Pool design and oligo recovery
<ol>
<li>The first step is to design the pre-mRNA pool to be studied. This can be done using the UCSC Genome Browser and downloading particular genes, splice junctions, or other areas of interest.</li>
<li>Once the windows of interest have been selected, tile across them computationally using the following conditions: read length should be 30 nucleotides with a 10 nucleotide overlap, therefore each oligo is shifted 20 nucleotides from the prior one. *Oligo overlap should be increased in proximity to splice sites to ensure adequate coverage; increasing the overlap from 10 to 20 nucleotides can accomplish this.</li>
<li>Flank each 30-nucleotide oligo with universal primer sequences, which will be used to amplify the pool downstream. Tiled oligo orders should be submitted to Agilent, where the sequences will be printed on a custom oligonucleotide microarray.</li>
<li>To recover the DNA oligos from the microarray surface begin by placing the array cover slide in an array hybridization chamber and gently pipetting 500 &mu;L of dH2O onto the slide</li>
<li>Sandwich the array on top of the cover slide, taking care to avoid air bubbles that can disrupt the process. Be sure to place the array face down so the side with the oligos is touching the surface of the water. A good rule of thumb is "Agilent touches Agilent" meaning that the Agilent label on the array faces the Agilent label on the cover slide</li>
<li>Close the hybridization chamber and rotate overnight at 99&deg;C in a hybridization oven</li>
<li>The following day, carefully remove the array from the chamber and draw off the water, which now contains the oligos liberated from the array surface. This is the oligo pool</li>
<li>Place the pool in a 1.5 mL Eppendorf tube and sonicate the pool at 50% amplitude for 3-5 second pulses</li>
<li>Next, amplify the pool by low cycle PCR, using the universal primers with a T7 tag appended to the end. For the first round, denature at 94&deg;C for 1 minute, anneal at 55&deg;C for 20 seconds, and elongate for 1 minute at 72&deg;C. For subsequent rounds, do 10 seconds, 20 seconds, and 10 seconds at each respective temperature, with a final elongation step of 5 minutes at 72&deg;C. Because the number of cycles necessary to amplify the pool depends on the efficiency of oligo recovery, it may be necessary to try multiple cycle numbers. Be careful not to over-amplify the pool, which is signified by a smeared band on an acrylamide gel. PCR amplified samples can be stored at 4&deg;C until needed.</li>
<li>The final step in oligo preparation is to transcribe RNA using the MEGAshortscript&trade; Kit from Ambion. *The following protocol has been adapted from Ambion
<ol class="lalpha">
 <li>Thaw the T7 10x Reaction Buffer, four ribonucleotide solutions and water at room temperature while keeping the T7 Enzyme Mix on ice</li>
 <li>Assemble the reaction mixture in an RNase-free microcentrifuge tube at room temperature. Components of the reaction buffer can cause template DNA to precipitate if the reaction is mixed on ice.</li>
 <li>In the following order, pipette 3 &mu;L nuclease-free water, 2 &mu;L of the 10x reaction buffer, 2 &mu;L of each 75 mM nTP solution, 5 &mu;L of template DNA (the PCR amplified oligo pool) and 2 &mu;L of T7 enzyme, for a final volume of 20 &mu;L</li>
 <li>Flick the tube gently to mix the reaction then briefly microfuge to collect the mixture at the bottom of the tube.</li>
 <li>Incubate the reaction in a thermocycler at 37&deg;C for 2 hours. Incubation time will be template-dependent, therefore it may be necessary to use a time-course experiment to determine the optimal incubation time for maximal yield</li>
 <li>After 2 hours, remove the template DNA by adding 1 &mu;L of Turbo DNase to the reaction and incubate at 37&deg;C for an additional 15 minutes</li>
 <li>Terminate the reaction and recover the RNA by phenol/chloroform extraction and ethanol precipitation.
 <ol class="lalpha">
 <li>For a 20 &mu;L reaction volume, add 115 &mu;L of dH2O and 15 &mu;L of 3 M sodium acetate to the reaction. </li>
 <li>Mix thoroughly then add 75 &mu;L of acidic phenol and 75 &mu;L of chloroform. Mix the solution by vortexing, then microfuge for one minute at 13,000 rpm at room temperature. Transfer the aqueous layer to a new tube and repeat the extraction</li>
 <li>Transfer the aqueous layer to a new tube and add 150 &mu;L of chloroform. Vortex the mixture and centrifuge as before. Transfer the aqueous layer to a new tube. </li>
 </ol>
 </li>
 <li>Recover the RNA by adding two volumes of 100% cold ethanol to the aqueous layer and mixing well. Chill the mixture for a minimum of 15 minutes at -20&deg;C then centrifuge at 4&deg;C for 15 minutes at maximum speed to pellet the RNA. Carefully remove the supernatant and resuspend the pellet in 105 &mu;L of 0.1x Tris-EDTA (TE) Buffer..</li>
</ol>
</li>
<li>It is difficult to remove all of the free nucleotides from the T7 Kit reaction, so for the most accurate results, quantify the RNA using RiboGreen.
<ol class="lalpha">
 <li>To begin the quantitation, first dilute enough 1x TE, 2,300 &mu;L + 50&mu;L/sample.</li>
 <li>Next, make the RiboGreen. You will need 1000 &mu;L + 50 &mu;L/sample. For high range samples (1 &mu;g/mL to 20 ng/mL) mix the RiboGreen 1:200 in 1x TE; for the low range samples (50 ng/mL to 1 ng/mL) mix the RiboGreen 1:2000 in 1x TE; Pipette 50 &mu;L of RiboGreen solution into each well of an opaque plate, starting with A1 and moving down, not across</li>
 <li>Dilute an RNA stock solution to 2 &mu;g/mL for high range samples or 100 ng/&mu;L for low range samples</li>
 <li>Using the stock solutions, create a standard curve by six subsequent 1:2 dilutions of the original 2 &mu;g/mL or 100 ng/mL stock; Pipette 240 &mu;L of the stock into a PCR strip tube. In the remaining 7 tubes, pipette 120 &mu;L of 1x TE. Take 120 &mu;L from the RNA stock well, and mix with tube 2. Take 120 &mu;L from tube 2 and mix with tube 3. Repeat until tube 7. Tube 8 will only have TE and will be your blank. Pipette 50 &mu;L of the standard curve solutions into the opaque plate, A1:H2 (run the curve twice)</li>
 <li>In a new tube, mix 45 &mu;L of TE with 5 &mu;L of the oligo pool and add all 50 &mu;L to a well that contains 50 &mu;L of RiboGreen.</li>
 <li>Using a plate reader set the machine to read fluorescence from the top after a one-minute mix with an excitation wavelength of 485 nm and emission wavelength of 530 nm and read the plate</li>
 <li>Use the standard curve generated to quantify the recovered RNA, which should be stored at -20&deg;C.</li>
</ol>
</li>
</ol>
Co-immunoprecipitation of the oligo pool with an RNP of interest
<ol>
<li>Prior to beginning the co-immunoprecipitation, prepare the Protein A and Protein G Dynabeads from Invitrogen by mixing them in a 1:1 ratio up to a final volume of 50 &mu;L per reaction. For example, for two reactions you would add 50 &mu;L each of each bead. </li>
<li>Use a Magnetic Separation Stand like this one from Novagen to hold the beads in place while you remove the supernatant and wash the beads twice with an equal volume of cold 1x PBS</li>
<li>After the second wash, resuspend the beads in an equal volume of cold 1x PBS and add 2 ng of the antibody per reaction. This amount may vary depending on the antibody, so experimentation is necessary to find the optimal conditions.</li>
<li>Incubate the antibody/bead mixture overnight at 4&deg;C on a rotating platform</li>
<li>In the morning, add 2 &mu;g per reaction of sonicated yeast total RNA to block non-specific binding, and rotate for an additional 30 minutes at 4&deg;C.</li>
<li>Using the magnetic rack to hold the beads, remove the supernatant and wash the beads twice with cold 1x PBS, leaving the PBS from the second wash in the tube.</li>
<li>Take out new tubes, one for each reaction and aliquot 50 &mu;L of the bead mixture into each tube. Let the beads sit while preparing a master mix</li>
<li>For each reaction add 200 ng of the oligo pool, an equimolar ratio of protection primers, 2 &mu;g of sonicated yeast total RNA, 20 &mu;L of HeLa nuclear extract (this is the RNP source), and buffer E to bring the volume up to 120 &mu;L. The protection primers are RNA versions of the universal primers. Order transcription templates of the universal primers with a T7 tag appended to the end, then transcribe RNA from the templates using the MEGAshortscript&trade; Kit .</li>
<li>Remove the supernatant from the bead aliquots and resuspend beads in 120 &mu;L of the master mix. Rock the reactions at 4&deg;C for 1-2 hours.</li>
<li>After the incubation, for each reaction, remove a small aliquot (including the beads) for a "total" RNA sample</li>
<li>Place the remainder of the reactions on the magnetic separation stand and remove the supernatant. Although it is not necessary to analyze this flow-through sample downstream, it may be useful to save.</li>
<li>Wash the beads 1-2 times with 50 &mu;L of cold 1x PBS. The RNA bound to the antibody on the beads is the "IP" sample.</li>
<li>Resuspend the "total" and the "IP" beads in 50 &mu;L of 1% SDS in 0.1x TE buffer and heat the samples at 65&deg;C for 15 minutes to elute the RNA.</li>
<li>Remove the supernatant, which now contains the RNA that was previously bound to the beads and clean by phenol/chloroform extraction followed by an ethanol precipitation. Dry the pellet and resuspend it in 50 &mu;L 0.1x TE buffer.</li>
</ol>
Reverse transcription and amplification of co-IP samples in preparation for aminoallyl labeling
<ol>
<li>Next, the RNA samples must be reverse transcribed and PCR amplified. For the reverse transcription reaction, use the reverse universal primers.</li>
<li>For each sample, begin by mixing 1 &mu;L of the reverse universal primer, with 1 &mu;L of the template RNA (from the immunoprecipitation reaction) and 10 &mu;L of dH2O. Heat the reactions at 70&deg;C for 5 minutes then chill on ice or keep at 4&deg;C.</li>
<li>While the samples cool, make the following master mix. For each reaction combine 4 &mu;L of 10 mM dNTPs, 2 &mu;L of 10x Arrayscript buffer, 1 &mu;L RNase inhibitor, and 1 &mu;L reverse transcription enzyme. Mix thoroughly and add 8 &mu;L of the master mix to each reverse transcription reaction.</li>
<li>Use a PCR machine to incubate the samples at 42&deg;C for 2 hours, followed by 95&deg;C for 5 minutes, then 4&deg;C until the samples are needed for amplification.</li>
<li>After reverse transcribing the RNA samples, PCR amplify them using the universal primers, with one of the primers containing a T7 tag. The number of cycles needed to amplify the pool will depend on the efficiency of the co-IP so it may be necessary to try multiple cycles. Each reaction requires 5 &mu;L of 10x Taq reaction buffer, 0.2 &mu;L of each primer (100 mM), 1 &mu;L of the cDNA template (from the reverse transcription reaction), 1 &mu;L of 10 mM dNTPs, 42.2 &mu;L of dH2O, and 0.4 &mu;L of Taq enzyme. Begin the PCR by incubating the samples at 94&deg;C for 3 minutes. Next, for each cycle, denature at 94&deg;C for 45 seconds, anneal at 55&deg;C for 30 seconds, and elongate at 72&deg;C for one minute, with a final elongation at 72&deg;C for 10 minutes.</li>
<li>Next, transcribe the samples again using the MEGAshortscript&trade; Kit, however, this time add 2 &mu;L of aminoallyl UTP, instead of the T7 UTP. The aminoallyl UTP will be used to label the RNA with the Cy dyes.</li>
<li>Phenol/chloroform and ethanol precipitate the samples. Do NOT use Glycoblue or ammonium acetate, both of which may interfere with hybridization of the samples to the array. Dry the pellet and resuspend in 4.5 &mu;L of 0.1 M sodium carbonate.</li>
</ol>
Amino-allyl labeling of RNA with Cy Dyes
<ol>
<li>To begin the labeling protocol dissolve each of the fluorescent dyes, cy3 and cy5, in 45 &mu;L of DMSO. To avoid light, wrap the tubes in aluminum foil and store at -20&deg;C in the dark</li>
<li>Add 2.5 &mu;L of nuclease-free water to RNA samples and mix thoroughly</li>
<li>Next, add 3 &mu;L of Cy3 to the "total" sample, and 3 &mu;L of Cy5 to the "IP" sample. Incubate the samples in the dark for 1 hour at room temperature; The Cy-3 sample will turn red or pink while the Cy-5 will turn blue.</li>
<li>To terminate the reaction, add 6 uL of 4 M hydroxylamine-HCl to each sample, mix thoroughly and spin briefly. Incubate for 15 minutes at room temperature.</li>
<li>Purify the labeled samples by phenol/chloroform extraction followed by ethanol precipitation and resuspend pellet in 50 &mu;L of 0.1x TE</li>
</ol>
Hybridize samples to the microarray using the Agilent Array Hybridization Kit
<ol>
<li>Begin the hybridization by gently mixing 50 &mu;L of 10x blocking buffer, 140 &mu;L of nuclease-free H2O, 25 &mu;L of the Cy3-labeled "total" pool, 25 &mu;L of the Cy5-labeled "IP" pool, 10 &mu;L of the 25x fragmentation buffer and 250 &mu;L of the 2x hybridization buffer, taking care to avoid introducing bubbles. Microfuge the sample briefly to collect the solution in the bottom of the tube and also to reduce bubbles.</li>
<li>Place the o-ring cover slide in the hybridization chamber and carefully pipette the solution onto the slide.</li>
<li>Place the detection array on top of the cover slide to "sandwich" the solution. Remember to orient the array slide so that the Agilent label on the array faces the Agilent label on the cover slide.</li>
<li>Clamp the slide/array sandwich in the hybridization chamber and rotate at 50&deg;C for 3 hours.</li>
<li>While the array is rotating, make the following 50 mL solutions in 50-mL conical tubes: one conical filled with 50 mL of dH2O, two 1x SCC buffer solutions, each in a separate tube, and one 2x SCC buffer.</li>
<li>After the three-hour incubation, carefully unclamp the array/slide sandwich and place it in the 50 mL conical containing the 2x SCC buffer. Using a pair of tweezers, gently separate the array from the cover slide and pull out the cover slide. Be careful not to touch the array as the slide is pulled out. Close the conical and gently invert for 60 seconds.</li>
<li>Using the tweezers, transfer the array to the 1x SCC solution, cap the tube, and gently invert for 1 minute then transfer the array to the second 1x SCC solution and again gently invert for another minute.</li>
<li>Lastly, transfer the array to the tube of dH2O and gently invert for 30 seconds.</li>
<li>Dry the array by placing the edges of the slide on a Kimwipe. Continue to rotate the array on the wipe making sure to not disrupt the side containing the oligos.</li>
<li>Place the dry array in an empty 50-mL conical tube and scan with a Microarray scanner</li>
</ol>
Interpreting the Microarray
<ol>
<li>The microarray scanner will create a TIF image of the array. Run this image through the Agilent Feature Extraction software, using the grid file (supplied by Agilent) which is unique to your specific pool.</li>
<li>The Agilent software will interpret the signal at each feature to create a file, which is parsed with a Perl script. The signal at each feature is normalize to the total intensity at that channel. For example at a particular feature in the green channel, the intensity of the green channel for the individual feature is divided by the total intensity of the green channel over all features. The final output is the log ratio of the red channel over the green channel at each feature on the array.</li>
<li>The oligo pool data from the microarray can be re-mapped back to the genome to get a picture of where, and how strongly, the RNP of interest bound.</li>
</ol>
Figure 1. Experimental schematic
<img src="/files/ftp_upload/1622/1622fig1a.jpg" alt="Figure 1a" /> A. Tiling scheme. After selecting and downloading pre-mRNA sequences of interest, the selecting area is tiled through in 30 nucleotide windows that shift in 10 nucleotide increments. Universal primer binding sites flank each side of the window.
<img src="/files/ftp_upload/1622/1622fig1b.jpg" alt="Figure 1b" /> B. Synthesis Array. Tiled oligo orders should be submitted to Agilent, where the sequences will be printed on a custom oligonucleotide microarray. 
<img src="/files/ftp_upload/1622/1622fig1c.jpg" alt="Figure 1c" /> C. Co-immunoprecipitation. After boiling the oligos off the array, incubate in HeLa nuclear extract, then add the extract to magnetic beads that are prepared with an antibody against the RNP of interest. Label the starting oligo pool with Cy3 and the bound IP pool with Cy5 and apply to the detection array and scan.
Figure 2. Annotating the genome with and analysis of enrichment data.
<img src="/files/ftp_upload/1622/1622fig2.jpg" alt="Figure 2" /> A. Convert the average score at each genomic coordinate from the array data to base-ten log and map that score to the given coordinates. An illustration of this averaging step is given for 3 overlapping 30-nt oligos with scores of 2, 4, and 0.5, where the average enrichment score for each 10-nt window is graphed above. The selected genomic regions can be visualized using a Custom Track in the USCS Genome Browser. The example browser window given is in the clta gene, where gene features (exons/introns/alternative splicing etc.) are given along the bottom and log average enrichment scores from the PTB-bound oligos are represented by dark red vertical bars. PTB strongly binds just upstream of exon 2 in the polypyrimidine tract.

When doing this procedure it&#8217;s important to remember that creation of the pool is flexible and open to modification at either the RNA or protein level. At the RNA level orthologous regions, disease mutations, polymorphisms, or random mutations can be introduced into the oligonucleotide sequence. At the protein level the RNA binding factor can be modified by phosphorylation or other post translational modifications. Additionally, the binding environment can be manipulated to either increase or decrease the levels ...

a rapid high-throughput method for mapping ribonucleoproteins (rnps) on human pre-mrna

Sequencing RNAs that co-immunoprecipitate (co-IP) with RNA binding proteins has increased our understanding of splicing by demonstrating that binding location often influences function of a splicing factor.  However, as with any sampling strategy the chance of identifying an RNA bound to a splicing factor is proportional to its cellular abundance.  We have developed a novel in vitro approach for surveying binding specificity on otherwise transient pre-mRNA. This approach utilizes a specifically designed oligonucleotide pool that tiles across introns, exons, splice junctions, or other pre-mRNA.  The pool is subjected to some kind of molecular selection. Here, we demonstrate the method by separating the oligonucleotide into a bound and unbound fraction and utilize a two color array strategy to record the enrichment of each oligonucleotide in the bound fraction. The array data generates high-resolution maps with the ability to identify sequence-specific and structural determinates of ribonucleoprotein (RNP) binding on pre-mRNA.  A unique advantage to this method is its ability to avoid the sampling bias towards mRNA associated with current IP and SELEX techniques, as the pool is specifically designed and synthesized from pre-mRNA sequence. The flexibility of the oligonucleotide pool is another advantage since the experimenter chooses which regions to study and tile across, tailoring the pool to their individual needs.  Using this technique, one can assay the effects of polymorphisms or mutations on binding on a large scale or clone the library into a functional splicing reporter and identify oligonucleotides that are enriched in the included fraction.  This novel in vitro high-resolution mapping scheme provides a unique way to study RNP interactions with transient pre-mRNA species, whose low abundance makes them difficult to study with current in vivo techniques.  



Watch this Scientific Journal Video about A Rapid High-throughput Method for Mapping Ribonucleoproteins RNPs on Human pre-mRNA at JoVE.com

A Rapid High-throughput Method for Mapping Ribonucleoproteins RNPs on Human pre-mRNA

Due to the transient nature of pre-mRNA, it can be difficult to isolate and study in vivo. Here, we present a novel in vitro approach to investigate RNA-protein interactions using a synthetic oligo pool that tiles across selected regions of pre-mRNA.

A Rapid High-throughput Method for Mapping Ribonucleoproteins (RNPs) on Human pre-mRNA

Sequencing RNAs that co-immunoprecipitate (co-IP) with RNA binding proteins has increased our understanding of splicing by demonstrating that binding ...

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11.7K Views.  Brown University. نظرا لطبيعة عابرة للمرنا المسبق ، يمكن أن يكون من الصعب عزل ودراسة في الجسم الحي. هنا ، فإننا نقدم رواية في المختبر نهج لتحقيق RNA البروتين التفاعلات باستخدام مجموعة جزئية الاصطناعية التي البلاط عبر مناطق مختارة من مرنا المسبق.

Video: A Rapid High-throughput Method for Mapping Ribonucleoproteins RNPs on Human pre-mRNA

A high-throughput method to globally study the organelle morphology in S. cerevisiae

High-throughput methods to examine protein localization or organelle morphology is an effective tool for studying protein interactions and can help achieve an comprehensive understanding of molecular pathways. In Saccharomyces cerevisiae, with the development of the non-essential gene deletion array, we can globally study the morphology of different organelles like the endoplasmic reticulum (ER) and the mitochondria using GFP (or variant)-markers in different gene backgrounds. However, incorporating GFP markers in each single mutant individually is a labor-intensive process. Here, we describe a procedure that is routinely used in our laboratory. By using a robotic system to handle high-density yeast arrays and drug selection techniques, we can significantly shorten the time required and improve reproducibility. In brief, we cross a GFP-tagged mitochondrial marker (Apc1-GFP) to a high-density array of 4,672 nonessential gene deletion mutants by robotic replica pinning.  Through diploid selection, sporulation, germination and dual marker selection, we recover both alleles. As a result, each haploid single mutant contains Apc1-GFP incorporated at its genomic locus. Now, we can study the morphology of mitochondria in all non-essential mutant background. Using this high-throughput approach, we can conveniently study and delineate the pathways and genes involved in the inheritance and the formation of organelles in a genome-wide setting.

GFP-fusion proteins are widely used to visualize organelles by confocal microscopy. However, screening for mutations that affect the morphology of organelles generally requires individual mutagenesis and is time consuming. Here, we demonstrate a method to simultaneously incorporate organelle-GFP markers in almost 5,000 non-essential genes in yeast.

High-throughput methods to examine protein localization or organelle morphology is an effective tool for studying protein interactions and can help ...

A High-Throughput Method For Zebrafish Sperm Cryopreservation and In Vitro Fertilization

This is a method for zebrafish sperm cryopreservation that is an adaptation of the Harvey method (Harvey et al., 1982).  We have introduced two changes to the original protocol that both streamline the procedure and increase sample uniformity.  First, we normalize all sperm volumes using freezing media that does not contain the cryoprotectant. Second, cryopreserved sperm are stored in cryovials instead of capillary tubes.  The rates of sperm freezing and thawing (&delta;&deg;C/time) are probably the two most critical variables to control in this procedure.  For this reason, do not substitute different tubes for those specified.  Working in teams of 2 it is possible to freeze the sperm of 100 males per team in ~2 hrs.  Sperm cryopreserved using this protocol has an average of 25% fertility (measured as the number of viable embryos generated in an in vitro fertilization divided by the total number of eggs fertilized) and this percent fertility is stable over many years.

A High-Throughput Method For Zebrafish Sperm Cryopreservation and In Vitro Fertilization

This is a high-throughput sperm cryopreservation protocol for zebrafish. Sperm cryopreserved using this protocol has an average of 25% fertility in subsequent vitro fertilization and is stable over many years.

This is a method for zebrafish sperm cryopreservation that is an adaptation of the Harvey method (Harvey et al., 1982).  We have introduced two changes to ...

Ice-Cap: A Method for Growing Arabidopsis and Tomato Plants in 96-well Plates for High-Throughput Genotyping

It is becoming common for plant scientists to develop projects that require the genotyping of large numbers of plants. The first step in any genotyping project is to collect a tissue sample from each individual plant. The traditional approach to this task is to sample plants one-at-a-time. If one wishes to genotype hundreds or thousands of individuals, however, using this strategy results in a significant bottleneck in the genotyping pipeline. The Ice-Cap method that we describe here provides a high-throughput solution to this challenge by allowing one scientist to collect tissue from several thousand seedlings in a single day 1,2. This level of throughput is made possible by the fact that tissue is harvested from plants 96-at-a-time, rather than one-at-a-time.
The Ice-Cap method provides an integrated platform for performing seedling growth, tissue harvest, and DNA extraction. The basis for Ice-Cap is the growth of seedlings in a stacked pair of 96-well plates. The wells of the upper plate contain plugs of agar growth media on which individual seedlings germinate. The roots grow down through the agar media, exit the upper plate through a hole, and pass into a lower plate containing water. To harvest tissue for DNA extraction, the water in the lower plate containing root tissue is rapidly frozen while the seedlings in the upper plate remain at room temperature. The upper plate is then peeled away from the lower plate, yielding one plate with 96 root tissue samples frozen in ice and one plate with 96 viable seedlings. The technique is named "Ice-Cap" because it uses ice to capture the root tissue. The 96-well plate containing the seedlings can then wrapped in foil and transferred to low temperature. This process suspends further growth of the seedlings, but does not affect their viability. Once genotype analysis has been completed, seedlings with the desired genotype can be transferred from the 96-well plate to soil for further propagation. We have demonstrated the utility of the Ice-Cap method using Arabidopsis thaliana, tomato, and rice seedlings. We expect that the method should also be applicable to other species of plants with seeds small enough to fit into the wells of 96-well plates.

Ice-Cap: A Method for Growing Arabidopsis and Tomato Plants in 96-well Plates for High-Throughput Genotyping

The Ice-Cap method allows one to grow plants in 96-well plates and non-destructively harvest root tissue from each seedling. DNA extracted from this root tissue can be used for genotyping reactions. We have found that Ice-Cap works well for Arabidopsis thaliana, tomato, and rice seedlings.

It is becoming common for plant scientists to develop projects that require the genotyping of large numbers of plants. The first step in any genotyping ...

High-throughput Crystallization of Membrane Proteins Using the Lipidic Bicelle Method

Membrane proteins (MPs) play a critical role in many physiological processes such as pumping specific molecules across the otherwise impermeable membrane bilayer that surrounds all cells and organelles. Alterations in the function of MPs result in many human diseases and disorders; thus, an intricate understanding of their structures remains a critical objective for biological research. However, structure determination of MPs remains a significant challenge often stemming from their hydrophobicity. 
MPs have substantial hydrophobic regions embedded within the bilayer. Detergents are frequently used to solubilize these proteins from the bilayer generating a protein-detergent micelle that can then be manipulated in a similar manner as soluble proteins. Traditionally, crystallization trials proceed using a protein-detergent mixture, but they often resist crystallization or produce crystals of poor quality. These problems arise due to the detergent&prime;s inability to adequately mimic the bilayer resulting in poor stability and heterogeneity. In addition, the detergent shields the hydrophobic surface of the MP reducing the surface area available for crystal contacts. To circumvent these drawbacks MPs can be crystallized in lipidic media, which more closely simulates their endogenous environment, and has recently become a de novo technique for MP crystallization.
Lipidic cubic phase (LCP) is a three-dimensional lipid bilayer penetrated by an interconnected system of aqueous channels1. Although monoolein is the lipid of choice, related lipids such as monopalmitolein and monovaccenin have also been used to make LCP2. MPs are incorporated into the LCP where they diffuse in three dimensions and feed crystal nuclei. A great advantage of the LCP is that the protein remains in a more native environment, but the method has a number of technical disadvantages including high viscosity (requiring specialized apparatuses) and difficulties in crystal visualization and manipulation3,4. Because of these technical difficulties, we utilized another lipidic medium for crystallization-bicelles5,6 (Figure 1). Bicelles are lipid/amphiphile mixtures formed by blending a phosphatidylcholine lipid (DMPC) with an amphiphile (CHAPSO) or a short-chain lipid (DHPC). Within each bicelle disc, the lipid molecules generate a bilayer while the amphiphile molecules line the apolar edges providing beneficial properties of both bilayers and detergents. Importantly, below their transition temperature, protein-bicelle mixtures have a reduced viscosity and are manipulated in a similar manner as detergent-solubilized MPs, making bicelles compatible with crystallization robots. 
Bicelles have been successfully used to crystallize several membrane proteins5,7-11 (Table 1). This growing collection of proteins demonstrates the versatility of bicelles for crystallizing both alpha helical and beta sheet MPs from prokaryotic and eukaryotic sources. Because of these successes and the simplicity of high-throughput implementation, bicelles should be part of every membrane protein crystallographer&prime;s arsenal. In this video, we describe the bicelle methodology and provide a step-by-step protocol for setting up high-throughput crystallization trials of purified MPs using standard robotics.

Bicelles are lipid/amphiphile mixtures that maintain membrane proteins (MPs) within a lipid bilayer but have unique phase behavior that facilitates high-throughput screening by crystallization robots. This technique has successfully produced a number of high-resolution structures from both prokaryotic and eukaryotic sources. This video describes protocols for generating the lipidic bicelle mixture, incorporating MPs into the bicelle mixture, setting up crystallizations trials (manually as well as robotically) and harvesting crystals from the medium.

Membrane proteins (MPs) play a critical role in many physiological processes such as pumping specific molecules across the otherwise impermeable membrane ...

High-throughput Physical Mapping of Chromosomes using Automated in situ Hybridization

Projects to obtain whole-genome sequences for 10,000 vertebrate species1 and for 5,000 insect and related arthropod species2 are expected to take place over the next 5 years. For example, the sequencing of the genomes for 15 malaria mosquitospecies is currently being done using an Illumina platform3,4. This Anopheles species cluster includes both vectors and non-vectors of malaria. When the genome assemblies become available, researchers will have the unique opportunity to perform comparative analysis for inferring evolutionary changes relevant to vector ability. However, it has proven difficult to use next-generation sequencing reads to generate high-quality de novo genome assemblies5. Moreover, the existing genome assemblies for Anopheles gambiae, although obtained using the Sanger method, are gapped or fragmented4,6.
Success of comparative genomic analyses will be limited if researchers deal with numerous sequencing contigs, rather than with chromosome-based genome assemblies. Fragmented, unmapped sequences create problems for genomic analyses because: (i) unidentified gaps cause incorrect or incomplete annotation of genomic sequences; (ii) unmapped sequences lead to confusion between paralogous genes and genes from different haplotypes; and (iii) the lack of chromosome assignment and orientation of the sequencing contigs does not allow for reconstructing rearrangement phylogeny and studying chromosome evolution. Developing high-resolution physical maps for species with newly sequenced genomes is a timely and cost-effective investment that will facilitate genome annotation, evolutionary analysis, and re-sequencing of individual genomes from natural populations7,8.
Here, we present innovative approaches to chromosome preparation, fluorescent in situ hybridization (FISH), and imaging that facilitate rapid development of physical maps. Using An. gambiae as an example, we demonstrate that the development of physical chromosome maps can potentially improve genome assemblies and, thus, the quality of genomic analyses. First, we use a high-pressure method to prepare polytene chromosome spreads. This method, originally developed for Drosophila9, allows the user to visualize more details on chromosomes than the regular squashing technique10. Second, a fully automated, front-end system for FISH is used for high-throughput physical genome mapping. The automated slide staining system runs multiple assays simultaneously and dramatically reduces hands-on time11. Third, an automatic fluorescent imaging system, which includes a motorized slide stage, automatically scans and photographs labeled chromosomes after FISH12. This system is especially useful for identifying and visualizing multiple chromosomal plates on the same slide. In addition, the scanning process captures a more uniform FISH result. Overall, the automated high-throughput physical mapping protocol is more efficient than a standard manual protocol.

High-throughput Physical Mapping of Chromosomes using Automated in situ Hybridization

Genome assemblies based on massively parallel DNA sequencing technologies are usually highly fragmented. The development of physical chromosome maps can potentially improve genome assemblies. Here, we demonstrate innovative approaches to chromosome preparation, fluorescent in situ hybridization, and imaging that significantly increase throughput of the physical map development.

Projects to obtain whole-genome sequences for 10,000 vertebrate species1 and for 5,000 insect and related arthropod species2 are expected to take place ...

Biochemical and High Throughput Microscopic Assessment of Fat Mass in Caenorhabditis Elegans

The nematode C. elegans has emerged as an important model for the study of conserved genetic pathways regulating fat metabolism as it relates to human obesity and its associated pathologies. Several previous methodologies developed for the visualization of C. elegans triglyceride-rich fat stores have proven to be erroneous, highlighting cellular compartments other than lipid droplets. Other methods require specialized equipment, are time-consuming, or yield inconsistent results. We introduce a rapid, reproducible, fixative-based Nile red staining method for the accurate and rapid detection of neutral lipid droplets in C. elegans. A short fixation step in 40% isopropanol makes animals completely permeable to Nile red, which is then used to stain animals. Spectral properties of this lipophilic dye allow it to strongly and selectively fluoresce in the yellow-green spectrum only when in a lipid-rich environment, but not in more polar environments. Thus, lipid droplets can be visualized on a fluorescent microscope equipped with simple GFP imaging capability after only a brief Nile red staining step in isopropanol. The speed, affordability, and reproducibility of this protocol make it ideally suited for high throughput screens. We also demonstrate a paired method for the biochemical determination of triglycerides and phospholipids using gas chromatography mass-spectrometry. This more rigorous protocol should be used as confirmation of results obtained from the Nile red microscopic lipid determination. We anticipate that these techniques will become new standards in the field of C. elegans metabolic research.

Biochemical and High Throughput Microscopic Assessment of Fat Mass in Caenorhabditis Elegans

We present robust biochemical and microscopic methods for studying Caenorhabditis elegans lipid stores. A rapid, simple, fixing-staining procedure for fluorescent lipid droplet imaging leverages the spectral properties of the lipophilic dye Nile red. We then present biochemical measurement of triglycerides and phospholipids using solid phase extraction and gas chromatography-mass spectrometry.

The nematode C. elegans has emerged as an important  model for the study of conserved genetic pathways regulating fat metabolism as  it relates to human ...

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.

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.

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

A High-throughput Method for Measurement of Glomerular Filtration Rate in Conscious Mice

The measurement of glomerular filtration rate (GFR) is the gold standard in kidney function assessment. Currently, investigators determine GFR by measuring the level of the endogenous biomarker creatinine or exogenously applied radioactive labeled inulin (3H or 14C). Creatinine has the substantial drawback that proximal tubular secretion accounts for ~50% of total renal creatinine excretion and therefore creatinine is not a reliable GFR marker. Depending on the experiment performed, inulin clearance can be determined by an intravenous single bolus injection or continuous infusion (intravenous or osmotic minipump). Both approaches require the collection of plasma or plasma and urine, respectively. Other drawbacks of radioactive labeled inulin include usage of isotopes, time consuming surgical preparation of the animals, and the requirement of a terminal experiment. Here we describe a method which uses a single bolus injection of fluorescein isothiocyanate-(FITC) labeled inulin and the measurement of its fluorescence in 1-2 &mu;l of diluted plasma. By applying a two-compartment model, with 8 blood collections per mouse, it is possible to measure GFR in up to 24 mice per day using a special work-flow protocol. This method only requires brief isoflurane anesthesia with all the blood samples being collected in a non-restrained and awake mouse. Another advantage is that it is possible to follow mice over a period of several months and treatments (i.e. doing paired experiments with dietary changes or drug applications). We hope that this technique of measuring GFR is useful to other investigators studying mouse kidney function and will replace less accurate methods of estimating kidney function, such as plasma creatinine and blood urea nitrogen.

Measurement of glomerular filtration rate (GFR) is the gold standard for kidney function assessment. Here we describe a high-throughput method which allows the determination of GFR in conscious mice by using a single bolus injection, determination of fluorescein isothiocyanate (FITC)-inulin in plasma and calculation of GFR by a two-phase exponential decay model.

The measurement of  glomerular filtration rate (GFR) is the gold standard in kidney function  assessment. Currently, investigators determine GFR by ...

High Throughput Microinjections of Sea Urchin Zygotes

Microinjection into cells and embryos is a common technique that is used to study a wide range of biological processes. In this method a small amount of treatment solution is loaded into a microinjection needle that is used to physically inject individual immobilized cells or embryos. Despite the need for initial training to perform this procedure for high-throughput delivery, microinjection offers maximum efficiency and reproducible delivery of a wide variety of treatment solutions (including complex mixtures of samples) into cells, eggs or embryos. Applications to microinjections include delivery of DNA constructs, mRNAs, recombinant proteins, gain of function, and loss of function reagents. Fluorescent or colorimetric dye is added to the injected solution to enable instant visualization of efficient delivery as well as a tool for reliable normalization of the amount of the delivered solution. The described method enables microinjection of 100-400 sea urchin zygotes within 10-15 min.

Microinjection is a common technique used to deliver DNA constructs, mRNAs, morpholino antisense oligonucleotides or other treatments into eggs, embryos, and cells of various species.

Microinjection into cells and embryos is a common technique that is used to study a wide range of biological processes. In this method a small amount of ...

A Method for Selecting Structure-switching Aptamers Applied to a Colorimetric Gold Nanoparticle Assay

Small molecules provide rich targets for biosensing applications due to their physiological implications as biomarkers of various aspects of human health and performance. Nucleic acid aptamers have been increasingly applied as recognition elements on biosensor platforms, but selecting aptamers toward small molecule targets requires special design considerations. This work describes modification and critical steps of a method designed to select structure-switching aptamers to small molecule targets. Binding sequences from a DNA library hybridized to complementary DNA capture probes on magnetic beads are separated from nonbinders via a target-induced change in conformation. This method is advantageous because sequences binding the support matrix (beads) will not be further amplified, and it does not require immobilization of the target molecule. However, the melting temperature of the capture probe and library is kept at or slightly above RT, such that sequences that dehybridize based on thermodynamics will also be present in the supernatant solution. This effectively limits the partitioning efficiency (ability to separate target binding sequences from nonbinders), and therefore many selection rounds will be required to remove background sequences. The reported method differs from previous structure-switching aptamer selections due to implementation of negative selection steps, simplified enrichment monitoring, and extension of the length of the capture probe following selection enrichment to provide enhanced stringency. The selected structure-switching aptamers are advantageous in a gold nanoparticle assay platform that reports the presence of a target molecule by the conformational change of the aptamer. The gold nanoparticle assay was applied because it provides a simple, rapid colorimetric readout that is beneficial in a clinical or deployed environment. Design and optimization considerations are presented for the assay as proof-of-principle work in buffer to provide a foundation for further extension of the work toward small molecule biosensing in physiological fluids.

A protocol is provided to select structure-switching aptamers for small molecule targets based on a tunable stringency magnetic bead selection method. Aptamers selected with structure-switching properties are beneficial for assays that require a conformational change to signal the presence of a target, such as the described gold nanoparticle assay.

Small molecules provide rich targets for biosensing applications due to their physiological implications as biomarkers of various aspects of human health ...