The authors would like to thanks members of the Houston lab for critical reading of the manuscript. Research is supported by the National Institutes of Health (GM083999) awarded to D.W.H.

1. Surgical Removal of Ovary Tissue
<ol class="lalpha">
	<li>Prepare a fresh batch of oocyte culture medium (OCM; Materials), between 400-800 mL depending on the size of the experiment. Adjust the pH to 7.6-7.8, until the phenol red turns a dark red color (six or so small drops of 5N NaOH). Pour out a small amount to check the pH in a separate tube since the pH electrode may contaminate the media. OCM should be stored at 18&deg;C and used within a week.</li>
	<li>Select 3-5 females and place one in ~ 2L buffered anesthetic solution (Materials). We try to avoid using frogs that were fed on the day of or prior to surgery. While the frog is succumbing to the anesthetic, disinfect the surgical area with 70% EtOH and prepare the surgical instruments.</li>
	<li>Sterilize the surgical instruments by 20-second immersion in a hot bead sterilizer at 250&deg;C. The following instruments should be on hand: scalpel handle (#3) and blade (#10 or 11), several pairs of Dumont forceps, Bonn iris scissors (curved or straight), Halsey or Olsen-Hegar micro needle holder, and sutures. Place sterilized instruments on a sterile Petri dish or on the inside of the suture package to keep sterile. Grasp the needle in the needle holder in the desired orientation before beginning. We prefer Olsen-Hegar needle holders since the handle contains scissor blades, which can be used to trim sutures without switching instruments.</li>
	<li>After 10 minutes, check that a surgical plane of anesthesia has been achieved. Anesthetized frogs will stop moving and will not respond to a toe pinch or being turned over. Anesthesia should last 15 minutes, which is more than enough time to perform the operations. Don appropriate surgical attire. This can include clean gloves (non-latex), lab coats, and surgical masks, depending on personal preference and institutional animal care guidelines. </li>
	<li>Note on aseptic technique: Frog skin secretions contain antimicrobial peptides that generally reduce the incidence of contamination during surgery, although aseptic technique should be followed as much as possible. Surgical drapes can be used, but may not be required by institutional IACUCs and may not provide much benefit. Generally "tip technique" used, where the hands do touch the sterile tips of the instruments, suture material or incision site. Instruments can be resterilized (in the bead sterilizer) following incision of the skin, and a sterile area should be maintained on the benchtop for placing instruments (such as a Petri dish or suture package). Institutional requirements can differ significantly, so please follow your institutions' guidelines and receive appropriate instruction prior to performing surgeries. </li>
	<li>Remove the frog from anesthetic and place in dorsal recumbency on a damp surgical wipe. Note. The skin can be cleaned by rinsing in diluted povidine iodine (1:20) or chlorhexidine (0.75%). However, avoid commercial surgical scrubs, soaps and 70% isopropyl alcohol, as these will damage the delicate skin of amphibians.</li>
	<li>Using a scalpel or small iris scissors, make a small (1- 1.5 cm) incision in the skin in the lower part of the abdomen, lateral to the midline. Subsequent incisions on the same frog should be performed on alternate sides, well spaced from each other. Incisions can be made either parallel or perpendicular to the midline. </li>
	<li>Make the skin and abdominal muscle incisions in two stages. After cutting the skin, lift the muscle layer and surrounding white-colored fascia with forceps and make the incision in the raised muscle. Use scissors and cut straight down, try to make one swift cut to expose the body cavity leaving clean wound edges. If just the fascia is cut, it will retract and will make suturing more difficult. Lifting the muscle will help avoid inadvertently wounding any internal organs. Also, try to avoid cutting through any visible lymph hearts or blood vessels within the muscle layers. Extend the length with scissors of the incision so that ovary can be easily pulled through.</li>
	<li>Ovary should be clearly visible once the incision is made. To procure the ovary, grasp with forceps and externalize the ovary tissue. The desired amount of tissue is removed, and trimmed off at the level of the body wall. Do not stuff the tissue back into the coelomic cavity, as this could be a source of infection or other complications.</li>
	<li>Immediately observe a small piece of ovary under a dissecting microscope and defolliculate a few oocytes to test their quality. They should be firm and of uniform size and coloration. If they are acceptable, remove the desired amount of ovary tissue and suture the incision. Enough to fill a 90 mm Petri dish is usually sufficient for most experiments. If the oocytes are of questionable quality, suture the frog and repeat with a different female. </li>
	<li>Close the incision by suturing the muscle and fascia layer first. Insert the needle a few millimeters lateral to the incision, making sure to pass through the fascia layer as well. Sutures just through the muscle may tear the tissue and come loose. Use forceps to help insert the needle from below on the opposite side of the incision. Release and re-grasp the needle from outside the body and pull the suture through until several inches or so of the tail remains, taking care not to pull it all the way through.</li>
	<li>Close the wound with a simple interrupted pattern of suturing. We use a basic instrument tie to make surgeon's square knots (reef knots). Loop the long end of the suture around the needle holder, grasp the tail and pull it through the loop towards you and tighten. Loop and pull through again, this time pull the needle holder away from you. A third throw can be used for added security (in the same direction as the first). Trim the suture, leaving short tail and do another suture in the muscle a few millimeters away. Two sutures are sufficient for small sutures; three for larger ones. Repeat for the skin layer. Video demonstrations of basic suturing technique are available on YouTube -- search for "instrument tie"; there are quite a few to choose from, although the basic technique is the same.</li>
	<li>Rinse off the female with deionized water and place her in a recovery bucket filled with cool (18&deg;C) water. While in the lab, frogs are kept in tap water treated with Amquel to remove chloramines. Every few minutes, gently lift the frog from the water and look for gulping or eye-bulging movements, indicative of recovery from anesthesia. </li>
	<li>Note: Survival surgery is not strictly necessary to obtain ovary tissue, although the females will continue to produce good oocytes after a number of operations. Performing survival surgery will also cut down on the numbers of animals used. Be sure to follow your animal care unit's guidelines on vertebrate survival surgery methods, post-op care and monitoring and for number of allowed surgeries on an individual.</li>
</ol>
2. Culturing and Defolliculating Oocytes
<ol class="lalpha">
	<li>Right after the surgery is finished, subdivide the ovary into small pieces (~2 cm2) and store in fresh OCM, using 5-6 pieces per 90 mm dish. Individual lobes of ovary are cut open, flattened out and cut into square pieces. These are rinsed in clean OCM and placed in dishes for culture. Flattening and dividing up the ovary will extend its life in culture and make defolliculation easier.</li>
	<li>Move a lobe to a small dish of OCM and manually defolliculate 300-500 oocytes. Transfer oocytes using a sterile, fire-polished pipette. We have found that using the cotton-plugged variety is essential for minimizing contamination of the cultures. Store the oocytes at 18&deg;C in 8 mL OCM (in medium dishes (Falcon 1007)) in groups of 100-150. With practice, you should be able to defolliculate enough oocytes in 1-2 hours. Collagenased oocytes are not suitable for host transfer since they are not be fertilizable.</li>
	<li>Defolliculating method. We defolliculate using watchmakers' forceps (Dumont #4s or #5s, Fine Science Tools), following the basic methods described in Smith et al., (1991). Using one pair of forceps (in your dominant hand), grasp the theca connective tissue layer surrounding a fully-grown stage VI oocyte near where the follicle is attached to the ovary (the stalk). With the other pair of forceps, grasp the ovary adjacent to the first pair. Lightly tear open the follicle layer by pulling across the oocyte. Gently tease the oocyte out of the tissue. Successfully defolliculated oocytes are floppier than oocytes simply pulled off without removing the follicle layer and will be devoid of visible blood vessels. </li>
	<li>Note: It is essential to maintain very light pressure on the tips of the forceps, barely enough for the tips to meet. Gripping too tightly is a common among beginners and will result in the oocyte being pulled away with the follicle intact. Keep a good pair of forceps exclusively for defolliculating. The tips should meet precisely and should be slightly blunter than forceps for making explants from Xenopus embryos (Figure 1).</li>
	<li>We routinely test mature about 20 with progesterone and abort the experiment if the maturation rate is poor (&lt;50% after 6-7 hours at room temperature).</li>
	<li>At this point, the oocytes can be manipulated and cultured as desired, such as microinjection of oligo, morpholino or mRNA. Injection procedures vary per lab, so we will not describe those here. Oocytes can be maintained up to a week in OCM, but generally the health declines so it is best to use them for transfer within 96 hours of isolation. </li>
	<li>Keep in mind that only five-to-six groups (including a control) can be transferred to a single female, owing to limitations on vital dyes, so plan your experiments accordingly.</li>
</ol>
3. Oocyte Maturation and Stimulation of Prospective Host Females
<ol class="lalpha">
	<li>If performing a rescue of oligo depletion, inject your mRNA (usually 50-300 pg of RNA) ~24-48 hours after the oligo. The oligos will have degraded by this time and should not affect the injected RNA.</li>
	<li>On the evening prior to performing the transfer, add 16 &mu;L of progesterone working solution (1mM in EtOH) to oocytes in 8 mL OCM in medium dishes (final progesterone concentration is ~2 &mu;M). Incubate 10-12 hours at 18&deg;C to allow for oocyte maturation.</li>
	<li>Inject 3-5 females with 1000 units per frog of hCG to induce egg laying. Our experiments seem to work better if oocyte maturation and hCG injection are done around the same time, about 12 hours prior to implantation into the host (e.g., 10 PM -10 AM). Leave females at RT or 18&deg;C overnight.</li>
</ol>
4. Preparation and Vital Dye Staining of Oocytes
<ol class="lalpha">
	<li>On the morning of the transfer, allow ~ 45-60' to set up and perform the procedure. At this point, mature oocytes can be frozen for later analysis of the efficacy of mRNA knockdown or protein expression. Also, check that the oocytes have not become infected and that there was a good maturation rate (&gt;50%). Infected oocytes will not fertilize. Thaw vital dyes (Materials) and spin at max speed for 5 min. </li>
	<li>Color the different experimental groups by adding 80 &mu;L of the appropriate vital dye(s) to the oocyte dishes and incubate with rocking for 15'. During this time, select a host female and begin anesthesia. Choose a frog that is slowly laying eggs normally or one that produces abundant eggs with mild squeezing. Eggs from the host should be of good quality. Avoid females that are laying strings of eggs or crushed eggs. </li>
	<li>Transfer the colored eggs to a large dish of fresh OCM, swirl briefly to wash and set them aside prior to transfer. </li>
	<li>Prepare a sterile Pasteur pipette with a bore just wide enough to accommodate an oocyte (~ 1.5 mm) for use in transplanting the oocytes. Score the pipette with a diamond pencil and break cleanly. Fire polish until the edges are smooth, taking care not to melt the opening closed.</li>
</ol>
5. Performing the Oocyte Transplantation
<ol class="lalpha">
	<li>Oocyte transplantation uses the same surgical procedures as described in Part I, although instead of removing ovary, manipulated oocytes are introduced into the body cavity with a pipette. Ideally, the transfer procedure should be performed as rapidly as possible and the size of the incision should be kept as small as possible.</li>
	<li>Anesthesize the host female and prepare surgical instruments as described in Part I. Make and incision as described in Part I. The incision can be smaller, but needs to be large enough to fit the bore of the transfer pipette (Figure 2A).</li>
	<li>To transplant the oocytes, grasp and elevate one flap of muscle and fascia with forceps and introduce the experimental oocytes using the Pasteur pipette prepared above. Swirl the dish of oocytes to collect them into the center and suck up as many as possible. Allow the oocytes to settle into the end of the pipette to prevent excess liquid from being introduced and insert the tip of the pipette into the incision and slowly allow the oocytes to trickle into the body cavity (Figure 2B). Repeat until all oocytes have been transferred. Do not release the muscle layer at any time, until you have begun suturing, or the oocytes will spill out of the incision. </li>
	<li>Once the oocytes have been transferred, grasp both sides of the incision with forceps and bring them together, allowing the oocytes to settle into the body cavity. Begin suturing as described, taking care to keep upward tension on the muscle and fascia until the first knot is tied (Figure 2C). This will keep the oocytes from being expelled out and will allow the edges of the incision to heal evenly. Add another suture in the muscle/fascia layer and then suture the skin. </li>
	<li>Rinse off the female with deionized water and place her in a recovery bucket filled with cooled (18&deg;C) Amquel-treated water. Monitor her recovery from anesthesia as above. She should begin laying eggs normally after the operation.</li>
	<li>Obtain testes from a male frog using normal procedures. Store the testes in OCM or L15 until needed. We prefer fresh testis when fertilizing transfer experiments.</li>
</ol>
6. Recovery of Oocytes and in vitro Fertilization
<ol class="lalpha">
	<li>Peritoneal cilia will drive the transferred eggs (and normal coelomic eggs of the host) to the openings of the oviducts in the anterior of the body cavity. Colored eggs can be recovered beginning 2-3 hours after the transfer by normal squeezing. Eggs can be obtained every half-hour or so until the female stops laying. </li>
	<li>Fertilize in a sperm suspension (in ~ 4 mL 1x MMR) for 4 minutes. Flood and rinse extensively with 0.1x MMR. Eggs should activate normally and will begin to cleave ~ 1.5-2 hrs after fertilization. Transferred eggs will often cleave slightly later than the host eggs. </li>
	<li>Remove the jelly coats with cysteine as normal and sort embryos into separate dishes. This can be done at any stage, depending on the needs of the experiment. It usually easiest to identify the various colors around the 4-cell stage and most difficult during late blastula stages. Culture the eggs at a low density (&lt; 50 per medium dish) in clean 0.1x MMR. From this point, embryos can be treated and manipulated normally. </li>
	<li>Return the frogs to the animal facility and monitor for complications over the next few days.</li>
	<li>Alternate fertilization method; egg-laying in high salt buffer
	<ol class="lroman">
		<li>After the frog has recovered from anesthesia, place her into ~ 1L high-salt MMR (1.2x). Allow the female to lay eggs into the buffer for about 4-6 hours. Squeeze remaining eggs out, drain 1.2x MMR and sort colored eggs into clean dish; remove all possible liquid.</li>
		<li>Wash extensively with 0.3x MMR and fertilize in minimal volume of 0.3x MMR/sperm suspension until eggs activate (10 minutes). Flood with 0.1x MMR and culture as above. This option is useful if many eggs are needed at the same stage or if manual expression of eggs damages the donor oocytes.</li>
</ol>
</li>
</ol>

<ol>
	<li>Brun, R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Oocyte+maturation+in+vitro:+contribution+of+the+oviduct+to+total+maturation+in+Xenopus+laevis.">Oocyte maturation in vitro: contribution of the oviduct to total maturation in Xenopus laevis.</a> Experientia. 31, 1275-1276 (1975).</li><li>Elinson, R. P., Pasceri, P. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Two+UV-sensitive+targets+in+dorsoanterior+specification+of+frog+embryos.">Two UV-sensitive targets in dorsoanterior specification of frog embryos.</a> Development. 106, 511-518 (1989).</li><li>Heasman, J., Holwill, S., Wylie, C. C., C, C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Fertilization+of+cultured+Xenopus+oocytes+and+use+in+studies+of+maternally+inherited+molecules.">Fertilization of cultured Xenopus oocytes and use in studies of maternally inherited molecules.</a> Methods Cell Biol. 36, 213-230 (1991).</li><li>Holwill, S., Heasman, J., Crawley, C., Wylie, C. C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Axis+and+germ+line+deficiencies+caused+by+u.v+irradiation+of+Xenopus+oocytes+cultured+in+vitro.">Axis and germ line deficiencies caused by u.v irradiation of Xenopus oocytes cultured in vitro.</a> Development. 100, 735-743 (1987).</li><li>Houston, D. W., King, M. L. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+critical+role+for+Xdazl,+a+germ+plasm-localized+RNA,+in+the+differentiation+of+primordial+germ+cells+in+Xenopus.">A critical role for Xdazl, a germ plasm-localized RNA, in the differentiation of primordial germ cells in Xenopus.</a> Development. 127, 447-456 (2000).</li><li>Mir, A., Heasman, J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=How+the+mother+can+help:+studying+maternal+Wnt+signaling+by+anti-sense-mediated+depletion+of+maternal+mRNAs+and+the+host+transfer+technique.">How the mother can help: studying maternal Wnt signaling by anti-sense-mediated depletion of maternal mRNAs and the host transfer technique.</a> Methods Mol Biol. 469, 417-429 (2008).</li><li>Rugh, R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=.">.</a> Experimental Embryology: Techniques and procedures. , (1962).</li><li>Smith, L. 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A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+stability,+toxicity+and+effectiveness+of+unmodified+and+phosphorothioate+antisense+oligodeoxynucleotides+in+Xenopus+oocytes+and+embryos.">The stability, toxicity and effectiveness of unmodified and phosphorothioate antisense oligodeoxynucleotides in Xenopus oocytes and embryos.</a> Nucleic Acids Res. 18, 1763-1769 (1990).</li><li>Wylie, C., Kofron, M., Payne, C., Anderson, R., Hosobuchi, M., Joseph, E., Heasman, J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Maternal+beta-catenin+establishes+a+'dorsal+signal'+in+early+Xenopus+embryos.">Maternal beta-catenin establishes a 'dorsal signal' in early Xenopus embryos.</a> Development. 122, 2987-2996 (1996).</li><li>Zhang, J., Houston, D. W., King, M. 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Experiments on animals were performed in accordance with the guidelines and regulations set forth by The University of Iowa IACUC Committee.

A successful transfer will result in fertilization and normal cleavage of &gt; 50% of the oocytes (Figure 3). Generally between 30-60% of the transferred oocytes will survive past the neurula stages. We usually transfer 75-150 oocytes per experimental group, which will yield enough embryos for several types of analysis (in situ, RT-PCR) as well as visualizing the phenotype. Implanting substantially more oocytes does not seem to greatly increase the yield. Also, at least 30 oocytes per group should be transferred to guara...

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<html xmlns="http://www.w3.org/1999/xhtml" xml:lang="en" lang="en">	
			<div>
			Solutions and recipes:
OCM (Oocyte culture medium; modified from Wylie et al., 1996)
<table border="0">
 <tbody>
 <tr>
 <td>70% Leibovitz's L-15 medium (containing l-glutamine)</td>
 <td>(Invitrogen 11415-064)</td>
 </tr>
 <tr>
 <td>0.4 mg/ml Bovine serum albumin (BSA; fraction V)</td>
 <td>(Sigma A-9418-50g)</td>
 </tr>
 <tr>
 <td>1x Penicillin-Streptomycin</td>
 <td>(Sigma P-0781)</td>
 </tr>
 <tr>
 <td colspan="2">Adjust to pH 7.6-7.8 with 5N NaOH</td>
 </tr>
 <tr>
 <td colspan="2">Make fresh weekly, store at 18&deg;C.</td>
 </tr>
 </tbody>
</table>
10X MMR (Marc's Modified Ringer's Solution; Zuck et al., 1998)
<table border="0">
 <tbody>
 <tr>
 <td>1M NaCl</td>
 </tr>
 <tr>
 <td>20 mM KCl</td>
 </tr>
 <tr>
 <td>20 mM CaCl2</td>
 </tr>
 <tr>
 <td>10 mM MgCl2</td>
 </tr>
 <tr>
 <td>150 mM HEPES</td>
 </tr>
 <tr>
 <td>adjust to pH 7.6-7.8</td>
 </tr>
 <tr>
 <td>Store at 4&deg;C</td>
 </tr>
 </tbody>
</table>
Anesthetic solution
<table border="0">
 <tbody>
 <tr>
 <td>1 g/L 3-Aminobenzoic Acid Ethyl Ester (MS222)</td>
 <td>(Sigma A-5040)</td>
 </tr>
 <tr>
 <td colspan="2">0.7 g/L sodium bicarbonate</td>
 </tr>
 <tr>
 <td colspan="2">Dissolve in Amquel-treated water</td>
 </tr>
 <tr>
 <td colspan="2">Make fresh weekly</td>
 </tr>
 </tbody>
</table>
Progesterone stocks
<table border="0">
 <tbody>
 <tr>
 <td>10 mM Progesterone (Sigma P- 0130) in 100% Ethanol</td>
 </tr>
 <tr>
 <td>Dilute to 1 mM in 100% EtOH for a 500x stock/working solution</td>
 </tr>
 <tr>
 <td>Store at -20&deg;C</td>
 </tr>
 </tbody>
</table>
Vital dyes (modified from Heasman et al., 1991)
<table border="0">
 <tbody>
 <tr>
 <td>Blue: 0.1% Nile Blue A</td>
 <td>(Aldrich 121479-5G),</td>
 </tr>
 <tr>
 <td>Red: 0.25% Neutral Red</td>
 <td>(Sigma N-6634),</td>
 </tr>
 <tr>
 <td>Brown: 1% Bismarck Brown</td>
 <td>(Sigma B-2759),</td>
 </tr>
 <tr>
 <td colspan="2">Green (80 &mu;l blue + 80 &mu;l brown),</td>
 </tr>
 <tr>
 <td colspan="2">Mauve (80 &mu;l blue + 80 &mu;l red).</td>
 </tr>
 <tr>
 <td colspan="2">A sixth color, Orange (80 &mu;l red+ 80 &mu;l brown), can also be used but is often hard to distinguish from brown.</td>
 </tr>
 </tbody>
</table>
Stocks of Blue, Red and Brown are made up in deionized water in 50 ml tubes, incubated for 20 minutes with rocking and spun in a clinical centrifuge. The supernatants are aliquotted into microfuge tubes and stored at -20&deg;C.		</div>

fertilization of xenopus oocytes using the host transfer method

Studying the contribution of maternally inherited molecules to vertebrate early development is often hampered by the time and expense necessary to generate maternal-effect mutant animals. Additionally, many of the techniques to overexpress or inhibit gene function in organisms such as Xenopus and zebrafish fail to sufficiently target critical maternal signaling pathways, such as Wnt signaling. In Xenopus, manipulating gene function in cultured oocytes and subsequently fertilizing them can ameliorate these problems to some extent. Oocytes are manually defolliculated from donor ovary tissue, injected or treated in culture as desired, and then stimulated with progesterone to induce maturation. Next, the oocytes are introduced into the body cavity of an ovulating host female frog, whereupon they will be translocated through the host's oviduct and acquire modifications and jelly coats necessary for fertilization. The resulting embryos can then be raised to the desired stage and analyzed for the effects of any experimental perturbations. This host-transfer method has been highly effective in uncovering basic mechanisms of early development and allows a wide range of experimental possibilities not available in any other vertebrate model organism.

Watch this Scientific Journal Video about Fertilization of Xenopus oocytes using the Host Transfer Method at JoVE.com

Fertilization of Xenopus oocytes using the Host Transfer Method

Procedure for fertilizing Xenopus oocytes by the host transfer method.

Fertilization of Xenopus oocytes using the Host Transfer Method

Studying the contribution of maternally inherited molecules to vertebrate early development is often hampered by the time and expense necessary to ...

fertilization-of-xenopus-oocytes-using-the-host-transfer-method

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14.5K Views.  University of Iowa. Procedure for fertilizing Xenopus oocytes by the host transfer method.

Video: Fertilization of Xenopus oocytes using the Host Transfer Method

Nuclear Transfer into Mouse Oocytes

Nuclear transfer into an unfertilized oocyte can restore developmental potential to a differentiated cell. This demonstrates that the processes underlying development, differentiation and aging are epigenetic rather than genetic processes. The reversibility of these processes opens exciting perspectives in basic research, and in the more distant future, in regenerative medicine. In the mouse, embryonic stem cells can be derived from cloned preimplantation stage embryos. Such embryonic stem cells have the ability to give rise to all cell types of the adult organism. Importantly, these cells are genetically identical to the donor. If applicable to human, this would allow the derivation of stem cells from a patient. These cells could then be differentiated into the affected cell type of the patient and studied in vitro, or used to replace the damaged or missing cells. The study of nuclear transfer in the mouse remains important as it can inform us about the principles of nuclear reprogramming. This movie and the accompanying protocol are intended to help learning nuclear transfer in the mouse, a method initially developed in the group of Prof. Yanagimachi (WAKAYAMA et al. 1998).

This movie and the protocol are intended to help learning nuclear transfer.

Nuclear transfer into an unfertilized oocyte can restore developmental potential to a differentiated cell. This demonstrates that the processes underlying ...

Retrieval of Mouse Oocytes

To date, only a few studies have reported successful manipulations of Peromyscus embryogenesis or reproductive biology.  Together with the Peromyscus Genetic Stock Center (http://stkctr.biol.sc.edu), we are characterizing the salient differences needed to develop this system.  A primary goal has been to optimize oocyte/early embryo retrieval.

This protocol illustrates the technique for extracting oocytes or early-stage fertilized embryos from the oviduct of mice. The ability to identify the infindibulum and insert a blunt end needle into it is essential to correctly performing the procedure.

To date, only a few studies have reported successful manipulations of Peromyscus embryogenesis or reproductive biology.  Together with the Peromyscus ...

In Vitro Nuclear Assembly Using Fractionated Xenopus Egg Extracts

Nuclear membrane assembly is an essential step in the cell division cycle; this process can be replicated in the test tube by combining Xenopus sperm chromatin, cytosol, and light membrane fractions. Complete nuclei are formed, including nuclear membranes with pore complexes, and these reconstituted nuclei are capable of normal nuclear processes.

Nuclear membrane assembly is an essential step in the cell division cycle; this process can be replicated in the test tube by combining Xenopus sperm chromatin, cytosol, and light membrane fractions. Complete nuclei are formed, including nuclear membranes with pore complexes, and these reconstituted nuclei are capable of normal nuclear processes.

Nuclear membrane assembly is an essential step in the cell division cycle; this process can be replicated in the test tube by combining Xenopus sperm ...

Patch Clamp Recording of Ion Channels Expressed in  Xenopus Oocytes

Since its development by Sakmann and Neher 1, 2, the patch clamp has become established as an extremely useful technique for electrophysiological measurement of single or multiple ion channels in cells. This technique can be applied to ion channels in both their native environment and expressed in heterologous cells, such as oocytes harvested from the African clawed frog, Xenopus laevis. Here, we describe the well-established technique of patch clamp recording from Xenopus oocytes. This technique is used to measure the properties of expressed ion channels either in populations (macropatch) or individually (single-channel recording). We focus on techniques to maximize the quality of oocyte preparation and seal generation. With all factors optimized, this technique gives a probability of successful seal generation over 90 percent. The process may be optimized differently by every researcher based on the factors he or she finds most important, and we present the approach that have lead to the greatest success in our hands.

This is intended as an introduction to patch clamp recording from Xenopus laevis oocytes. It covers vitelline membrane removal, formation of a gigaohm seal (gigaseal), and the optional conversion of the patch to the outside-out topology.

Since its development by Sakmann and Neher 1, 2, the patch clamp has become established as an extremely useful technique for electrophysiological ...

Microinjection of Xenopus Laevis Oocytes

Microinjection of Xenopus laevis oocytes followed by thin-sectioning electron microscopy (EM) is an excellent system for studying nucleocytoplasmic transport. Because of its large nucleus and high density of nuclear pore complexes (NPCs), nuclear transport can be easily visualized in the Xenopus oocyte. Much insight into the mechanisms of nuclear import and export has been gained through use of this system (reviewed by Pant&eacute;, 2006). In addition, we have used microinjection of Xenopus oocytes to dissect the nuclear import pathways of several viruses that replicate in the host nucleus. 
Here we demonstrate the cytoplasmic microinjection of Xenopus oocytes with a nuclear import substrate. We also show preparation of the injected oocytes for visualization by thin-sectioning EM, including dissection, dehydration, and embedding of the oocytes into an epoxy embedding resin. Finally, we provide representative results for oocytes that have been microinjected with the capsid of the baculovirus Autographa californica nucleopolyhedrovirus (AcMNPV) or the parvovirus Minute Virus of Mice (MVM), and discuss potential applications of the technique.

Here we demonstrate cytoplasmic microinjection of Xenopus laevis oocytes with a nuclear import substrate, as well as preparation of the injected oocytes for visualization by thin-sectioning electron microscopy.

Microinjection of Xenopus laevis oocytes followed by thin-sectioning electron microscopy (EM) is an excellent system for studying nucleocytoplasmic ...

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

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

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

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

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

Visualizing RNA Localization in Xenopus Oocytes

RNA localization is a conserved mechanism of establishing cell polarity. Vg1 mRNA localizes to the vegetal pole of Xenopus laevis oocytes and acts to spatially restrict gene expression of Vg1 protein. Tight control of Vg1 distribution in this manner is required for proper germ layer specification in the developing embryo. RNA sequence elements in the 3' UTR of the mRNA, the Vg1 localization element (VLE) are required and sufficient to direct transport. To study the recognition and transport of Vg1 mRNA in vivo, we have developed an imaging technique that allows extensive analysis of trans-factor directed transport mechanisms via a simple visual readout. 
To visualize RNA localization, we synthesize fluorescently labeled VLE RNA and microinject this transcript into individual oocytes. After oocyte culture to allow transport of the injected RNA, oocytes are fixed and dehydrated prior to imaging by confocal microscopy. Visualization of mRNA localization patterns provides a readout for monitoring the complete pathway of RNA transport and for identifying roles in directing RNA transport for cis-acting elements within the transcript and trans-acting factors that bind to the VLE (Lewis et al., 2008, Messitt et al., 2008). We have extended this technique through co-localization with additional RNAs and proteins (Gagnon and Mowry, 2009, Messitt et al., 2008), and in combination with disruption of motor proteins and the cytoskeleton (Messitt et al., 2008) to probe mechanisms underlying mRNA localization.

Visualizing RNA Localization in Xenopus Oocytes

Visualization of in vivo RNA transport is accomplished by microinjection of fluorescently labeled RNA transcripts into Xenopus oocytes, followed by confocal microscopy.

RNA localization is a conserved mechanism of establishing cell polarity.  Vg1 mRNA localizes to the vegetal pole of Xenopus laevis oocytes and acts to ...

Tissue Determination Using the Animal Cap Transplant (ACT) Assay in Xenopus laevis

Many proteins play a dual role in embryonic development. Those that regulate cell fate determination in a specific tissue can also affect the development of a larger region of the embryo. This makes defining its role in a particular tissue difficult to analyze. For example, noggin overexpression in Xenopus laevis embryos causes the expansion of the entire anterior region, including the eye1,2. From this result, it is not known if Noggin plays a direct role in eye determination or that by causing an expansion of neural tissue, Noggin indirectly affects eye formation. Having this complex phenotype makes studying its eye-specific role in cell fate determination difficult to analyze. We have developed an assay that overcomes this problem. Taking advantage of the pluripotent nature of the Xenopus laevis animal cap 3, we have developed an assay to test the ability of gene product(s), like noggin or the eye field transcription factors (EFTFs), to transform caps into particular tissue or cell types by transplanting this tissue onto the side of the embryo 4. While we have found either Noggin protein treatment or a collection of transcription factors can determine retinal cell fate in animal caps, this procedure could be used to identify gene product(s) involved in specifying other tissues as well.

Tissue Determination Using the Animal Cap Transplant ACT Assay in Xenopus laevis

Animal caps overexpressing gene product(s) are transplanted to the flank of developing Xenopus laevis embryos in order to establish whether tissue is determined.

Many proteins play a dual role in embryonic development. Those that regulate cell fate determination in a specific tissue can also affect the development ...

Patch Clamp and Perfusion Techniques for Studying Ion Channels Expressed in Xenopus oocytes

The protocol presented here is designed to study the activation of the large conductance, voltage- and Ca2+-activated K+ (BK) channels. The protocol may also be used to study the structure-function relationship for other ion channels and neurotransmitter receptors1. BK channels are widely expressed in different tissues and have been implicated in many physiological functions, including regulation of smooth muscle contraction, frequency tuning of inner hair cells and regulation of neurotransmitter release2-6. BK channels are activated by membrane depolarization and by intracellular Ca2+ and Mg2+ 6-9. Therefore, the protocol is designed to control both the membrane voltage and the intracellular solution. In this protocol, messenger RNA of BK channels is injected into Xenopus laevis oocytes (stage V-VI) followed by 2-5 days of incubation at 18&deg;C10-13. Membrane patches that contain single or multiple BK channels are excised with the inside-out configuration using patch clamp techniques10-13. The intracellular side of the patch is perfused with desired solutions during recording so that the channel activation under different conditions can be examined. To summarize, the mRNA of BK channels is injected into Xenopus laevis oocytes to express channel proteins on the oocyte membrane; patch clamp techniques are used to record currents flowing through the channels under controlled voltage and intracellular solutions.

Patch Clamp and Perfusion Techniques for Studying Ion Channels Expressed in Xenopus oocytes

Ionic current of BK channels is recorded using patch clamp techniques. BK channels are expressed in Xenopus oocytes by injecting messenger RNA. The intracellular solution during patch clamp recordings is controlled by a perfusion system.

The protocol presented here is designed to study the activation of the large conductance, voltage- and Ca2+-activated K+ (BK) channels. The protocol may ...

Study of the DNA Damage Checkpoint using Xenopus Egg Extracts

On a daily basis, cells are subjected to a variety of endogenous and environmental insults. To combat these insults, cells have evolved DNA damage checkpoint signaling as a surveillance mechanism to sense DNA damage and direct cellular responses to DNA damage. There are several groups of proteins called sensors, transducers and effectors involved in DNA damage checkpoint signaling (Figure 1). In this complex signaling pathway, ATR (ATM and Rad3-related) is one of the major kinases that can respond to DNA damage and replication stress. Activated ATR can phosphorylate its downstream substrates such as Chk1 (Checkpoint kinase 1). Consequently, phosphorylated and activated Chk1 leads to many downstream effects in the DNA damage checkpoint including cell cycle arrest, transcription activation, DNA damage repair, and apoptosis or senescence (Figure 1). When DNA is damaged, failing to activate the DNA damage checkpoint results in unrepaired damage and, subsequently, genomic instability. The study of the DNA damage checkpoint will elucidate how cells maintain genomic integrity and provide a better understanding of how human diseases, such as cancer, develop.
Xenopus laevis egg extracts are emerging as a powerful cell-free extract model system in DNA damage checkpoint research. Low-speed extract (LSE) was initially described by the Masui group1. The addition of demembranated sperm chromatin to LSE results in nuclei formation where DNA is replicated in a semiconservative fashion once per cell cycle.
The ATR/Chk1-mediated checkpoint signaling pathway is triggered by DNA damage or replication stress 2. Two methods are currently used to induce the DNA damage checkpoint: DNA damaging approaches and DNA damage-mimicking structures 3. DNA damage can be induced by ultraviolet (UV) irradiation, &gamma;-irradiation, methyl methanesulfonate (MMS), mitomycin C (MMC), 4-nitroquinoline-1-oxide (4-NQO), or aphidicolin3, 4. MMS is an alkylating agent that inhibits DNA replication and activates the ATR/Chk1-mediated DNA damage checkpoint 4-7. UV irradiation also triggers the ATR/Chk1-dependent DNA damage checkpoint 8. The DNA damage-mimicking structure AT70 is an annealed complex of two oligonucleotides poly-(dA)70 and poly-(dT)70. The AT70 system was developed in Bill Dunphy's laboratory and is widely used to induce ATR/Chk1 checkpoint signaling 9-12.
Here, we describe protocols (1) to prepare cell-free egg extracts (LSE), (2) to treat Xenopus sperm chromatin with two different DNA damaging approaches (MMS and UV), (3) to prepare the DNA damage-mimicking structure AT70, and (4) to trigger the ATR/Chk1-mediated DNA damage checkpoint in LSE with damaged sperm chromatin or a DNA damage-mimicking structure.

Study of the DNA Damage Checkpoint using Xenopus Egg Extracts

Xenopus egg extract is a useful model system to investigate the DNA damage checkpoint. This protocol is for the preparation of Xenopus egg extracts and DNA damage checkpoint inducing reagents. These techniques are adaptable to a variety of DNA damaging approaches in the study of the DNA damage checkpoint signaling.

Fertilization of Xenopus oocytes using the Host Transfer Method

Summary

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Chapters in this video

Nuclear Transfer into Mouse Oocytes

Retrieval of Mouse Oocytes

In Vitro Nuclear Assembly Using Fractionated Xenopus Egg Extracts

Patch Clamp Recording of Ion Channels Expressed in Xenopus Oocytes

Microinjection of Xenopus Laevis Oocytes

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

Visualizing RNA Localization in Xenopus Oocytes

Tissue Determination Using the Animal Cap Transplant (ACT) Assay in Xenopus laevis

Patch Clamp and Perfusion Techniques for Studying Ion Channels Expressed in Xenopus oocytes

Study of the DNA Damage Checkpoint using Xenopus Egg Extracts