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 inverted microscope. Completely non-motile sperm should never be used, and the concentration should be adjusted to 80,000 motile cells/mL. Too much sperm in the activation step can also prevent normal egg development. Sperm pits, which appear as small, dark indentations on the surface of the egg, are a physical indication that a sperm cell has penetrated the egg and typically appear 20 to 60 min after applying sperm. Eggs with ten or more sperm pits may not activate properly.
Unfertilized eggs can be activated if collected within 15 min after being laid under water, placed in a dry petri dish, and dried thoroughly with paper towels prior to sperm application. Alternatively, as shown in the video, eggs may be squeezed from a hormone-injected female. We find that these &#34;dry&#34; eggs give more consistent results, though we regularly employ both methods.
Successful grafting requires the proper coupling of diploid embryos with appropriately staged haploid limb bud tissue. This protocol contains a number of measures to ensure stage-matching. As natural mating does not always result in egg laying, HCG injection to obtain haploid oocytes should not be performed until the naturally mated female begins to lay diploid eggs. After this injection, the induced female should be housed at reduced temperature (8 to 12 &#176;C), which will reduce the rate of egg laying. Housing the hormone-stimulated female at normal temperatures may result in most of the eggs being laid in a short period of time. The onset of egg laying in a chilled female indicates that the female is ready for anesthesia and oocyte extraction. Oocyte activation must occur within 15 min of oocyte extraction. Because activated haploid embryos will be several days behind the naturally produced diploid embryos in development, we recommend incubating diploid embryos at 12 &#176;C while increasing the relative rate of development of the haploids by incubating them at 18 &#176;C. Because there will be some variation in the interval between HCG-injection and egg laying of induced females, slight adjustments in the incubation temperatures of either haploids or diploids may be necessary so that the staging of host and donor embryos are paired at the time of grafting. Embryos should be chilled to 4 &#176;C for several hours prior to grafting, and this nearly arrests development. Differing the time of onset of chilling haploid and diploid embryos prior to grafting can enable stage-matching. While haploid embryos differ morphologically from diploids, both haploids and diploids reach stage 21 after the neural folds close and the embryos lie one their sides. Like diploids, stage 25 haploid embryos have prominent gill and pronephric bulges. The large head protrudes at an angle relative to the body in haploids, though this is greatly reduced relative to stage 25 diploid embryos, whose heads protrude from the body at nearly a right angle30.
Sterile technique is absolutely critical for successful tissue grafting. Sterile conditions should be maintained through the entirety of the experiment from haploid generation through the chimeras&#39; embryonic development. We recommend keeping all parent animals in clean, non-system water conditions during the egg laying period to minimize the amount of contaminating organic material at the start of the procedure. Consistent, daily removal of deceased or dying embryos through the entire process is important for maintaining the health of the other embryos. We recommend individually housing all haploid and chimera embryos to minimize cross contamination.
Embryonic microdissection and tissue grafting skills are acquired through practice. During the grafting process, it is important not to puncture or tear the limb bud itself. Tissues being removed from the host and donor embryos should extend beyond the limb bud, as pictured, to provide a buffering zone. If too small of an area of tissue is grafted, the haploid limbs will be infiltrated by GFP+ diploid skin and other tissues.
One of the limitations of this grafting technique is that the neural tissue and blood in the limbs are derived from the host body. In a few rare cases, we have been able to generate grafted haploid limbs which completely lack all signs of GFP+ expressing nerves. In these cases, we were able to replace enough of the neural tissue in the host animal with that of the donor. However, such extensive grafting is difficult, unreliable, and often produces developmental abnormalities outside of the limb.
Haploidy is embryonic lethal in the axolotl. Similarly, mutations in many genes potentially essential for limb development and regeneration are also early embryonic lethal. Our protocol bypasses the embryonic lethality of haploidy for the production of experimental limbs and could also be used in cases where mutation of a candidate regeneration gene produces an embryonic lethal phenotype. Our limb bud grafting technique provides an advantage for accomplishing this versus previously described methods of limb bud and contributing tissue grafting, as it is performed during early embryonic development and involves the transplantation of complete ectoderm and mesoderm layers1,2.
Haploid limb bud grafting has been previously described; however, in this earlier study, haploid limb buds were grafted ectopically31. Microscopic nuclear analysis of the ectopic limbs derived from these grafts revealed extensive contributions of diploid tissue. While these limbs developed abnormally, they were able to regenerate31. Rather than grafting ectopic limb buds, we replace the entire endogenous limb bud with equivalent haploid tissues. Through the use of fluorescent markers, we are able to visually identify haploid limbs that are free of diploid contributions, with the exception of nerve and blood cells, without sacrificing the haploid limb itself. We find that haploid limbs develop and regenerate normally. However, limbs in which GFP+ hosts contribute to tissues other than neural and blood-derived cells often display abnormalities. Thus, when investigating gene function in haploid limbs, those with excessive host contributions must be excluded from analysis before a genotype can be associated with any phenotype observed in mutant haploid limbs.
Recent innovations, like the assembly of the axolotl genome and recent CRISPR/Cas-based insertional methods15,16,32, have dramatically expanded the genetic malleability of the axolotl. Methods to restrict genetic manipulations temporally and spatially hold great promise, but their current applications are restricted by the resources needed to generate, house, and distribute animal lines. The protocol detailed here accelerates the process by which the consequences of genetic perturbations of genes may be investigated in limb development and regeneration. There has been little characterization of many of the genes found to be upregulated in regenerating limbs, and these genes may be involved in a variety of essential developmental and cellular processes. While we anticipate that many genes required for limb regeneration will also be required for limb development, this method may uncover whether any genes implicated in regeneration have loss-of-function phenotypes that are regeneration-specific. CRISPR/Cas mutagenesis in axolotls typically produces allelic mosaicism that includes retained wildtype alleles, and this mosaicism itself may be regarded as a quantifiable phenotype20. Using Next Generation Sequencing of amputated original and regenerated limbs, researchers may quantify whether haploid cells with mutations in a gene of interest are preferentially lost after regeneration. This approach may permit the investigation of regeneration phenotypes produced by perturbation of otherwise essential developmental genes.
Haploid loss-of-function genetic screens facilitate the investigation of the genetic origins of many biological processes without the need to establish biallelic mutant cell lines. By combining haplogenesis, CRISPR/Cas9 mutagenesis, and limb bud grafting, we provide a novel platform for a haploid genetic screen exploring limb development and regeneration in a living animal.

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 3B).
When grafts are clean, GFP expression should be limited primarily to the brachial plexus, the neural network derived from the host&#39;s spinal cord, as seen in Figure 2. Punctate GFP expression will also be present in single cells that appear to be sensory neurons and blood-derived host cells, which migrate into the developing limb. When RFP+ donors are used, haploid graft limbs will show universal expression of RFP (Figure 1B&#39;). Throughout development, non-mutagenized haploid limbs are significantly shorter than the opposing diploid forelimbs in size-matched chimeric animals (Figure 1B, Figure 1B&#39;, and Figure 5, n = 16 limb pairs, haploid mean = 0.522 cm, SD = &#177; 0.087 cm, diploid mean = 0.667 cm, SD = &#177; 0.069 m, paired T-test p-value &#60; 0.0001, mean ratio = 0.784, SD = &#177; 0.113). Non-mutagenized haploid limbs also fully regenerate (4/4 haploid and 4/4 diploid complete regeneration, Figure 6), though they show a slight delay in reaching the digital outgrowth stage compared to diploids (n = 4 haploid, n = 4 diploid; 23 days after amputation: haploids = 3 palette and 1 digital outgrowth stage limbs; diploids = 4 digital outgrowth stage limbs).
Successful grafting requires practice, and consistency will vary depending on the technician&#39;s micromanipulation skills and sterile technique. Failed and impure grafts produce a variety of phenotypes, as seen in Figure 7 and Figure 8. In our hands, approximately 38.7% (SD = &#177; 8.78%) of oocytes develop into normally developed haploid embryos and 11.1% (SD = &#177; 5.46%) of all haploid-diploid grafts produce viable animals with normally developed, cleanly grafted haploid limbs (Figure 9).
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/60156/60156fig1.jpg" /> 
Figure 1: Overview of protocol and an example of a chimeric axolotl. (A) Schematic outlining the relative timing of steps taken to obtain diploid host embryos, sperm, and eggs in order to generate haploids and chimeric embryos. (B) Composite bright image of a juvenile axolotl produced by embryonic limb bud grafting from an RFP+ haploid embryo to a GFP+ diploid host (B&#39;) Overlay of green and red fluorescent images of the same 8 cm juvenile axolotl. The haploid limb is grossly normal, but shorter than the opposing diploid limb. Scale bar = 1 mm. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/60156/60156fig1large.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/60156/60156fig2.jpg" /> 
Figure 2: Fluorescent image of a normally developed and cleanly grafted haploid limb. GFP- haploid limb grafted to a GFP+ diploid host shows GFP expression pattern that appears to be restricted to spinal nerves innervating the limb (yellow arrow) and individual sensory neurons and blood-derived cells (white arrows). The limb pictured belonged to a 4 cm juvenile. Scale bar = 1 mm. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/60156/60156fig2large.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/60156/60156fig3.jpg" /> 
Figure 3: Comparison of diploid and haploid embryos. (A) Light image of diploids (left) and haploids (right). Note the reduced AP-axis curvature and the protruding endoderm of the haploid embryos (white arrows). (B) Green fluorescent image of the same embryos. GFP- haploids were generated using eggs from a GFP- female and irradiated sperm from a GFP+. Diploids are GFP+. Embryos pictured are approximately stage 2530. Staging series. Scale bar = 1 mm. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/60156/60156fig3large.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/60156/60156fig4.jpg" /> 
Figure 4: Schematic outlining the extent of a haploid limb bud graft. (A) Lateral view of a stage 25 haploid embryo. The dotted lines show the approximate area that should be transplanted, relative to the gill bulge and limb bud. (B) Transverse schematic of stage 25. The dotted lines show the approximate area and tissue types that should be removed from the host and replaced with the corresponding haploid donor tissue. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/60156/60156fig4large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 5" class="xfigimg" src="/files/ftp_upload/60156/60156fig5.jpg" /> 
Figure 5: Comparison of haploid and diploid limb lengths. (A) Scatter plot of the lengths of haploid limbs (pink) and opposing diploid limbs (green) of 16 similarly sized chimeric animals (haploid mean = 0.522 cm, SD = &#177; 0.087 cm, diploid mean = 0.667 cm, SD = &#177; 0.069 cm). Limbs were measured from the base of the zeugopod to the junction of digits 2 and 3. (B) Scatter plot of the haploid limb length to diploid limb length ratios of the 16 animals (mean ratio = 0.784, SD = &#177; 0.113 cm). (C) Scatter plot of the body lengths (cm) of the 16 measured chimeras (mean animal length = 8.72 cm &#177; 0.456 cm). Animals were measured from the tip of the snout to the tail tip. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/60156/60156fig5large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 6" class="xfigimg" src="/files/ftp_upload/60156/60156fig6.jpg" /> 
Figure 6: Comparison of regeneration in haploid and diploid limbs. (A) Haploid limb pre-amputation. (B) The same haploid limb after regeneration. (C) Diploid limb pre-amputation. (D) The same diploid limb after regeneration. 4/4 haploid limbs and 4/4 diploid limbs regenerated with normal morphology. Black dotted lines indicate the amputation plane. Scale bars = 1 mm. Amputations were performed on size-matched juvenile animals (8.5 cm). <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/60156/60156fig6large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 7" class="xfigimg" src="/files/ftp_upload/60156/60156fig7.jpg" /> 
Figure 7: Examples of normal and abnormal development of haploid limbs. (A) A haploid limb with grossly normal morphology. (B-E) Examples of grafts that failed to support complete development of haploid. (B) Oligodactyly. (C) More severe oligodactyly. (D) No distinct digital structures are present. (E) No limb. The limbs pictured belong to similarly sized juveniles (8.0 to 10 cm). Scale bars = 1 mm. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/60156/60156fig7large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 8" class="xfigimg" src="/files/ftp_upload/60156/60156fig8.jpg" /> 
Figure 8: Examples of GFP+ cells in impure haploid limb grafts. (A,A&#39;) Morphologically normal haploid graft limb with GFP+ diploid cells in the skin and dermis (white dashed outline) revealed by fluorescent microscopy. (B,B&#39;) A grafted abnormal haploid limb with GFP+ diploid cells contributing extensively to the dermis and skin (white outline). (C,C&#39;) A grafted abnormal haploid limb in which the epidermis has been replaced with GFP+ diploid cells (white outline). The limbs pictured belong to similarly sized juveniles (8.0 to 10.0 cm). Scale bar = 1 mm. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/60156/60156fig8large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 9" class="xfigimg" src="/files/ftp_upload/60156/60156fig9.jpg" /> 
Figure 9: Success rates of haploid embryo and haploid limb generation. (A) Percent of viable haploid embryos generated using in vitro activation. Data were collected from five independent in vitro activation experiments. Green indicates the fraction of eggs that produced stage 25 haploid embryos that could be used for grafting (mean = 38.7%, SD = &#177; 8.78%). Grey indicates the fraction of eggs that did not show signs of cleavage, were non-viable, or were normal in development. (B) Percent of normally developed, cleanly grafted haploid limbs. Purple indicates the fraction of grafts that yielded clean, normally developed haploid limbs (mean = 11.1%, SD = &#177; 5.46%). Green indicates limbs that developed normally but had contaminating GFP+ host tissues (mean = 11.3%, SD = &#177; 8.64%). Red indicates graft limbs that did not develop normally (mean = 27.1%, SD = &#177; 30.3%). Blue indicates all grafted embryos and animals that did not survive to complete limb development (mean = 50.5%, SD = &#177; 31.2%). Loss of grafted animals was distributed across late embryonic and larval stages. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/60156/60156fig9large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

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 ...

generation-chimeric-axolotls-with-mutant-haploid-limbs-through

Nanyang Technological University

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JoVE Journal

Genetics

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

The Terroir Concept Interpreted through Grape Berry Metabolomics and Transcriptomics

Terroir refers to the combination of environmental factors that affect the characteristics of crops such as grapevine (Vitis vinifera) according to particular habitats and management practices. This article shows how certain terroir signatures can be detected in the berry metabolome and transcriptome of the grapevine cultivar Corvina using multivariate statistical analysis. The method first requires an appropriate sampling plan. In this case study, a specific clone of the Corvina cultivar was selected to minimize genetic differences, and samples were collected from seven vineyards representing three different macro-zones during three different growing seasons. An untargeted LC-MS metabolomics approach is recommended due to its high sensitivity, accompanied by efficient data processing using MZmine software and a metabolite identification strategy based on fragmentation tree analysis. Comprehensive transcriptome analysis can be achieved using microarrays containing probes covering ~99% of all predicted grapevine genes, allowing the simultaneous analysis of all differentially expressed genes in the context of different terroirs. Finally, multivariate data analysis based on projection methods can be used to overcome the strong vintage-specific effect, allowing the metabolomics and transcriptomics data to be integrated and analyzed in detail to identify informative correlations.

This article describes the application of untargeted metabolomics, transcriptomics and multivariate statistical analysis to grape berry transcripts and metabolites in order to gain insight into the terroir concept, i.e., the impact of the environment on berry quality traits.

Terroir refers to the combination of environmental factors that affect the characteristics of crops such as grapevine (Vitis vinifera) according to ...

Delivery of Nucleic Acids through Embryo Microinjection in the Worldwide Agricultural Pest Insect, Ceratitis capitata

The Mediterranean fruit fly (medfly) Ceratitis capitata (Wiedemann) (Diptera: Tephritidae) is a pest species with extremely high agricultural relevance. This is due to its reproductive behavior: females damage the external surface of fruits and vegetables when they lay eggs and the hatched larvae feed on their pulp. Wild C. capitata populations are traditionally controlled through insecticide spraying and/or eco-friendly approaches, the most successful being the Sterile Insect Technique (SIT). The SIT relies on mass-rearing, radiation-based sterilization and field release of males that retain their capacity to mate but are not able to generate fertile progeny. The advent and the subsequent rapid development of biotechnological tools, together with the availability of the medfly genome sequence, has greatly boosted our understanding of the biology of this species. This favored the proliferation of new strategies for genome manipulation, which can be applied to population control.
In this context, embryo microinjection plays a dual role in expanding the toolbox for medfly control. The ability to interfere with the function of genes that regulate key biological processes, indeed, expands our understanding of the molecular machinery underlying medfly invasiveness. Furthermore, the ability to achieve germ-line transformation facilitates the production of multiple transgenic strains that can be tested for future field applications in novel SIT settings. Indeed, genetic manipulation can be used to confer desirable traits that can, for example, be used to monitor sterile male performance in the field, or that can result in early life-stage lethality. Here we describe a method to microinject nucleic acids into medfly embryos to achieve these two main goals.

Delivery of Nucleic Acids through Embryo Microinjection in the Worldwide Agricultural Pest Insect, Ceratitis capitata

The Mediterranean fruit fly (medfly) Ceratitis capitata (Diptera: Tephritidae) is a worldwide pest of agriculture. A deeper understanding of its biology is key to control medfly populations and thus reduce economic impact. Embryo microinjection is a fundamental tool allowing both germ-line transformation and reverse genetics studies in this species.

The Mediterranean fruit fly (medfly) Ceratitis capitata (Wiedemann) (Diptera: Tephritidae) is a pest species with extremely high agricultural relevance. ...

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 ...

Targeted Next-generation Sequencing and Bioinformatics Pipeline to Evaluate Genetic Determinants of Constitutional Disease

Next-generation sequencing (NGS) is quickly revolutionizing how research into the genetic determinants of constitutional disease is performed. The technique is highly efficient with millions of sequencing reads being produced in a short time span and at relatively low cost. Specifically, targeted NGS is able to focus investigations to genomic regions of particular interest based on the disease of study. Not only does this further reduce costs and increase the speed of the process, but it lessens the computational burden that often accompanies NGS. Although targeted NGS is restricted to certain regions of the genome, preventing identification of potential novel loci of interest, it can be an excellent technique when faced with a phenotypically and genetically heterogeneous disease, for which there are previously known genetic associations. Because of the complex nature of the sequencing technique, it is important to closely adhere to protocols and methodologies in order to achieve sequencing reads of high coverage and quality. Further, once sequencing reads are obtained, a sophisticated bioinformatics workflow is utilized to accurately map reads to a reference genome, to call variants, and to ensure the variants pass quality metrics. Variants must also be annotated and curated based on their clinical significance, which can be standardized by applying the American College of Medical Genetics and Genomics Pathogenicity Guidelines. The methods presented herein will display the steps involved in generating and analyzing NGS data from a targeted sequencing panel, using the ONDRISeq neurodegenerative disease panel as a model, to identify variants that may be of clinical significance.

Targeted next-generation sequencing is a time- and cost-efficient approach that is becoming increasingly popular in both disease research and clinical diagnostics. The protocol described here presents the complex workflow required for sequencing and the bioinformatics process used to identify genetic variants that contribute to disease.

Next-generation sequencing (NGS) is quickly revolutionizing how research into the genetic determinants of constitutional disease is performed. The ...

Amplification of Near Full-length HIV-1 Proviruses for Next-Generation Sequencing

The Full-Length Individual Proviral Sequencing (FLIPS) assay is an efficient and high-throughput method designed to amplify and sequence single, near full-length (intact and defective), HIV-1 proviruses. FLIPS allows determination of the genetic composition of integrated HIV-1 within a cell population. Through identifying defects within HIV-1 proviral sequences that arise during reverse transcription, such as large internal deletions, deleterious stop codons/hypermutation, frameshift mutations, and mutations/deletions in cis acting elements required for virion maturation, FLIPS can identify integrated proviruses incapable of replication. The FLIPS assay can be utilized to identify HIV-1 proviruses that lack these defects and are&#160;therefore potentially replication-competent. The FLIPS protocol involves: lysis of HIV-1 infected cells, nested PCR of near full-length HIV-1 proviruses (using primers targeted to the HIV-1 5&#39; and 3&#39; LTR), DNA purification and quantification, library preparation for Next-generation Sequencing (NGS), NGS, de novo assembly of proviral contigs, and a simple process of elimination for identifying replication-competent proviruses. FLIPS provides advantages over traditional methods designed to sequence integrated HIV-1 proviruses, such as single-proviral sequencing. FLIPS amplifies and sequences near full-length proviruses enabling replication competency to be determined, and also uses fewer amplification primers, preventing the consequences of primer mismatches. FLIPS is a useful tool for understanding the genetic landscape of integrated HIV-1 proviruses, especially within the latent reservoir, however, its utilization can extend to any application in which the genetic composition of integrated HIV-1 is required.

Full-length individual proviral sequencing (FLIPS) provides an efficient and high-throughput method for the amplification and sequencing of single, near full-length (intact and defective) HIV-1 proviruses and allows for determination of their potential replication-competency. FLIPS overcomes limitations of previous assays designed to sequence the latent HIV-1 reservoir.

The Full-Length Individual Proviral Sequencing (FLIPS) assay is an efficient and high-throughput method designed to amplify and sequence single, near ...

Inducible, Cell Type-Specific Expression in Arabidopsis thaliana Through LhGR-Mediated Trans-Activation

Inducible, tissue-specific expression is an important and powerful tool to study the spatio-temporal dynamics of genetic perturbation. Combining the flexible and efficient GreenGate cloning system with the proven and benchmarked LhGR system (here termed GR-LhG4) for the inducible expression, we have generated a set of transgenic Arabidopsis lines that can drive the expression of an effector cassette in a range of specific cell types in the three main plant meristems. To this end, we chose the previously developed GR-LhG4 system based on a chimeric transcription factor and a cognate pOp-type promoter ensuring tight control over a wide range of expression levels. In addition, to visualize the expression domain where the synthetic transcription factor is active, an ER-localized mTurquoise2 fluorescent reporter under control of the pOp4 or pOp6 promoter is encoded in driver lines. Here, we describe the steps necessary to generate a driver or effector line and demonstrate how cell type specific expression can be induced and followed in the shoot apical meristem, the root apical meristem and the cambium of Arabidopsis. By using several or all driver lines, the context specific effect of expressing one or multiple factors (effectors) under control of the synthetic pOp promoter can be assessed rapidly, for example in F1 plants of a cross between one effector and multiple driver lines. This approach is exemplified by the ectopic expression of VND7, a NAC transcription factor capable of inducing ectopic secondary cell wall deposition in a cell autonomous manner.

Inducible, Cell Type-Specific Expression in Arabidopsis thaliana Through LhGR-Mediated Trans-Activation

Here, we describe the generation and application of a set of transgenic Arabidopsis thaliana lines enabling inducible, tissue-specific expression in the three main meristems, the shoot apical meristem, the root apical meristem, and the cambium.

Inducible, tissue-specific expression is an important and powerful tool to study the spatio-temporal dynamics of genetic perturbation. Combining the ...

Generation of Chimeric Axolotls with Mutant Haploid Limbs Through Embryonic Grafting

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Chapters in this video

The Terroir Concept Interpreted through Grape Berry Metabolomics and Transcriptomics

Delivery of Nucleic Acids through Embryo Microinjection in the Worldwide Agricultural Pest Insect, Ceratitis capitata

Generation of Fluorescent Protein Fusions in Candida Species

Controllable Ion Channel Expression through Inducible Transient Transfection

Generation of Native Chromatin Immunoprecipitation Sequencing Libraries for Nucleosome Density Analysis

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

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

Targeted Next-generation Sequencing and Bioinformatics Pipeline to Evaluate Genetic Determinants of Constitutional Disease

Amplification of Near Full-length HIV-1 Proviruses for Next-Generation Sequencing

Inducible, Cell Type-Specific Expression in Arabidopsis thaliana Through LhGR-Mediated Trans-Activation