The authors would like to thank the Rare Disease Models and Mechanism (RDMM) Network (funded by the Canadian Institutes of Health Research (CIHR) and Genome Canada). CK is supported by a UROP scholarship award (University of Ottawa). DLJ is supported by a Vanier Canada Graduate Scholarship. IAP is supported by a Canadian Institutes of Health Research (CIHR) postdoctoral fellowship award. The authors thank the Genetics Society of America for granting permission to publish this protocol and to use an adaptation of the Supplemental Figure 5 from Pena, et al.

Procedures involving animal subjects have been approved and are in accordance with animal care guidelines provided by the Canadian Council on Animal Care and the University of Ottawa animal care committees.
1. Preparation
<ol>
	<li>Prepare the dissection surface by taping a 9 cm Petri dish lid with autoclave tape across its interior surface. Position it under a stereo-microscope.</li>
	<li>Place 3 - 5 days post-fertilization (dpf) zebrafish larvae in a Petri dish and anesthetize in &#732;1.5 mM Tricaine in 1x E3 embryo media (5 mM NaCl, 0.17 mM KCl, 0.33 mM CaCl2, 0.33 mM MgSO4, pH 7.2).</li>
	<li>Cut off the end of a P1000 pipette tip with a pair of scissors or a razor blade to a diameter of 2 mm to accommodate a 3 dpf larva with minimal stress.</li>
</ol>
2. Fin Clipping of Zebrafish Larvae
<ol>
	<li>Pick up an anesthetized larva using the modified P1000 micropipette and place the zebrafish larva onto the positioned autoclave tape on the Petri dish lid.</li>
	<li>Remove excess Tricaine solution surrounding the larva using a micropipette. The area should be as dry as possible to successfully section and pick up the fin but still wet to ensure survival.</li>
	<li>Using a micro scalpel, section the caudal fin under a stereo-microscope as indicated in Figure 1, as described in Wilkinson&#160;et al16. During the incision, apply a steady downward pressure within the pigment gap site of the caudal fin, distal to the limit of the blood circulation (Figure 1A). Apply gentle pressure to avoid damaging the notochord.</li>
	<li>Visualize the sectioned fin under the microscope and position the piece on top of the tip of the microscalpel blade (Figure 2A). Place the microscalpel containing the fin onto the surface of a small piece of filter paper. 
	NOTE: Due to the presence of melanocytes in the transected tissue, this step allows visualization of the fin as a small black spot (Figure 2B).</li>
	<li>Using scissors and tweezers (Figure 2C), transfer the filter paper containing the fin to a 96-well PCR plate, containing 25 &#956;L of 50 mM NaOH solution per well (Figure 2D).</li>
	<li>Prepare a flat-bottom 96-well plate, numbering the well according to the label of the PCR plate. Using the modified P1000 pipette filled with 200 &#181;L of fresh 1x E3 embryo media, carefully collect the zebrafish larva using gentle pressure. Dispense the larva in the 96-well tissue culture plate, in the same well number as the PCR plate (Figure 2E).</li>
	<li>Make sure that the filter paper is well submerged in the solution (Figure 2F). Clean the blade of the microscapel and tweezers between each cut to prevent cross-contamination by dipping it into 70% EtOH solution. Wipe the blade with clean paper tissue/wipe.</li>
	<li>Repeat steps 2.1 - 2.7 until all embryos have been clipped.</li>
	<li>Store the larvae in the 96-well plate at 28 &#176;C until the genotypes have been identified. 
	NOTE: The embryo media does not need to be changed, since the protocol can be easily completed in less than 2 days. If more time is needed, a media change is recommended.</li>
</ol>
3. Genomic DNA Extraction
<ol>
	<li>Seal the 96-well PCR plate and centrifuge the samples at 1,000 x g for 1 min to make sure all the filter papers are submerged within the NaOH solution.</li>
	<li>For tissue lysis, heat the samples in a thermocycler at 95 &#176;C for 5 min followed by cooling to 4 &#176;C for 10 min.</li>
	<li>Add 6 &#956;L of 500 mM Tris-HCl, pH 8.0 to each sample. Vortex.</li>
	<li>Briefly centrifuge the plate at 1,500 x g, for 5 min at room temperature.</li>
	<li>Use 1.5 &#956;L of the DNA supernatant per PCR reaction.</li>
	<li>Prepare a PCR to genotype the larvae, as shown in Table 1-2. 
	NOTE: This protocol was tested for two applications: multiplex PCR reactions revealing the genotype using agarose gels9, and heteroduplex melting assay (HMA) revealing the genotypes using polyacrylamide gel electrophoresis (PAGE)17.</li>
</ol>
4. Alternative Genomic Extraction Using a Chelating Resin
<ol>
	<li>Complete step 2.5 by adding filter paper with the clipped fin to a 96-well PCR plate, containing 30 &#956;L of 5% chelating resin (styrene-divinylbenzene copolymer containing paired iminodiacetate ions). 
	NOTE: This type of resin is commonly used for tissue lysis and preparation of PCR-ready DNA in a fast and efficient manner18.</li>
	<li>Follow steps 2.6 - 2.8 as indicated.</li>
	<li>Seal the 96-well PCR plate and briefly centrifuge the samples to make sure all filter paper is submerged within the chelating resin.</li>
	<li>For tissue lysis, heat the samples in a thermocycler at 95 &#176;C for 15 min followed by cooling to 4 &#176;C for 10 min. 
	NOTE: This step allows lysis and subsequent binding of the polar resin beads to cellular components while DNA and RNA remain in solution.</li>
	<li>Briefly centrifuge the samples to pellet the resin beads and obtain the DNA in the suspension.</li>
	<li>Follow steps 3.5 - 3.6 to complete genotyping procedure.</li>
</ol>
5. Application 1: Multiplex PCR Followed by Agarose Gel Analysis
<ol>
	<li>Following Table 1, set up a multiplex PCR reaction to enable the discrimination between homozygous mutants, heterozygotes, and WT larvae. Use a 96-well thermal cycler to perform the PCR reactions using the cycling conditions described in Table 3. 
	NOTE: The PCR reactions can also be performed in any other conventional PCR thermal cycler. This example describes the PCR strategy used by Pena, et al.9 to identify aldh7a1 WT and 5-bp insertion mutant alleles in the same multiplex reaction.</li>
	<li>Prepare a 1% agarose gel in sodium borate buffer, stained with 1x GelRed. 
	NOTE: 20x sodium borate buffer stock is made of 40 mL of 10 M NaOH, 1800 mL of ddH2O, pH adjusted to 8.5 with 76 g of boric acid and then completed to 2 L final volume using ddH2O.</li>
	<li>Run the gel in 1x sodium borate buffer for 15 min at 250 V. To facilitate the process, if possible, use a gel system that is compatible with multichannel pipets to enable higher throughput capabilities by reducing the time to load samples.</li>
	<li>Image the gel under ultraviolet (UV) light to allow discrimination of the different genotypes.</li>
</ol>
6. Application 2: Heteroduplex Melting Assay
<ol>
	<li>Prepare PAGE 12% gels using the 1.5 mm spacer plate and the 15-samples combs. Prepare two PAGE gels using 12.21 mL of ddH2O, 2.52 mL of 10x Tris/Borate/EDTA (TBE), 10 mL of 30% acrylamide, 250 &#956;L of 10% ammonium persulfate (APS) and 20 &#956;L of tetramethylethylenediamine (TEMED). Use 1x TBE buffer (0.089 M Tris-HCl, 0.089 M boric acid and 0.002 M ethylenediaminetetraacetic acid (EDTA)) as running buffer.</li>
	<li>Heat the PCR products at 94 &#176;C for 5 min.</li>
	<li>Cool the tubes on ice or in the thermocycler for 10 min at 4 &#176;C.</li>
	<li>Load 10 &#956;L per PCR sample and 8 &#956;L of molecular weight marker.</li>
	<li>Run the PAGE at 150 V in 1x TBE using a vertical electrophoresis system for 1 h or until the molecular weight marker almost reaches the foot line of the glass plate.</li>
	<li>Image the gel under UV light to allow discrimination of the different genotypes.</li>
</ol>

<ol>
	<li>Howe, K., 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=The+zebrafish+reference+genome+sequence+and+its+relationship+to+the+human+genome.">The zebrafish reference genome sequence and its relationship to the human genome.</a> Nature. 496 (7446), (2013).</li><li>Stewart, A. M., Kalueff, A. V. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Developing+better+and+more+valid+animal+models+of+brain+disorders.">Developing better and more valid animal models of brain disorders.</a> Behav Brain Res. 276, 28-31 (2015).</li><li>Grone, B. P., Baraban, S. C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Animal+models+in+epilepsy+research:+legacies+and+new+directions.">Animal models in epilepsy research: legacies and new directions.</a> Nat Neurosci. 18 (3), 339-343 (2015).</li><li>Rosen, J. N., Sweeney, M. F., Mably, J. D. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Microinjection+of+Zebrafish+Embryos+to+Analyze+Gene+Function.">Microinjection of Zebrafish Embryos to Analyze Gene Function.</a> J Vis Exp. (25), e1115 (2009).</li><li>Doyon, Y., 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=Heritable+targeted+gene+disruption+in+zebrafish+using+designed+zinc-finger+nucleases.">Heritable targeted gene disruption in zebrafish using designed zinc-finger nucleases.</a> Nat Biotechnol. 26 (6), 702-708 (2008).</li><li>Bedell, V. M., 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=In+vivo+Genome+Editing+Using+High+Efficieny+TALENs.">In vivo Genome Editing Using High Efficieny TALENs.</a> Nature. 491 (7422), 114-118 (2012).</li><li>Sander, J. D., 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=Targeted+gene+disruption+in+somatic+zebrafish+cells+using+engineered+TALENs.">Targeted gene disruption in somatic zebrafish cells using engineered TALENs.</a> Nat Biotechnol. 29 (8), 697-698 (2012).</li><li>Hwang, W. Y., 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=Efficient+genome+editing+in+zebrafish+using+a+CRISPR-Cas+system.">Efficient genome editing in zebrafish using a CRISPR-Cas system.</a> Nat Biotechnol. 31 (3), 227-229 (2013).</li><li>Pena, I. 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=Pyridoxine-Dependent+Epilepsy+in+Zebrafish+Caused+by+Aldh7a1+Deficiency.">Pyridoxine-Dependent Epilepsy in Zebrafish Caused by Aldh7a1 Deficiency.</a> Genetics. 207, 1501-1518 (2017).</li><li>Baraban, S. C., Dinday, M. T., Hortopan, G. A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Drug+screening+in+Scn1a+zebrafish+mutant+identifies+clemizole+as+a+potential+Dravet+syndrome+treatment.">Drug screening in Scn1a zebrafish mutant identifies clemizole as a potential Dravet syndrome treatment.</a> Nat Commun. 4, 2410 (2013).</li><li>Godoy, R., Noble, S., Yoon, K., Anisman, H., Ekker, M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Chemogenetic+ablation+of+dopaminergic+neurons+leads+to+transient+locomotor+impairments+in+zebrafish+larvae.">Chemogenetic ablation of dopaminergic neurons leads to transient locomotor impairments in zebrafish larvae.</a> J Neurochem. 135, 249-260 (2015).</li><li>Scheldeman, 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=Neurobiology+of+Disease+mTOR-related+neuropathology+in+mutant+tsc2+zebra+fish+Phenotypic,+transcriptomic+and+pharmacological+analysis.">Neurobiology of Disease mTOR-related neuropathology in mutant tsc2 zebra fish Phenotypic, transcriptomic and pharmacological analysis.</a> Neurobiol Dis. 108 (September), 225-237 (2017).</li><li>Zabinyakov, 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=Characterization+of+the+first+knock-out+aldh7a1+zebrafish+model+for+pyridoxine-dependent+epilepsy+using+CRISPR-Cas9+technology.">Characterization of the first knock-out aldh7a1 zebrafish model for pyridoxine-dependent epilepsy using CRISPR-Cas9 technology.</a> PLoS One. 12 (10), 1-14 (2017).</li><li>Grone, B. P., 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=Epilepsy,+behavioral+abnormalities,+and+physiological+comorbidities+in+syntaxin-binding+protein+1+(STXBP1)+mutant+zebrafish.">Epilepsy, behavioral abnormalities, and physiological comorbidities in syntaxin-binding protein 1 (STXBP1) mutant zebrafish.</a> PLoS One. 11 (3), (2016).</li><li>Kawakami, K., Hopkins, N. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Rapid+identification+of+transgenic+zebrafish.">Rapid identification of transgenic zebrafish.</a> Trends Genet. 12 (1), 9-10 (1996).</li><li>Wilkinson, R. N., Elworthy, S., Ingham, P. W., van Eeden, F. J. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+method+for+high-throughput+PCR-based+genotyping+of+larval+zebrafish+tail+biopsies.">A method for high-throughput PCR-based genotyping of larval zebrafish tail biopsies.</a> Biotechniques. 55 (6), 314-316 (2013).</li><li>Zhu, X., 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=An+efficient+genotyping+method+for+genome-modified+animals+and+human+cells+generated+with+CRISPR/Cas9+system.">An efficient genotyping method for genome-modified animals and human cells generated with CRISPR/Cas9 system.</a> Sci Rep. 4, 6420 (2014).</li><li>Walsh, P. S., Metzger, D. 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The authors declare that they have no competing financial interests.

Gene editing tools have been employed to precisely introduce mutations and transgenes in zebrafish over the past years5,6,7,8. When performing disease modeling in zebrafish, early larval or juvenile phenotypes are often observed9,12,13,14. The protocol presented here describes the extraction of PCR-ready DNA from small zebrafish larval tail biopsies, enabling genotyping of larvae as young as 3 dpf. Early identification of genotypes is crucial for several laboratory applications, especially when post hoc genotyping is not possible9. Knowing genotypes from an early age also facilitates survival studies and phenotypic analysis of mutants. This protocol is also particularly useful for the raising of larvae of specific genotypes to maturity. Larval genotyping is not yet widely used in zebrafish laboratories across the world and this protocol aims to facilitate the dissemination of this technique. While manipulating the transected fins, the use of this method provides confidence as to the presence of DNA within the sample; the fin biopsy can be readily visualized as a dark spot on the filter paper throughout the handling process. This represents an important improvement of the protocol published by Wilkinson et al.16.
Several critical steps within the protocol must be correctly followed to obtain quality results. First and foremost, the transection of the larval fin must occur within the pigment gap, distal to the limit of the blood circulation, to avoid bleeding. This ensures survival of the larvae during the fin biopsy. Moreover, a black dot on the filter paper, marking the placement of the fin, must be observed before placing the filter paper in the NaOH solution. This assures the presence of fin tissue (and therefore, DNA) in each sample well. The filter paper must also be immersed within the NaOH solution to guarantee cell lysis and successful DNA extraction.&#160;This protocol was not tested for younger ages than 3 dpf as only hatched larvae were used. To genotype younger embryos (unhatched&#160;&#60;3 dpf) chorion removal would be required. In case that insufficient PCR amplification occurs, standard PCR optimization strategies should be applied (e.g., varying primer concentration, annealing temperature, and/or DNA concentration). Although sufficient DNA is extracted to successfully perform several types of PCR reactions (as shown for aldh7a1 and plpbp genotyping examples), the concentration of DNA obtained following this fin clipping procedure is small (~5 ng/&#181;L per sample) and thus, increasing the concentration of DNA in the PCR reaction may allow for greater success of amplification.
The larval fin DNA extracted with this protocol can be used for several amplification reactions, allowing accurate identification of larval zebrafish genotypes and in a high-throughput manner. A heteroduplex melting assay in PAGE gels can be used to identify different mutations17. Given that ~30 &#956;L of DNA is obtained per sample, and ~1 - 1.5 &#956;L of DNA is used per PCR reaction, approximately 30 PCR reactions can be run per extracted sample. The PCR products are also suitable for Sanger sequencing applications; high quality of reads were obtained, allowing accurate confirmation of WT and homozygous mutant sequences for aldh7a1. Genomic-DNA extraction from larval zebrafish has many applications in research. The original goal for developing this protocol was to effectively genotype larvae that cannot reach adulthood due to specific mutations in the aldh7a19. In addition, this protocol enabled the segregation of genotypes and the observation of epilepsy-associated phenotypes specifically in aldh7a1-/- fish from the larval stage onward9. The pooling of larvae of each genotype for approaches such as RNA extraction or metabolite extraction can be accurately executed after genotyping using this larval fin clipping procedure9. In conclusion, after a short period of training, an experienced investigator can use this simple and time-efficient protocol, requiring only minimal and inexpensive reagents, to routinely perform hundreds of fin clips per day. The use of this protocol made possible the research described by Pena et al. and will hopefully serve a similar purpose for the broader Zebrafish community.

The zebrafish (Danio rerio) is a vertebrate organism widely used as a model for disease investigation as well as for preclinical testing of therapeutic hypotheses1,2,3. These animals have a short generation time, are easy to handle, display a high reproductive rate, low cost, and are easily amenable to genetic manipulations by microinjection of DNA in embryos4. Through the increasing development of novel transgenic and gene editing technologies such as Zinc-Finger nucleases (ZFN)5, TAL-like Effector Nucleases (TALENs)6,7, and the Clustered Regularly Interspaced Short Palindromic Repeats (CRISPR)/CRISPR-associated 9 (Cas9) system8, the zebrafish is poised to substantially improve the understanding of several pathologic conditions. These technologies have been used to create targeted knockouts in both somatic and germline cells in zebrafish, efficiently engendering genetically modified animals8. Recent examples in the literature include the successful development of genetic models of epilepsy and metabolic disorders in zebrafish3,9,10,11,12.
With the progress made in zebrafish genome editing, rapid and reliable genotyping has become an undeniable rate-limiting step. Identification of specific DNA mutations adjacent to the predicted genome-editing target site is an essential stage of the protocol. Traditionally, genotyping techniques by fin clipping are usually only performed in juvenile or adult zebrafish. However, the time spent raising zebrafish to adulthood (&#62;2 months) considerably delays advancements in research. In many cases, adulthood cannot be achieved due to the knockout of essential genes (e.g., in disease modeling) or gain-of-function mutations leading to detrimental phenotypic effects. In addition, several assays require pooling of larvae, such as in RNA or metabolite extraction, and thus involve prior identification of the genotypes, which can be challenging when only post hoc genotyping techniques are available. These aforementioned examples highlight the importance of larval genotyping techniques which are at the same time precise and enable full recovery and normal development of the larvae. Early stage zebrafish genotyping does not currently appear to be widely used; this is reflected by recent publications of disease models in which larval genotypes are identified via post-mortem genotyping rather than genotyping prior to the execution of the experiment12,13,14. In this paper, a larval fin clip strategy is presented aiming to improve previously established genotyping procedures.
In the past, strategies employing proteinase K digestion to genotype live zebrafish larvae have led to variable polymerase chain reaction (PCR) efficiency15. Although a more recently published microscopic tail biopsy technique provided more promising results16, we and other groups were not able to reproduce the high efficiency and DNA recovery reported by the authors. A major setback of the protocol described by Wilkinson&#160;et al.16 could be the transfer of the tail tissue to the PCR tube. Due to its microscopic size, it is hard to ensure that the fin was properly collected and dispensed by pipetting. To address this barrier, we have developed an improved protocol for genotyping large numbers of live zebrafish larvae with nearly 100% efficiency9. Using a microscalpel, the tip of the fin tail is removed and placed onto a piece of filter paper. The piece of filter paper containing the tissue can be readily visualized and correctly placed into a PCR tube. Genomic DNA extraction is then performed, followed by PCR amplification of the region of interest to genotype the specimen. The advantages of this method include a high PCR efficiency, a low false positive rate and a low mortality rate. Additionally, genotyping of large numbers of embryos is possible using this protocol. The protocol described herein allows fin-clipping of hundreds of larvae within 2 - 3 h using this approach and correct identification of genotyped fish and tail regeneration within two days.

<table>
	<tbody>
		<tr>
			<td>Petri dish</td>
			<td>Corning</td>
			<td>430167</td>
			<td></td>
		</tr>
		<tr>
			<td>Microscalpel</td>
			<td>World Precision Instruments</td>
			<td>500249</td>
			<td></td>
		</tr>
		<tr>
			<td>Stereomicroscope</td>
			<td>Nikon</td>
			<td>SMZ1500</td>
			<td></td>
		</tr>
		<tr>
			<td>Tricaine/Ethyl 3-aminobenzoate methanesulfonate</td>
			<td>Sigma-Aldrich</td>
			<td>E10521</td>
			<td></td>
		</tr>
		<tr>
			<td>96-well Tissue-culture plate</td>
			<td>Costar</td>
			<td>3595</td>
			<td></td>
		</tr>
		<tr>
			<td>96-well PCR plate</td>
			<td>Fisher</td>
			<td>14230232</td>
			<td></td>
		</tr>
		<tr>
			<td>NaOH</td>
			<td>Fisher</td>
			<td>S25548A&#160;</td>
			<td></td>
		</tr>
		<tr>
			<td>Tris base&#160;</td>
			<td>Sigma-Aldrich</td>
			<td>T1503</td>
			<td></td>
		</tr>
		<tr>
			<td>NaCl</td>
			<td>Fisher</td>
			<td>BP358-10</td>
			<td></td>
		</tr>
		<tr>
			<td>KCl</td>
			<td>Sigma-Aldrich</td>
			<td>P3911</td>
			<td></td>
		</tr>
		<tr>
			<td>MgSO4</td>
			<td>Sigma-Aldrich</td>
			<td>230391</td>
			<td></td>
		</tr>
		<tr>
			<td>5% Chelex 100 sodium form</td>
			<td>Sigma-Aldrich</td>
			<td>C7901</td>
			<td></td>
		</tr>
		<tr>
			<td>GelRed 1x</td>
			<td>Biotium</td>
			<td>41003</td>
			<td></td>
		</tr>
		<tr>
			<td>Boric Acid</td>
			<td>Sigma-Aldrich</td>
			<td>B6768</td>
			<td></td>
		</tr>
		<tr>
			<td>Sub-Cell Model 192 system</td>
			<td>Bio-Rad&#160;</td>
			<td>1704508</td>
			<td></td>
		</tr>
		<tr>
			<td>30 % Acrylamide/Bis Solution, 37.5: 1</td>
			<td>Bio-Rad&#160;</td>
			<td>1610158</td>
			<td></td>
		</tr>
		<tr>
			<td>GoTaq Green Master Mix, 2X</td>
			<td>Promega</td>
			<td>M7123</td>
			<td></td>
		</tr>
		<tr>
			<td>paper wipes (Kimwipes&#174; disposable wipers)</td>
			<td>Sigma-Aldrich</td>
			<td>Z188956</td>
			<td></td>
		</tr>
		<tr>
			<td>1kb-plus molecylar weight marker&#160;</td>
			<td>Thermo Scientific</td>
			<td>SM1343</td>
			<td></td>
		</tr>
		<tr>
			<td>Nuclease free water&#160;</td>
			<td>Sigma-Aldrich</td>
			<td>W4502</td>
			<td></td>
		</tr>
		<tr>
			<td>Sub-Cell Model 192 system (high-throughput agarose gel system)</td>
			<td>Bio-Rad&#160;</td>
			<td>1704508</td>
			<td></td>
		</tr>
		<tr>
			<td>vertical gel electrophoresis system Mini-PROTEAN Tetra Cell&#160;</td>
			<td>Bio-Rad&#160;</td>
			<td>1658004</td>
			<td></td>
		</tr>
	</tbody>
</table>

high-throughput dna extraction and genotyping of 3dpf zebrafish larvae by fin clipping

Zebrafish (Danio rerio) possess orthologues for 84% of the genes known to be associated with human diseases. In addition, these animals have a short generation time, are easy to handle, display a high reproductive rate, low cost, and are easily amenable to genetic manipulations by microinjection of DNA in embryos. Recent advances in gene editing tools are enabling precise introduction of mutations and transgenes in zebrafish. Disease modeling in zebrafish often leads to larval phenotypes and early death which can be challenging to interpret if genotypes are unknown. This early identification of genotypes is also needed in experiments requiring sample pooling, such as in gene expression or mass spectrometry studies. However, extensive genotypic screening is limited by traditional methods, which in most labs are performed only on adult zebrafish or in postmortem larvae. We addressed this problem by adapting a method for the isolation of PCR-ready genomic DNA from live zebrafish larvae that can be achieved as early as 72 h post-fertilization (hpf). This time and cost-effective technique, improved from a previously published genotyping protocol, allows the identification of genotypes from microscopic fin biopsies. The fins quickly regenerate as the larvae develop. Researchers are then able to select and raise the desired genotypes to adulthood by utilizing this high-throughput PCR-based genotyping procedure.

The efficiency of the protocoll was demonstrated by genotyping the offspring of an heterozygous cross (aldh7a1+/ot100, here shown as aldh7a1+/-) (Figure 3A-C)9. The aldh7a1ot100 mutant allele has a 5-base pair (bp) insertion in the first coding exon of the zebrafish aldh7a1 gene that leads to frameshift and loss-of-function due to an early stop codon9. Four primers were used in a multiplex reaction to obtain three distinctive bands to differentiate between WT, heterozygous (aldh7a1+/-), and homozygous mutant (aldh7a1-/-) genotypes (from Pena, et al.9): Gen1_FW 5&#39;-ATGATGCAGCGCGTGCTGAC-3&#39;, &#34;Gen2_RV&#34;:5&#39; -CCCTTTGAACCTCACAGGAGTT-3&#39;, &#34;5 nt-ins-specific_FW&#34;:5&#39;-TGTTTTCAACGGTTCAACGG-3&#39;, and &#34;WT-specific_RV&#34;:5&#39;-TCCCTGTCCTCCCCAAGAAC-3&#39;). The amplification of both alleles using the Gen1_FW and Gen1_RV primers resulted in a 434-bp amplicon. A 293-bp band arose from specific amplification of the mutant allele, and a 195-bp band was obtained specifically for the WT allele, as shown in Figure 3A. Following PCR, 8 &#181;L of each 20 &#181;L reaction volume was analyzed on a 1% agarose/sodium borate gel, as shown in Figure 3B. Sufficient larval DNA was recovered from 98% (n = 576) of the samples, resulting in a high PCR efficiency rate. DNA extraction using the NaOH technique obtains on average 4.7 0.5 ng/&#181;L of larval DNA per sample following the fin biopsy procedure, in ~30 &#181;L volume. DNA extraction using chelating resin results in a similar yield, on average 3.8 0.5 ng/&#181;L of DNA per sample. This amount of DNA collected from larval fin transections generates PCR-ready genomic DNA of sufficient quality to allow for the identification of genotypes. Each DNA sample can be used in &#732;30 PCR amplification reactions. Amplification products obtained using primers Gen1_FW and Gen2_RV can also be used for sequencing applications9, which successfully identified the 5-bp insertion in the homozygous mutants (aldh7a1-/-) (Figure 3C). Sanger sequencing was performed via an external service to test if the PCR products were suitable for this application. DNA samples from agarose-gel confirmed homozygous mutants and WTs (n = 3 each) were used. High Phred19 scores (&#62;30) were obtained indicating high quality reads, except for the first and last 40 base pairs.
This protocol was subsequently used to genotype various other zebrafish mutations by heteroduplex melting analysis. The plpbpot101 and plpbpot102 mutant alleles of the plpbp gene (Figure 4A-D) each lead to a frameshift and early stop codon. In order to identify plpbp-null zebrafish larvae, two primers were used to amplify the first coding exon including the mutation site (plpbp-F 5&#39;-GCACTCTGGCTATGTGGAGA-3&#39; and plpbp-R 5&#39;-AGCTGTCACTCATCCCTCGT-3&#39;). The amplification of the plpbp fragment in WT animals results in a 272-bp amplicon (homoduplex). The amplicon obtained from plpbpot101 (4-bp deletion allele, Figure 4C) consists of a fragment of 268-bp and that of the plpbpot102 (5bp-substitution, 2-bp deletion allele, Figure 4D), 270-bp, as seen in Figure 4A-B. After denaturation and annealing, the PCR fragments from heterozygous animals would contain heteroduplex and homoduplex DNA17. Homoduplex and heteroduplex bands can be easily separated and visualized using polyacrylamide gel electrophoresis (PAGE). The presence of an open angle between matched and mismatched DNA strands caused by imperfect annealing due to the occurrence of mutations cause the heteroduplex DNA to migrate at a significantly slower pace than homoduplex DNA17. Distinct migration patterns are produced by distinct mutations. The heterozygous genotypes (plpbp+/ot101, plpbp+/ot102) would therefore display heteroduplex DNA fragments, as well as the compound heterozygous (plpbpot101/ot102) (Figure 4A-B). Precise genotyping is obtained by this heteroduplex melting assay as a unique PAGE pattern is obtained for each genotype and distinct heterozygous mutations can be visualized (Figure 4A-B). Homozygous mutants would display a homoduplex pattern in the PAGE gel and therefore be indistinguishable to WTs. To distinguish these genotypes, another round of PAGE gel analyses should be performed in which PCR products from WT alleles need to be mixed with the samples to be studied, as described elsewhere17.
About 90% (88/98) of WT and heterozygous embryos were successfully raised to adulthood (&#62;2 months). As aldh7a1 and plpbp are loss-of-function mutants that display an early death phenotype, untreated animals could not be raised to adulthood. However, the genotypes of all surviving WT and heterozygous adults were identical to larval fin clipping results, indicating a 100% accuracy rate for the identification of genotypes using the larval fin biopsy procedure.
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/58024/58024fig1.jpg" /> 
Figure 1. Larval fin clip. (A). Non-severed larval fin at three days post-fertilization (dpf). The line represents the site of transection, located within the pigment gap. The black arrowhead demonstrates the limit of the caudal blood circulation. (B). Larva following fin clipping procedure at 3 dpf. (C). Larval zebrafish fin regrowth at 5 dpf. Fin regeneration from a blastemal formation is observed only 2 days post-fin transection. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/58024/58024fig1large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 2" class="xfigimg" src="/files/ftp_upload/58024/58024fig2.jpg" /> 
Figure 2. Procedure for larval fin clipping. (A). The larvae are placed onto the tape and excess embryo media is removed. Under the microscope, the fin is transected at the region of the pigment gap as shown in Figure 1A. Using a microscalpel, the fin is collected and placed onto a filter paper surface simply by touch (B), and can be readily visualized as a black spot. Hold the piece of filter paper containing the fin and cut a small square surrounding the desired region (C). Place the small piece of filter paper containing the fin into a well of a 96 well plate (D). The corresponding larva should be placed into the corresponding well of a flat-bottom 96-well plate in 200 &#181;L of embryo media (E). The filter paper piece should be submerged in the lysis solution (F). <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/58024/58024fig2large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 3" class="xfigimg" src="/files/ftp_upload/58024/58024fig3.jpg" /> 
Figure 3. Genotyping of aldh7a1 heterozygous cross from larval fin biopsies. (A). Expected result where amplification of both WT and aldh7a1ot100 alleles results in a 434-bp amplicon. A 293-bp band arises from amplification of the mutant allele (aldh7a1ot100, 5-bp insertion), and a 195-bp band is obtained for the WT allele. (B). Example of PCR amplification from aldh7a1 larval fin tissues. A 1 kb molecular weight marker is shown in lane 7. Lanes 1 - 6 each represent a single fin biopsy. PCR results identify aldh7a1-/- (aldh7a1ot100/ot100) genotypes (lane 1), aldh7a1+/- heterozygous genotypes (aldh7a1+/ot100) (lanes 3, 4, and 5) and WT (aldh7a1+/+) genotypes (lanes 2 and 6). (C). Alignment between the sequenced WT and aldh7a1ot100 allele showing the position of the 5-bp insertion mutation. This figure was adapted from the Supplemental Figure 5 of Pena&#160;et al.9 with permission granted from the Genetics Society of America. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/58024/58024fig3large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 4" class="xfigimg" src="/files/ftp_upload/58024/58024fig4.jpg" /> 
Figure 4. Genotyping of plpbp compound heterozygous (2 bp deletion /4 bp deletion) from larval fin biopsies. (A). Expected result where amplification of the WT allele resulted in a 272-bp amplicon. The 4-bp deletion allele (plpbpot101) and the 5-bp substitution with 2-bp deletion allele (plpbpot102) produce amplicon lengths of 268-bp and 270-bp respectively. A heteroduplex is formed between a WT allele and the mutant allele in a heterozygous genotype (plpbp+/-). Formation of a heteroduplex due to mismatching alleles will cause retardation of the band within the gel compared to the WT homoduplex. (B). PCR amplification resulting from larval fin clippings of the offspring of a plpbp+/ot101x plpbp+/ot102 cross. A 1 kb molecular weight marker is shown in lane 1. Lanes 2 - 10 each represent a single fin biopsy. PCR results identify mutant genotypes (plpbp-null compound heterozygous plpbpot101/ot102, lanes 2, 4, and 8), heterozygous genotypes (lanes 3, 6, 7, 9, and 10) and WT genotypes (lane 5). (C). Alignment between the sequenced WT and plpbpot101 allele showing the position of the 4-bp deletion mutation. (D). Alignment between the sequenced WT and plpbpot102 allele showing the position of the 2-bp deletion and 5-bp substitution (bold). <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/58024/58024fig4large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>Components&#160;</td>
			<td>20 &#956;l reaction</td>
			<td>100 reactions</td>
			<td>Final Conc.</td>
		</tr>
		<tr>
			<td>Nuclease-free H2O&#160;</td>
			<td>7.45 &#956;L</td>
			<td>745 &#956;L</td>
			<td></td>
		</tr>
		<tr>
			<td>Go-Taq 2x (Promega, M7122)</td>
			<td>10 &#956;L</td>
			<td>1000 &#956;L</td>
			<td>1x</td>
		</tr>
		<tr>
			<td>primer Gen1_FW&#160;</td>
			<td>0.25 &#956;L</td>
			<td>25 &#956;L</td>
			<td>0.125 &#956;M</td>
		</tr>
		<tr>
			<td>primer 5 nt-ins-specific_FW</td>
			<td>0.3 &#956;L</td>
			<td>30 &#956;L</td>
			<td>0.15 &#956;M&#160;</td>
		</tr>
		<tr>
			<td>Primer Gen2_RV</td>
			<td>0.25 &#956;L</td>
			<td>25 &#956;L</td>
			<td>0.125 &#956;M</td>
		</tr>
		<tr>
			<td>Primer WT-specific_RV</td>
			<td>0.25 &#956;L</td>
			<td>25 &#956;L</td>
			<td>0.125 &#956;M</td>
		</tr>
		<tr>
			<td>DNA</td>
			<td>1.5 &#956;L</td>
			<td>--</td>
			<td></td>
		</tr>
	</tbody>
</table>
Table 1. Reaction conditions for multiplex PCR used for the aldh7a1 allele in this protocol. The primers used are as follows: Gen1_FW 5&#39;-ATGATGCAGCGCGTGCTGAC-3&#39;, &#34;Gen2_RV&#34;:5&#39; -CCCTTTGAACCTCACAGGAGTT-3&#39;, &#34;5 nt-ins-specific_FW&#34;:5&#39;-TGTTTTCAACGGTTCAACGG-3&#39;, and &#34;WT-specific_RV&#34;:5&#39;-TCCCTGTCCTCCCCAAGAAC-3&#39;. Primer stocks were 10 &#181;M.
<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>Components&#160;</td>
			<td>13 &#956;l reaction</td>
			<td>100 reactions</td>
			<td>Final Conc.</td>
		</tr>
		<tr>
			<td>H2O&#160;</td>
			<td>3.25 &#956;L</td>
			<td>325 &#956;L</td>
			<td></td>
		</tr>
		<tr>
			<td>Go-Taq 2x (Promega, M7122)</td>
			<td>6.25 &#956;L</td>
			<td>625 &#956;L</td>
			<td>1x</td>
		</tr>
		<tr>
			<td>primer FW&#160;</td>
			<td>1 &#956;L</td>
			<td>100 &#956;L</td>
			<td>0.77 &#956;M</td>
		</tr>
		<tr>
			<td>Primer RV&#160;</td>
			<td>1 &#956;L</td>
			<td>100 &#956;L</td>
			<td>0.77 &#956;M</td>
		</tr>
		<tr>
			<td>DNA</td>
			<td>1.5 &#956;L</td>
			<td>--</td>
			<td></td>
		</tr>
	</tbody>
</table>
Table 2. Reaction conditions for PCR used for plpbp allele in this protocol. The forward primer used was PLPBP-F 5&#39;-GCACTCTGGCTATGTGGAGA-3&#39; and the reverse, PLPBP-R 5&#39;-AGCTGTCACTCATCCCTCGT-3&#39;. Primer stocks were 10 &#181;M.
<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>Cycle Step&#160;</td>
			<td>Temp.</td>
			<td>Time</td>
			<td>Cycles</td>
		</tr>
		<tr>
			<td>Initial denaturation&#160;</td>
			<td>95 &#176;C</td>
			<td>3 min</td>
			<td>1</td>
		</tr>
		<tr>
			<td>Denaturing&#160;</td>
			<td>95 &#176;C</td>
			<td>30 sec</td>
			<td rowspan="3">36</td>
		</tr>
		<tr>
			<td>Annealing*</td>
			<td>X &#176;C</td>
			<td>30 sec</td>
		</tr>
		<tr>
			<td>Extension</td>
			<td>72 &#176;C</td>
			<td>20 sec/kb</td>
		</tr>
		<tr>
			<td rowspan="2">Final Extension&#160;</td>
			<td>72 &#176;C</td>
			<td>5 min</td>
			<td rowspan="2">1</td>
		</tr>
		<tr>
			<td>4 &#176;C</td>
			<td>hold</td>
		</tr>
	</tbody>
</table>
Table 3. PCR conditions used in this protocol.

Watch this Scientific Journal Video about High-throughput DNA Extraction and Genotyping of 3dpf Zebrafish Larvae by Fin Clipping at JoVE.com

High-throughput DNA Extraction and Genotyping of 3dpf Zebrafish Larvae by Fin Clipping

Zebrafish have been used as reliable genetic model organisms in biomedical research, especially with the advent of gene-editing technologies. When larval phenotypes are expected, DNA extraction and genotype identification can be challenging. Here, we describe an efficient genotyping procedure for zebrafish larvae, by tail clipping, as early as 72-h post-fertilization.

Zebrafish (Danio rerio) possess orthologues for 84% of the genes known to be associated with human diseases. In addition, these animals have a short ...

high-throughput-dna-extraction-genotyping-3dpf-zebrafish-larvae-fin

Nanyang Technological University

Research

JoVE Journal

Genetics

13.9K Views.  Children's Hospital of Eastern Ontario Research Institute. Zebrafish have been used as reliable genetic model organisms in biomedical research, especially with the advent of gene-editing technologies. When larval phenotypes are expected, DNA extraction and genotype identification can be challenging. Here, we describe an efficient genotyping procedure for zebrafish larvae, by tail clipping, as early as 72-h post-fertilization.

Video: High-throughput DNA Extraction and Genotyping of 3dpf Zebrafish Larvae by Fin Clipping

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 ...

Improved Polymerase Chain Reaction-restriction Fragment Length Polymorphism Genotyping of Toxic Pufferfish by Liquid Chromatography/Mass Spectrometry

An improved version of a polymerase chain reaction (PCR)-restriction fragment length polymorphism (RFLP) method for genotyping toxic pufferfish species by liquid chromatography/electrospray ionization mass spectrometry (LC/ESI-MS) is described. DNA extraction is carried out using a silica membrane-based DNA extraction kit. After the PCR amplification using a detergent-free PCR buffer, restriction enzymes are added to the solution without purifying the reaction solution. A reverse-phase silica monolith column and a Fourier transform high resolution mass spectrometer having a modified Kingdon trap analyzer are employed for separation and detection, respectively. The mobile phase, consisting of 400 mM 1,1,1,3,3,3-hexafluoro-2-propanol, 15 mM triethylamine (pH 7.9) and methanol, is delivered at a flow rate of 0.4 ml/min. The cycle time for LC/ESI-MS analysis is 8 min including equilibration of the column. Deconvolution software having an isotope distribution model of the oligonucleotide is used to calculate the corresponding monoisotopic mass from the mass spectrum. For analysis of oligonucleotides (range 26-79 nucleotides), mass accuracy was 0.62 &#177; 0.74 ppm (n = 280) and excellent accuracy and precision were sustained for 180 hr without use of a lock mass standard.

An improved polymerase chain reaction-restriction fragment length polymorphism method for genotyping pufferfish species by liquid chromatography/mass spectrometry is described. A reverse-phase silica monolith column is employed for separating digested amplicons. This method can elucidate the monoisotopic masses of oligonucleotides, which is useful for identifying base composition.

An improved version of a polymerase chain reaction (PCR)-restriction fragment length polymorphism (RFLP) method for genotyping toxic pufferfish species by ...

Genome-wide Determination of Mammalian Replication Timing by DNA Content Measurement

Replication of the genome occurs during S phase of the cell cycle in a highly regulated process that ensures the fidelity of DNA duplication. Each genomic region is replicated at a distinct time during S phase through the simultaneous activation of multiple origins of replication. Time of replication (ToR) correlates with many genomic and epigenetic features and is linked to mutation rates and cancer. Comprehending the full genomic view of the replication program, in health and disease is a major future goal and challenge.
This article describes in detail the &#34;Copy Number Ratio of S/G1 for mapping genomic Time of Replication&#34; method (herein called: CNR-ToR), a simple approach to map the genome wide ToR of mammalian cells. The method is based on the copy number differences between S phase cells and G1 phase cells. The CNR-ToR method is performed in 6 steps: 1. Preparation of cells and staining with propidium iodide (PI); 2. Sorting G1 and S phase cells using fluorescence-activated cell sorting (FACS); 3. DNA purification; 4. Sonication; 5. Library preparation and sequencing; and 6. Bioinformatic analysis. The CNR-ToR method is a fast and easy approach that results in detailed replication maps.

We describe here a relatively fast and simple approach for mapping genome-wide mammalian replication timing, from cell isolation to the basic analysis of the sequencing results. A genomic map of a representative replication program will be provided following the protocol.

Replication of the genome occurs during S phase of the cell cycle in a highly regulated process that ensures the fidelity of DNA duplication. Each genomic ...

High Fat Diet Feeding and High Throughput Triacylglyceride Assay in Drosophila Melanogaster

Heart disease is the number one cause of human death worldwide. Numerous studies have shown strong connections between obesity and cardiac malfunction in humans, but more tools and research efforts are needed to better elucidate the mechanisms involved. For over a century, the genetically highly tractable model of Drosophila has been instrumental in the discovery of key genes and molecular pathways that proved to be highly conserved across species. Many biological processes and disease mechanisms are functionally conserved in the fly, such as development (e.g., body plan, heart), cancer, and neurodegenerative disease. Recently, the study of obesity and secondary pathologies, such as heart disease in model organisms, has played a highly critical role in the identification of key regulators involved in metabolic syndrome in humans.
Here, we propose to use this model organism as an efficient tool to induce obesity, i.e., excessive fat accumulation, and develop an efficient protocol to monitor fat content in the form of TAGs accumulation. In addition to the highly conserved, but less complex genome, the fly also has a short lifespan for rapid experimentation, combined with cost-effectiveness. This paper provides a detailed protocol for High Fat Diet (HFD) feeding in Drosophila to induce obesity and a high throughput triacylglyceride (TAG) assay for measuring the associated increase in fat content, with the aim to be highly reproducible and efficient for large-scale genetic or chemical screening. These protocols offer new opportunities to efficiently investigate regulatory mechanisms involved in obesity, as well as provide a standardized platform for drug discovery research for rapid testing of the effect of drug candidates on the development or prevention of obesity, diabetes and related metabolic diseases.

High Fat Diet Feeding and High Throughput Triacylglyceride Assay in Drosophila Melanogaster

This is a high fat diet feeding protocol to induce obesity in Drosophila, a model for understanding fundamental molecular mechanisms implicated in lipotoxicity. It also provides a high throughput triacylglyceride assay for measuring fat accumulation in Drosophila and potentially other (insect) models under various dietary, environmental, genetic or physiological conditions.

Heart disease is the number one cause of human death worldwide. Numerous studies have shown strong connections between obesity and cardiac malfunction in ...

Generation of Native Chromatin Immunoprecipitation Sequencing Libraries for Nucleosome Density Analysis

We present a modified native chromatin immunoprecipitation sequencing (ChIP-seq) experimental protocol compatible with a Gaussian mixture distribution based analysis methodology (nucleosome density ChIP-seq; ndChIP-seq) that enables the generation of combined measurements of micrococcal nuclease (MNase) accessibility with histone modification genome-wide. Nucleosome position and local density, and the posttranslational modification of their histone subunits, act in concert to regulate local transcription states. Combinatorial measurements of nucleosome accessibility with histone modification generated by ndChIP-seq allows for the simultaneous interrogation of these features. The ndChIP-seq methodology is applicable to small numbers of primary cells inaccessible to cross-linking based ChIP-seq protocols. Taken together, ndChIP-seq enables the measurement of histone modification in combination with local nucleosome density to obtain new insights into shared mechanisms that regulate RNA transcription within rare primary cell populations.

We present a modified native chromatin immunoprecipitation sequencing (ChIP-seq) methodology for the generation of sequence datasets suitable for a nucleosome density ChIP-seq analytical framework integrating micrococcal nuclease (MNase) accessibility with histone modification measurements.

We present a modified native chromatin immunoprecipitation sequencing (ChIP-seq) experimental protocol compatible with a Gaussian mixture distribution ...

G2-seq: A High Throughput Sequencing-based Technique for Identifying Late Replicating Regions of the Genome

Numerous techniques have been developed to follow the progress of DNA replication through the S phase of the cell cycle. Most of these techniques have been directed toward elucidation of the location and timing of initiation of genome duplication rather than its completion. However, it is critical that we understand regions of the genome that are last to complete replication, because these regions suffer elevated levels of chromosomal breakage and mutation, and they have been associated with both disease and aging. Here we describe how we have extended a technique that has been used to monitor replication initiation to instead identify those regions of the genome last to complete replication. This approach is based on a combination of flow cytometry and high throughput sequencing. Although this report focuses on the application of this technique to yeast, the approach can be used with any cells that can be sorted in a flow cytometer according to DNA content.

We describe a technique for combining flow cytometry and high throughput sequencing to identify late replicating regions of the genome.

Numerous techniques have been developed to follow the progress of DNA replication through the S phase of the cell cycle. Most of these techniques have ...

Robust DNA Isolation and High-throughput Sequencing Library Construction for Herbarium Specimens

Herbaria are an invaluable source of plant material that can be used in a variety of biological studies. The use of herbarium specimens is associated with a number of challenges including sample preservation quality, degraded DNA, and destructive sampling of rare specimens. In order to more effectively use herbarium material in large sequencing projects, a dependable and scalable method of DNA isolation and library preparation is needed. This paper demonstrates a robust, beginning-to-end protocol for DNA isolation and high-throughput library construction from herbarium specimens that does not require modification for individual samples. This protocol is tailored for low quality dried plant material and takes advantage of existing methods by optimizing tissue grinding, modifying library size selection, and introducing an optional reamplification step for low yield libraries. Reamplification of low yield DNA libraries can rescue samples derived from irreplaceable and potentially valuable herbarium specimens, negating the need for additional destructive sampling and without introducing discernible sequencing bias for common phylogenetic applications. The protocol has been tested on hundreds of grass species, but is expected to be adaptable for use in other plant lineages after verification. This protocol can be limited by extremely degraded DNA, where fragments do not exist in the desired size range, and by secondary metabolites present in some plant material that inhibit clean DNA isolation. Overall, this protocol introduces a fast and comprehensive method that allows for DNA isolation and library preparation of 24 samples in less than 13 h, with only 8 h of active hands-on time with minimal modifications.

This article demonstrates a detailed protocol for DNA isolation and high-throughput sequencing library construction from herbarium material including rescue of exceptionally poor-quality DNA.

Herbaria are an invaluable source of plant material that can be used in a variety of biological studies. The use of herbarium specimens is associated with ...

Profiling DNA Replication Timing Using Zebrafish as an In Vivo Model System

DNA replication timing is an important cellular characteristic, exhibiting significant relationships with chromatin structure, transcription, and DNA mutation rates. Changes in replication timing occur during development and in cancer, but the role replication timing plays in development and disease is not known. Zebrafish were recently established as an in vivo model system to study replication timing. Here is detailed the protocols for using the zebrafish to determine DNA replication timing. After sorting cells from embryos and adult zebrafish, high-resolution genome-wide DNA replication timing patterns can be constructed by determining changes in DNA copy number through analysis of next generation sequencing data. The zebrafish model system allows for evaluation of the replication timing changes that occur in vivo throughout development, and can also be used to assess changes in individual cell types, disease models, or mutant lines. These methods will enable studies investigating the mechanisms and determinants of replication timing establishment and maintenance during development, the role replication timing plays in mutations and tumorigenesis, and the effects of perturbing replication timing on development and disease.

Profiling DNA Replication Timing Using Zebrafish as an In Vivo Model System

Zebrafish were recently used as an in vivo model system to study DNA replication timing during development. Here is detailed the protocols for using zebrafish embryos to profile replication timing. This protocol can be easily adapted to study replication timing in mutants, individual cell types, disease models, and other species.

DNA replication timing is an important cellular characteristic, exhibiting significant relationships with chromatin structure, transcription, and DNA ...

High-throughput Identification of Synergistic Drug Combinations by the Overlap2 Method

Although antimicrobial drugs have dramatically increased the lifespan and quality of life in the 20th century, antimicrobial resistance threatens our entire society's ability to treat systemic infections. In the United States alone, antibiotic-resistant infections kill approximately 23,000 people a year and cost around 20 billion USD in additional healthcare. One approach to combat antimicrobial resistance is combination therapy, which is particularly useful in the critical early stage of infection, before the infecting organism and its drug resistance profile have been identified. Many antimicrobial treatments use combination therapies. However, most of these combinations are additive, meaning that the combined efficacy is the same as the sum of the individual antibiotic efficacy. Some combination therapies are synergistic: the combined efficacy is much greater than additive. Synergistic combinations are particularly useful because they can inhibit the growth of antimicrobial drug resistant strains. However, these combinations are rare and difficult to identify. This is due to the sheer number of molecules needed to be tested in a pairwise manner: a library of 1,000 molecules has 1 million potential combinations. Thus, efforts have been made to predict molecules for synergy. This article describes our high-throughput method for predicting synergistic small molecule pairs known as the Overlap2 Method (O2M). O2M uses patterns from chemical-genetic datasets to identify mutants that are hypersensitive to each molecule in a synergistic pair but not to other molecules. The Brown lab exploits this growth difference by performing a high-throughput screen for molecules that inhibit the growth of mutant but not wild-type cells. The lab's work previously identified molecules that synergize with the antibiotic trimethoprim and the antifungal drug fluconazole using this strategy. Here, the authors present a method to screen for novel synergistic combinations, which can be altered for multiple microorganisms.

High-throughput Identification of Synergistic Drug Combinations by the Overlap2 Method

Synergistic drug combinations are difficult and time-consuming to identify empirically. Here, we describe a method for identifying and validating synergistic small molecules.

Although antimicrobial drugs have dramatically increased the lifespan and quality of life in the 20th century, antimicrobial resistance threatens our ...

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 ...