This research was supported by the Microbial Ecology Collaborative with funding from NSF award #EPS-1655726.

1. Laboratory bench and tool preparation for loading a 96-well plate
<ol>
	<li>Clear bench top by misting with 70% ethanol. Wipe and let air dry before spraying bench top with 10% bleach. Wipe the bench top dry.</li>
	<li>Sterilize micro-scoopula, spatula, and surgical scissors (curved) by dipping in 95% ethanol and then expose to a flame. Dip each in 10% bleach and allow to air dry prior to use. 
	NOTE: Tools should be sterilized between each sample.</li>
</ol>
2. Sub-sampling and sample preparation
<ol>
	<li>Sterilize gloves using ethanol prior to sub-sampling.</li>
	<li>Homogenize soil samples thoroughly prior to sub-sampling.</li>
	<li>Using sterilized tools, load a labeled 2 mL centrifuge tube with sample until approximately&#160;half full.</li>
	<li>Repeat step 2.2 until all 95 samples have been sub-sampled into labeled 2 mL centrifuge tubes. The 96th well should be used as an extraction blank. 
	NOTE: Steps 2.3 and 2.4 are done to minimize required storage space and to prevent over or double sampling.</li>
	<li>Store 2 mL sub-sample tubes on ice until loading of plate.</li>
	<li>Label 96 sterile 200 &#181;L flat-capped PCR tubes A1&#8210;A12, B1&#8210;B12, &#8230;. H1&#8210;H12. Place these tubes in order into a 96-well rack.</li>
	<li>Assign a sample ID with a well location (A1&#8210;H12) and record it.</li>
</ol>
3. Plate preparation
<ol>
	<li>Remove the rubber cover from 96-well plate (Figure 1A&#8211;1C) and place it into a sterile plastic bag (see Table of Materials). Seal the plastic bag to prevent contamination.</li>
	<li>Cover the 96-well plate with sealing film (Figure 1D&#8211;1E); for example, use a precut pierceable sealing film (see the Table of Materials). Ensure seal by rolling with a rubber roller. 
	NOTE: Pierceable silicone mats could potentially offer a reusable option, provided appropriate cleaning between uses, but the silicone mats that we tested easily split along the pierces and would not offer equal protection to any following plates.</li>
	<li>Place plate in refrigerator (4 &#176;C) to keep cool.</li>
</ol>
4. Transfer of sub-samples
NOTE: See step 4.13 for modifications when soil sample is very small.
<ol>
	<li>Place 24 of the 2 mL sub-samples into an ice block for cold storage.</li>
	<li>Using the sample name and well location sheet choose the correct 200 &#181;L flat-capped PCR tube.</li>
	<li>Take a 2 mL sub-sample tube and vortex for ~5 s to ensure homogenization.</li>
	<li>Using flame and bleach sterilized tools load ~200 &#181;L of the first sample into the correct flat-capped PCR tube (Figure 1F).</li>
	<li>Repeat steps 4.2&#8210;4.4 until all 24 samples have been loaded. Then move to step 4.6.</li>
	<li>Using a bleach-dipped paper wipe, clean the outside of the 200 &#181;L flat-capped PCR tube.</li>
	<li>Invert the tube and tap on the bench to move sample to top of the tube. With bleached and flame-sterilized scissors, clip the bottom of the PCR tube to create an opening for sample to fall into the 96-well plate (Figure 1G).</li>
	<li>Locate the correct well on a 96-well plate and pass the tube across plate with cut end facing up until reaching that well. Tilt the plate slightly to facilitate puncturing of precut pierceable sealing film, and only when tube is directly above the correct well carefully invert it so that cut tip fits into the well. Using sterilized tools, tap the top of the PCR tube until all soil has fallen from the tube into the well. Leave the tube within the well with lid closed (Figure 1H&#8211;1J).</li>
	<li>Remove one flat-capped 200 &#956;L PCR tube at a time and add 750 &#956;L of bead solution to that well&#160;(Figure 1K). Push the 200 &#956;L flat-capped PCR tube all the way down into the well and mark the top with sharpie to indicate that this well has been loaded. Repeat this for each sample.</li>
	<li>Once all 24 samples have been loaded and bead solution added, remove the 2 mL sub-sample tubes and replace with 24 unsampled tubes.&#160;</li>
	<li>Repeat steps 4.1&#8210;4.10 until all 95 wells have been loaded with sample or blank and bead solution.&#160;Remove tubes without passing them over open wells (Figure 1L).</li>
	<li>Carefully remove pierceable film and replace the rubber cover to protect the plate until beginning extraction (Figure 1M&#8211;1O).</li>
	<li>For very small quantities of soil that are also fine-grained, add 750 &#181;L of bead solution to the sample tube and use a wide-bore pipette tip to transfer all contents to the appropriate well. Place PCR tube into the opening to prevent double loading. 
	NOTE: Be careful when implementing this modification as particulate organic matter or small rock fragments larger than the bore of the pipette tip can clog the pipette and make transfer of the sample slurry difficult.</li>
	<li>As needed freeze at -20 &#176;C and store plates loaded with sample and bead solution until planned extraction.</li>
</ol>
5. Comparison of plate loading methods
NOTE: In order to verify our novel method for loading of 96-well DNA extraction plates, we divided a single 96-well plate into three sections. We used three different methods of plate loading to compare potential for unintentional loading of soil and cross contamination. The three methods we used were the methods outlined in McPherson et al.9, Qiagen&#8217;s default loading protocol10, and the protocol we outline in this publication.
<ol>
	<li>Loading soil into the plate
	<ol>
		<li>Section a 96-well plate into three four-column blocks. Load each block using one of the three method mentioned above.</li>
		<li>Load soil into every other well so that sample and blank wells are staggered. This arrangement allows for maximum opportunity for inadvertent sample loading and cross contamination.</li>
		<li>Freeze the 96-well plate at -20 &#176;C until DNA extraction.</li>
	</ol>
	</li>
	<li>DNA extraction
	<ol>
		<li>Extract DNA according to manufacturer&#8217;s 96-well plate extraction protocol10.</li>
		<li>Store the DNA extracts at -20 &#176;C until further analysis.</li>
	</ol>
	</li>
	<li>Quantification of DNA in blank wells
	<ol>
		<li>Thaw DNA extracts at room temperature, vortex and spin down using a plate spinner.</li>
		<li>Measure and record DNA concentrations of blank wells using a spectrophotometer.</li>
		<li>Use ANVOA and Tukey&#8217;s post-hoc test to analyze differences in mean DNA concentration in blank wells. 
		NOTE: Significant differences were reported at &#945; = 0.05.</li>
	</ol>
	</li>
</ol>

<ol>
	<li>Thompson, L. R., 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=A+communal+catalogue+reveals+Earth's+multiscale+microbial+diversity.">A communal catalogue reveals Earth's multiscale microbial diversity.</a> Nature. 551 (7681), 457-463 (2017).</li><li>Davis, A., 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=Improved+yield+and+accuracy+for+DNA+extraction+in+microbiome+studies+with+variation+in+microbial+biomass.">Improved yield and accuracy for DNA extraction in microbiome studies with variation in microbial biomass.</a> BioTechniques. 66 (6), 285-289 (2019).</li><li>Marotz, C., 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=Triplicate+PCR+reactions+for+16S+rRNA+gene+amplicon+sequencing+are+unnecessary.">Triplicate PCR reactions for 16S rRNA gene amplicon sequencing are unnecessary.</a> BioTechniques. 67 (1), 29-32 (2019).</li><li>Romdhane, S., 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=Cover+crop+management+practices+rather+than+composition+of+cover+crop+mixtures+affect+bacterial+communities+in+no-till+agroecosystems.">Cover crop management practices rather than composition of cover crop mixtures affect bacterial communities in no-till agroecosystems.</a> Frontiers in Microbiology. 10, 1618 (2019).</li><li>Rochefort, A., 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=Influence+of+environment+and+host+plant+genotype+on+the+structure+and+diversity+of+the+Brassica+napus+seed+microbiota.">Influence of environment and host plant genotype on the structure and diversity of the Brassica napus seed microbiota.</a> Phytobiomes Journal. 3 (4), 326-336 (2019).</li><li>Minich, J. J., 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=Quantifying+and+understanding+well-to-well+contamination+in+microbiome+research.">Quantifying and understanding well-to-well contamination in microbiome research.</a> mSystems. 4 (4), 00186 (2019).</li><li>Walker, A. W. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+lot+on+your+plate?+Well-to-well+contamination+as+an+additional+confounder+in+microbiome+sequence+analyses.">A lot on your plate? Well-to-well contamination as an additional confounder in microbiome sequence analyses.</a> mSystems. 4 (4), 00362 (2019).</li><li>Eisenhofer, R., 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=Contamination+in+low+microbial+biomass+microbiome+studies:+issues+and+recommendations.">Contamination in low microbial biomass microbiome studies: issues and recommendations.</a> Trends in Microbiology. 27 (2), 105-117 (2019).</li><li>McPherson, M. R., 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=Isolation+and+analysis+of+microbial+communities+in+soil,+rhizosphere,+and+roots+in+perennial+grass+Experiments.">Isolation and analysis of microbial communities in soil, rhizosphere, and roots in perennial grass Experiments.</a> Journal of Visualized Experiments. (137), e57932 (2018).</li><li>Qiagen, D. N., 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=.">.</a> easy PowerSoil HTP 96 Kit Handbook. , (2019).</li></ol>

The authors report no conflicts of interest and have nothing to disclose.

This method reduces opportunities for well-to-well cross-contamination while loading high-throughput sample extraction plates and offers a more controlled means to load 96-well plates beyond existing plate loading strategies9,10. Contamination among wells can be more pervasive in 96-well plate extractions than in single-tube extractions, especially when automated6,7, and, though not specifically tested in...

Recent advances in high-throughput sequencing of microbial communities are providing unparalleled sequencing depth and, consequently, an unprecedent glimpse into the functioning and diversity of Earth&#39;s microbiome1. As the ability to multiplex more and more samples onto a single sequencing lane increases, single tube DNA extraction has the potential to become a rate-limiting step in the generation of ecological data. However, new methods in high-throughput DNA extractions hold promise for processing large quantities of environmental samples with greater efficiency than has previously been possible2. These methods often involve using 96-well plates instead of single-tubes, thereby increasing the possible number of extractions that can occur simultaneously. As such, the practicality and efficiency of high-throughput extraction methods are evident and have been implemented for processing of environmental samples ranging from soil3,4 and plant tissues3,5 to human fecal matter2.
While these methods can dramatically speed along sample processing and DNA extraction, the initial step of loading soil and other ecological samples into 96-well plates is susceptible to cross-sample contamination. This type of well-to-well contamination can occur during DNA extractions6,7,8, and wells are particularly vulnerable in this first step before samples are suspended in a buffer solution. McPherson et al.9 demonstrate a method to load rhizosphere soils into 96-well plates using funnels and 8-well PCR strip covers, but while their method is a more controlled approach to loading plates, it still provides ample opportunity for contamination of neighboring wells when loading each sample. Additionally, the open wells allow the chance for a distracted researcher to place a sample into the incorrect well or add a sample into a well that already has been loaded. In addition, a variety of sample types prove to be ill-suited for loading with this method; wet samples often stick to the funnels and dry samples &#8216;jump&#8217; between wells due to static electricity.
To reduce opportunities for contamination among wells in the first step of high-throughput DNA extractions, we developed a new approach to loading soil samples into 96-well plates. Our methods both protect wells from environmental exposure and prevent us from accidentally loading multiple samples into one well (double-loading). We believe the method reported below offers promise to reduce the potential for contamination and as such provides a more controlled means to load 96-well plates for subsequent DNA extraction.

<table><tbody><tr><td>18 oz WhirlPak</td><td>Nasco</td><td>B01065</td><td></td></tr><tr><td>2 mL centrifuge tubes</td><td>Fisherbrand</td><td>05-408-129</td><td></td></tr><tr><td>200 &amp;micro;L micro-PCR tubes</td><td>Thermo Scientific</td><td>AB 2000</td><td></td></tr><tr><td>96-well PowerSoil DNA extraction kit</td><td>Qiagen</td><td>12955-4</td><td>We used a soil extraction kit but any 96-well format kit would work.</td></tr><tr><td>Ice block for 2 mL centrifuge tubes</td><td>Any</td><td>Any</td><td>Any ice block made for 2 mL tubes will work</td></tr><tr><td>Ice block for 200 &amp;micro;L micro-PCR tubes</td><td>Any</td><td>Any</td><td>Any ice block made for 200 &amp;micro;L tubes will work</td></tr><tr><td>Micro scoopula</td><td>Any</td><td>Any</td><td></td></tr><tr><td>Precut Pierceable Sealing Film</td><td>Excel Scientific</td><td>XPS25</td><td></td></tr><tr><td>Spatula</td><td>Any</td><td>Any</td><td></td></tr><tr><td>Surgical scissors</td><td>Any</td><td>Any</td><td></td></tr></tbody></table>

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.

This novel method was used successfully to load 96-well DNA extraction plates. Comparison of plate loading methods showed our method to have the lowest DNA concentration in the blank wells. The DNA concentration in the blank wells was significantly lower than the method proposed my McPherson et al.9 (p &#60; 0.05), though DNA concentrations in our method were not statistically different from the Qiagen default method (Figure 2). All three methods produced mean DNA con...

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

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Research

JoVE Journal

Biology

5.1K Views.  University of Wyoming. Dit protocol biedt een techniek voor het verhogen van de efficiëntie van 96-well DNA-extractieplaat laden, terwijl tegelijkertijd het verminderen van het risico van kruisbesmetting van monsters. Deze techniek vermindert de kans op besmetting tijdens de kritische eerste stap van DNA-extractie door de individuele toevoeging van monsters aan elke put in de 96-put plaat. Deze methodologie kan worden toegepast op een van de verschillende gebieden van microbioom onderzoek.Voor het begin van...

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

A Robotic Platform for High-throughput Protoplast Isolation and Transformation

Over the last decade there has been a resurgence in the use of plant protoplasts that range from model species to crop species, for analysis of signal transduction pathways, transcriptional regulatory networks, gene expression, genome-editing, and gene-silencing. Furthermore, significant progress has been made in the regeneration of plants from protoplasts, which has generated even more interest in the use of these systems for plant genomics. In this work, a protocol has been developed for automation of protoplast isolation and transformation from a &#39;Bright Yellow&#39; 2 (BY-2) tobacco suspension culture using a robotic platform. The transformation procedures were validated using an orange fluorescent protein (OFP) reporter gene (pporRFP) under the control of the Cauliflower mosaic virus 35S promoter (35S). OFP expression in protoplasts was confirmed by epifluorescence microscopy. Analyses also included protoplast production efficiency methods using propidium iodide. Finally, low-cost food-grade enzymes were used for the protoplast isolation procedure, circumventing the need for lab-grade enzymes that are cost-prohibitive in high-throughput automated protoplast isolation and analysis. Based on the protocol developed in this work, the complete procedure from protoplast isolation to transformation can be conducted in under 4 hr, without any input from the operator. While the protocol developed in this work was validated with the BY-2 cell culture, the procedures and methods should be translatable to any plant suspension culture/protoplast system, which should enable acceleration of crop genomics research.

A high-throughput, automated, tobacco protoplast production and transformation methodology is described. The robotic system enables massively parallel gene expression and discovery in the model BY-2 system that should be translatable to non-model crops.

Over the last decade there has been a resurgence in the use of plant protoplasts that range from model species to crop species, for analysis of signal ...

Genetics

Exploring the Root Microbiome: Extracting Bacterial Community Data from the Soil, Rhizosphere, and Root Endosphere

The intimate interaction between plant host and associated microorganisms is crucial in determining plant fitness, and can foster improved tolerance to abiotic stresses and diseases. As the plant microbiome can be highly complex, low-cost, high-throughput methods such as amplicon-based sequencing of the 16S rRNA gene are often preferred for characterizing its microbial composition and diversity. However, the selection of appropriate methodology when conducting such experiments is critical for reducing biases that can make analysis and comparisons between samples and studies difficult. This protocol describes in detail a standardized methodology for the collection and extraction of DNA from soil, rhizosphere, and root samples. Additionally, we highlight a well-established 16S rRNA amplicon sequencing pipeline that allows for the exploration of the composition of bacterial communities in these samples, and can easily be adapted for other marker genes. This pipeline has been validated for a variety of plant species, including sorghum, maize, wheat, strawberry, and agave, and can help overcome issues associated with the contamination from plant organelles.

Here, we describe a protocol to obtain amplicon sequence data of soil, rhizosphere, and root endosphere microbiomes. This information can be used to investigate the composition and diversity of plant-associated microbial communities, and is suitable for the use with a wide range of plant species.

The intimate interaction between plant host and associated microorganisms is crucial in determining plant fitness, and can foster improved tolerance to ...

High-throughput, Automated Extraction of DNA and RNA from Clinical Samples using TruTip Technology on Common Liquid Handling Robots

TruTip is a simple nucleic acid extraction technology whereby a porous, monolithic binding matrix is inserted into a pipette tip. The geometry of the monolith can be adapted for specific pipette tips ranging in volume from 1.0 to 5.0 ml. The large porosity of the monolith enables viscous or complex samples to readily pass through it with minimal fluidic backpressure. Bi-directional flow maximizes residence time between the monolith and sample, and enables large sample volumes to be processed within a single TruTip. The fundamental steps, irrespective of sample volume or TruTip geometry, include cell lysis, nucleic acid binding to the inner pores of the TruTip monolith, washing away unbound sample components and lysis buffers, and eluting purified and concentrated nucleic acids into an appropriate buffer. The attributes and adaptability of TruTip are demonstrated in three automated clinical sample processing protocols using an Eppendorf epMotion 5070, Hamilton STAR and STARplus liquid handling robots, including RNA isolation from nasopharyngeal aspirate, genomic DNA isolation from whole blood, and fetal DNA extraction and enrichment from large volumes of maternal plasma (respectively).

TruTip is a simple nucleic acid extraction technology whereby a porous, monolithic binding matrix is inserted into a pipette tip. Consequently, the sample preparation format is compatible with most liquid handling instruments, and can be used for many medium to high-throughput clinical applications and sample types.

TruTip is a simple nucleic acid extraction  technology whereby a porous, monolithic binding matrix is inserted into a  pipette tip. The geometry of the ...

Bioengineering

High-Throughput DNA Plasmid Multiplexing and Transfection Using Acoustic Nanodispensing Technology

Cell transfection, indispensable for many biological studies, requires controlling many parameters for an accurate and successful achievement. Most often performed at low throughput, it is moreover time-consuming and error-prone, even more so when multiplexing several plasmids. We developed an easy, fast, and accurate method to perform cell transfection in a 384-well plate layout using acoustic droplet ejection (ADE) technology. The nanodispenser device used in this study&#160;is based on this technology and allows precise nanovolume delivery at high speed from a source well plate to a destination one. It can dispense and multiplex DNA and transfection reagent according to a predesigned spreadsheet. Here we present an optimal protocol to perform ADE-based high-throughput plasmid transfection which makes it possible to reach an efficiency of up to 90% and a nearly 100% cotransfection in cotransfection experiments. We extend initial work by proposing a user-friendly spreadsheet-based macro, able to manage up to four plasmids/wells from a library containing up to 1,536 different plasmids, and a tablet-based pipetting guide application. The macro designs the necessary template(s) of the source plate(s) and generates the ready-to-use files for the nanodispenser and tablet-based application. The four-steps transfection protocol involves i) a diluent dispense with a classical liquid handler, ii) plasmid distribution and multiplexing, iii) a transfection reagent dispense by the nanodispenser, and iv) cell plating on the prefilled wells. The described software-based control of ADE plasmid multiplexing and transfection allows even nonspecialists in the field to perform a reliable cell transfection in a fast and safe way. This method enables rapid identification of optimal settings for a given cell type and can be transposed to higher-scale and manual approaches. The protocol eases applications, such as human ORFeome protein (set of open reading frames [ORFs] in a genome) expression or CRISPR-Cas9-based gene function validation, in nonpooled screening strategies.

This protocol describes high-throughput plasmid transfection of mammalian cells in a 384-well plate using acoustic droplet ejection technology. The time-consuming, error-prone DNA dispensing and multiplexing, but also the transfection reagent dispensing, are software-driven and performed by a nanodispenser device. The cells are then seeded in these prefilled wells.

Cell transfection, indispensable for many biological studies, requires controlling many parameters for an accurate and successful achievement. Most often ...

Kinase Inhibitor Screening In Self-assembled Human Protein Microarrays

The screening of kinase inhibitors is crucial for better understanding properties of a drug and for the identification of potentially new targets with clinical implications. Several methodologies have been reported to accomplish such screening. However, each has its own limitations (e.g., the screening of only ATP analogues, restriction to using purified kinase domains, significant costs associated with testing more than a few kinases at a time, and lack of flexibility in screening protein kinases with novel mutations). Here, a new protocol that overcomes some of these limitations and can be used for the unbiased screening of kinase inhibitors is presented. A strength of this method is its ability to compare the activity of kinase inhibitors across multiple proteins, either between different kinases or different variants of the same kinase. Self-assembled protein microarrays generated through the expression of protein kinases by a human-based in vitro transcription and translation system (IVTT) are employed. The proteins displayed on the microarray are active, allowing for measurement of the effects of kinase inhibitors. The following procedure describes the protocol steps in detail, from the microarray generation and screening to the data analysis.

A detailed protocol for the generation of self-assembled human protein microarrays for the screening of kinase inhibitors is presented.

The screening of kinase inhibitors is crucial for better understanding properties of a drug and for the identification of potentially new targets with ...

Biochemistry

Streamlined Purification of Plasmid DNA From Prokaryotic Cultures

We describe the complete process of AcroPrep Advance Filter Plates for 96 plasmid preparations, starting from prokaryotic culture and ending with high purity DNA. Based on multi-well filtration for bacterial lysate clearance and DNA purification, this method creates a streamlined process for plasmid preparation. Filter plates containing silica-based media can easily be processed by vacuum filtration or centrifuge to yield appreciable quantities of plasmid DNA. Quantitative analyses determine the purified plasmid DNA is consistently of high quality with average OD260/280 ratios of 1.97. Overall, plasmid yields offer more pure DNA for downstream applications, such as sequencing and cloning. This streamlined method of using AcroPrep Advance Filter Plates allows for manual, semi-automated or fully-automated processing.

This protocol is a cost effective alternative for efficient parallel clarification and plasmid DNA purification from E. coli cultures. The AcroPrep Advance process starts with an optimized lysate clarification filter plate followed by purification on a high binding capacity DNA binding filter plate.

We describe the complete process of AcroPrep Advance Filter Plates for 96 plasmid preparations, starting from prokaryotic culture and ending with high ...

Integration of Wet and Dry Bench Processes Optimizes Targeted Next-generation Sequencing of Low-quality and Low-quantity Tumor Biopsies

All next-generation sequencing (NGS) procedures include assays performed at the laboratory bench (&#34;wet bench&#34;) and data analyses conducted using bioinformatics pipelines (&#34;dry bench&#34;). Both elements are essential to produce accurate and reliable results, which are particularly critical for clinical laboratories. Targeted NGS technologies have increasingly found favor in oncology applications to help advance precision medicine objectives, yet the methods often involve disconnected and variable wet and dry bench workflows and uncoordinated reagent sets. In this report, we describe a method for sequencing challenging cancer specimens with a 21-gene panel as an example of a comprehensive targeted NGS system. The system integrates functional DNA quantification and qualification, single-tube multiplexed PCR enrichment, and library purification and normalization using analytically-verified, single-source reagents with a standalone bioinformatics suite. As a result, accurate variant calls from low-quality and low-quantity formalin-fixed, paraffin-embedded (FFPE) and fine-needle aspiration (FNA) tumor biopsies can be achieved. The method can routinely assess cancer-associated variants from an input of 400 amplifiable DNA copies, and is modular in design to accommodate new gene content. Two different types of analytically-defined controls provide quality assurance and help safeguard call accuracy with clinically-relevant samples. A flexible &#34;tag&#34; PCR step embeds platform-specific adaptors and index codes to allow sample barcoding and compatibility with common benchtop NGS instruments. Importantly, the protocol is streamlined and can produce 24 sequence-ready libraries in a single day. Finally, the approach links wet and dry bench processes by incorporating pre-analytical sample quality control results directly into the variant calling algorithms to improve mutation detection accuracy and differentiate false-negative and indeterminate calls. This targeted NGS method uses advances in both wetware and software to achieve high-depth, multiplexed sequencing and sensitive analysis of heterogeneous cancer samples for diagnostic applications.

An integrated system for targeted next-generation sequencing of oncology specimens is described. This cross-platform system is optimized for low-quality and low-quantity tumor biopsies, accommodates low DNA inputs, includes well-characterized multi-variant controls, and features a novel variant caller that is informed by quantitative pre-analytical quality control measures.

All next-generation sequencing (NGS) procedures include assays performed at the laboratory bench (&#34;wet bench&#34;) and data analyses conducted using ...

Medicine

gDNA Enrichment by a Transposase-based Technology for NGS Analysis of the Whole Sequence of BRCA1, BRCA2, and 9 Genes Involved in DNA Damage Repair

The widespread use of Next Generation Sequencing has opened up new avenues for cancer research and diagnosis. NGS will bring huge amounts of new data on cancer, and especially cancer genetics. Current knowledge and future discoveries will make it necessary to study a huge number of genes that could be involved in a genetic predisposition to cancer. In this regard, we developed a Nextera design to study 11 complete genes involved in DNA damage repair. This protocol was developed to safely study 11 genes (ATM, BARD1, BRCA1, BRCA2, BRIP1, CHEK2, PALB2, RAD50, RAD51C, RAD80, and TP53) from promoter to 3&#39;-UTR in 24 patients simultaneously. This protocol, based on transposase technology and gDNA enrichment, gives a great advantage in terms of time for the genetic diagnosis thanks to sample multiplexing. This protocol can be safely used with blood gDNA.

gDNA Enrichment by a Transposase-based Technology for NGS Analysis of the Whole Sequence of BRCA1, BRCA2, and 9 Genes Involved in DNA Damage Repair

gDNA enrichment for NGS sequencing is an easy and powerful tool for the study of constitutional mutations. In this article, we present the procedure to analyse simply the complete sequence of 11 genes involved in DNA damage repair.

The widespread use of Next Generation Sequencing has opened up new avenues for cancer research and diagnosis. NGS will bring huge amounts of new data on ...

Efficient Nucleic Acid Extraction and 16S rRNA Gene Sequencing for Bacterial Community Characterization

There is a growing appreciation for the role of microbial communities as critical modulators of human health and disease. High throughput sequencing technologies have allowed for the rapid and efficient characterization of bacterial communities using 16S rRNA gene sequencing from a variety of sources. Although readily available tools for 16S rRNA sequence analysis have standardized computational workflows, sample processing for DNA extraction remains a continued source of variability across studies. Here we describe an efficient, robust, and cost effective method for extracting nucleic acid from swabs. We also delineate downstream methods for 16S rRNA gene sequencing, including generation of sequencing libraries, data quality control, and sequence analysis. The workflow can accommodate multiple samples types, including stool and swabs collected from a variety of anatomical locations and host species. Additionally, recovered DNA and RNA can be separated and used for other applications, including whole genome sequencing or RNA-seq. The method described allows for a common processing approach for multiple sample types and accommodates downstream analysis of genomic, metagenomic and transcriptional information.

We describe an efficient, robust, and cost effective method for extracting nucleic acid from swabs for characterization of bacterial communities using 16S rRNA gene amplicon sequencing. The method allows for a common processing approach for multiple sample types and accommodates a number of downstream analytic processes.

There is a growing appreciation for the role of microbial communities as critical modulators of human health and disease. High throughput sequencing ...

Single-Cell Factor Localization on Chromatin using Ultra-Low Input Cleavage Under Targets and Release using Nuclease

Determining the binding locations of a protein on chromatin is essential for understanding its function and potential regulatory targets. Chromatin Immunoprecipitation (ChIP) has been the gold standard for determining protein localization for over 30 years and is defined by the use of an antibody to pull out the protein of interest from sonicated or enzymatically digested chromatin. More recently, antibody tethering techniques have become popular for assessing protein localization on chromatin due to their increased sensitivity. Cleavage Under Targets &amp; Release Under Nuclease (CUT&amp;RUN) is the genome-wide derivative of Chromatin Immunocleavage (ChIC) and utilizes recombinant Protein A tethered to micrococcal nuclease (pA-MNase) to identify the IgG constant region of the antibody targeting a protein of interest, therefore enabling site-specific cleavage of the DNA flanking the protein of interest. CUT&amp;RUN can be used to profile histone modifications, transcription factors, and other chromatin-binding proteins such as nucleosome remodeling factors. Importantly, CUT&amp;RUN can be used to assess the localization of either euchromatic- or heterochromatic-associated proteins and histone modifications. For these reasons, CUT&amp;RUN is a powerful method for determining the binding profiles of a wide range of proteins. Recently, CUT&amp;RUN has been optimized for transcription factor profiling in low populations of cells and single cells and the optimized protocol has been termed ultra-low input CUT&amp;RUN (uliCUT&amp;RUN). Here, a detailed protocol is presented for single-cell factor profiling using uliCUT&amp;RUN in a manual 96-well format.

CUT&#38;RUN and its variants can be used to determine protein occupancy on chromatin. This protocol describes how to determine protein localization on chromatin using single-cell uliCUT&#38;RUN.

Determining the binding locations of a protein on chromatin is essential for understanding its function and potential regulatory targets. Chromatin ...

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

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