Research reported in this manuscript was supported by the National Institute of General Medical Sciences of the National Institutes of Health (P20GM130448) (NAW and RP); National Cancer Institute of the National Institutes of Health (NCI R15 CA242057 01A1); Johnson Cancer Research Center in Kansas State University; and the U.S. Department of Defense (CMDRP PRCRP CA160224 (NAW)). We appreciate KSU-CVM Confocal Core and Joel Sanneman for our immunofluorescence microscopy. The content is solely the responsibility of the authors and does not necessarily represent the official views of these funding agencies.

1. Cell plating
<ol>
	<li>Grow HFK LXSN and HFK 8E6 cells in 10 cm plates in keratinocyte culture media (10 mL/plate) with human keratinocyte growth supplement (HKGS) and 1% penicillin/streptomycin. Grow cells to about 80% confluence at 37 &#176;C in a jacket incubator with 5% CO2.</li>
	<li>Replace culture media with 3 mL of trypsin-EDTA (0.05%, ethylenediamine tetraacetic acid). Incubate at 37 &#176;C for 3 min. Neutralize trypsin with equal volume of fetal bovine serum (FBS) supplemented media and transfer cells to a 15 mL centrifuge tube. Centrifuge at 300 x g for 5 min.</li>
	<li>Resuspend cells with 10 mL of keratinocyte culture media with HKGS. Determine the concentration of cells with a hemocytometer.</li>
	<li>Plate 4 x 105 cells/6-cm plates (seed two plates for HFK LXSN and two plates for HFK 8E6) in 4mL of keratinocyte culture media with HKGS and 1% penicillin and streptomycin. Grow at 37 &#176;C in a jacket incubator with 5% CO2. 
	NOTE: Analysis of HFK cells was chosen for this protocol for two reasons. First, HFK is&#160;a difficult to transfect cell line. Thus, by demonstrating that the protocol works in this cell line, evidence is provided that it will likely work in more commonly used and more readily transfected cells. Secondly, previously published data demonstrate that a viral protein (8E6) hinders the repair of double strand breaks in DNA20,21,22. Thus, comparing HFK LXSN and HFK 8E6 allows us to demonstrate the ability of the assay to detect increases in mutations associated with a reduction in cellular repair capacity.</li>
</ol>
2. Transfection
<ol>
	<li>On the day of transfection (24 h after plating), replace media with 3 mL of antibiotic-free supplemented media. Incubate for 2 h at 37 &#176;C in a jacket incubator.</li>
	<li>Transfect cells with appropriate lipid-based transfection reagents according to the manufacturer&#39;s instructions.
	<ol>
		<li>Warm transfection reagents to room temperature and pipette gently before using.</li>
		<li>For each cell line (HFK LXSN and HFK 8E6), place appropriate amount of transfection buffer (as directed by the manufacturer) in a sterile 1.5 mL centrifuge tube (Tube 1). Include another tube with the same amount of transfection buffer (mock transfection or Tube 2).</li>
		<li>Add 2 &#181;g of plasmid DNA expressing CAS9/sgRNA targeting human CD4 (px330-CD4, 5&#39;- GGCGTATCTGTGTGAGGACT) to Tube 1 from step 2.2.2. Pipette gently to mix completely. Add equal volume of sterile water to Tube 2.</li>
		<li>Include a control plate with transfection reagents alone (no plasmid) for each cell line. 
		NOTE: The second plate serves as a negative control in the experiment, allowing the user to confirm that transfection with the CAS9/sgRNA are not responsible for any mutations.</li>
		<li>Add appropriate amount of the transfection reagent (as directed by the manufacturer) to the tube with DNA mixture (Tube 1) from step 2.2.3 and the mock transfection (Tube 2) from step 2.2.2. Pipette gently to mix completely. Incubate at room temperature for 15-30 min to allow sufficient time for complexes to form.</li>
		<li>Add the transfection mixture drop-wise to the plate. Gently rock the culture for 1 min to evenly distribute the transfection mixture.</li>
	</ol>
	</li>
	<li>Incubate for 48 h after the transfection to allow CAS9 expression.</li>
	<li>Harvest cells by trypsinization.
	<ol>
		<li>Replace culture media with 1 mL of trypsin-EDTA (0.05%, ethylenediamine tetraacetic acid). Incubate at 37 &#176;C for 3 min. Neutralize trypsin with equal volume of FBS supplemented media.</li>
		<li>For each plate of cells, transfer the cell suspension to two microcentrifuge tubes with equal aliquots. Centrifuge at 300 x g for 5 min.</li>
		<li>Resuspend the cell pellet from one tube in step 2.4.2 in 1 mL of phosphate buffered saline (PBS) for sequencing. Resuspend the other tube from step 2.4.2 with ice-cold PBS for immunoblot.</li>
	</ol>
	</li>
	<li>Harvest the whole cell lysates for immunoblot.
	<ol>
		<li>Centrifuge the tube at 300 x g for 5 min. Discard the supernatant.</li>
		<li>Add 100 &#181;L radioimmunoprecipitation assay buffer (RIPA lysis buffer) mixed with 1 % protease inhibitor and 1% phosphatase inhibitor into the tube, mix thoroughly with a pipette and incubate for 10 min on ice. 
		NOTE: RIPA lysis buffer contains 10 mM Tris-HCl, pH 8.0; 1 mM EDTA; 0.5 mM EGTA; 1% Triton X-100; 0.1% Sodium Deoxycholate; 0.1% SDS; 140 mM NaCl, and deionized water.</li>
		<li>Centrifuge lysates at 13,000 x g for 10 min. Collect supernatants for immunoblot.</li>
	</ol>
	</li>
</ol>
3. Measuring CAS9 expression via immunoblot
<ol>
	<li>Determine the protein concentration with a bicinchoninic acid (BCA) assay according to the manufacturer&#39;s instructions.</li>
	<li>Run 20 &#181;g of protein of each sample in the wells of a 3%-8% Tris-acetate gel for 150 min and semidry transfer (10 V for 30 min and then 25 V for 12 min) to polyvinylidene difluoride membrane.</li>
	<li>After blocking the membrane in 5% nonfat dry milk in PBS with 0.1% tween (PBST) for 1 h at room temperature, add anti-CAS9 (1:1000) and anti-GAPDH (1:1000) antibodies. Incubate at 4 &#176;C overnight.</li>
	<li>After washing the membrane with PBST, incubate the membrane with secondary antibody in 5% nonfat dry milk in PBST for 1 h at room temperature.</li>
	<li>Image the blot and determine CAS9 level by densitometry23. See Figure 1 for a representative blot. 
	NOTE: Detecting phosphorylated H2AX (S139) foci formation by immunofluorescence microscopy can be used to validate CAS9 activity14. A low number of distinct foci (typically 1-4 foci) are expected depending on the cell cycle position, whether mutations in the CAS9 target site prevent further cutting, and how many copies of the CAS9 cutting site exist in the genome of interest. A representative image is shown in Figure 2.</li>
</ol>
4. Nucleic acid extraction and amplicon generation
<ol>
	<li>Extract DNA from cell samples from step 2.4.3 using a high-molecular weight DNA extraction kit, as specified by the manufacturer.</li>
	<li>Resuspend primers with indicated solvent according to the datasheet. Dilute with the same reagent to 20 &#181;M and pool 20 &#181;M primers into the indicated pools. 
	NOTE: Primer pool is listed in Supplemental Table 1.</li>
	<li>Create a PCR Master mix using a long amplification Taq polymerase for each 20 &#181;M primer pool as specified in Table 1.
	<ol>
		<li>Add 21 &#181;L of the mastermixes to separate PCR tubes.</li>
	</ol>
	</li>
	<li>Add 4 &#181;L of the target sample (100 ng/&#181;L) from step 4.1 to PCR assay tubes containing mastermix and cap assay tubes. Ensure separate reactions for each primer pool.
	<ol>
		<li>Vortex to mix PCR assay tubes and centrifuge (quick spin) to remove droplets from tube lids.</li>
		<li>Place the PCR tubes on a conventional thermal cycler machine.</li>
		<li>Program PCR machine as specified in Table 2.</li>
		<li>Run the program on a thermal cycler.</li>
	</ol>
	</li>
</ol>
5. PCR clean-up
<ol>
	<li>Remove primers from PCR reactions using a bead-based PCR cleanup system.
	<ol>
		<li>Remove clean-up beads from the refrigerator 30 min prior to use.</li>
		<li>Vortex beads well prior to use and ensure all beads are resuspended.</li>
		<li>Add 30 &#181;L (1.2x) of resuspended beads to each well of a deep well 96-well plate.</li>
		<li>Add 25 &#181;L of PCR reaction to wells containing beads.</li>
		<li>Place the plate on a plate shaker at 2000 rpm for 2 min.</li>
		<li>Allow the plate to remain at room temperature for 5 min following shaking.</li>
		<li>Place the deep well plate on a 96-well plate magnet and incubate for 2 min.</li>
		<li>Remove and discard the supernatant without the disturbing beads.</li>
		<li>While the plate remains on the magnet, add 180 &#181;L of 80% ethanol and incubate for 30 s. Remove and discard the supernatant.</li>
		<li>Repeat step 5.1.9.</li>
		<li>Using a 10 &#181;L pipette, remove and discard any remaining liquid from the wells.</li>
		<li>Allow beads to dry at room temperature for 10 min.</li>
		<li>Add 20 &#181;L of nuclease free water to the wells containing beads and remove the plate from the magnet.</li>
		<li>Shake the plate at 2000 rpm for 2 min at room temperature.</li>
		<li>Incubate the plate at room temperature for 5 min.</li>
		<li>Place the plate on a magnet stand and incubate for 2 min at room temperature.</li>
		<li>Remove the supernatant into a second, labeled PCR plate. This contains the cleaned-up DNA.</li>
	</ol>
	</li>
	<li>Measure the concentration of each reaction with a Fluorometer.
	<ol>
		<li>Ensure that dsDNA Fluorometer reagents are at room temperature.</li>
		<li>Set up Fluorometric assay tubes plus two additional tubes for standards.</li>
		<li>Add 199 &#181;L of 1x dsDNA working solution to all but two tubes. Add 190 &#181;L of the working solution to the last two tubes.</li>
		<li>Add 10 &#181;L of the two standards (included in the Table of Materials) to separate the assay tubes.</li>
		<li>Add 1 &#181;L of each PCR reaction to Fluorometer mastermix tubes.</li>
		<li>Vortex tubes to mix and incubate at room temperature for 2 min.</li>
		<li>On the home screen of the fluorometer, select the button with the assay kitin use (1x dsDNA) then select Read Standards and Run Samples.</li>
		<li>Insert standard 1 tube, select the Read button and then repeat for standard 2.</li>
		<li>Following step 5.2.6, repeat for one sample, select a sample volume of 1 &#181;L and the resulting concentration will be provided.</li>
		<li>Repeat step 5.2.7 for the remaining samples.</li>
	</ol>
	</li>
	<li>Calculate the projected molarity of all reactions and pool equal concentrations of reactions from each individual sample separately (one final pool per sample) using the equation below. 
	<img alt="Equation 1" src="/files/ftp_upload/62583/62583eq01.jpg" style="margin: auto;" />
	<ol>
		<li>Repeat steps 5.2.1 to 5.2.7 to obtain the final pool concentration.</li>
	</ol>
	</li>
	<li>Check the amplicon pool on a capillary electrophoresis machine/agarose gel as specified by the manufacturer.
	<ol>
		<li>Prepare capillary electrophoresis tubes for the appropriate number of samples. Add 7 &#181;L of the DNA buffer as specified by the manufacturer.</li>
		<li>Add 4 &#181;L of the amplicon pool to the tube containing DNA buffer.</li>
		<li>Place tubes on the electrophoresis machine and run the machine as specified by the manufacturer for dsDNA.</li>
		<li>View the electrophoresis gel pictures ensuring the bands localize to ~5 kb (size of amplicons).</li>
	</ol>
	</li>
	<li>Calculate the projected molarity of all reactions and pool equal concentrations of reactions from each individual sample separately (one final pool per sample) using the equation below. 
	<img alt="Equation 2" src="/files/ftp_upload/62583/62583eq01.jpg" style="margin: auto;" /></li>
</ol>
6. Library preparation
<ol>
	<li>Dilute sample pools from step 5.3.1 to 0.2 ng/&#181;L for library preparation.
	<ol>
		<li>Using a low-input library preparation kit compatible with short sequences (300bp) prepare libraries using unique index combinations for each sample pool created in step 5.3 following the manufacturer&#39;s instructions. 
		NOTE: Follow the manufacturer&#39;s instructions to select index sequences. All indexes amenable to the library prep kit will work for the samples.</li>
		<li>Following library preparation, pool all samples according to the manufacturer&#39;s instructions. 
		NOTE: Before creating the library pool, calculate the number of reads necessary for 250x coverage of your target sequence and ensure that the selected sequencing cartridge can provide adequate coverage for each included sample. For 0.5Mb total, this will equate to 1M reads.</li>
	</ol>
	</li>
	<li>Prepare the library pool for sequencing.
	<ol>
		<li>Thaw and prepare a 300-cycle cartridge and sequencing reagents.</li>
		<li>Denature and dilute the sequencing pool created in step 5.1.1 according to the sequencer&#39;s manufacturer&#39;s instructions.</li>
		<li>Add denatured and diluted library pool to sequencing reagents and Run the sequencing machine as specified by the manufacturer. 
		NOTE: See attached Table 3 for trouble shootings.</li>
	</ol>
	</li>
</ol>
7. Data analysis
NOTE: All data steps are performed in the genomic data analysis software. Parentheses indicate user input. Greater than sign indicates the order of mouse clicks for any given step (e.g., 1st mouse click&#62;2nd mouse click)
<ol>
	<li>Import the reads by clicking on Open Software &#62; Import &#62; Illumina &#62; Select Files &#62; Next &#62; Select location to Save &#62; Finish. The reads will now appear in the software.</li>
	<li>Trim and filter the reads.
	<ol>
		<li>In the deep sequence data analysis software, trim the raw reads default parameters.</li>
		<li>Highlight the reads and click Toolbox &#62; Prepare Sequencing Data &#62; Trim Reads &#62; Next &#62; Next &#62; Next &#62; Next &#62; Save &#62; Next &#62; (Select location to save) &#62; Finish</li>
	</ol>
	</li>
	<li>Map the trimmed reads to reference.
	<ol>
		<li>Map trimmed reads to the reference sequence used in step 4.1 using a match score of 2, mismatch cost of 3 and insertion/deletion costs of 2. Ensure that the length fraction is above 0.7 and similarity fraction is at or above 0.8.</li>
		<li>Highlight the trimmed read file and click Toolbox &#62; Resequencing Analysis &#62; Map Reads to Reference &#62; Next &#62; (Select reference sequence) &#62; Next &#62; (Ensure parameters are indicated as above) &#62; Next &#62; Save &#62; (Select location to save) &#62; Finish.</li>
	</ol>
	</li>
	<li>Extract variants and indels.
	<ol>
		<li>Using an appropriate indel caller, extract indels using a p-value threshold of 0.005 or lower and a maximum number of mismatches of 3.</li>
		<li>Highlight mapped read file and click Toolbox &#62; Resequencing Analysis &#62; Variant detection &#62; Indels and Structural Variants &#62; Next &#62; (Ensure required significance is input &#62; Next &#62; Save &#62; (Select location to save) &#62; Finish.</li>
		<li>Using an appropriate variant caller, call variants from the read mapping using a significance of 5%.</li>
		<li>Highlight mapped read file and click Toolbox &#62; Resequencing Analysis &#62; Variant Detection &#62; Low frequency variant detection &#62; Next &#62; (Ensure required significance is input &#62; Next &#62; Next &#62; Save &#62; (Select location to save) &#62; Finish. 
		NOTE: Ensure to account for the ploidy of the host genome in the indel and variant callers. Do not extract mutants below 5%. This threshold accounts for PCR and sequencing errors associated with the assay. Normalization (based on immunoblot detection of CAS9) should be done by adjusting sequencing coverage. For example, if sample A has twice the transfection efficiency of sample B, then 50% of the reads from sample A should be used for analysis. This should be done by random sampling and not reduce the coverage for any sample below 100x.</li>
	</ol>
	</li>
</ol>

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T., Van Eynde, B., Yu, N., Ma, S., Smagghe, G. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=CRISPR/Cas9+in+insects:+Applications,+best+practices+and+biosafety+concerns.">CRISPR/Cas9 in insects: Applications, best practices and biosafety concerns.</a> Journal of Insect Physiology. 98, 245-257 (2017).</li><li>Ghezraoui, H., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Chromosomal+translocations+in+human+cells+are+generated+by+canonical+nonhomologous+end-joining.">Chromosomal translocations in human cells are generated by canonical nonhomologous end-joining.</a> Molecular Cell. 55 (6), 829-842 (2014).</li></ol>

The authors have nothing to disclose.

In addition to the depth of information provided, there are several advantages to this method. First, DSB repair, in theory, can be assessed at any genomic loci without modifying the genome of the cell of interest. Second, access to NGS analysis of repair is increased by the reduced cost and computational effort afforded by making and analyzing a single DSB targeted to a defined area. Finally, with the genomes of additional organisms routinely becoming available and multiple publications demonstrating successful transfec...

Maintaining genomic stability is critical for all living organisms. Accurate DNA replication and a robust DNA damage response (DDR) are necessary to faithfully propagate the genetic material1,2. DNA damages occur regularly in most cells2,3. When these damages are sensed, cell cycle progression is halted, and DNA repair mechanisms are activated. Double strand breaks in DNA or DSBs are the most toxic and mutagenic type of DNA damage3,4.
While several DDR signaling pathways can repair these lesions, the most thoroughly studied DSB repair pathways are homologous recombination (HR) and non-homologous end joining (NHEJ). HR is a largely error-free pathway that repairs a DSB using a sister chromatid as a homologous template. This tends to happen in the S phase and G2 phase of a cell cycle5,6,7. NHEJ is more error-prone, but it can happen throughout the cell cycle8,9. Various reporter assays have been developed to measure the efficiency of specific repair mechanisms10,11,12. These assays tend to rely on flow cytometry for a high throughput measurement of DSB repair pathway activity using GFP or mCherry as a readout11,13. While highly efficient, they rely on canonical repair occurring at an artificially introduced DSB.
There are a variety of other methods used to study DSB repair. Many of these rely on immunofluorescence (IF) microscopy1,14. IF microscopy detects discrete nuclear foci representative of repair complexes after DSBs are induced by exposure to genotoxic chemicals or ionizing radiation15,16. Tracking the formation and resolution of these foci provides an indication of repair initiation and completion, respectively14,17. However, these methods of DSB induction (i.e., chemicals or ionizing radiation) do not cause DSBs at defined locations in the genome. It is also functionally impossible to use them to consistently induce only a small number (e.g., 2-4) of DSBs. As a result, the most commonly used methods of inducing DSBs cause a multitude of lesions randomly distributed throughout the genome18. A small number of DSBs can be introduced by inserting the recognition site for a rare-cutting endonuclease and expressing the pertinent endonuclease, such as I-Sce119. Unfortunately, the required integration of a target site prevents the examination of DSB at endogenous genomic loci.
This manuscript describes a method to detect mutations associated with the repair of a DSB generated at a user-defined locus. We provide a representative example of the approach applied to assess the ability of a viral protein to increase the number of mutations associated with a DSB. Specifically, this manuscript describes the use of a single guide RNA (sgRNA) to direct a CAS9 endonuclease to induce a DSB at human CD4 open reading frame in human foreskin keratinocytes expressing vector control (HFK LXSN) and HFK that expresses the E6 protein of human papilloma virus type 8 (HFK 8E6). Targeted next-generation sequencing (NGS) of the region surrounding the break allows mutations associated with the repair of the lesion to be rigorously defined. These data demonstrate that the viral protein causes an approximately 20-fold increase in mutations during DSB repair. It also provides an unbiased characterization of the mutagenic consequences of DSBs at a single locus without the need for whole-genome sequencing. In principle, the protocol could be readily adapted to compare the relative risk of mutations between genome loci or cell lines.

<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>6 Well Tissue Culture Plate</td>
			<td>Celltreat</td>
			<td>229106</td>
			<td>Cell culture plate</td>
		</tr>
		<tr>
			<td>BCA Kit</td>
			<td>VWR</td>
			<td>89167-794</td>
			<td>BCA assay kit</td>
		</tr>
		<tr>
			<td>Centrifuge 5910 R</td>
			<td>Eppendorf</td>
			<td>2231000772</td>
			<td>Tabletop Centrifuge</td>
		</tr>
		<tr>
			<td>CLC Genomic Workbench</td>
			<td>Qiagen</td>
			<td>832001</td>
			<td>deep sequence data analysis software/indel caller/variant caller</td>
		</tr>
		<tr>
			<td>Digital Microplate Genie pulse</td>
			<td>Scientific industries</td>
			<td>SI-400A</td>
			<td>Plate shaker</td>
		</tr>
		<tr>
			<td>DYKDDDDK Tag Monoclonal Antibody (FG4R)</td>
			<td>ThermoFisher Scientific</td>
			<td>MA191878</td>
			<td>Anti-FLAG antibody</td>
		</tr>
		<tr>
			<td>Epilife CF Kit</td>
			<td>ThermoFisher Scientific</td>
			<td>MEPICF500</td>
			<td>Cell cultrue media and supplements</td>
		</tr>
		<tr>
			<td>Fetal Bovine Serum (FBS)</td>
			<td>VWR</td>
			<td>89510-194</td>
			<td>Cell culture supplement</td>
		</tr>
		<tr>
			<td>Goat anti-Rabbit IgG</td>
			<td>ThermoFisher Scientific</td>
			<td>A-11012</td>
			<td>Secondary antibody</td>
		</tr>
		<tr>
			<td>HighPrep PCR Clean-up system</td>
			<td>MagBio</td>
			<td>AC-60005</td>
			<td>Bead-based PCR cleanup kit</td>
		</tr>
		<tr>
			<td>KAPA HiFi HotStart ReadyMix PCR Kit</td>
			<td>KAPA Biosystems</td>
			<td>KK2600</td>
			<td>PCR mastermix/PCR assay</td>
		</tr>
		<tr>
			<td>MagAttract HMW DNA kit</td>
			<td>Qiagen</td>
			<td>67563</td>
			<td>High Molecular Weight DNA extraction kit</td>
		</tr>
		<tr>
			<td>Magnetic Stand-96</td>
			<td>Thermo Fisher Scientific</td>
			<td>AM10027</td>
			<td>96-Well Magnetic Rack</td>
		</tr>
		<tr>
			<td>MiniAmp Thermal Cycler</td>
			<td>Applied Biosystems</td>
			<td>A37834</td>
			<td>Thermal Cycler</td>
		</tr>
		<tr>
			<td>Miseq</td>
			<td>Illumina</td>
			<td>SY-410-1003</td>
			<td>Sequencer</td>
		</tr>
		<tr>
			<td>Miseq v2 300 cycle reagent kit</td>
			<td>Illumina</td>
			<td>MS-102-2002</td>
			<td>300-cycle cartridge/sequencing reagents</td>
		</tr>
		<tr>
			<td>Nextera XT DNA Library Prep kit</td>
			<td>Illumina</td>
			<td>FC-131-1024</td>
			<td>Library preparation kit</td>
		</tr>
		<tr>
			<td>Nextera XT Kit v2 Set A</td>
			<td>Illumina</td>
			<td>20027215</td>
			<td>Indexes</td>
		</tr>
		<tr>
			<td>Nunc 96-well polypropylene DeepWell Stroage plates</td>
			<td>Thermo Fisher Scientific</td>
			<td>260251</td>
			<td>deep well 96-well plates</td>
		</tr>
		<tr>
			<td>Penicillin-Streptomycin Solution (100X)</td>
			<td>Calsson Labs</td>
			<td>PSL02-6X100ML</td>
			<td>Antibiotics for cell culture</td>
		</tr>
		<tr>
			<td>Phosphate Buffered Saline (PBS)</td>
			<td>Bio Basic</td>
			<td>PD8117</td>
			<td>PBS</td>
		</tr>
		<tr>
			<td>px330-CD4</td>
			<td>Addgen</td>
			<td>136938</td>
			<td>SgRNA/CAS9 plasmids targeting 5&#8217;- GGCGTATCTGTGTGAGGACT</td>
		</tr>
		<tr>
			<td>QIAxcel Advanced System</td>
			<td>Qiagen</td>
			<td>9001941</td>
			<td>capillary electrophersis machine</td>
		</tr>
		<tr>
			<td>QIAxcel DNA screening kit</td>
			<td>Qiagen</td>
			<td>929004</td>
			<td>DNA buffer/ capillary electrophersis tubes</td>
		</tr>
		<tr>
			<td>Qubit 1x ds HS Assay Kit</td>
			<td>ThermoFisher Scientific</td>
			<td>Q23851</td>
			<td>Fluorometer reagents/1x dsDNA solution</td>
		</tr>
		<tr>
			<td>Qubit 4 Fluorometer</td>
			<td>ThermoFisher Scientific</td>
			<td>Q33238</td>
			<td>Fluorometer</td>
		</tr>
		<tr>
			<td>Qubit Assay Tubes</td>
			<td>Thermo Fisher Scientific</td>
			<td>Q32856</td>
			<td>Fluorometer assay tubes</td>
		</tr>
		<tr>
			<td>RIPA Lysis Buffer</td>
			<td>VWR</td>
			<td>VWRVN653-100ML</td>
			<td>Lysis buffer for protein extraction</td>
		</tr>
		<tr>
			<td>Trypsin-EDTA (0.05%), phenol red</td>
			<td>ThermoFisher Scientific</td>
			<td>25300054</td>
			<td>Trypsin</td>
		</tr>
		<tr>
			<td>Vortex-Genie 2</td>
			<td>Scientific industries</td>
			<td>SI-0236</td>
			<td>Vortex</td>
		</tr>
		<tr>
			<td>Xfect Transfection Reagent</td>
			<td>Takara Bio</td>
			<td>631318</td>
			<td>Transfection reagent</td>
		</tr>
		<tr>
			<td>genomic data analysis software</td>
			<td>QIAGEN</td>
			<td>CLC Workbench v21.0.</td>
			<td>Data analysis software</td>
		</tr>
	</tbody>
</table>

using next generation sequencing to identify mutations associated with repair of a cas9-induced double strand break near the cd4 promoter

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

Three representative results are presented for this protocol. Figure 1 is an immunoblot confirming expression of CAS9 in HFK control (LXSN) and HFK expressing beta-HPV 8E6 (8E6). 48 h after transfection, whole cell lysates were harvested and subsequently probed with an anti-CAS9 antibody (or GAPDH as a loading control). The result shows that HFK LXSN and HFK 8E6 are expressing similar amount of CAS9 indicating that transfection efficiency is similar between t...

Watch this Scientific Journal Video about Using Next Generation Sequencing to Identify Mutations Associated with Repair of a CAS9-induced Double Strand Break Near the CD4 Promoter at JoVE.com

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

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

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

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2.3K Views.  Kansas State University. Presented here is sgRNA/CAS9 endonuclease and next-generation sequencing protocol that can be used to identify the mutations associated with double strand break repair near the CD4 promoter.

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

Wild-type Blocking PCR Combined with Direct Sequencing as a Highly Sensitive Method for Detection of Low-Frequency Somatic Mutations

Accurate detection and identification of low frequency mutations can be problematic when assessing residual disease after therapy, screening for emerging resistance mutations during therapy, or when patients have few circulating tumor cells. Wild-type blocking PCR followed by sequencing analysis offers high sensitivity, flexibility, and simplicity as a methodology for detecting these low frequency mutations. By adding a custom designed locked nucleic acid oligonucleotide to a new or previously established conventional PCR based sequencing assay, sensitivities of approximately 1 mutant allele in a background of 1,000 WT alleles can be achieved (1:1,000). Sequencing artifacts associated with deamination events commonly found in formalin fixed paraffin embedded tissues can be partially remedied by the use of uracil DNA glycosylase during extraction steps. The optimized protocol here is specific for detecting MYD88 mutation, but can serve as a template to design any WTB-PCR assay. Advantages of the WTB-PCR assay over other commonly utilized assays for the detection of low frequency mutations including allele specific PCR and real-time quantitative PCR include fewer occurrences of false positives, greater flexibility and ease of implementation, and the ability to detect both known and unknown mutations.

Wild-type blocking PCR followed by direct sequencing offers a highly sensitive method of detection for low frequency somatic mutations in a variety of sample types.

Accurate detection and identification of low frequency mutations can be problematic when assessing residual disease after therapy, screening for emerging ...

Detection of Copy Number Alterations Using Single Cell Sequencing

Detection of genomic changes at single cell resolution is important for characterizing genetic heterogeneity and evolution in normal tissues, cancers, and microbial populations. Traditional methods for assessing genetic heterogeneity have been limited by low resolution, low sensitivity, and/or low specificity. Single cell sequencing has emerged as a powerful tool for detecting genetic heterogeneity with high resolution, high sensitivity and, when appropriately analyzed, high specificity. Here we provide a protocol for the isolation, whole genome amplification, sequencing, and analysis of single cells. Our approach allows for the reliable identification of megabase-scale copy number variants in single cells. However, aspects of this protocol can also be applied to investigate other types of genetic alterations in single cells.

Single cell sequencing is an increasingly popular and accessible tool for addressing genomic changes at high resolution. We provide a protocol that uses single cell sequencing to identify copy number alterations in single cells.

Detection of genomic changes at single cell resolution is important for characterizing genetic heterogeneity and evolution in normal tissues, cancers, and ...

Optimization and Comparative Analysis of Plant Organellar DNA Enrichment Methods Suitable for Next-generation Sequencing

Plant organellar genomes contain large, repetitive elements that may undergo pairing or recombination to form complex structures and/or sub-genomic fragments. Organellar genomes also exist in admixtures within a given cell or tissue type (heteroplasmy), and an abundance of subtypes may change throughout development or when under stress (sub-stoichiometric shifting). Next-generation sequencing (NGS) technologies are required to obtain deeper understanding of organellar genome structure and function. Traditional sequencing studies use several methods to obtain organellar DNA: (1) If a large amount of starting tissue is used, it is homogenized and subjected to differential centrifugation and/or gradient purification. (2) If a smaller amount of tissue is used (i.e., if seeds, material, or space is limited), the same process is performed as in (1), followed by whole-genome amplification to obtain sufficient DNA. (3) Bioinformatics analysis can be used to sequence the total genomic DNA and to parse out organellar reads. All these methods have inherent challenges and tradeoffs. In (1), it may be difficult to obtain such a large amount of starting tissue; in (2), whole-genome amplification could introduce a sequencing bias; and in (3), homology between nuclear and organellar genomes could interfere with assembly and analysis. In plants with large nuclear genomes, it is advantageous to enrich for organellar DNA to reduce sequencing costs and sequence complexity for bioinformatics analyses. Here, we compare a traditional differential centrifugation method with a fourth method, an adapted CpG-methyl pulldown approach, to separate the total genomic DNA into nuclear and organellar fractions. Both methods yield sufficient DNA for NGS, DNA that is highly enriched for organellar sequences, albeit at different ratios in mitochondria and chloroplasts. We present the optimization of these methods for wheat leaf tissue and discuss major advantages and disadvantages of each approach in the context of sample input, protocol ease, and downstream application.

The comparison and optimization of two plant organellar DNA enrichment methods are presented: traditional differential centrifugation and fractionation of the total gDNA based on methylation status. We assess the resulting DNA quantity and quality, demonstrate performance in short-read next-generation sequencing, and discuss the potential for use in long-read single-molecule sequencing.

Plant organellar genomes contain large, repetitive elements that may undergo pairing or recombination to form complex structures and/or sub-genomic ...

Analysis of the c-KIT Ligand Promoter Using Chromatin Immunoprecipitation

Multiple cellular processes, including DNA replication and repair, DNA recombination, and gene expression, require interactions between proteins and DNA. Therefore, DNA-protein interactions regulate multiple physiological, pathophysiological, and biological functions, such as cell differentiation, cell proliferation, cell cycle control, chromosome stability, epigenetic gene regulation, and cell transformation. In eukaryotic cells, the DNA interacts with histone and nonhistone proteins and is condensed into chromatin. Several technical tools can be used to analyze DNA-protein interactions, such as the Electrophoresis (gel) Mobility Shift Assay (EMSA) and DNase I footprinting. However, these techniques analyze the protein-DNA interaction in vitro, not within the cellular context. Chromatin immunoprecipitation (ChIP) is a technique that captures proteins at their specific DNA binding sites, thereby allowing for the identification of DNA-protein interactions within their chromatin context. It is done by fixation&#160;of the DNA-protein interaction, followed by&#160;immunoprecipitation of the protein of interest. Subsequently, the genomic site that the protein was bound to is characterized. Here, we describe and discuss ChIP and demonstrate its analytical value for the identification of the Transforming Growth Factor-&#946; (TGF-&#946;)-induced binding of the transcription factor SMAD2 to SMAD Binding Elements (SBE) within the promoter region of the tyrosine-protein kinase Kit (c-KIT) receptor ligand Stem Cell Factor (SCF).

DNA-protein interactions are essential for multiple biological processes. During the evaluation of cellular functions, the analysis of DNA-protein interactions is indispensable for understanding gene regulation. Chromatin immunoprecipitation (ChIP) is a powerful tool to analyze such interactions in vivo.

Multiple cellular processes, including DNA replication and repair, DNA recombination, and gene expression, require interactions between proteins and DNA. ...

Generation of a Gene-disrupted Streptococcus mutans Strain Without Gene Cloning

Typical methods for the elucidation of the function of a particular gene involve comparative phenotypic analyses of the wild-type strain and a strain in which the gene of interest has been disrupted. A gene-disruption DNA construct containing a suitable antibiotic resistance marker gene is useful for the generation of gene-disrupted strains in bacteria. However, conventional construction methods, which require gene cloning steps, involve complex and time-consuming protocols. Here, a relatively facile, rapid, and cost-effective method for targeted gene disruption in Streptococcus mutans is described. The method utilizes a 2-step fusion polymerase chain reaction (PCR) to generate the disruption construct and electroporation for genetic transformation. This method does not require an enzymatic reaction, other than PCR, and additionally offers greater flexibility in terms of the design of the disruption construct. Employment of electroporation facilitates the preparation of competent cells and improves the transformation efficiency. The present method may be adapted for the generation of gene-disrupted strains of various species.

Generation of a Gene-disrupted Streptococcus mutans Strain Without Gene Cloning

We describe a facile, rapid, and relatively inexpensive method for the generation of gene-disrupted Streptococcus mutans strains; this technique may be adapted for the generation of gene-disrupted strains of various species.

Typical methods for the elucidation of the function of a particular gene involve comparative phenotypic analyses of the wild-type strain and a strain in ...

Targeted Next-generation Sequencing and Bioinformatics Pipeline to Evaluate Genetic Determinants of Constitutional Disease

Next-generation sequencing (NGS) is quickly revolutionizing how research into the genetic determinants of constitutional disease is performed. The technique is highly efficient with millions of sequencing reads being produced in a short time span and at relatively low cost. Specifically, targeted NGS is able to focus investigations to genomic regions of particular interest based on the disease of study. Not only does this further reduce costs and increase the speed of the process, but it lessens the computational burden that often accompanies NGS. Although targeted NGS is restricted to certain regions of the genome, preventing identification of potential novel loci of interest, it can be an excellent technique when faced with a phenotypically and genetically heterogeneous disease, for which there are previously known genetic associations. Because of the complex nature of the sequencing technique, it is important to closely adhere to protocols and methodologies in order to achieve sequencing reads of high coverage and quality. Further, once sequencing reads are obtained, a sophisticated bioinformatics workflow is utilized to accurately map reads to a reference genome, to call variants, and to ensure the variants pass quality metrics. Variants must also be annotated and curated based on their clinical significance, which can be standardized by applying the American College of Medical Genetics and Genomics Pathogenicity Guidelines. The methods presented herein will display the steps involved in generating and analyzing NGS data from a targeted sequencing panel, using the ONDRISeq neurodegenerative disease panel as a model, to identify variants that may be of clinical significance.

Targeted next-generation sequencing is a time- and cost-efficient approach that is becoming increasingly popular in both disease research and clinical diagnostics. The protocol described here presents the complex workflow required for sequencing and the bioinformatics process used to identify genetic variants that contribute to disease.

Next-generation sequencing (NGS) is quickly revolutionizing how research into the genetic determinants of constitutional disease is performed. The ...

Promoter Capture Hi-C: High-resolution, Genome-wide Profiling of Promoter Interactions

The three-dimensional organization of the genome is linked to its function. For example, regulatory elements such as transcriptional enhancers control the spatio-temporal expression of their target genes through physical contact, often bridging considerable (in some cases hundreds of kilobases) genomic distances and bypassing nearby genes. The human genome harbors an estimated one million enhancers, the vast majority of which have unknown gene targets. Assigning distal regulatory regions to their target genes is thus crucial to understand gene expression control. We developed Promoter Capture Hi-C (PCHi-C) to enable the genome-wide detection of distal promoter-interacting regions (PIRs), for all promoters in a single experiment. In PCHi-C, highly complex Hi-C libraries are specifically enriched for promoter sequences through in-solution hybrid selection with thousands of biotinylated RNA baits complementary to the ends of all promoter-containing restriction fragments. The aim is to then pull-down promoter sequences and their frequent interaction partners such as enhancers and other potential regulatory elements. After high-throughput paired-end sequencing, a statistical test is applied to each promoter-ligated restriction fragment to identify significant PIRs at the restriction fragment level. We have used PCHi-C to generate an atlas of long-range promoter interactions in dozens of human and mouse cell types. These promoter interactome maps have contributed to a greater understanding of mammalian gene expression control by assigning putative regulatory regions to their target genes and revealing preferential spatial promoter-promoter interaction networks. This information also has high relevance to understanding human genetic disease and the identification of potential disease genes, by linking non-coding disease-associated sequence variants in or near control sequences to their target genes.

DNA regulatory elements, such as enhancers, control gene expression by physically contacting target gene promoters, often through long-range chromosomal interactions spanning large genomic distances. Promoter Capture Hi-C (PCHi-C) identifies significant interactions between promoters and distal regions, enabling the assignment of potential regulatory sequences to their target genes.

The three-dimensional organization of the genome is linked to its function. For example, regulatory elements such as transcriptional enhancers control the ...

Amplification of Near Full-length HIV-1 Proviruses for Next-Generation Sequencing

The Full-Length Individual Proviral Sequencing (FLIPS) assay is an efficient and high-throughput method designed to amplify and sequence single, near full-length (intact and defective), HIV-1 proviruses. FLIPS allows determination of the genetic composition of integrated HIV-1 within a cell population. Through identifying defects within HIV-1 proviral sequences that arise during reverse transcription, such as large internal deletions, deleterious stop codons/hypermutation, frameshift mutations, and mutations/deletions in cis acting elements required for virion maturation, FLIPS can identify integrated proviruses incapable of replication. The FLIPS assay can be utilized to identify HIV-1 proviruses that lack these defects and are&#160;therefore potentially replication-competent. The FLIPS protocol involves: lysis of HIV-1 infected cells, nested PCR of near full-length HIV-1 proviruses (using primers targeted to the HIV-1 5&#39; and 3&#39; LTR), DNA purification and quantification, library preparation for Next-generation Sequencing (NGS), NGS, de novo assembly of proviral contigs, and a simple process of elimination for identifying replication-competent proviruses. FLIPS provides advantages over traditional methods designed to sequence integrated HIV-1 proviruses, such as single-proviral sequencing. FLIPS amplifies and sequences near full-length proviruses enabling replication competency to be determined, and also uses fewer amplification primers, preventing the consequences of primer mismatches. FLIPS is a useful tool for understanding the genetic landscape of integrated HIV-1 proviruses, especially within the latent reservoir, however, its utilization can extend to any application in which the genetic composition of integrated HIV-1 is required.

Full-length individual proviral sequencing (FLIPS) provides an efficient and high-throughput method for the amplification and sequencing of single, near full-length (intact and defective) HIV-1 proviruses and allows for determination of their potential replication-competency. FLIPS overcomes limitations of previous assays designed to sequence the latent HIV-1 reservoir.

The Full-Length Individual Proviral Sequencing (FLIPS) assay is an efficient and high-throughput method designed to amplify and sequence single, near ...

High-throughput Identification of Gene Regulatory Sequences Using Next-generation Sequencing of Circular Chromosome Conformation Capture (4C-seq)

The identification of regulatory elements for a given target gene poses a significant technical challenge owing to the variability in the positioning and effect sizes of regulatory elements to a target gene. Some progress has been made with the bioinformatic prediction of the existence and function of proximal epigenetic modifications associated with activated gene expression using conserved transcription factor binding sites. Chromatin conformation capture studies have revolutionized our ability to discover physical chromatin contacts between sequences and even within an entire genome. Circular chromatin conformation capture coupled with next-generation sequencing (4C-seq), in particular, is designed to discover all possible physical chromatin interactions for a given sequence of interest (viewpoint), such as a target gene or a regulatory enhancer. Current 4C-seq strategies directly sequence from within the viewpoint but require numerous and diverse viewpoints to be simultaneously sequenced to avoid the technical challenges of uniform base calling (imaging) with next generation sequencing platforms. This volume of experiments may not be practical for many laboratories. Here, we report a modified approach to the 4C-seq protocol that incorporates both an additional restriction enzyme digest and qPCR-based amplification steps that are designed to facilitate a greater capture of diverse sequence reads and mitigate the potential for PCR bias, respectively. Our modified 4C method is amenable to the standard molecular biology lab for assessing chromatin architecture.

High-throughput Identification of Gene Regulatory Sequences Using Next-generation Sequencing of Circular Chromosome Conformation Capture 4C-seq

The identification of physical interactions between genes and regulatory elements is challenging but has been facilitated by chromosome conformation capture methods. This modification to the 4C-seq protocol mitigates PCR bias by minimizing over-amplification of PCR templates and maximizes the mappability of reads by incorporating an addition restriction enzyme digest step.

The identification of regulatory elements for a given target gene poses a significant technical challenge owing to the variability in the positioning and ...

Sequencing of mRNA from Whole Blood using Nanopore Sequencing

Sequencing in remote locations and resource-poor settings presents unique challenges. Nanopore sequencing can be successfully used under such conditions, and was deployed to West Africa during the recent Ebola virus epidemic, highlighting this possibility. In addition to its practical advantages (low cost, ease of equipment transport and use), this technology also provides fundamental advantages over second-generation sequencing approaches, particularly the very long read length, ability to directly sequence RNA, and real-time availability of data. Raw read accuracy is lower than with other sequencing platforms, which represents the main limitation of this technology; however, this can be partially mitigated by the high read depth generated. Here, we present a field-compatible protocol for sequencing of the mRNAs encoding for Niemann-Pick C1, which is the cellular receptor for ebolaviruses. This protocol encompasses extraction of RNA from animal blood samples, followed by RT-PCR for target enrichment, barcoding, library preparation, and the sequencing run itself, and can be easily adapted for use with other DNA or RNA targets.

Nanopore sequencing is a novel technology that allows cost-effective sequencing in remote locations and resource-poor settings. Here, we present a protocol for sequencing of mRNAs from whole blood that is compatible with such conditions.

Sequencing in remote locations and resource-poor settings presents unique challenges. Nanopore sequencing can be successfully used under such conditions, ...