The work is supported by an Institute Development Award from The Children's Hospital of Philadelphia.

1. Cross-link Chromatin
<ol>
 <li>Grow cells in 100x20mm cell culture dishes. The amount of cells can range from 1 to 10 million cells per dish depending on cell type. Approximately 2 million cells is sufficient for one immunoprecipitation.</li>
 <li>Cross-link cells in 1% Formaldehyde for 10 min at room temperature with occasional rocking.</li>
 <li>Quench cross-linking by adding a final concentration of 125 mM Glycine and incubate for 5 min at room temperature.</li>
 <li>Wash cells with 1X Phosphate Buffered Saline (PBS) twice, decant PBS, and then add 0.2 ml of PBS.</li>
 <li>Harvest cells with a plastic cell scraper into a microcentrifuge tube.</li>
 <li>Spin down cells at 2,000 rpm for 5 min at 4 &deg;C.</li>
 <li>Aspirate the supernatant. Resuspend cells in SDS Lysis buffer (1% SDS, 10mM EDTA, 50mM Tris-HCl pH 8.1) for whole cell lysate or keep them as a pellet for nuclear extraction.</li>
 <li>Cells can be saved at -80 &deg;C or one can proceed immediately with sonication.</li>
</ol>
2. Prepare Nuclei (Proceed to Step 3.5 for Whole Cell Lysate)
<ol>
 <li>Supplement Cell Lysis Buffer (5 mM PIPES pH 8.0, 85 mM KCl, 0.5% NP-40) with 1X Proteinase Inhibitor each experiment.</li>
 <li>Resuspend thawed cell pellet in approximately 10 times the pellet volume with Cell Lysis Buffer.</li>
 <li>Dounce-homogenize 10 times with pestle then incubate on ice for 10 min.</li>
 <li>Centrifuge sample at 4,000 rpm for 5 min at 4 &deg;C, discard supernatant, and save nuclear pellet.</li>
</ol>
3. Sonication*
<ol>
 <li>Warm up SDS Lysis Buffer and supplement amount of buffer to be used with Proteinase Inhibitor.</li>
 <li>Resuspend nuclear pellet in SDS Lysis Buffer (approximately 0.5 ml of buffer per 1-10 million cells)</li>
 <li>Incubate on ice for 10 min.</li>
 <li>Add 0.5 ml aliquots of samples to microcentrifuge tubes.</li>
 <li>Sonicate on wet ice using Misonix sonicator with 30 sec on and 45 sec off at an amplitude setting of 2. Number of cycles for ideal fragment size can be determined by first trying out various cycle numbers (ex. 2, 4, 8, 12, 16, and 20 or more cycles). A different brand of sonicator may be used, however, the conditions will vary. Experimentation with the number of cycles and amount of time on and off must be carried out in order to determine the ideal conditions.</li>
 <li>Collect 20 &mu;l of each sample to check sonication results and to do quantification,. The rest of the sample can be stored at -80 &deg;C.</li>
 <li>Dilute the 20 &mu;l of sample by adding 30 &mu;l of 0.1X TE Buffer.</li>
 <li>Treat sample with 1 &mu;l of RNase A at 37 &deg;C for 1 hr then add 1 &mu;l of Proteinase K and incubate at 62 &deg;C for 2 hr.</li>
 <li>Run 20 &mu;l of the sample on a 2% agarose gel.</li>
 <li>Purify the remaining amount of sample with QIAquick PCR Purification kit then quantify using NanoDrop spectrophotometer.</li>
</ol>
*For native ChIP, microccocal nuclease digestion may be used alternatively to shear the DNA.
4. Block Agarose Beads*
<ol>
 <li>If beads are already blocked, proceed to step 5.1.</li>
 <li>Use Protein A or Protein G agarose. For 5 immunoprecipitations (IPs), use 600 &mu;l of 50% bead slurry (300 &mu;l bead pellet)</li>
 <li>To wash the beads, spin them down at 800 rpm for 1 min at 4 &deg;C and discard supernatant. Add slightly more than 2 ml ChIP Dilution Buffer (0.01% SDS, 1.2 mM EDTA, 167 mM NaCl, 1.1% Triton X-100, 16.7 mM Tris-HCl pH 8.1) and mix by slowly inverting the tube 10X. Spin down again at 800 rpm for 1 min at 4 &deg;C and discard supernatant. Repeat the wash 2 more times.</li>
 <li>Block beads by rotating at 4 &deg;C overnight in blocking solution. Refer to Table 1 for the recipe of the blocking solution.</li>
</ol>
5. Pre-clear Chromatin
<ol>
 <li>Thaw sonicated chromatin on ice.</li>
 <li>Spin down at 12,000 rpm for 10 min at 4 &deg;C then put on ice right away to remove SDS (white pellet).</li>
 <li>Collect supernatant, discard pellet, and combine samples if necessary.</li>
 <li>Take out the amounts needed for the experiment based on calculations (1-10 ug of chromatin per IP).</li>
 <li>Dilute chromatin 10X in ChIP Dilution Buffer supplemented with proteinase inhibitor.</li>
 <li>Add 100 &mu;l of blocked beads per IP.</li>
 <li>Rotate at 4 &deg;C for 1 hr.</li>
</ol>
6. Immunoprecipitation
<ol>
 <li>Spin down samples at 800 rpm for 1 min and transfer the supernatant to a fresh tube.</li>
 <li>Spin down supernatant at 800 rpm for 1 min and transfer to another clean tube.</li>
 <li>Save 20 &mu;l of the supernatant, to serve as the input control, at -20 &deg;C.</li>
 <li>Aliquot the chromatin to the number of IPs to be done in the experiment.</li>
 <li>Add 2 ug of antibody per 1-10 ug of chromatin to each sample.</li>
 <li>Incubate overnight 4 &deg;C with rotation.</li>
 <li>Add 100 &mu;l of blocked beads to each IP sample.</li>
 <li>Incubate for 1 hr at 4 &deg;C with rotation.</li>
 <li>Pellet the beads by spinning down at 800 rpm for 1 min and discard as much of the supernatant as possible.</li>
 <li>Wash beads once with Low Salt Immune Complex wash buffer. Add 1 ml of buffer to each tube; rotate at room temperature for 5-8 min; spin down at 800 rpm for 1 min, then discard supernatant. Repeat wash once with High Salt Immune Complex wash buffer and LiCl Immune Complex wash buffer and twice with TE Buffer for a total of 5 washes (Table 2).</li>
</ol>
7. Elution
<ol>
 <li>Thaw input samples from previous day to be processed with the eluants.</li>
 <li>Make Elution Buffer fresh (Table 3).</li>
 <li>Make a master mix of enough Elution Buffer needed for IPs and input control samples plus 1-2 extra samples.</li>
 <li>Add 100 &mu;l Elution Buffer to each IP sample and incubate at room temp for 15 min with rotation.</li>
 <li>Spin down at 800 rpm for 1 min and add supernatant to a fresh tube.</li>
 <li>Add another 100 &mu;l of Elution buffer to each tube of beads and incubate at room temp for 15 min with rotation.</li>
 <li>Vortex for 15 sec after incubation; spin down at 5,000 rpm for 1 min, then combine supernatant with the supernatant from the first elution. (Make sure there are no left over beads in the supernatants. If unsure, spin down supernatant again at 5,000 rpm for 1 min and collect the supernatant in a new tube.</li>
 <li>Add 180 &mu;l of Elution Buffer to the 20 &mu;l of input control samples.</li>
</ol>
8. Reverse Cross-link
<ol>
 <li>To the 200 &mu;l of eluants and input controls, add 8 &mu;l of 5 M NaCl.</li>
 <li>Seal tubes with parafilm and incubate in water bath at 65 &deg;C overnight.</li>
</ol>
9. DNA Purification
<ol>
 <li>Treat each sample with 1 &mu;l of RNase A for 1 hr at 37 &deg;C.</li>
 <li>Add 4 &mu;l of 0.5 M EDTA, 8 &mu;l 1M Tris-HCl, mix, then add 1 &mu;l of Proteinase K to each sample and incubate at 45 &deg;C for 2 hr.</li>
 <li>Purify the samples using QIAquick PCR Purification kit. The samples can be saved at -20 &deg;C and PCR check can be done on a later date.</li>
</ol>
*Alternatively, ChIP-grade magnetic beads can be used in place of agarose for the immunoprecipitation portion.
10. PCR check
<ol>
 <li>For the PCR check, use primers for regions known to be bound by the protein of interest. Also, use primers for non-binding regions as negative controls.</li>
 <li>Mix the reagents for the reaction. Dilute Input sample at 1:100 (Table 3).</li>
 <li>Run reaction. PCR Program:</li>
</ol>
Step 1: 94 &deg;C 3 min
Step 2: 94 &deg;C 20 sec 
59 &deg;C 30 sec 
72 &deg;C 30 sec 
(Repeat Step 2 for at least 30 cycles)
Step 3: 72 &deg;C 2 min
<ol start="4">
 <li>Run samples on 1% agarose gel.</li>
 <li>Enrichment can also be determined quantitatively with real-time PCR.</li>
</ol>

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The authors declare that they have no competing financial interests.

It is now feasible to carry out a genome-wide profile of protein-DNA interactions association using ChIP-seq, as has been very recently demonstrated with other transcription factors2,3. The key to a successful sequencing outcome is the generation of a high quality chromatin immunoprecipitation DNA template.
Once the DNA template has been generated and ascertained to be adequately enriched, one can then take it in library preparation for subsequent sequencing. For example, one...

For many years there has been an unmet need to identify the set of genes bound and regulated by a given protein genome wide, in particular those in the transcription factor class.
Odom et al.1 used chromatin immunoprecipitation (ChIP) combined with promoter microarrays to systematically identify the genes occupied by pre-specified transcriptional regulators in human liver and pancreatic islets. Subsequently, Johnson et al.2 developed a large-scale chromatin immunoprecipitation assay based on direct ultra high-throughput DNA sequencing (ChIP-seq) in order to comprehensively map protein-DNA interactions across entire mammalian genomes. As a test case, they mapped in vivo the binding of the neuron-restrictive silencer factor (NRSF) to 1946 locations in the human genome. The data displayed sharp resolution of binding position (+50 base pairs), which facilitated both the isolation of motifs and the identification of NRSF-binding motifs. These ChIP-seq data also had high sensitivity and specificity and statistical confidence (P &lt; 10&minus;4), properties that are important for inferring new candidate interactions.
Robertson et al.3 also used ChIP-seq in order to map STAT1 targets in interferon-&gamma; (IFN-&gamma;)-stimulated and unstimulated human HeLa S3 cells in vivo. By ChIP-seq, using 15.1 and 12.9 million uniquely mapped sequence reads, and an estimated false discovery rate of less than 0.001, they identified 41,582 and 11,004 putative STAT1-binding regions in stimulated and unstimulated cells, respectively. Of the 34 loci known to contain STAT1 interferon-responsive binding sites4-8, ChIP-seq found 24 (71%). ChIP-seq targets were enriched in sequences similar to known STAT1 binding motifs. Comparisons with two existing ChIP-PCR data sets suggested that ChIP-seq sensitivity was between 70% and 92% and specificity was at least 95%. Additionally, it was clear that ChIP-seq offers both low analytical complexity and sensitivity that increases with sequencing depth.
As such, "next-generation" genome sequencing technologies provide 1-2 orders of magnitude increase in the amount of sequence that can be cost-effectively generated over older technologies9. ChIP-seq methods therefore directly provide whole-genome coverage for effective profiling of mammalian protein-DNA interactions3.
In 2006, a strong association of variants in the transcription factor 7-like 2 (TCF7L2) gene with type 2 diabetes was discovered10. Other investigators have already independently replicated this finding in different ethnicities and, interestingly, from the first genome wide association studies of type 2 diabetes published in Nature11,12, Science13-15 and elsewhere16,17, the strongest association was indeed with TCF7L2; this is now considered the most significant genetic finding in type 2 diabetes to date18-20. In addition, TCF7L2 has been linked to cancer risk 21,22; indeed, this connection became more obvious when the 8q24 locus revealed by genome wide association studies of a number of cancers, including colorectal carcinomas, was shown to be due to an extreme upstream TCF7L2-binding element driving the transcription of MYC23,24. As such, there is great interest in determining the downstream genes regulated by this key transcription factor.
Based on experience with TCF7L2 as an example of the methodology, this paper outlines how to generate high quality ChIP DNA template. ChIP was carried out in the colorectal carcinoma cell line, HCT116, for subsequent sequencing in order build a high-resolution map of the genes bound by TCF7L225 in an endeavor to yield further insight in to its key role in the pathogenesis of complex traits.

<table border="1">
 <tbody>
 <tr>
 <td>Name of the reagent</td>
 <td>Company</td>
 <td>Catalogue number</td>
 <td></td>
 </tr>
 <tr>
 <td>QIAquick PCR Purification Kit</td>
 <td>Qiagen</td>
 <td>28104 </td>
 <td></td>
 </tr>
 <tr>
 <td>EZ-ChIP Kit </td>
 <td>Millipore </td>
 <td>17-371 </td>
 <td></td>
 </tr>
 <tr>
 <td>GoTaq Hot Start Polymerase </td>
 <td>Promega</td>
 <td>M5001</td>
 <td></td>
 </tr>
 <tr>
 <td>Misonix Sonicator</td>
 <td>Qsonica </td>
 <td>XL-2000 </td>
 <td></td>
 </tr>
 <tr>
 <td>
 NanoDrop 1000 Spectrophotometer </td>
 <td>
 Thermo-Scientific</td>
 <td></td>
 <td></td>
 </tr>
 <tr>
 <td></td>
 <td></td>
 <td></td>
 <td>Positive control primer sequences (TCF7L2-1) 
 Forward- 5'-TCGCCCTGTCAATAATCTCC-3' 
 Reverse- 5'-GCTCACCTCCTGTATCTTCG-3' 
 Negative control primer sequences (CTRL-1) 
 Forward-5'-ATGTGGTGTGGCTGTGATGGGAAC-3'
 
 Reverse- 5'-CGAGCAATCGGTAAATAGGTCTGG-3'</td>
 </tr>
 </tbody>
</table>

generation of high quality chromatin immunoprecipitation dna template for high-throughput sequencing (chip-seq)

ChIP-sequencing (ChIP-seq) methods directly offer whole-genome coverage, where combining chromatin immunoprecipitation (ChIP) and massively parallel sequencing can be utilized to identify the repertoire of mammalian DNA sequences bound by transcription factors in vivo. "Next-generation" genome sequencing technologies provide 1-2 orders of magnitude increase in the amount of sequence that can be cost-effectively generated over older technologies thus allowing for ChIP-seq methods to directly provide whole-genome coverage for effective profiling of mammalian protein-DNA interactions.
For successful ChIP-seq approaches, one must generate high quality ChIP DNA template to obtain the best sequencing outcomes. The description is based around experience with the protein product of the gene most strongly implicated in the pathogenesis of type 2 diabetes, namely the transcription factor transcription factor 7-like 2 (TCF7L2). This factor has also been implicated in various cancers.
Outlined is how to generate high quality ChIP DNA template derived from the colorectal carcinoma cell line, HCT116, in order to build a high-resolution map through sequencing to determine the genes bound by TCF7L2, giving further insight in to its key role in the pathogenesis of complex traits.

Once the chromatin has been sonicated and have been treated with RNase and Proteinase, the samples run on the 2% agarose gel should present a smear with the bulk of the DNA at the desired size. If several different cycles are tested, a gradual decrease in size should be seen as the number of cycles increase (Figure 2).
After completing the immunoprecipitation portion of the protocol the enrichment can either be checked by PCR or real-time PCR. For PCR samples run on an a...

Watch this Scientific Journal Video about Generation of High Quality Chromatin Immunoprecipitation DNA Template for High-throughput Sequencing ChIP-seq at JoVE.com

Generation of High Quality Chromatin Immunoprecipitation DNA Template for High-throughput Sequencing ChIP-seq

The combination of chromatin immunoprecipitation and ultra-high-throughput sequencing (ChIP-seq) can identify and map protein-DNA interactions in a given tissue or cell line. Outlined is how to generate a high quality ChIP template for subsequent sequencing, using experience with the transcription factor TCF7L2 as an example.

Generation of High Quality Chromatin Immunoprecipitation DNA Template for High-throughput Sequencing (ChIP-seq)

ChIP-sequencing (ChIP-seq) methods directly  offer whole-genome coverage, where combining chromatin immunoprecipitation  (ChIP) and massively parallel ...

generation-high-quality-chromatin-immunoprecipitation-dna-template

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24.3K 조회수.  Children's Hospital of Philadelphia Research Institute. 염색질의 immunoprecipitation과 매우 높은 처리량 시퀀싱 (칩 SEQ)의 조합은 특정 조직이나 세포주에서 단백질 DNA 상호 작용을 파악하고 매핑 할 수 있습니다. 설명은 예를 들어 전사 인자 TCF7L2 경험을 사용하여 다음 순서를위한 고품질 칩 템플릿을 생성하는 방법이다.

비디오: Generation of High Quality Chromatin Immunoprecipitation DNA Template for High-throughput Sequencing ChIP-seq

Highly Efficient Ligation of Small RNA Molecules for MicroRNA Quantitation by High-Throughput Sequencing

MiRNA cloning and high-throughput sequencing, termed miR-Seq, stands alone as a transcriptome-wide approach to quantify miRNAs with single nucleotide resolution. This technique captures miRNAs by attaching 3&#8217; and 5&#8217; oligonucleotide adapters to miRNA molecules and allows de novo miRNA discovery. Coupling with powerful next-generation sequencing platforms, miR-Seq has been instrumental in the study of miRNA biology. However, significant biases introduced by oligonucleotide ligation steps have prevented miR-Seq from being employed as an accurate quantitation tool. Previous studies demonstrate that biases in current miR-Seq methods often lead to inaccurate miRNA quantification with errors up to 1,000-fold for some miRNAs1,2. To resolve these biases imparted by RNA ligation, we have developed a small RNA ligation method that results in ligation efficiencies of over 95% for both 3&#8217; and 5&#8242; ligation steps. Benchmarking this improved library construction method using equimolar or differentially mixed synthetic miRNAs, consistently yields reads numbers with less than two-fold deviation from the expected value. Furthermore, this high-efficiency miR-Seq method permits accurate genome-wide miRNA profiling from in vivo total RNA samples2.

MicroRNAs (miRNAs) are a widely conserved class of regulatory molecules. Here we describe a miRNA cloning method that relies upon two potent ligation steps followed by high-throughput sequencing. Our method permits accurate genome-wide quantitation of miRNAs.

MiRNA cloning and high-throughput sequencing, termed miR-Seq, stands alone as a transcriptome-wide approach to quantify miRNAs with single nucleotide ...

Targeted DNA Methylation Analysis by Next-generation Sequencing

The role of epigenetic processes in the control of gene expression has been known for a number of years. DNA methylation at cytosine residues is of particular interest for epigenetic studies as it has been demonstrated to be both a long lasting and a dynamic regulator of gene expression. Efforts to examine epigenetic changes in health and disease have been hindered by the lack of high-throughput, quantitatively accurate methods. With the advent and popularization of next-generation sequencing (NGS) technologies, these tools are now being applied to epigenomics in addition to existing genomic and transcriptomic methodologies. For epigenetic investigations of cytosine methylation where regions of interest, such as specific gene promoters or CpG islands, have been identified and there is a need to examine significant numbers of samples with high quantitative accuracy, we have developed a method called Bisulfite Amplicon Sequencing (BSAS). This method combines bisulfite conversion with targeted amplification of regions of interest, transposome-mediated library construction and benchtop NGS. BSAS offers a rapid and efficient method for analysis of up to 10 kb of targeted regions in up to 96 samples at a time that can be performed by most research groups with basic molecular biology skills. The results provide absolute quantitation of cytosine methylation with base specificity. BSAS can be applied to any genomic region from any DNA source. This method is useful for hypothesis testing studies of target regions of interest as well as confirmation of regions identified in genome-wide methylation analyses such as whole genome bisulfite sequencing, reduced representation bisulfite sequencing, and methylated DNA immunoprecipitation sequencing.

Bisulfite amplicon sequencing (BSAS) is a method for quantifying cytosine methylation in targeted genomic regions of interest. This method uses bisulfite conversion paired with PCR amplification of target regions prior to next-generation sequencing to produce absolute quantitation of DNA methylation at a base-specific level.

The role of epigenetic processes in the control of gene expression has been known for a number of years. DNA methylation at cytosine residues is of ...

High-throughput Screening for Chemical Modulators of Post-transcriptionally Regulated Genes

Both transcriptional and post-transcriptional regulation have a profound impact on genes expression. However, commonly adopted cell-based screening assays focus on transcriptional regulation, being essentially aimed at the identification of promoter-targeting molecules. As a result, post-transcriptional mechanisms are largely uncovered by gene expression targeted drug development. Here we describe a cell-based assay aimed at investigating the role of the 3&#39; untranslated region (3&#8217; UTR) in the modulation of the fate of its mRNA, and at identifying compounds able to modify it. The assay is based on the use of a luciferase reporter construct containing the 3&#8217; UTR of a gene of interest stably integrated into a disease-relevant cell line. The protocol is divided into two parts, with the initial focus on the primary screening aimed at the identification of molecules affecting luciferase activity after 24 hr of treatment. The second part of the protocol describes the counter-screening necessary to discriminate compounds modulating luciferase activity specifically through the 3&#8217; UTR. In addition to the detailed protocol and representative results, we provide important considerations about the assay development and the validation of the hit(s) on the endogenous target. The described cell-based reporter gene assay will allow scientists to identify molecules modulating protein levels via post-transcriptional mechanisms dependent on a 3&#8217; UTR.

Here we describe a cell-based reporter gene assay as a valuable tool to screen chemical libraries for compounds modulating post-transcriptional control mechanisms exerted through 3&#8217; UTR.

Both transcriptional and post-transcriptional regulation have a profound impact on genes expression. However, commonly adopted cell-based screening assays ...

Chromatin Immunoprecipitation Assay for the Identification of Arabidopsis Protein-DNA Interactions In Vivo

Intricate gene regulatory networks orchestrate biological processes and developmental transitions in plants. Selective transcriptional activation and silencing of genes mediate the response of plants to environmental signals and developmental cues. Therefore, insights into the mechanisms that control plant gene expression are essential to gain a deep understanding of how biological processes are regulated in plants. The chromatin immunoprecipitation (ChIP) technique described here is a procedure to identify the DNA-binding sites of proteins in genes or genomic regions of the model species Arabidopsis thaliana. The interactions with DNA of proteins of interest such as transcription factors, chromatin proteins or posttranslationally modified versions of histones can be efficiently analyzed with the ChIP protocol. This method is based on the fixation of protein-DNA interactions in vivo, random fragmentation of chromatin, immunoprecipitation of protein-DNA complexes with specific antibodies, and quantification of the DNA associated with the protein of interest by PCR techniques. The use of this methodology in Arabidopsis has contributed significantly to unveil transcriptional regulatory mechanisms that control a variety of plant biological processes. This approach allowed the identification of the binding sites of the Arabidopsis chromatin protein EBS to regulatory regions of the master gene of flowering FT. The impact of this protein in the accumulation of particular histone marks in the genomic region of FT was also revealed through ChIP analysis.

Chromatin Immunoprecipitation Assay for the Identification of Arabidopsis Protein-DNA Interactions In Vivo

Chromatin immunoprecipitation is a powerful technique for the identification of DNA binding sites of Arabidopsis proteins in vivo. This procedure includes chromatin cross-linking and fragmentation, immunoprecipitation with selective antibodies against the protein of interest, and qPCR analysis of bound DNA. We describe a simple ChIP assay optimized for Arabidopsis plants.

Intricate gene regulatory networks orchestrate biological processes and developmental transitions in plants. Selective transcriptional activation and ...

DamID-seq: Genome-wide Mapping of Protein-DNA Interactions by High Throughput Sequencing of Adenine-methylated DNA Fragments

The DNA adenine methyltransferase identification (DamID) assay is a powerful method to detect protein-DNA interactions both locally and genome-wide. It is an alternative approach to chromatin immunoprecipitation (ChIP). An expressed fusion protein consisting of the protein of interest and the E. coli DNA adenine methyltransferase can methylate the adenine base in GATC motifs near the sites of protein-DNA interactions. Adenine-methylated DNA fragments can then be specifically amplified and detected. The original DamID assay detects the genomic locations of methylated DNA fragments by hybridization to DNA microarrays, which is limited by the availability of microarrays and the density of predetermined probes. In this paper, we report the detailed protocol of integrating high throughput DNA sequencing into DamID (DamID-seq). The large number of short reads generated from DamID-seq enables detecting and localizing protein-DNA interactions genome-wide with high precision and sensitivity. We have used the DamID-seq assay to study genome-nuclear lamina (NL) interactions in mammalian cells, and have noticed that DamID-seq provides a high resolution and a wide dynamic range in detecting genome-NL interactions. The DamID-seq approach enables probing NL associations within gene structures and allows comparing genome-NL interaction maps with other functional genomic data, such as ChIP-seq and RNA-seq.

We describe herein an assay by coupling DNA adenine methyltransferase identification (DamID) to high throughput sequencing (DamID-seq). This improved method provides a higher resolution and a wider dynamic range, and allows analyzing DamID-seq data in conjunction with other high throughput sequencing data such as ChIP-seq, RNA-seq, etc.

The DNA adenine methyltransferase identification (DamID) assay is a powerful method to detect protein-DNA interactions both locally and genome-wide. It is ...

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 ...

High-throughput CRISPR Vector Construction and Characterization of DNA Modifications by Generation of Tomato Hairy Roots

Targeted DNA mutations generated by vectors with clustered regularly interspaced short palindromic repeats (CRISPR)/Cas9 technology have proven useful for functional genomics studies. While most cloning strategies are simple to perform, they generally use multiple steps and can require several days to generate the ultimate constructs. The method presented here is based on DNA assembly and can produce fully functional CRISPR vectors in a single cloning reaction. Vector construction can also be pooled, further increasing the efficiency and utility of the process. A modification of the method is used to create CRISPR vectors with multiple gene targets. CRISPR vectors are then transformed into tomato hairy roots to generate transgenic materials with targeted DNA modifications. Hairy roots are a useful system for testing vector functionality as they are technically simple to generate and amenable to large-scale production. The methods presented here will have wide application as they can be used to generate a variety of CRISPR vectors and be used in a wide range of plant species.

Using DNA assembly, multiple CRISPR vectors can be constructed in parallel in a single cloning reaction, making the construction of large numbers of CRISPR vectors a simple task. Tomato hairy roots are an excellent model system to validate CRISPR vectors and generate mutant materials.

Targeted DNA mutations generated by vectors with clustered regularly interspaced short palindromic repeats (CRISPR)/Cas9 technology have proven useful for ...

A Protocol for Functional Assessment of Whole-Protein Saturation Mutagenesis Libraries Utilizing High-Throughput Sequencing

Site-directed mutagenesis has long been used as a method to interrogate protein structure, function and evolution. Recent advances in massively-parallel sequencing technology have opened up the possibility of assessing the functional or fitness effects of large numbers of mutations simultaneously. Here, we present a protocol for experimentally determining the effects of all possible single amino acid mutations in a protein of interest utilizing high-throughput sequencing technology, using the 263 amino acid antibiotic resistance enzyme TEM-1 &#946;-lactamase as an example. In this approach, a whole-protein saturation mutagenesis library is constructed by site-directed mutagenic PCR, randomizing each position individually to all possible amino acids. The library is then transformed into bacteria, and selected for the ability to confer resistance to &#946;-lactam antibiotics. The fitness effect of each mutation is then determined by deep sequencing of the library before and after selection. Importantly, this protocol introduces methods which maximize sequencing read depth and permit the simultaneous selection of the entire mutation library, by mixing adjacent positions into groups of length accommodated by high-throughput sequencing read length and utilizing orthogonal primers to barcode each group. Representative results using this protocol are provided by assessing the fitness effects of all single amino acid mutations in TEM-1 at a clinically relevant dosage of ampicillin. The method should be easily extendable to other proteins for which a high-throughput selection assay is in place.

We present a protocol for the functional assessment of comprehensive single-site saturation mutagenesis libraries of proteins utilizing high-throughput sequencing. Importantly, this approach uses orthogonal primer pairs to multiplex library construction and sequencing. Representative results using TEM-1 &#946;-lactamase selected at a clinically relevant dosage of ampicillin are provided.

Site-directed mutagenesis has long been used as a method to interrogate protein structure, function and evolution. Recent advances in massively-parallel ...

Parallel High Throughput Single Molecule Kinetic Assay for Site-Specific DNA Cleavage

Site-specific DNA cleavage (SSDC) is a key step in many cellular processes, and it is crucial to gene editing. This work describes a kinetic assay capable of measuring SSDC in many single DNA molecules simultaneously. Bead-tethered substrate DNAs, each containing a single copy of the target sequence, are prepared in a microfluidic flow channel. An external magnet applies a weak force to the paramagnetic beads. The integrity of up to 1,000 individual DNAs can be monitored by visualizing the microbeads under darkfield imaging using a wide-field, low magnification objective. Injecting of a restriction endonuclease, NdeI, initiates the cleavage reaction. Video microscopy is used to record the exact moment of each DNA cleavage by observing the frame in which the associated bead moves up and out of the focal plane of the objective. Frame-by-frame bead counting quantifies the reaction, and an exponential fit determines the reaction rate. This method allows collection of quantitative and statistically significant data on single molecule SSDC reactions in a single experiment.

A highly parallel method for measuring the site-specific cleavage of DNA at the single molecule level is described. This protocol demonstrates the technique using the restriction endonuclease NdeI. The method can easily be modified to study any process that results in site-specific DNA cleavage.

Site-specific DNA cleavage (SSDC) is a key step in many cellular processes, and it is crucial to gene editing. This work describes a kinetic assay capable ...

Modified Methods for Loading of High-Throughput DNA Extraction Plates Reduce Potential for Contamination

High-throughput DNA sequencing techniques have contributed substantially to advances in our understanding of relationships among microbial communities, host characteristics, and broader ecosystem functions. With this rapid increase in breadth and depth of sequencing capabilities have come methods to extract, amplify, analyze, and interpret environmental DNA successfully with maximum efficiency. Unfortunately, performing DNA extractions quickly can come at the cost of increasing the risk of contamination among samples. In particular, high-throughput extractions that are based on samples contained in a 96-well plate offer a relatively quick method, compared to single-tube extractions, but also increase opportunities for well-to-well cross-contamination. To minimize the risk of cross-contamination among samples, while retaining the benefits of high-throughput extraction techniques, we developed a new method for loading environmental samples into 96-well plates. We used pierceable PCR sealing films to cover each plate while loading samples and added samples first to PCR tubes before moving them into wells; together, these practices reduce the risk of sample drift and unintended double loading of wells. The method outlined in this paper provides researchers with an approach to maximize available high-throughput extraction techniques while reducing the risk of cross-contamination inherent to 96-well plates. We provide a detailed step by step outline of how to move from sample collection to DNA extraction while minimizing the risk of unwanted cross-contamination.

Current methods for loading 96-well plates for DNA extractions can be prone to contamination. We detail a new method for loading 96-well plates that reduces risk of cross-contamination among wells. Our method will help other researchers capitalize on the efficiency of high-throughput DNA extraction techniques and minimize risk of contamination.

High-throughput DNA sequencing techniques have contributed substantially to advances in our understanding of relationships among microbial communities, ...