progressively+inactivated+and+require+two+different+RAD51-mediated+pathways+for+restart+and+repair.">Hydroxyurea-stalled replication forks become progressively inactivated and require two different RAD51-mediated pathways for restart and repair.</a> Mol. Cell. 37, 492-502 (2010).</li><li>Edmunds, C. E., Simpson, L. J., Sale, J. E. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=PCNA+ubiquitination+and+REV1+define+temporally+distinct+mechanisms+for+controlling+translesion+synthesis+in+the+avian+cell+line+DT40.">PCNA ubiquitination and REV1 define temporally distinct mechanisms for controlling translesion synthesis in the avian cell line DT40.</a> Mol. Cell. 30, 519-529 (2008).</li></ol>

No conflicts of interest declared.

We describe a method that allows for quantitative analysis of parameters that influence DNA synthesis during S-phase at a single molecule level. Over the past ten years, different versions of the DNA fiber fluorography technique were developed to visualize the movement of individual replication forks within living cells. The critical parameter in all these techniques is the procedure used to achieve the best possible stretching of DNA on glass surfaces providing well-separated DNA fibers. A crucial step to achieve this is the incubation time of the cell suspension on the microscope slide as well as lysis time. These parameters depend on the temperature and airflow in a particular lab and should be determined empirically. The practical part of a DNA fiber experiment is typically accomplished within one day. However, subsequent picture acquisition and analysis is more time consuming and requires practice.
The labeling protocol can be adapted to answer various scientific problems: using an agent that interferes with replication during the second pulse labeling monitors fork elongation/stalling in response to replicative stress. In this scenario, the first label does not need to be washed out as the second nucleotide is added in excess. Alternatively, drugs that impede replication can be used in combination or just after addition of the first label. This requires the first label and the drug to be removed before the second nucleotide analogue is applied. This procedure allows one to determine the importance of various DNA repair factors on replicative fork stability/recovery under conditions of replicative stress (i.e fork stalling and/or collapse). Noteworthy, a more detailed analysis of particulars of the DNA replication program could be performed with the use of an alternative approaches combining DNA combing and the fluorescence in situ hybridization. This however, is technically more challenging and requires expensive equipment. 
The ability to qualitatively and quantitatively analyze particulars of DNA replication provides an essential tool for investigating defects in DNA replication itself as well as the pathways that suppress genomic instability associated with replication forks. Undoubtedly, this assay will further help to shed light on the mechanisms of replication-mediated DNA repair that allow normal cells to respond/adapt to environmental changes as well as how cancer cells utilize these mechanisms to resist the toxicity of drugs targeting replication forks.

<table border="1">
	<tbody>
		<tr>
 	<th>Name of the reagent</th>
 <th>Company</th>
 <th>Catalogue number</th>
 <th>Comments (optional)</th>
 </tr>
 <tr>
 	<td>Fetal bovine serum gold (FBS)</td>
 <td>PAA</td>
 <td>A15-151</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 	<td>Chicken serum</td>
 <td>Sigma</td>
 <td>C5405</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 	<td>Penicillin/Streptomycin</td>
 <td>PAA</td>
 <td>P11-010</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 	<td>RPMI 1640</td>
 <td>Gibco</td>
 <td>21875</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 	<td>5-Iodo-2'-deoxyuridine (IdU)</td>
 <td>Sigma-Aldrich</td>
 <td>I7125</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 	<td>5-Chloro-2'-deoxy-uridine (CldU)</td>
 <td>Sigma</td>
 <td>C6891</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 	<td>Microscope slides SuperFrost</td>
 <td>VWR</td>
 <td>631-0910</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 	<td>Cover slips No1</td>
 <td>Scientific lab suppplies</td>
 <td>MIC3234</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 	<td>Vectashield mounting medium	</td>
 <td>H-1000 Vector lab</td>
 <td>H-1000</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 	<td>Anti-BrdU antibody (mouse)</td>
 <td>BD Biosciences</td>
 <td>347580</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 	<td>Anti-BrdU antibody (rat)</td>
 <td>Abcam</td>
 <td>ab6326</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 	<td>sheep anti-mouse Cy3</td>
 <td>Sigma</td>
 <td>C2181</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 	<td>goat anti-rat Alexa Fluor 488 antibody</td>
 <td>Invitrogen</td>
 <td>A110060</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 	<td>&nbsp;</td>
 <td>&nbsp;</td>
 <td>&nbsp;</td>
 <td>&nbsp;</td>
 </tr>
 </tbody>
</table>

visualization of dna replication in the vertebrate model system dt40 using the dna fiber technique

Maintenance of replication fork stability is of utmost importance for dividing cells to preserve viability and prevent disease. The processes involved not only ensure faithful genome duplication in the face of endogenous and exogenous DNA damage but also prevent genomic instability, a recognized causative factor in tumor development.
Here, we describe a simple and cost-effective fluorescence microscopy-based method to visualize DNA replication in the avian B-cell line DT40. This cell line provides a powerful tool to investigate protein function in vivo by reverse genetics in vertebrate cells1. DNA fiber fluorography in DT40 cells lacking a specific gene allows one to elucidate the function of this gene product in DNA replication and genome stability. Traditional methods to analyze replication fork dynamics in vertebrate cells rely on measuring the overall rate of DNA synthesis in a population of pulse-labeled cells. This is a quantitative approach and does not allow for qualitative analysis of parameters that influence DNA synthesis. In contrast, the rate of movement of active forks can be followed directly when using the DNA fiber technique2-4. In this approach, nascent DNA is labeled in vivo by incorporation of halogenated nucleotides (Fig 1A). Subsequently, individual fibers are stretched onto a microscope slide, and the labeled DNA replication tracts are stained with specific antibodies and visualized by fluorescence microscopy (Fig 1B). Initiation of replication as well as fork directionality is determined by the consecutive use of two differently modified analogues. Furthermore, the dual-labeling approach allows for quantitative analysis of parameters that influence DNA synthesis during the S-phase, i.e. replication structures such as ongoing and stalled forks, replication origin density as well as fork terminations. Finally, the experimental procedure can be accomplished within a day, and requires only general laboratory equipment and a fluorescence microscope.

Watch this Scientific Journal Video about Visualization of DNA Replication in the Vertebrate Model System DT40 using the DNA Fiber Technique at JoVE.com

Visualization of DNA Replication in the Vertebrate Model System DT40 using the DNA Fiber Technique

DT40, a model vertebrate genetic system, provides a powerful tool to analyze protein function. Here we describe a simple method that allows qualitative analysis of parameters that influence DNA synthesis during the S-phase in DT40 cells at the single molecule level.

Maintenance of replication fork stability is of utmost importance for dividing cells to preserve viability and prevent disease. The processes involved not ...

visualization-dna-replication-vertebrate-model-system-dt40-using-dna

Research

JoVE Journal

Biology

Visualizing Single-molecule DNA Replication with Fluorescence Microscopy

We describe a simple fluorescence microscopy-based real-time method for observing DNA replication at the single-molecule level. A circular, forked DNA template is attached to a functionalized glass coverslip and replicated extensively after introduction of replication proteins and nucleotides (Figure 1). The growing product double-strand DNA (dsDNA) is extended with laminar flow and visualized by using an intercalating dye. Measuring the position of the growing DNA end in real time allows precise determination of replication rate (Figure 2). Furthermore, the length of completed DNA products reports on the processivity of replication. This experiment can be performed very easily and rapidly and requires only a fluorescence microscope with a reasonably sensitive camera.


This protocol demonstrates a simple single-molecule fluorescence microscopy technique for visualizing DNA replication by individual replisomes in real time.

We describe a simple fluorescence microscopy-based real-time method for observing DNA replication at the single-molecule level. A circular, forked DNA ...

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 ...

Visualization of Caenorhabditis elegans Cuticular Structures Using the Lipophilic Vital Dye DiI

The cuticle of C. elegans is a highly resistant structure that surrounds the exterior of the animal1-4. The cuticle not only protects the animal from the environment, but also determines body shape and plays a role in motility4-6. Several layers secreted by epidermal cells comprise the cuticle, including an outermost lipid layer7.
Circumferential ridges in the cuticle called annuli pattern the length of the animal and are present during all stages of development8. Alae are longitudinal ridges that are present during specific stages of development, including L1, dauer, and adult stages2,9. Mutations in genes that affect cuticular collagen organization can alter cuticular structure and animal body morphology5,6,10,11. While cuticular imaging using compound microscopy with DIC optics is possible, current methods that highlight cuticular structures include fluorescent transgene expression12, antibody staining13, and electron microscopy1. Labeled wheat germ agglutinin (WGA) has also been used to visualize cuticular glycoproteins, but is limited in resolving finer cuticular structures14. Staining of cuticular surface using fluorescent dye has been observed, but never characterized in detail15. We present a method to visualize cuticle in live C. elegans using the red fluorescent lipophilic dye DiI (1,1'-dioctadecyl-3,3,3',3'-tetramethylindocarbocyanine perchlorate), which is commonly used in C. elegans to visualize environmentally exposed neurons. This optimized protocol for DiI staining is a simple, robust method for high resolution fluorescent visualization of annuli, alae, vulva, male tail, and hermaphrodite tail spike in C. elegans.

Visualization of Caenorhabditis elegans Cuticular Structures Using the Lipophilic Vital Dye DiI

We present a method to visualize cuticle in live C. elegans using the red fluorescent lipophilic dye DiI (1,1'-dioctadecyl-3,3,3',3'-tetramethylindocarbocyanine perchlorate), which is commonly used in C. elegans to visualize environmentally exposed neurons. With this optimized protocol, alae and annular cuticular structures are stained by DiI and observed using compound microscopy.

The cuticle of C. elegans is a highly resistant structure that surrounds the exterior of the animal1-4. The cuticle not only protects the animal from the ...

Study of the DNA Damage Checkpoint using Xenopus Egg Extracts

On a daily basis, cells are subjected to a variety of endogenous and environmental insults. To combat these insults, cells have evolved DNA damage checkpoint signaling as a surveillance mechanism to sense DNA damage and direct cellular responses to DNA damage. There are several groups of proteins called sensors, transducers and effectors involved in DNA damage checkpoint signaling (Figure 1). In this complex signaling pathway, ATR (ATM and Rad3-related) is one of the major kinases that can respond to DNA damage and replication stress. Activated ATR can phosphorylate its downstream substrates such as Chk1 (Checkpoint kinase 1). Consequently, phosphorylated and activated Chk1 leads to many downstream effects in the DNA damage checkpoint including cell cycle arrest, transcription activation, DNA damage repair, and apoptosis or senescence (Figure 1). When DNA is damaged, failing to activate the DNA damage checkpoint results in unrepaired damage and, subsequently, genomic instability. The study of the DNA damage checkpoint will elucidate how cells maintain genomic integrity and provide a better understanding of how human diseases, such as cancer, develop.
Xenopus laevis egg extracts are emerging as a powerful cell-free extract model system in DNA damage checkpoint research. Low-speed extract (LSE) was initially described by the Masui group1. The addition of demembranated sperm chromatin to LSE results in nuclei formation where DNA is replicated in a semiconservative fashion once per cell cycle.
The ATR/Chk1-mediated checkpoint signaling pathway is triggered by DNA damage or replication stress 2. Two methods are currently used to induce the DNA damage checkpoint: DNA damaging approaches and DNA damage-mimicking structures 3. DNA damage can be induced by ultraviolet (UV) irradiation, &gamma;-irradiation, methyl methanesulfonate (MMS), mitomycin C (MMC), 4-nitroquinoline-1-oxide (4-NQO), or aphidicolin3, 4. MMS is an alkylating agent that inhibits DNA replication and activates the ATR/Chk1-mediated DNA damage checkpoint 4-7. UV irradiation also triggers the ATR/Chk1-dependent DNA damage checkpoint 8. The DNA damage-mimicking structure AT70 is an annealed complex of two oligonucleotides poly-(dA)70 and poly-(dT)70. The AT70 system was developed in Bill Dunphy's laboratory and is widely used to induce ATR/Chk1 checkpoint signaling 9-12.
Here, we describe protocols (1) to prepare cell-free egg extracts (LSE), (2) to treat Xenopus sperm chromatin with two different DNA damaging approaches (MMS and UV), (3) to prepare the DNA damage-mimicking structure AT70, and (4) to trigger the ATR/Chk1-mediated DNA damage checkpoint in LSE with damaged sperm chromatin or a DNA damage-mimicking structure.

Study of the DNA Damage Checkpoint using Xenopus Egg Extracts

Xenopus egg extract is a useful model system to investigate the DNA damage checkpoint. This protocol is for the preparation of Xenopus egg extracts and DNA damage checkpoint inducing reagents. These techniques are adaptable to a variety of DNA damaging approaches in the study of the DNA damage checkpoint signaling.

On a daily basis, cells are subjected to a variety of endogenous and environmental insults. To combat these insults, cells have evolved DNA damage ...

Robust 3D DNA FISH Using Directly Labeled Probes

3D DNA FISH has become a major tool for analyzing three-dimensional organization of the nucleus, and several variations of the technique have been published. In this article we describe a protocol which has been optimized for robustness, reproducibility, and ease of use. Brightly fluorescent directly labeled probes are generated by nick-translation with amino-allyldUTP followed by chemical coupling of the dye. 3D DNA FISH is performed using a freeze-thaw step for cell permeabilization and a heating step for simultaneous denaturation of probe and nuclear DNA. The protocol is applicable to a range of cell types and a variety of probes (BACs, plasmids, fosmids, or Whole Chromosome Paints) and allows for high-throughput automated imaging. With this method we routinely investigate nuclear localization of up to three chromosomal regions.

We describe a robust and versatile protocol for analyzing nuclear architecture by 3D DNA FISH using directly labeled fluorescent probes.

3D DNA FISH has become a major tool for  analyzing three-dimensional organization of the nucleus, and several variations  of the technique have been ...

Using Caenorhabditis elegans as a Model System to Study Protein Homeostasis in a Multicellular Organism

The folding and assembly of proteins is essential for protein function, the long-term health of the cell, and longevity of the organism. Historically, the function and regulation of protein folding was studied in vitro, in isolated tissue culture cells and in unicellular organisms. Recent studies have uncovered links between protein homeostasis (proteostasis), metabolism, development, aging, and temperature-sensing. These findings have led to the development of new tools for monitoring protein folding in the model metazoan organism Caenorhabditis elegans. In our laboratory, we combine behavioral assays, imaging and biochemical approaches using temperature-sensitive or naturally occurring metastable proteins as sensors of the folding environment to monitor protein misfolding. Behavioral assays that are associated with the misfolding of a specific protein provide a simple and powerful readout for protein folding, allowing for the fast screening of genes and conditions that modulate folding. Likewise, such misfolding can be associated with protein mislocalization in the cell. Monitoring protein localization can, therefore, highlight changes in cellular folding capacity occurring in different tissues, at various stages of development and in the face of changing conditions. Finally, using biochemical tools ex vivo, we can directly monitor protein stability and conformation. Thus, by combining behavioral assays, imaging and biochemical techniques, we are able to monitor protein misfolding at the resolution of the organism, the cell, and the protein, respectively.

Using Caenorhabditis elegans as a Model System to Study Protein Homeostasis in a Multicellular Organism

To study the relationship between protein homeostasis, stress and aging, we monitored changes in protein folding by following protein dysfunction, protein localization in the cell and protein stability at the organismal, cellular and protein levels, using the genetically tractable metazoan Caenorhabditis elegans as a model system.

The folding and assembly of proteins is essential for protein function, the long-term health of the cell, and longevity of the organism. Historically, the ...

Real-time Imaging of Single Engineered RNA Transcripts in Living Cells Using Ratiometric Bimolecular Beacons

The growing realization that both the temporal and spatial regulation of gene expression can have important consequences on cell function has led to the development of diverse techniques to visualize individual RNA transcripts in single living cells. One promising technique that has recently been described utilizes an oligonucleotide-based optical probe, ratiometric bimolecular beacon (RBMB), to detect RNA transcripts that were engineered to contain at least four tandem repeats of the RBMB target sequence in the 3&#8217;-untranslated region. RBMBs are specifically designed to emit a bright fluorescent signal upon hybridization to complementary RNA, but otherwise remain quenched. The use of a synthetic probe in this approach allows photostable, red-shifted, and highly emissive organic dyes to be used for imaging. Binding of multiple RBMBs to the engineered RNA transcripts results in discrete fluorescence spots when viewed under a wide-field fluorescent microscope. Consequently, the movement of individual RNA transcripts can be readily visualized in real-time by taking a time series of fluorescent images. Here we describe the preparation and purification of RBMBs, delivery into cells by microporation and live-cell imaging of single RNA transcripts.

Ratiometric bimolecular beacons (RBMBs) can be used to image single engineered RNA transcripts in living cells. Here, we describe the preparation and purification of RBMBs, delivery of RBMBs into cells by microporation and fluorescent imaging of single RNA transcripts in real-time.

The growing realization that both the temporal and spatial regulation of gene expression can have important consequences on cell function has led to the ...

Amplification, Next-generation Sequencing, and Genomic DNA Mapping of Retroviral Integration Sites

Retroviruses exhibit signature integration preferences on both the local and global scales. Here, we present a detailed protocol for (1) generation of diverse libraries of retroviral integration sites using ligation-mediated PCR (LM-PCR) amplification and next-generation sequencing (NGS), (2) mapping the genomic location of each virus-host junction using BEDTools, and (3) analyzing the data for statistical relevance. Genomic DNA extracted from infected cells is fragmented by digestion with restriction enzymes or by sonication. After suitable DNA end-repair, double-stranded linkers are ligated onto the DNA ends, and semi-nested PCR is conducted using primers complementary to both the long terminal repeat (LTR) end of the virus and the ligated linker DNA. The PCR primers carry sequences required for DNA clustering during NGS, negating the requirement for separate adapter ligation. Quality control (QC) is conducted to assess DNA fragment size distribution and adapter DNA incorporation prior to NGS. Sequence output files are filtered for LTR-containing reads, and the sequences defining the LTR and the linker are cropped away. Trimmed host cell sequences are mapped to a reference genome using BLAT and are filtered for minimally 97% identity to a unique point in the reference genome. Unique integration sites are scrutinized for adjacent nucleotide (nt) sequence and distribution relative to various genomic features. Using this protocol, integration site libraries of high complexity can be constructed from genomic DNA in three days. The entire protocol that encompasses exogenous viral infection of susceptible tissue culture cells to integration site analysis can therefore be conducted in approximately one to two weeks. Recent applications of this technology pertain to longitudinal analysis of integration sites from HIV-infected patients.

We describe a protocol for amplifying retroviral integration sites from the genomic DNA of infected cells, sequencing the amplified virus-host junctions, and then mapping these sequences to a reference genome. We also describe techniques to quantify the distribution of integration sites relative to various genomic annotations using BEDTools.

Retroviruses exhibit signature integration preferences on both the local and global scales. Here, we present a detailed protocol for (1) generation of ...

Visualization of Surface-tethered Large DNA Molecules with a Fluorescent Protein DNA Binding Peptide

Large DNA molecules tethered on the functionalized glass surface have been utilized in polymer physics and biochemistry particularly for investigating interactions between DNA and its binding proteins. Here, we report a method that uses fluorescent microscopy for visualizing large DNA molecules tethered on the surface. First, glass coverslips are biotinylated and passivated by coating with biotinylated polyethylene glycol, which specifically binds biotinylated DNA via avidin protein linkers and significantly reduces undesirable binding from non-specific interactions of proteins or DNA molecules on the surface. Second, the DNA molecules are biotinylated by two different methods depending on their terminals. The blunt ended DNA is tagged with biotinylated dUTP at its 3' hydroxyl terminus, by terminal transferase, while the sticky ended DNA is hybridized with biotinylated complimentary oligonucleotides by DNA ligase. Finally, a microfluidic shear flow makes single DNA molecules stretch to their full contour lengths after being stained with fluorescent protein-DNA binding peptide (FP-DBP).

We present an approach for visualizing fluorescent protein DNA binding peptide (FP-DBP)-stained large DNA molecules tethered on the polyethylene glycol (PEG) and avidin-coated glass surface and stretched with microfluidic shear flows.

Large DNA molecules tethered on the functionalized glass surface have been utilized in polymer physics and biochemistry particularly for investigating ...

Filtration Isolation of Nucleic Acids:  A Simple and Rapid DNA Extraction Method

FINA, filtration isolation of nucleic acids, is a novel extraction method which utilizes vertical filtration via a separation membrane and absorbent pad to extract cellular DNA from whole blood in less than 2 min. The blood specimen is treated with detergent, mixed briefly and applied by pipet to the separation membrane. The lysate wicks into the blotting pad due to capillary action, capturing the genomic DNA on the surface of the separation membrane. The extracted DNA is retained on the membrane during a simple wash step wherein PCR inhibitors are wicked into the absorbent blotting pad. The membrane containing the entrapped DNA is then added to the PCR reaction without further purification. This simple method does not require laboratory equipment and can be easily implemented with inexpensive laboratory supplies. Here we describe a protocol for highly sensitive detection and quantitation of HIV-1 proviral DNA from 100 &#181;l whole blood as a model for early infant diagnosis of HIV that could readily be adapted to other genetic targets.

We describe here a simple and rapid paper-based DNA extraction method of HIV proviral DNA from whole blood detected by quantitative PCR. This protocol can be extended for use in detecting other genetic markers or using alternative amplification methods.

FINA, filtration isolation of nucleic acids, is a novel extraction method which utilizes vertical filtration via a separation membrane and absorbent pad ...