We would like to thank Katherine Roberts for her care of the axolotl colony. Funding for this work was provided by the Connecticut Innovations Regenerative Medicine Research Fund (15RMA-YALE-09 and 15-RMB-YALE-01) and the Eunice Kennedy Shriver National Institute of Child Health and Human Development (Individual Postdoctoral Fellowship F32HD086942).

Experimental procedures used in this protocol were approved by the Yale University Institutional Animal Care and Use Committee (IACUC, 2017&#8211;10557) and were in accordance with all federal policies and guidelines governing the use of vertebrate animals. All animal experiments were carried out on Ambystoma mexicanum (axolotls) in facilities at Yale University.
1. Diploid Embryo Generation
<ol>
	<li>Obtain GFP+ diploid embryos to serve as graft hosts through natural mating using one or two gfp parents24.</li>
	<li>Collect freshly laid diploid eggs and place them in a metal sieve.</li>
	<li>Rinse the eggs thoroughly with 40% Holtfreter&#39;s solution (20 mM NaCl, 0.2 mM KCl, 0.8 mM NaHCO3, 0.2 mM CaCl2, 4 mM MgSO4, pH to 7.4).</li>
	<li>Place the eggs in fresh 40% Holtfreter&#39;s solution and store at 12 &#176;C. 
	NOTE: Diploid embryos should always be obtained before moving on to haploid embryo generation.</li>
</ol>
2. Haploid Embryo Generation
<ol>
	<li>Female gamete donor preparation
	<ol>
		<li>48 h before conducting in vitro activation, anesthetize a sexually mature white or white/RFP female axolotl by immersion in 1 g/L HEPES-buffered MS-222 in 40% Holtfreter&#39;s solution25.</li>
		<li>Ensure that the animal is fully anesthetized after approximately 30 min of immersion by firmly pinching its tail between the thumb and forefinger. Fully anesthetized animals will not physically respond to any pinch.</li>
		<li>Prepare a solution of human chorionic gonadotropin (HCG) containing 10,000 U/mL in sterile saline.</li>
		<li>Using a 30 G insulin syringe, inject 0.15 CC of HCG (1,500 units) into the musculature dorsal to the hind limb of the anesthetized female at 45&#176; angle to the midline to avoid contact with the spinal cord.</li>
		<li>Return the female to fresh 40% Holtfreter&#39;s solution and place in an 8&#8722;12 &#176;C refrigerator.</li>
		<li>After 48 h, return the female to room temperature. Place a few stones or plastic plants in her container as surfaces for egg laying. 
		NOTE: Females injected with HCG will often deposit empty jelly cases for several hours before laying eggs.</li>
		<li>Remove the stones or plastic plants after the female has begun consistently laying eggs.</li>
		<li>Allow the female to sit in the empty tank for 30 min to 1 h. 
		NOTE: Withholding materials for the animal to lay eggs on will allow tighter control of oocyte collection.</li>
	</ol>
	</li>
	<li>Male gamete collection
	<ol>
		<li>While the female&#39;s egg laying is stalled, anesthetize a sexually mature GFP+ male axolotl to serve as a sperm donor, as in step 2.1.1.</li>
		<li>Place the fully anesthetized male on his back on damp paper towels under a dissecting microscope.</li>
		<li>If the experimenter is righthanded, position the tip of a P1000 pipette at the base of the cloaca with the right hand.</li>
		<li>Place the left forefinger and thumb of the left hand 2&#8722;3 cm rostral to the pelvis. Gently squeeze the animal while moving the fingers towards the hind legs to flush out spermic urine samples.</li>
		<li>Collect each that is flushed out of the cloaca into an individual microcentrifuge tube. Repeat this process to collect 6 to 10 samples.</li>
		<li>Pipette 5.0 &#181;L from each sample onto a petri dish to inspect the quality of the sperm using an inverted microscope. 
		NOTE: Concentrated sperm samples are a milky white color, typically range from 5 to 20 &#181;L, and are usually retrieved after several, higher volume spermic urine samples are extracted. Sperm will collect at the bottom of the tubes in higher volume samples when left undisturbed. Concentrated samples of healthy sperm are highly motile and lose activity as their concentration decreases. Healthy males can produce up to 50 &#181;L of concentrated sperm.</li>
	</ol>
	</li>
	<li>Female gamete collection
	<ol>
		<li>After obtaining and confirming a healthy sperm sample, anesthetize the HCG-injected female axolotl as in step 2.1.1.</li>
		<li>Place the fully anesthetized female on damp paper towels on her back.</li>
		<li>Extract unfertilized eggs from the female using a hand motion similar to that in step 2.2.4.</li>
		<li>Collect the eggs using wet forceps and transfer them to a 10 cm petri dish. Treat eggs with irradiated sperm within 15 min of collection.</li>
	</ol>
	</li>
	<li>Male gamete preparation and in vitro activation
	<ol>
		<li>Use a P10 or P20 pipette to pipette the sperm up and down, breaking apart the clumps to form a homogenous suspension.</li>
		<li>Approximate the percentage of motile sperm by placing a 0.5 &#181;L drop of undiluted sperm suspension on a petri dish lid or glass slide and examining with an inverted microscope.</li>
		<li>Add 9.5 &#181;L of 0.1x Marc&#39;s modified Ringer&#39;s solution (MMR; Table of Materials) to this sample to make a 20x sperm dilution. Gently pipette up and down to mix.</li>
		<li>With an inverted microscope, count the sperm in three 1.0 &#181;L drops of the 20x diluted aliquot on a petri dish cover or hemocytometer to estimate the sperm concentration of the undiluted suspension.</li>
		<li>Prepare the sperm for irradiation by diluting an aliquot of the original sperm sample to about 80,000 motile cells/mL in sterile 0.1x MMR.</li>
		<li>Count the number of eggs obtained within the past 15 min. Add 0.5 &#181;L of the freshly diluted sperm from step 2.4.5 per egg to a petri dish. Use the pipette tip to spread this suspension into a 1 mm thick layer.</li>
		<li>Using a plastic lift, place the sample 4 cm from the bulbs of a 254 nm crosslinker. Genetically inactivate the sperm by irradiating the sample with 800,000 uJ/mm2.</li>
		<li>Using a P10 pipette, pipette the suspension onto the unfertilized eggs, coating each egg with 0.25&#8722;0.5 &#181;L of irradiated sperm. Allow the eggs to sit at room temperature for 30 min.</li>
		<li>After 30 min, flood the eggs with 0.1x MMR. Immediately place the eggs in an 8&#8722;10 &#176;C incubator for injections the next day or at 18 &#176;C for injections within 7 hours post activation (hpa).</li>
		<li>Dejelly haploids using sharp forceps 30 min after hydration. Immediately place the eggs in an 8&#8722;10 &#176;C incubator for injections the next day or at 18 &#176;C for injections the same day.</li>
	</ol>
	</li>
</ol>
3. Haploid Mutagenesis and Maintenance
<ol>
	<li>CRISPR/Cas9 microinjections
	<ol>
		<li>Design sgRNAs using CRISPRscan and synthesize26,27.</li>
		<li>For multiplex mutagenesis, prepare a stock of 5 sgRNAs (10 ng/&#181;L for each sgRNA) and Cas9 protein (1 &#181;g/&#181;L). Prepare a 100-fold dilution of this stock (0.1 ng/&#181;L per sgRNA, 10 ng/&#181;L Cas9) for injection. For single gene, high mutation frequency mutagenesis, follow the protocol outlined previously28.</li>
		<li>Inject haploid embryos at the single cell stage 7 hpa if stored at 18 &#176;C or inject embryos the next day at the 2&#8722;8 cell stage if they are stored at 8&#8722;10 &#176;C.</li>
		<li>Transfer the embryos to 1.0x MMR with 20% polysucrose 400.</li>
		<li>If single cell, inject each embryo with a 5 nL drop of the injection solution (approximately &#188; radius of the egg) containing a total mass of 0.5 pg/sgRNA and 50 pg Cas9 protein. If multicellular, distribute this mass by injecting smaller volumes into multiple cells.</li>
		<li>Allow the embryos to heal in the polysucrose 400 for a minimum of 4 h and a maximum of 18 h at 18 &#176;C.</li>
	</ol>
	</li>
	<li>Haploid embryo housing
	<ol>
		<li>Transfer the embryos to 0.1x MMR with antibiotic-antimycotic.</li>
		<li>House each embryo in an individual well of a 24-well plate, as some will die or develop abnormally. Maintain at 16&#8722;18 &#176;C. Lower temperatures can be used to prolong development, if necessary.</li>
		<li>Replace the media with fresh 0.1x MMR with antibiotic-antimycotic every other day.</li>
	</ol>
	</li>
</ol>
4. Diploid Host Embryo Preparation
<ol>
	<li>Maintain embryos in the jelly coating at 12 to 16 &#176;C until they reach stage 22 to 26.</li>
	<li>Collect the embryos that are ready for grafting, place them within a sieve (4 mm mesh size), and gently rinse them with 40% Holtfreter&#39;s solution.</li>
	<li>Transfer the embryos into filter-sterilized 0.1x MMR containing 1.5% bleach for up to 2 min. Completely submerge the embryos in the bleach solution and swirl them gently to ensure that the jelly coating makes full contact with the bleach solution to kill the microbes present on the embryos.</li>
	<li>After 2 min, dilute the bleach solution containing the embryos with an equal volume of filter-sterilized 0.1x MMR.</li>
	<li>Gently pour the embryos into a bleach-sanitized sieve (4 mm mesh size) and rinse the embryos five times with filter-sterilized 0.1x MMR.</li>
	<li>Place the embryos into sterile 10 cm petri dishes with 0.1x MMR with antibiotics for dejellying (penicillin 100 units/mL, streptomycin 100 &#181;g/&#181;L, 0.25 &#181;g/mL, gentamicin 25 &#181;g/mL).</li>
	<li>Under a fluorescent stereomicroscope, remove the jelly coats and vitelline membranes from GFP+ embryos using sharp forceps (tip dimensions 0.05 x 0.01 mm2).</li>
	<li>Transfer the GFP+ embryos to a new petri dish. Minimize the amount of liquid transferred from the petri dish where they were dejellied.</li>
	<li>Rinse the GFP+ host embryos with sterile 0.1x MMR with antibiotics four to six times in order to remove contaminants.</li>
	<li>Place the clean embryos at 4 &#176;C overnight before grafting. 
	NOTE: Cooling the embryos makes them more rigid and clean separation of the mesoderm from the endoderm feasible.</li>
</ol>
5. Haploid-diploid Chimera Generation
<ol>
	<li>Surgical dish and media preparation
	<ol>
		<li>Prepare sterile surgical operating dishes by pouring autoclaved 2% agarose in 0.1x MMR into sterile 35 mm easy-grip petri dishes. Fill the petri dishes halfway with agarose.</li>
		<li>After the agarose cools, use a sterile scalpel to cut a 25 mm long, slanted trough in the agarose to hold the embryos in place.</li>
		<li>Fill the dish with sterile surgical media (0.1x MMR with anti-mycoplasma 2.5 &#181;g/mL, amphotericin B 0.25 &#181;g/mL, and ciprofloxacin 10.0 &#181;g/mL) and refrigerate at 4 &#176;C.</li>
	</ol>
	</li>
	<li>Embryo grafting procedure
	<ol>
		<li>Place one healthy haploid donor with one or two stage-matched GFP+ diploid host embryos inside the trough of the prechilled operating dish containing surgical media (Figure 3). 
		NOTE: Perform the procedure on a cooling stage (10 &#176;C or lower) if possible.</li>
		<li>Use two ultra-fine, autoclaved forceps (straight tip, tip dimensions 0.05 x 0.02 mm2) to remove the ectoderm and mesoderm layers from the host with the limb bud near the center (Figure 4). 
		NOTE: The rectangular tissue graft region encompasses the limb bud and extend from the ninth somite to the posterior half of the gill bulge, about 2 mm, along the anterior posterior axis. Along the dorsoventral axis, the grafted region spans approximately 1.5 mm, including the somites to just beyond the ventral edge of the gill bulge. The grafted region includes all lateral plate mesoderm and the lateral halves of the somites, without disturbing underlying endoderm. See Figure 4 and the accompanying video for details.</li>
		<li>Set aside the host tissue and remove an equivalently sized tissue sheet from the haploid donor using the same methods.</li>
		<li>Place the haploid donor tissue sheet onto the corresponding region of the donor embryo.</li>
		<li>Secure the tissue by covering it with an autoclaved, rectangular glass shard from a crushed microscope cover glass and gently pressing it into the host embryo body.</li>
		<li>Flip the haploid donor embryo onto its other side to harvest the limb bud for the second host embryo. Repeat steps 5.2.2 through 5.2.5.</li>
		<li>Carefully remove the remaining excess host tissues and the donor embryo from the dish.</li>
		<li>Leave the grafts with the glass shard anchors in place for 60&#8722;75 min, checking every 20 min to ensure that the glass has not slipped off.</li>
		<li>After the tissue grafts fully adhere, use the forceps to slowly peel off the glass shard anchors.</li>
	</ol>
	</li>
	<li>Chimera maintenance
	<ol>
		<li>Transfer the engrafted embryos to fresh surgical media and maintain them at 8&#8722;12 &#176;C overnight to heal. Individually house engrafted embryos in 12 or 24-well plates.</li>
		<li>After 36&#8722;48 h, transfer the engrafted embryos to sterile 0.1x MMR antibiotic-antimycotic. The strong antibiotics in the surgical media cause toxicity in engrafted embryos after 3 days.</li>
		<li>Replace the media with fresh 0.1x MMR and antibiotics every 2&#8722;3 days.</li>
		<li>Maintain the engrafted embryos at 18 &#176;C until they are able to feed28.</li>
		<li>After 1 to 2 months of development and care, haploid limbs can be scored for purity of the graft using a fluorescent dissection microscope. 
		NOTE: The presence of non-neural or non-blood host-derived GFP-tissue in limbs is an indicator of impure grafting and is often associated with abnormal limb development. These animals should be excluded from further analysis.</li>
	</ol>
	</li>
</ol>

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G. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Normal+stages+of+development+of+the+axolotl,+Ambystoma+mexicanum.">Normal stages of development of the axolotl, Ambystoma mexicanum.</a> Developmental Biology. 42 (2), 391-399 (1975).</li><li>Hertwig, G. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Beitrage+Zum+Determinations-+Und+Regenerationsproblem+Mittels+Der+Transplantation+Haploidkerniger+Zellen.">Beitrage Zum Determinations- Und Regenerationsproblem Mittels Der Transplantation Haploidkerniger Zellen.</a> Archiv f. Entwicklungsmechanik. 111, 292-316 (1927).</li><li>Fei, J. -. F., 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+gene+knockin+in+axolotl+and+its+use+to+test+the+role+of+satellite+cells+in+limb+regeneration.">Efficient gene knockin in axolotl and its use to test the role of satellite cells in limb regeneration.</a> Proceedings of the National Academy of Sciences of the United States of America. 114 (47), 12501-12506 (2017).</li></ol>

The authors have nothing to disclose.

There are a few critical steps in our protocol for generating haploid-diploid chimeras that the operating technician should consider for consistent grafting results.
The most likely reason for haploid generation to fail is due to poor in vitro activation conditions. The proper quantities of motile sperm must be used to activate eggs. To prolong motility, sperm samples should always be maintained at 4 &#176;C. Before applying any sperm sample to eggs, check the viability of the sperm using an i...

Historically, embryonic tissue grafting in amphibians has been an important technique for exploring fundamental mechanisms of developmental biology and regeneration. The axolotl, a species of salamander, possesses an impressive ability to regenerate tissues and complex structures such as limbs and organs after injury or amputation. Similarly impressively, they can receive, without rejection, tissue grafts from other individuals at embryonic, juvenile, and adult stages1,2,3. Regions of embryos that produce whole structures such as limbs, tails, eyes, and heads, and more specific tissues, such as neuroectoderm and somites, can be grafted between embryos to produce chimeric animals1,2,4,5,6. For nearly a century, studies of such chimeric animals have provided crucial insights into regeneration, tissue differentiation, size control, and patterning1,7,8.
In the last the decade, numerous transcriptional studies of regenerating tissues have produced insights into the genetic programs underlying salamander regeneration9,10,11,12,13. These studies have added to an expanding list of candidate genes that, to date, are largely uncharacterized in the context of regeneration. Targeted mutagenesis techniques, such as CRISPR/Cas, now permit the investigation of such genes, and such genetic approaches are greatly facilitated by the recent sequencing and assembly of the large axolotl genome14,15,16.
We sought to develop techniques that coupled classic developmental biology with new genetic technology for the purpose of dissecting the mechanisms of regeneration. Methods for generating haploid embryos of axolotls and other salamanders have been established for decades17. While these techniques have long been noted to be advantages of salamanders as genetic model organisms18, few subsequent genetic studies have incorporated haploid animals. We use in vitro activation in the axolotl to produce haploid embryos that serve as tissue donors for grafting19. Using embryos carrying fluorescent genetic markers, we have devised reliable methods for generating limbs derived almost entirely from donor tissues (Figure 1A). By combining these two techniques, we have bypassed the late embryonic lethality associated with haploidy, allowing for the production of fully developed, grafted haploid limbs (Figure 1B, Figure 1B&#39;, and Figure 2).
By conducting CRISPR/Cas-mediated mutagenesis in haploid embryos prior to grafting to create chimeric axolotls with mutant haploid limbs, we may investigate gene function specifically within the context of limb development and regeneration. This allows the rescue of limbs from potentially embryonic-lethal mutant phenotypes. While CRISPR/Cas microinjection can generate animals that are highly mutant, such animals are typically highly mosaic, with some degree of retention of wildtype alleles and a variety of distinct mutations at targeted sites14,20. CRISPR-based mutagenesis in haploid cells increases the penetrance of single allele loss-of-function mutations, as they cannot be masked by retained wildtype alleles. For this reason, CRISPR-based screening in haploid cell lines is increasingly used to investigate the genetic basis of many cellular processes21,22,23. By combining CRISPR-based lineage tracing with our haploid limb bud grafting protocols, the approach described here can serve as a platform for haploid genetic screens in living animals20.

<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>#55 Dumont Forceps</td>
			<td>Fine Science Tools</td>
			<td>11295-1</td>
			<td>Only use Dumostar material (can be autoclaved)</td>
		</tr>
		<tr>
			<td>Amphotericin B</td>
			<td>Sigma Aldrich</td>
			<td>A2942-20ML</td>
			<td>20 mL</td>
		</tr>
		<tr>
			<td>Antibiotic-Antimycotic 100x</td>
			<td>Thermo Fisher</td>
			<td>15240062</td>
			<td></td>
		</tr>
		<tr>
			<td>Ciprofloxacin</td>
			<td>Sigma Aldrich</td>
			<td>17850-5G-F</td>
			<td></td>
		</tr>
		<tr>
			<td>Ficoll 400 (polysucrose 400)</td>
			<td>bioworld</td>
			<td>40600032-3</td>
			<td>Ficoll 400</td>
		</tr>
		<tr>
			<td>Gentamicin</td>
			<td>Sigma Aldrich</td>
			<td>G1914-250MG</td>
			<td></td>
		</tr>
		<tr>
			<td>Heating/Cooling Incubator</td>
			<td>RevSci</td>
			<td>RS-IF-233</td>
			<td></td>
		</tr>
		<tr>
			<td>Human Chorionic Gonadotropin</td>
			<td>Merk</td>
			<td></td>
			<td>Chorulon</td>
		</tr>
		<tr>
			<td>Megascript T7 Transcription Kit</td>
			<td>Thermo Fisher</td>
			<td>AM1334</td>
			<td>40 reactions</td>
		</tr>
		<tr>
			<td>Miroscope Cooling Stage</td>
			<td>Brook Industries</td>
			<td>Custom</td>
			<td>Custom</td>
		</tr>
		<tr>
			<td>NLS Cas9 Protein</td>
			<td>PNAbio</td>
			<td>CP01-200</td>
			<td>4 vials of 50 &#181;g protein each</td>
		</tr>
		<tr>
			<td>Plasmocin</td>
			<td>Invivogen</td>
			<td>ant-mpt-1</td>
			<td>Treatment level</td>
		</tr>
		<tr>
			<td>Recipes</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>1.0x Marc&#39;s modified Ringer&#39;s solution (MMR)</td>
			<td></td>
			<td></td>
			<td>0.1 M NaCl, 2 mM KCl, 1 mM MgSO4, 2 mM CaCl2, 0.1 mM EDTA, 5 mM HEPES (pH 7.8), ph 7.4</td>
		</tr>
		<tr>
			<td>40% Holtfreter&#39;s solution</td>
			<td></td>
			<td></td>
			<td>20 mM NaCl, 0.2 mM KCl, 0.8 mM NaHCO3, 0.2 mM CaCl2, 4 mM MgSO4, pH to 7.4</td>
		</tr>
	</tbody>
</table>

generation of chimeric axolotls with mutant haploid limbs through embryonic grafting

A growing set of genetic techniques and resources enable researchers to probe the molecular origins of the ability of some species of salamanders, such as axolotls, to regenerate entire limbs as adults. Here, we outline techniques used to generate chimeric axolotls with Cas9-mutagenized haploid forelimbs that can be used for exploring gene function and the fidelity of limb regeneration. We combine several embryological and genetic techniques, including haploid generation via in vitro activation, CRISPR/Cas9 mutagenesis, and tissue grafting into one protocol to produce a unique system for haploid genetic screening in a model organism of regeneration. This strategy reduces the number of animals, space, and time required for the functional analysis of genes in limb regeneration. This also permits the investigation of regeneration-specific functions of genes that may be required for other essential processes, such as organogenesis, tissue morphogenesis, and other essential embryonic processes. The method described here is a unique platform for conducting haploid genetic screening in a vertebrate model system.

Developing haploid embryos can be distinguished from diploid embryos by their &#39;haploid syndrome&#39; phenotype29. At graft-stage, haploid embryos exhibit reduced curvature along the anterior-posterior axis and incomplete enclosure of the yolk plug (Figure 3A). A fluorescent microscope can be used to ensure that haploid embryos are free of paternally derived GFP expression (Figure 3<str...

Watch this Scientific Journal Video about Generation of Chimeric Axolotls with Mutant Haploid Limbs Through Embryonic Grafting at JoVE.com

Generation of Chimeric Axolotls with Mutant Haploid Limbs Through Embryonic Grafting

This goal of this protocol is to produce chimeric axolotls with haploid forelimbs derived from Cas9-mutagenized donor tissue using embryonic tissue grafting techniques.

A growing set of genetic techniques and resources enable researchers to probe the molecular origins of the ability of some species of salamanders, such as ...

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7.1K Views.  Yale University. This goal of this protocol is to produce chimeric axolotls with haploid forelimbs derived from Cas9-mutagenized donor tissue using embryonic tissue grafting techniques.

Video: Generation of Chimeric Axolotls with Mutant Haploid Limbs Through Embryonic Grafting

Generation of Fluorescent Protein Fusions in Candida Species

Candida species, prevalent colonizers of the intestinal and genitourinary tracts, are the cause of the majority of invasive fungal infections in humans. Thus, molecular and genetic tools are needed to facilitate the study of their pathogenesis mechanisms. PCR-mediated gene modification is a straightforward and quick approach to generate epitope-tagged proteins to facilitate their detection. In particular, fluorescent protein (FP) fusions are powerful tools that allow visualization and quantitation of both yeast cells and proteins by fluorescence microscopy and immunoblotting, respectively. Plasmids containing FP encoding sequences, along with nutritional marker genes that facilitate the transformation of Candida species, have been generated for the purpose of FP construction and expression in Candida. Herein, we present a strategy for constructing a FP fusion in a Candida species. Plasmids containing the nourseothricin resistance transformation marker gene (NAT1) along with sequences for either green, yellow, or cherry FPs (GFP, YFP, mCherry) are used along with primers that include gene-specific sequences in a polymerase chain reaction (PCR) to generate a FP cassette. This gene-specific cassette has the ability to integrate into the 3'-end of the corresponding gene locus via homologous recombination. Successful in-frame fusion of the FP sequence into the gene locus of interest is verified genetically, followed by analysis of fusion protein expression by microscopy and/or immuno-detection methods. In addition, for the case of highly expressed proteins, successful fusions can be screened for primarily by fluorescence imaging techniques.

Generation of Fluorescent Protein Fusions in Candida Species

PCR-mediated gene modification can be used to generate fluorescent protein fusions in Candida species, which facilitates visualization and quantitation of yeast cells and proteins. Herein, we present a strategy for constructing a fluorescent protein fusion (Eno1-FP) in Candida parapsilosis.

Candida species, prevalent colonizers of the intestinal and genitourinary tracts, are the cause of the majority of invasive fungal infections in humans. ...

Controllable Ion Channel Expression through Inducible Transient Transfection

Transfection, the delivery of foreign nucleic acids into a cell, is a powerful tool in protein research. Through this method, ion channels can be investigated through electrophysiological analysis, biochemical characterization, mutational studies, and their effects on cellular processes. Transient transfections offer a simple protocol in which the protein becomes available for analysis within a few hours to days. Although this method presents a relatively straightforward and time efficient protocol, one of the critical components is calibrating the expression of the gene of interest to physiological relevant levels or levels that are suitable for analysis. To this end, many different approaches that offer the ability to control the expression of the gene of interest have emerged. Several stable cell transfection protocols provide a way to permanently introduce a gene of interest into the cellular genome under the regulation of a tetracycline-controlled transcriptional activation. While this technique produces reliable expression levels, each gene of interest requires a few weeks of skilled work including calibration of a killing curve, selection of cell colonies, and overall more resources. Here we present a protocol that uses transient transfection of the Transient Receptor Potential cation channel subfamily V member 1 (TRPV1) gene in an inducible system as an efficient way to express a protein in a controlled manner which is essential in ion channel analysis. We demonstrate that using this technique, we are able to perform calcium imaging, whole cell, and single channel analysis with controlled channel levels required for each type of data collection with a single transfection. Overall, this provides a replicable technique that can be used to study ion channels structure and function.

Studying ion channels through a heterologously expressing system has become a core technique in biomedical research. In this manuscript, we present a time efficient method to achieve tightly controlled ion channel expression by performing transient transfection under the control of an inducible promoter.

Transfection, the delivery of foreign nucleic acids into a cell, is a powerful tool in protein research. Through this method, ion channels can be ...

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 ...

Chromatin Immunoprecipitation (ChIP) Protocol for Low-abundance Embryonic Samples

Chromatin immunoprecipitation (ChIP) is a widely-used technique for mapping the localization of post-translationally modified histones, histone variants, transcription factors, or chromatin-modifying enzymes at a given locus or on a genome-wide scale. The combination of ChIP assays with next-generation sequencing (i.e., ChIP-Seq) is a powerful approach to globally uncover gene regulatory networks and to improve the functional annotation of genomes, especially of non-coding regulatory sequences. ChIP protocols normally require large amounts of cellular material, thus precluding the applicability of this method to investigating rare cell types or small tissue biopsies. In order to make the ChIP assay compatible with the amount of biological material that can typically be obtained in vivo during early vertebrate embryogenesis, we describe here a simplified ChIP protocol in which the number of steps required to complete the assay were reduced to minimize sample loss. This ChIP protocol has been successfully used to investigate different histone modifications in various embryonic chicken and adult mouse tissues using low to medium cell numbers (5 x 104 - 5 x 105 cells). Importantly, this protocol is compatible with ChIP-seq technology using standard library preparation methods, thus providing global epigenomic maps in highly relevant embryonic tissues.

Chromatin Immunoprecipitation ChIP Protocol for Low-abundance Embryonic Samples

Here, we describe a chromatin immunoprecipitation (ChIP) and ChIP-seq library preparation protocol to generate global epigenomic profiles from low-abundance chicken embryonic samples.

Chromatin immunoprecipitation (ChIP) is a widely-used technique for mapping the localization of post-translationally modified histones, histone variants, ...

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

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

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

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

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

Generation of Tumor Organoids from Genetically Engineered Mouse Models of Prostate Cancer

Methods based on homologous recombination to modify genes have significantly furthered biological research. Genetically engineered mouse models (GEMMs) are a rigorous method for studying mammalian development and disease. Our laboratory has developed several GEMMs of prostate cancer (PCa) that lack expression of one or multiple tumor suppressor genes using the site-specific Cre-loxP recombinase system and a prostate-specific promoter. In this article, we describe our method for necropsy of these PCa GEMMs, primarily focusing on dissection of mouse prostate tumors. New methods developed over the last decade have facilitated the culture of epithelial-derived cells to model organ systems in vitro in three dimensions. We also detail a 3D cell culture method to generate tumor organoids from mouse PCa GEMMs. Pre-clinical cancer research has been dominated by 2D cell culture and cell line-derived or patient-derived xenograft models. These methods lack tumor microenvironment, a limitation of using these techniques in pre-clinical studies. GEMMs are more physiologically-relevant for understanding tumorigenesis and cancer progression. Tumor organoid culture is an in vitro model system that recapitulates tumor architecture and cell lineage characteristics. In addition, 3D cell culture methods allow for growth of normal cells for comparison to tumor cell cultures, rarely possible using 2D cell culture techniques. In combination, use of GEMMs and 3D cell culture in pre-clinical studies has the potential to improve our understanding of cancer biology.&#160;

We show a method for necropsy and dissection of mouse prostate cancer models, focusing on prostate tumor dissection. A step-by-step protocol for generation of mouse prostate tumor organoids is also presented.

Methods based on homologous recombination to modify genes have significantly furthered biological research. Genetically engineered mouse models (GEMMs) ...

Generation of Defined Genomic Modifications Using CRISPR-CAS9 in Human Pluripotent Stem Cells

Human pluripotent stem cells offer a powerful system to study gene function and model specific mutations relevant to disease. The generation of precise heterozygous genetic modifications is challenging due to CRISPR-CAS9 mediated indel formation in the second allele. Here, we demonstrate a protocol to help overcome this difficulty by using two repair templates in which only one expresses the desired sequence change, while both templates contain silent mutations to prevent re-cutting and indel formation. This methodology is most advantageous for gene editing coding regions of DNA to generate isogenic control and mutant human stem cell lines for studying human disease and biology. In addition, optimization of transfection and screening methodologies have been performed to reduce labor and cost of a gene editing experiment. Overall, this protocol is widely applicable to many genome editing projects utilizing the human pluripotent stem cell model.

This protocol provides a method to facilitate the generation of defined heterozygous or homozygous nucleotide changes using CRISPR-CAS9 in human pluripotent stem cells.

Human pluripotent stem cells offer a powerful system to study gene function and model specific mutations relevant to disease. The generation of precise ...

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

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

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

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

Measuring Embryonic Viability and Brood Size in Caenorhabditis elegans

Caenorhabditis elegans is an excellent model organism for the study of meiosis, fertilization, and embryonic development. C. elegans exist as self-fertilizing hermaphrodites, which produce large broods of progeny-when males are present, they can produce even larger broods of cross progeny. Errors in meiosis, fertilization, and embryogenesis can be rapidly assessed as phenotypes of sterility, reduced fertility, or embryonic lethality. This article describes a method to determine embryonic viability and brood size in C. elegans. We demonstrate how to set up this assay by picking a worm onto an individual Modified Youngren's, Only Bacto-peptone (MYOB) plate, establish the appropriate timeframe to count viable progeny and non-viable embryos, and explain how to accurately count live worm specimens. This technique can be used to determine viability in self-fertilizing hermaphrodites as well as cross-fertilization by mating pairs. These relatively simple experiments are easily adoptable for new researchers, such as undergraduate students and first-year graduate students.

Measuring Embryonic Viability and Brood Size in Caenorhabditis elegans

Here, we present a general method to determine the embryonic viability and total number of embryos produced (brood) using the model organism C. elegans.

Caenorhabditis elegans is an excellent model organism for the study of meiosis, fertilization, and embryonic development. C. elegans exist as ...

Epigenetic Inheritance and ChIP in Caenorhabditis elegans

Analysis of Transgenerational Epigenetic Inheritance in C. elegans Using a Fluorescent Reporter and Chromatin Immunoprecipitation (ChIP)

Transgenerational epigenetic inheritance (TEI) allows the transmission of information through the germline without changing the genome sequence, through factors such as non-coding RNAs and chromatin modifications. The phenomenon of RNA interference (RNAi) inheritance in the nematode Caenorhabditis elegans is an effective model to investigate TEI that takes advantage of this model organism's short life cycle, self-propagation, and transparency. In RNAi inheritance, exposure of animals to RNAi leads to gene silencing and altered chromatin signatures at the target locus that persist for multiple generations in the absence of the initial trigger. This protocol describes the analysis of RNAi inheritance in C. elegans using a germline-expressed nuclear green fluorescent protein (GFP) reporter. Reporter silencing is initiated by feeding the animals bacteria expressing double-stranded RNA targeting GFP. At each generation, animals are passaged to maintain synchronized development, and reporter gene silencing is determined by microscopy. At select generations, populations are collected and processed for chromatin immunoprecipitation (ChIP)-quantitative polymerase chain reaction (qPCR) to measure histone modification enrichment at the GFP reporter locus. This protocol for studying RNAi inheritance can be easily modified and combined with other analyses to further investigate TEI factors in small RNA and chromatin pathways.

Author Spotlight: RNAi Inheritance and ChIP in C. elegans 

This protocol describes an RNA interference and ChIP assay to study the epigenetic inheritance of RNAi-induced silencing and associated chromatin modifications in C. elegans.

Transgenerational epigenetic inheritance (TEI) allows the transmission of information through the germline without changing the genome sequence, through ...