This work was supported by funds from the Ministry of Economy and Competitiveness of Spain (AGL2017-85162-C2-1-R) and Generalitat Valenciana Research Programme (PrometeoII 2014/036). English text version revised by N. Macowan English Language Service

All experimental procedures used in this study were performed in accordance with Directive 2010/63/EU EEC for animal experiments and reviewed and approved by the Ethical Committee for Experimentation with Animals of the Universitat Polit&#232;cnica de Val&#232;ncia, Spain (research code: 2015/VSC/PEA/00170). XGD, FMJ, MPVC and JSV holds an authorization certificate issued by the Valencian governmental administration to experiment on animals. XGD is authorized to in situ supervise the welfare and care of the animals during the experiment.
1. Embryo Transfer
<ol>
	<li>Preparation of recipient females
	<ol>
		<li>Use only sexually mature females (&#62; 4.5 months old).</li>
		<li>One week before ET, adapt females to a 16 h light/8 h dark regime to initiate follicular growth and enhanced female receptivity.</li>
		<li>Select the recipient females, observing the turgidity and color of the vulva. If the vulva is turgid and reddish, the female is receptive.</li>
		<li>Induce pseudopregnancy (ovulation) by a single intramuscular injection of 1 &#181;g of buserelin acetate (synthetic analogue of Gonadotropin-releasing hormone) regardless of body weight. 
		NOTE: Normally, 0.8 &#181;g is a suitable dose for ovulation induction in medium-size rabbits (4-5 kg), so 1 &#181;g generally guarantees the ovulation.</li>
		<li>Induce ovulation as many days beforehand as the age of the embryos to be transferred (for example, 70-72 h before fresh morula ET).</li>
	</ol>
	</li>
	<li>Anesthesia and analgesia
	<ol>
		<li>Weigh the rabbit and load the following anesthetics and analgesics.
		<ol>
			<li>In a 1 mL syringe with a 30G needle: load xylazine (5mg/kg) and buprenorphine hydrochloride (0.03 mg/kg). In another 1 mL syringe with a 23G pericranial needle, load ketamine hydrochloride (35 mg/kg).</li>
		</ol>
		</li>
		<li>Hold the rabbit and inject the xylazine-buprenorphine mixture intramuscularly.</li>
		<li>Insert the pericranial needle with ketamine in the marginal ear vein, slowly introducing all the syringe contents intravenously.</li>
		<li>Fix the needle and leave it inserted throughout the remaining steps to administer more anesthesia if necessary.</li>
		<li>Leave the rabbit in the cage (clean and without any other animals) on a warm stage.</li>
		<li>Once unconscious, apply eye ointment to avoid dryness of the eye and check for the absence of the palpebral freflex. 
		NOTE: This protocol provides a surgical anesthesia plane for a minimum of 30 min. If a longer time is required, inject additional dosages with half of the amount of ketamine hydrochloride described in 1.2.1 after 30 min.</li>
		<li>Monitor the depth of anesthesia by checking the pedal reflex and breathing movement. Changes in the breathing pattern to an irregular and faster rate indicate loss of the proper plane of anesthesia.</li>
		<li>Monitor the color of the mucous membranes (eyes, lips, etc.), respiratory rate (30-60 breaths per minute), heart rate (120-325 beats per minute) and rectal temperature (38-39.6 &#176;C).</li>
		<li>Eight hours before transfer, withhold food from animals to avoid the greater gut size and activity until the ET process is finished. Leave free access to water.</li>
	</ol>
	</li>
	<li>Embryo preparation
	<ol>
		<li>Warm the embryo manipulation media to 25 &#176;C: Base Medium (BM), consisting of Dulbecco&#39;s Phosphate-Buffered Saline (DPBS) supplemented with 0.2% (w/v) of Bovine Serum Albumin.</li>
		<li>Working under a stereomicroscope, rinse fresh or thawed (Step 2) embryos with BM.</li>
		<li>Using sterile gloves, attach an appropriately configured 17G epidural catheter to a 1 mL syringe.</li>
		<li>Aspirate 1 cm of BM into the catheter, followed by a small air bubble.</li>
		<li>Aspirate 5-7embryos in a volume of 10 &#181;L of BM, followed by another small air bubble.</li>
		<li>Finish loading the catheter by aspirating 1 cm of BM.</li>
	</ol>
	</li>
	<li>Embryo transfer
	<ol>
		<li>Consider the use of sterile gloves, gown and mask to ensure an aseptic environment.</li>
		<li>Sterilize surgical instruments, clean the surfaces where surgery will be performed, and wipe them with 70% ethanol.</li>
		<li>Perform anesthesia as previously detailed (step 1.2), checking for loss of reflexes.</li>
		<li>Shave the fur from the ventral abdomen with an electric razor.</li>
		<li>Prepare the ventral abdomen aseptically.
		<ol>
			<li>Clean the surgical area and remove any remaining hair. Wash the surgical area with a chlorhexidine gluconate soap. Sanitize the area with chlorhexidine solution and ethanol 96&#186; (3 times).</li>
		</ol>
		</li>
		<li>Place the animal on a warm surgical table, in Trendelenburg&#39;s position (head down at 45&#176;) to ensure that the stomach and intestines are cranially located. Consider the evacuation of the bladder if it is turgid. If any viscera are damaged in the process, the animal may die. It is therefore important to have them properly located (Figure 1).</li>
		<li>Cover the area using a sterile towel, with a hole (fenestration) exposing the shaved area, to separate the surgical site from any potential contaminating areas.</li>
		<li>Insert one endoscopic trocar 5 cm into the abdominal cavity, 2 cm caudal to the xiphoid process, and insufflate through it the peritoneal cavity with a pressure-regulating mechanical insufflator. 
		NOTE: The intra-abdominal pressure should be 8-12 mmHg with CO2 (Figure 1A).</li>
		<li>Insert the endoscope camera through the endoscopic trocar (Figure 1B). 
		NOTE: Identify the reproductive tract, determining the status and position of the infundibulum and ampulla before ET to facilitate the next steps.</li>
		<li>Insert the 17-G epidural needle into the inguinal region between 2-3 cm from the infundibulum (Figure 1B).</li>
		<li>Identify the entrance of the infundibulum (Figure 2A, 2B).</li>
		<li>Insert the loaded catheter (step 1.3) through the epidural needle into the abdomen (Figure 1C).</li>
		<li>Locate the oviduct and insert 1-2 cm of the epidural catheter through the infundibulum in the ampulla (Figure 2A-2C). Do not progress very far into the oviduct to prevent damage and hemorrhage.</li>
		<li>Release the embryos into the oviduct by gently pressing the plunger of the syringe coupled to the catheter (Figure 2D-2F). Both air bubbles must exit the catheter.</li>
		<li>Remove the catheter just after the embryos have been released.</li>
		<li>Rinse the catheter, aspirating and releasing manipulating medium to check the absence of the embryos and confirm their successful transfer.</li>
		<li>Repeat steps 1.4.11 to 1.4.16 in the other side of the uterus, if desired.</li>
		<li>Remove the epidural needle and endoscope camera.</li>
		<li>Release CO2 through the endoscopic trocar. If excess gas remains in the abdomen of the animal, it will have pain and discomfort.</li>
		<li>Remove the endoscopic trocar from the abdominal cavity. Remove the surgical towel.</li>
		<li>Discontinue anesthesia.</li>
		<li>Clean the incision made by the trocar with chlorhexidine solution. Close the incision made by the trocar with a micronized aluminum and a plastic dressing.&#160;</li>
	</ol>
	</li>
	<li>Postoperative care
	<ol>
		<li>Treat the animals with antibiotics: 10 mg/kg of enrofloxacin, subcutaneously, every 24-h for 5 days.</li>
		<li>Administer analgesics: buprenorphine hydrochloride (0.03 mg/kg), intramuscularly, each 12 hours for 3 days; Meloxicam (0.2 mg/kg), subcutaneously, every 24-h for 3 days.</li>
		<li>Monitor the animals for at least 30 min after surgery (depending on the animal and the dose of anesthesia used) making sure they recover their physiological conditions.</li>
		<li>Identify the recipient (e.g., ear tattoo) and house animals individually in a clean cage with the appropriate environmental condition.</li>
	</ol>
	</li>
</ol>
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/58055/58055fig1.jpg" /> 
Figure 1: Laparoscopic embryo transfer assisted by laparoscopy (External view). A) Insertion of the endoscopic trocar (one port). B) Insertion of the endoscopic camera and the epidural needle (black arrow). C) Insertion of the embryo transfer catheter (white arrow) through the epidural needle. <a href="https://www.jove.com/files/ftp_upload/58055/58055fig1large.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/58055/58055fig2.jpg" /> 
Figure 2: Laparoscopic embryo transfer assisted by laparoscopy (Internal view). A: Insertion of the catheter through the epidural needle into the abdominal zone. Asterisk indicates the infundibulum. B, C, D: The catheter loaded with the embryos is inserted into ampulla region across the infundibulum. E, F: Release of the embryos, confirmed by the visualization of a swollen oviduct. This figure has been adapted from Marco-Jim&#233;nez et al.38. <a href="https://www.jove.com/files/ftp_upload/58055/58055fig2large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
2. Embryo Vitrification and Warming
<ol>
	<li>Perform all the manipulations at room temperature (around 22 &#176;C) to reduce the vitrification solution toxicity at warmer temperatures. 
	NOTE: Embryos can be moved using 0.1-2 &#181;L automatic pipette in this protocol, but other similar devices to move the embryos dragging the minimum volume can be suitable.</li>
	<li>Vitrify the embryos in a two-step addition procedure:
	<ol>
		<li>Place the embryos for 2 minutes in equilibrating solution consisting of 10% (v/v) ethylene glycol and 10% (v/v) dimethyl sulfoxide dissolved in BM.</li>
		<li>Move the embryos (from step 2.2.1) for 1 minute into vitrification solution consisting of 20% (v/v) ethylene glycol and 20% (v/v) dimethyl sulfoxide dissolved in BM.</li>
	</ol>
	</li>
	<li>Load the embryos into a 125 &#181;L plastic ministraw (which contains one closed end with a cotton plug and one open extreme). The process is schematized in Figure 3.
	<ol>
		<li>Couple the closed end of 0.125 &#956;L ministraw with the appropriate microdispenser (e.g. Captroll III&#174;).&#160;</li>
		<li>Aspirate BM until 1/3 of the straw length, following by a small air bubble.</li>
		<li>Aspirate the embryos in a volume of 40 &#181;L of vitrification solution, followed by another small air bubble.</li>
		<li>Aspirate BM until the first liquid fraction (step 2.3.2) reaches the cotton.</li>
		<li>Close the open end with a straw plug.</li>
	</ol>
	</li>
	<li>Perform step 2.2.2 while step 2.3 is being done to ensure that no more than one-minute elapses, which would be toxic to embryos.</li>
	<li>Plunge the ministraw directly into liquid nitrogen to achieve vitrification.</li>
	<li>Store the ministraw in a dewar for nitrogen for the desired time.</li>
	<li>Thaw the embryos in a single step.
	<ol>
		<li>Place the ministraw horizontally 10 cm from liquid nitrogen vapour for 20-30 s.</li>
		<li>When the crystallization process begins inside the ministraw, immerse the ministraw in a water bath at 25 &#176;C for 10-15 s.</li>
		<li>Remove the ministraw plug and cut the cotton plug.</li>
		<li>With a coupled microdispenser, expel all the ministraw content into a plate containing 0.33 M sucrose solution at 25 &#176;C in BM for 5 minutes. 
		NOTE: This step must be done quickly in order to reduce embryo exposure to the vitrification solution.&#160;</li>
		<li>Move the embryos to a new plate containing BM solution for another 5 min.</li>
		<li>Consider only non-damaged embryos (with intact mucin coat and zona pellucida) to continue with the ET. 
		NOTE: Take into account that in thawed embryos, asynchronous transfers (e.g., 60-62 h in morula transfers) may improve the results by allowing a resynchronization between the embryo and the maternal endometrium.</li>
	</ol>
	</li>
</ol>
<img alt="Figure 3" class="xfigimg" src="/files/ftp_upload/58055/58055fig3.jpg" /> 
Figure 3: Schematization of correctly loaded straw. A) BM refers to the embryo manipulating media employed during vitrification. Embryos must be loaded in vitrification solution. B) Macroscopic appearance of the loaded straw with a magnified detail of the embryo position. This large-volume device allows us to vitrify large number of embryos, unlike minimum volume devices. Furthermore, the handling of this device is easier compared with minimum volume devices, while the results are similar in rabbits41. <a href="https://www.jove.com/files/ftp_upload/58055/58055fig3large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

The authors have nothing to disclose.

Since the first documented live birth case from transferred embryos9, this technique and the rabbit species have become crucial in reproductive studies. Besides, embryo research studies involving manipulation, production, cryopreservation, etc. require as a last step the evaluation of embryo capacity to generate healthy full-term offspring. Therefore, embryo transfer technique is indispensable13,28. Over the years, the surgical me...

With the aims of bypassing human infertility or improving dissemination of livestock of high genetic value and preserving animal genetic resources, a set of techniques collectively termed assisted reproduction technologies, such as superovulation, in vitro fertilization, embryo culture, or cryopreservation, were developed1,2. Currently, hormonal treatments are given to stimulate the ovaries and produce a large number of antral ovarian follicles1. Oocytes collected from these follicles can be matured, fertilized, and developed in vitro until they are either cryopreserved or transferred to surrogate mothers3. However, during these treatments, gametes and zygotes are exposed to a series of non-physiological processes that could require embryo adaptation to survive in these conditions4,5. This adaptation is possible due to early embryo plasticity, which allows embryo changes in gene expression and developmental programming6. However, these modifications can influence the subsequent stages of embryo development until adulthood, and it is now widely accepted that methods, timing, cryopreservation procedure or culture conditions show different outcomes on embryo fate7,8. Therefore, to elucidate the specific induced effects of ARTs, the use of well-characterized animal models is inevitable.
The first documented live birth resulting from transfer of mammalian embryos took place in 18909. Today, embryo transfer (ET) to a surrogate female is a crucial step in studying the ART-induced effects during preimplantation on subsequent embryo development stages10. ET techniques depend on the size and anatomical structure of each animal. In the case of large-sized animal models, it has been possible to perform ET by transcervical nonsurgical ET techniques, but in smaller-size species catheterization of the cervix is more complex and surgical techniques are frequently used11. However, surgical ET can cause hemorrhaging that could impair implantation and embryo development, as blood can invade the uterine lumen, causing embryo death10. Transcervical nonsurgical ET techniques are still applied in humans, baboons, bovine, pigs and mice12,13,14,15,16,17, but surgical ETs are still being used in species such as goats, sheep or other animals which present additional difficulties10,18,19,20,21, such as rabbits (two independent cervices) or mice (small size). Nonetheless, surgical transfer methods tend to have gradually been replaced by less invasive methods. Endoscopy was used to transfer embryos, for example, in rabbits, pigs and small ruminants18,19,20. These minimally invasive endoscopy methods can be used to transfer embryos into the ampulla via the infundibulum, which is essential in rabbits and has demonstrated beneficial effects in some species20. This is based on the importance of the correct dialogue between embryo and mother during early embryo stages in the oviduct. As mentioned above, the embryo remodeling that takes place in rabbits during embryo migration through the oviduct is essential to achieve embryos able to implant22,23.
Larger-size animal models, such as bovine, are interesting because the biochemical and preimplantation features are similar to those in human species24. However, large animals are too expensive to use in preliminary trials, and rodents are considered an ideal model (76% model organisms are rodents) for laboratory research25. Nevertheless, the rabbit model provides some advantages over rodents in reproductive studies, as some reproductive biological processes exhibited by humans are more similar in rabbits than those in mice. Human and rabbits present a similar chronological embryonic genome activation, gastrulation and hemochorial placenta structure. In addition, using rabbits it is possible to know the exact timing of fertilization and pregnancy stages due to their induced ovulation25. Rabbit life cycles are short, completing gestation in 31 days and reaching puberty at about 4-5 months; the animal is easy to handle due to its docile and non-aggressive behavior, and its upkeep is very economical compared to the expense of larger animals. Moreover, it is crucial to mention that rabbits have a duplex uterus with two independent cervixes11,25. This places the rabbit in a preferential position, as embryos from the different experimental groups can be transferred into the same animal, but into a different uterine horn. This allows us to compare both experimental effects, reducing the maternal factor from the results.
Today, nonsurgical ET methods are not in use in rabbit. Some studies carried out in the late 90s using a transcervical ET technique resulted in low delivery rates ranging from 5.5% to 20.0%11,26 versus 50-65% by surgical methods, among them the laparoscopy procedure described by Besenfelder and Brem18. The low success rates of these nonsurgical ET methods in rabbits coincide with the lack of the necessary embryo remodeling in the oviduct, which is avoided in transcervical ET. Here, we describe an effective minimally invasive laparoscopic ET procedure using rabbits as a model organism. This technique provides a model for further reproductive research in large animals and humans.
Because rabbits have a particularly narrow time window for embryo implantation, ET in this species requires a high degree of synchrony between the developmental stage of the embryo at ET and the physiological status of the recipient27. In some cases, after a reproductive treatment that slows embryo development (such as in vitro culture) or alters the endometrial receptivity (such as superovulation treatments), there is no synchrony between the embryo and the maternal uterus. These situations can negatively affect outcome. To respond in these contexts, we describe an effective rabbit morula vitrification protocol that allows us to pause, organize and resume the experiments. This process is logistically desirable for reproductive studies and gives us the capacity for long-term storage of embryos, allowing their transport. The laparoscopic procedure and cryopreservation strategies allow better planning of studies with fewer animals. Thus, our methodology offers hygienic and economic advantages and conforms to the concept of the 3Rs (replacement, reduction and refinement) of animal research with the stated goal of improving human treatment of experimental animals. Thus, with these methods, rabbits constitute an ideal model organism for in vivo reproductive assays.

<table>
	<tbody>
		<tr>
			<td>Bovine Serum Albumin (BSA)</td>
			<td>VWR</td>
			<td>332</td>
			<td></td>
		</tr>
		<tr>
			<td>Buprenorphine hydrochloride</td>
			<td>Alvet Escart&#237;</td>
			<td>626</td>
			<td>To be ordered by a licensed veterinarian.</td>
		</tr>
		<tr>
			<td>Buserelin Acetate</td>
			<td>Sigma Aldrich</td>
			<td>B3303</td>
			<td></td>
		</tr>
		<tr>
			<td>Clorhexidine digluconate soap</td>
			<td>Alvet Escart&#237;</td>
			<td>0265DCCJ500B</td>
			<td></td>
		</tr>
		<tr>
			<td>Clorhexidine digluconate solution</td>
			<td>Alvet Escart&#237;</td>
			<td>0265DCCA500B</td>
			<td></td>
		</tr>
		<tr>
			<td>CO2</td>
			<td>Air Liquide</td>
			<td>99921</td>
			<td>CO2 N48.</td>
		</tr>
		<tr>
			<td>CO2 Incubator</td>
			<td>Fisher scientific</td>
			<td>15385194</td>
			<td></td>
		</tr>
		<tr>
			<td>Dimethyl Sulfoxide</td>
			<td>Sigma Aldrich</td>
			<td>W387509</td>
			<td></td>
		</tr>
		<tr>
			<td>Dulbecco&#8217;s phosphate-buffered saline (DPBS)</td>
			<td>Sigma Aldrich</td>
			<td>D5773</td>
			<td>Without calcium chloride.</td>
		</tr>
		<tr>
			<td>Electric razor</td>
			<td>Oster Golden A5</td>
			<td>078005-140-002</td>
			<td></td>
		</tr>
		<tr>
			<td>Endoscope camera</td>
			<td>Optomic Spain S.A</td>
			<td>OP-714</td>
			<td></td>
		</tr>
		<tr>
			<td>Endoscope trocar with silicone leaflet valve</td>
			<td>Karl Storz Endoscopia Ib&#233;rica S.A.</td>
			<td>30114GK</td>
			<td>Lightweight trocar model.</td>
		</tr>
		<tr>
			<td>Enrofloxacin</td>
			<td>Alvet Escart&#237;</td>
			<td>9993046</td>
			<td>To be ordered by a licensed veterinarian.</td>
		</tr>
		<tr>
			<td>Epicraneal needle 23G</td>
			<td>Alvet Escart&#237;</td>
			<td>514056353</td>
			<td>Smaller needles can be also used.</td>
		</tr>
		<tr>
			<td>Epidural catheter</td>
			<td>Vygon corporate</td>
			<td>187.10</td>
			<td></td>
		</tr>
		<tr>
			<td>Epidural needle</td>
			<td>Vygon corporate</td>
			<td>187.10</td>
			<td></td>
		</tr>
		<tr>
			<td>Ethylene Glycol</td>
			<td>Sigma Aldrich</td>
			<td>102466-M</td>
			<td></td>
		</tr>
		<tr>
			<td>Eye ointment</td>
			<td>Alvet Escart&#237;</td>
			<td>5273</td>
			<td></td>
		</tr>
		<tr>
			<td>Ketamine hydrochloride</td>
			<td>Alvet Escart&#237;</td>
			<td>184</td>
			<td>To be ordered by a licensed veterinarian.</td>
		</tr>
		<tr>
			<td>Laparoscopy equipment</td>
			<td>Karl Storz Endoscopia Ib&#233;rica S.A.</td>
			<td>26003 AA</td>
			<td>Hopkins&#174; Laparoscope, 0&#186;-mm straight-viewing laparoscope, 30-cm length, 5-mm working channel.</td>
		</tr>
		<tr>
			<td>Light source</td>
			<td>Optomic Spain S.A</td>
			<td>Fibrolux 250</td>
			<td></td>
		</tr>
		<tr>
			<td>Liquid Nitrogen</td>
			<td>Air Liquide</td>
			<td>P1505XXX</td>
			<td></td>
		</tr>
		<tr>
			<td>Mechanical CO2 insufflator</td>
			<td>Karl Storz Endoscopia Ib&#233;rica S.A.</td>
			<td>Endoflator&#174;</td>
			<td></td>
		</tr>
		<tr>
			<td>Meloxicam</td>
			<td>Alvet Escart&#237;</td>
			<td>9993501</td>
			<td>To be ordered by a licensed veterinarian.</td>
		</tr>
		<tr>
			<td>Petri dishes, 35-mm</td>
			<td>Sigma Aldrich</td>
			<td>CLS430165-500EA</td>
			<td></td>
		</tr>
		<tr>
			<td>Plastic dressing (Nobecutan)</td>
			<td>IBOR medica</td>
			<td>7140028</td>
			<td></td>
		</tr>
		<tr>
			<td>Plastic Straw 0.25 mL</td>
			<td>IMV - technologies</td>
			<td>6431</td>
			<td></td>
		</tr>
		<tr>
			<td>Povidone iodide solution</td>
			<td>Alvet Escart&#237;</td>
			<td>02656DPYS500S</td>
			<td></td>
		</tr>
		<tr>
			<td>Scissors</td>
			<td>ROBOZ</td>
			<td>RS-5880</td>
			<td>Any regular surgical grade steel small straight scissors will work.</td>
		</tr>
		<tr>
			<td>Silicone tube for insufflator</td>
			<td>Karl Storz Endoscopia Ib&#233;rica S.A.</td>
			<td>20400040</td>
			<td></td>
		</tr>
		<tr>
			<td>Stereomicroscope</td>
			<td>Leica</td>
			<td>MZ16F</td>
			<td>There are cheaper options such as Leica MZ8 or Nikon SMZ-10 or SMZ-2B, to name a few.</td>
		</tr>
		<tr>
			<td>Sterile Gloves</td>
			<td>Alvet Escart&#237;</td>
			<td>087GL010075</td>
			<td></td>
		</tr>
		<tr>
			<td>Sterile gown</td>
			<td>Alvet Escart&#237;</td>
			<td>12261501</td>
			<td></td>
		</tr>
		<tr>
			<td>Sterile mask</td>
			<td>Alvet Escart&#237;</td>
			<td>058B15924B</td>
			<td></td>
		</tr>
		<tr>
			<td>Straw Plug</td>
			<td>IMV - technologies</td>
			<td>6431</td>
			<td></td>
		</tr>
		<tr>
			<td>Sucrose</td>
			<td>Sigma Aldrich</td>
			<td>S7903</td>
			<td></td>
		</tr>
		<tr>
			<td>Syringe, 1-mL</td>
			<td>Fisher scientific</td>
			<td>11750425</td>
			<td></td>
		</tr>
		<tr>
			<td>Syringe, 5-mL</td>
			<td>Fisher scientific</td>
			<td>11773313</td>
			<td></td>
		</tr>
		<tr>
			<td>Urinary catheter</td>
			<td>IMV - technologies</td>
			<td>17722</td>
			<td></td>
		</tr>
		<tr>
			<td>Waterbath</td>
			<td>RAYPA</td>
			<td>BAE-4</td>
			<td></td>
		</tr>
		<tr>
			<td>Xylazine</td>
			<td>Alvet Escart&#237;</td>
			<td>525225</td>
			<td>To be ordered by a licensed veterinarian.</td>
		</tr>
		<tr>
			<td>Rabbits</td>
			<td>Universitat Polit&#232;cnica de Val&#232;ncia</td>
			<td>Line A</td>
			<td>Other maternal lines, such as Line V or Line HP can be used.</td>
		</tr>
	</tbody>
</table>

minimally invasive embryo transfer and embryo vitrification at the optimal embryo stage in rabbit model

Assisted reproductive techniques (ARTs), such as in vitro embryo culture or embryo cryopreservation, affect natural development patterns with perinatal and postnatal consequences. To ensure the innocuousness of ART applications, studies in animal models are necessary. In addition, as a last step, embryo development studies require evaluation of their capacity to develop full-term healthy offspring. Here, embryo transfer to the uterus is indispensable to perform any ARTs-related experiment.
The rabbit has been used as a model organism to study mammalian reproduction for over a century. In addition to its phylogenetic proximity to the human species and its small size and low maintenance cost, it has important reproductive characteristics such as induced ovulation, a chronology of early embryonic development similar to humans and a short gestation that allow us to study the consequences of ART application easily. Moreover, ARTs (such as intracytoplasmic sperm injection, embryo culture, or cryopreservation) are applied with suitable efficiency in this species.
Using the laparoscopic embryo transfer technique and the cryopreservation protocol presented in this article, we describe 1) how to transfer embryos through an easy, minimally invasive technique and 2) an effective protocol for long-term storage of rabbit embryos to provide time-flexible logistical capacities and the ability to transport the sample. The outcomes obtained after transferring rabbit embryos at different developmental stages indicate that morula is the ideal stage for rabbit embryo recovery and transfer. Thus, an oviductal embryo transfer is required, justifying the surgical procedure. Furthermore, rabbit morulae are successfully vitrified and laparoscopically transferred, proving the effectiveness of the described techniques.

Minimally invasive laparoscopic transfer of fresh or vitrified embryos places the rabbit among the best model animals for reproductive studies. Table 1 shows the results of fresh ET at different developmental stages (Figure 4) of transferred embryos. The survival rate at birth (percentage of embryos resulting in a pup) proved the efficacy of the laparoscopic technique described in this paper. The higher values were achieved when the ET was pe...

Watch this Scientific Journal Video about Minimally Invasive Embryo Transfer and Embryo Vitrification at the Optimal Embryo Stage in Rabbit Model at JoVE.com

Minimally Invasive Embryo Transfer and Embryo Vitrification at the Optimal Embryo Stage in Rabbit Model

Assisted reproductive techniques (ARTs) are in continuous evaluation to improve outcomes and reduce the associated risks. This manuscript describes a minimally invasive embryo transfer procedure with an efficient cryopreservation protocol that allows the use of rabbits as an ideal animal model of human reproduction.

Assisted reproductive techniques (ARTs), such as in vitro embryo culture or embryo cryopreservation, affect natural development patterns with perinatal ...

minimally-invasive-embryo-transfer-embryo-vitrification-at-optimal

Research

JoVE Journal

Developmental Biology

12.6K Views.  Universitat Politècnica de València (UPV). Assisted reproductive techniques (ARTs) are in continuous evaluation to improve outcomes and reduce the associated risks. This manuscript describes a minimally invasive embryo transfer procedure with an efficient cryopreservation protocol that allows the use of rabbits as an ideal animal model of human reproduction.

Video: Minimally Invasive Embryo Transfer and Embryo Vitrification at the Optimal Embryo Stage in Rabbit Model

Microbead Implantation in the Zebrafish Embryo

The zebrafish has emerged as a valuable genetic model system for the study of developmental biology and disease. Zebrafish share a high degree of genomic conservation, as well as similarities in cellular, molecular, and physiological processes, with other vertebrates including humans. During early ontogeny, zebrafish embryos are optically transparent, allowing researchers to visualize the dynamics of organogenesis using a simple stereomicroscope. Microbead implantation is a method that enables tissue manipulation through the alteration of factors in local environments. This allows researchers to assay the effects of any number of signaling molecules of interest, such as secreted peptides, at specific spatial and temporal points within the developing embryo. Here, we detail a protocol for how to manipulate and implant beads during early zebrafish development.

The zebrafish is an excellent model system for genetic and developmental studies. Bead implantation is a valuable tissue manipulation technique that can be used to interrogate developmental mechanisms by introducing alterations in local cellular environments. This protocol describes how to perform microbead implantation in the zebrafish embryo.

The zebrafish has emerged as a valuable genetic model system for the study of developmental biology and disease. Zebrafish share a high degree of genomic ...

Imaging Subcellular Structures in the Living Zebrafish Embryo

In vivo imaging provides unprecedented access to the dynamic behavior of cellular and subcellular structures in their natural context. Performing such imaging experiments in higher vertebrates such as mammals generally requires surgical access to the system under study. The optical accessibility of embryonic and larval zebrafish allows such invasive procedures to be circumvented and permits imaging in the intact organism. Indeed the zebrafish is now a well-established model to visualize dynamic cellular behaviors using in vivo microscopy in a wide range of developmental contexts from proliferation to migration and differentiation. A more recent development is the increasing use of zebrafish to study subcellular events including mitochondrial trafficking and centrosome dynamics. The relative ease with which these subcellular structures can be genetically labeled by fluorescent proteins and the use of light microscopy techniques to image them is transforming the zebrafish into an in vivo model of cell biology. Here we describe methods to generate genetic constructs that fluorescently label organelles, highlighting mitochondria and centrosomes as specific examples. We use the bipartite Gal4-UAS system in multiple configurations to restrict expression to specific cell-types and provide protocols to generate transiently expressing and stable transgenic fish. Finally, we provide guidelines for choosing light microscopy methods that are most suitable for imaging subcellular dynamics.

Imaging the dynamic behavior of organelles and other subcellular structures in vivo can shed light on their function in physiological and disease conditions. Here, we present methods for genetically tagging two organelles, centrosomes and mitochondria, and imaging their dynamics in living zebrafish embryos using wide-field and confocal microscopy.

In vivo imaging provides unprecedented access to the dynamic behavior of cellular and subcellular structures in their natural context. Performing such ...

Observing Mitotic Division and Dynamics in a Live Zebrafish Embryo

Mitosis is critical for organismal growth and differentiation. The process is highly dynamic and requires ordered events to accomplish proper chromatin condensation, microtubule-kinetochore attachment, chromosome segregation, and cytokinesis in a small time frame. Errors in the delicate process can result in human disease, including birth defects and cancer. Traditional approaches investigating human mitotic disease states often rely on cell culture systems, which lack the natural physiology and developmental/tissue-specific context advantageous when studying human disease. This protocol overcomes many obstacles by providing a way to visualize, with high resolution, chromosome dynamics in a vertebrate system, the zebrafish. This protocol will detail an approach that can be used to obtain dynamic images of dividing cells, which include: in vitro transcription, zebrafish breeding/collecting, embryo embedding, and time-lapse imaging. Optimization and modifications of this protocol are also explored. Using H2A.F/Z-EGFP (labels chromatin) and mCherry-CAAX (labels cell membrane) mRNA-injected embryos, mitosis in AB wild-type, auroraBhi1045, and esco2hi2865 mutant zebrafish is visualized. High resolution live imaging in zebrafish allows one to observe multiple mitoses to statistically quantify mitotic defects and timing of mitotic progression. In addition, observation of qualitative aspects that define improper mitotic processes (i.e., congression defects, missegregation of chromosomes, etc.) and improper chromosomal outcomes (i.e., aneuploidy, polyploidy, micronuclei, etc.) are observed. This assay can be applied to the observation of tissue differentiation/development and is amenable to the use of mutant zebrafish and pharmacological agents. Visualization of how defects in mitosis lead to cancer and developmental disorders will greatly enhance understanding of the pathogenesis of disease.

Mitosis is critical to every living organism and defects often lead to cancer and developmental disorders. Using this imaging protocol and zebrafish as a model system, researchers can visualize mitosis in a live vertebrate organism and the multitude of defects that arise when mitotic processes are defective.

Mitosis is critical for organismal growth and differentiation. The process is highly dynamic and requires ordered events to accomplish proper chromatin ...

Embryo Microinjection and Electroporation in the Chordate Ciona intestinalis

Simple model organisms are instrumental for in vivo studies of developmental and cellular differentiation processes. Currently, the evolutionary distance to man of conventional invertebrate model systems and the complexity of genomes in vertebrates are critical challenges to modeling human normal and pathological conditions. The chordate Ciona intestinalis is an invertebrate chordate that emerged from a common ancestor with the vertebrates and may represent features at the interface between invertebrates and vertebrates. A common body plan with much simpler cellular and genomic composition should unveil gene regulatory network (GRN) links and functional genomics readouts explaining phenomena in the vertebrate condition. The compact genome of Ciona, a fixed embryonic lineage with few divisions and large cells, combined with versatile community tools foster efficient gene functional analyses in this organism. Here, we present several crucial methods for this promising model organism, which belongs to the closest sister group to vertebrates. We present protocols for transient transgenesis by electroporation, along with microinjection-mediated gene knockdown, which together provide the means to study gene function and genomic regulatory elements. We extend our protocols to provide information on how community databases are utilized for in silico design of gene regulatory or gene functional experiments. An example study demonstrates how novel information can be gained on the interplay, and its quantification, of selected neural factors conserved between Ciona and man. Furthermore, we show examples of differential subcellular localization in embryonic cells, following DNA electroporation in Ciona zygotes. Finally, we discuss the potential of these protocols to be adapted for tissue specific gene interference with emerging gene editing methods. The in vivo approaches in Ciona overcome major shortcomings of classical model organisms in the quest of unraveling conserved mechanisms in the chordate developmental program, relevant to stem cell research, drug discovery, and subsequent clinical application.

Embryo Microinjection and Electroporation in the Chordate Ciona intestinalis

We present transient transgenesis and gene knockdown in Ciona intestinalis, a chordate sister group to vertebrates, using microinjection and electroporation techniques. Such methods facilitate functional genomics in this simple invertebrate that features rudimentary characteristics of vertebrates, including notochord and head sensory epithelia, and many orthologs of human disease associated genes.

Simple model organisms are instrumental for in vivo studies of developmental and cellular differentiation processes. Currently, the evolutionary distance ...

Use of Immunolabeling to Analyze Stable, Dynamic, and Nascent Microtubules in the Zebrafish Embryo

Microtubules (MTs) are dynamic and fragile structures that are challenging to image in vivo, particularly in vertebrate embryos. Immunolabeling methods are described here to analyze distinct populations of MTs in the developing neural tube of the zebrafish embryo. While the focus is on neural tissue, this methodology is broadly applicable to other tissues. The procedures are optimized for early to mid-somitogenesis-stage embryos (1 somite to 12 somites), however they can be adapted to a range of other stages with relatively minor adjustments. The first protocol provides a method to assess the spatial distribution of stable and dynamic MTs and perform a quantitative analysis of these populations with image-processing software. This approach complements existing tools to image microtubule dynamics and distribution in real-time, using transgenic lines or transient expression of tagged constructs. Indeed, such tools are very useful, however they do not readily distinguish between dynamic and stable MTs. The ability to image and analyze these distinct microtubule populations has important implications for understanding mechanisms underlying cell polarization and morphogenesis. The second protocol outlines a technique to analyze nascent MTs specifically. This is accomplished by capturing the de novo growth properties of MTs over time, following microtubule depolymerization with the drug nocodazole and a recovery period after drug washout. This technique has not yet been applied to the study of MTs in zebrafish embryos, but is a valuable assay for investigating the in vivo function of proteins implicated in microtubule assembly.

Immunolabeling methods to analyze distinct populations of microtubules in the developing zebrafish brain are described here, which are broadly applicable to other tissues. The first protocol outlines an optimized method for immunolabeling stable and dynamic microtubules. The second protocol provides a method to image and quantify nascent microtubules specifically.

Microtubules (MTs) are dynamic and fragile structures that are challenging to image in vivo, particularly in vertebrate embryos. Immunolabeling methods ...

AFM and Microrheology in the Zebrafish Embryo Yolk Cell

Elucidating the factors that direct the spatio-temporal organization of evolving tissues is one of the primary purposes in the study of development. Various propositions claim to have been important contributions to the understanding of the mechanical properties of cells and tissues in their spatiotemporal organization in different developmental and morphogenetic processes. However, due to the lack of reliable and accessible tools to measure material properties and tensional parameters in vivo, validating these hypotheses has been difficult. Here we present methods employing atomic force microscopy (AFM) and particle tracking with the aim of quantifying the mechanical properties of the intact zebrafish embryo yolk cell during epiboly. Epiboly is an early conserved developmental process whose study is facilitated by the transparency of the embryo. These methods are simple to implement, reliable, and widely applicable since they overcome intrusive interventions that could affect tissue mechanics. A simple strategy was applied for the mounting of specimens, AFM recording, and nanoparticle injections and tracking. This approach makes these methods easily adaptable to other developmental times or organisms.

The lack of tools to measure material properties and tensional parameters in vivo prevents validating their roles during development. We employed atomic force microscopy (AFM) and nanoparticle-tracking to quantify mechanical features on the intact zebrafish embryo yolk cell during epiboly. These methods are reliable and widely applicable avoiding intrusive interventions.

Elucidating the factors that direct the spatio-temporal organization of evolving tissues is one of the primary purposes in the study of development. ...

Effectiveness of the Air Stripping in Two Salmonid Fish, Rainbow Trout (Oncorhynchus Mykiss) and Brown Trout (Salmo Trutta Morpha fario)

Egg collection is one of the most crucial procedures during fish reproduction in salmonid hatcheries. Classic methods involve the use of hand massage on fish abdomens to expel the eggs. An alternative method uses the pressure of gas injected into the body cavity, which causes the subsequent release of the eggs. This method is believed to have less negative effects on both the welfare and egg quality of the broodstocks. Herein, we compare the results of air and hand stripping methods with respect to one-year survival and egg quantity and quality in two salmonid fish, rainbow trout (Oncorhynchus mykiss) and brown trout (Salmo trutta morpha fario). Our results indicate that air stripping yielded a better quality of eggs and higher one-year survival rate in rainbow trout. In addition, air stripping resulted in lower mortality rate than the group subjected to hand stripping (25% vs. 35%). The pH and hatching rate of the hand stripped group was lower than those of the air stripped group. In the case of brown trout, the quality of eggs obtained by both hand and air-stripping methods was similar; however, the one-year losses in fish were higher in air stripped group (15% compared to 0% in hand stripped fish). Although the advantages of air stripping method over hand stripping in terms of egg quality might not be observed in all salmonid species, the air-stripping procedure might be a promising option to be adopted in hatcheries as it ensures a high level of reproducibility and efficiency.

Effectiveness of the Air Stripping in Two Salmonid Fish, Rainbow Trout Oncorhynchus Mykiss and Brown Trout Salmo Trutta Morpha fario

The principal aim of this study is to standardize and test the pneumatic method (air stripping) of collecting eggs in rainbow trout and brown trout. This method allows effective and simple collection of the eggs without the necessity of fish abdomen massage.

Egg collection is one of the most crucial procedures during fish reproduction in salmonid hatcheries. Classic methods involve the use of hand massage on ...

Following Endocardial Tissue Movements via Cell Photoconversion in the Zebrafish Embryo

During embryogenesis, cells undergo dynamic changes in cell behavior, and deciphering the cellular logic behind these changes is a fundamental goal in the field of developmental biology. The discovery and development of photoconvertible proteins have greatly aided our understanding of these dynamic changes by providing a method to optically highlight cells and tissues. However, while photoconversion, time-lapse microscopy, and subsequent image analysis have proven to be very successful in uncovering cellular dynamics in organs such as the brain or the eye, this approach is generally not used in the developing heart due to challenges posed by the rapid movement of the heart during the cardiac cycle. This protocol consists of two parts. The first part describes a method for photoconverting and subsequently tracking endocardial cells (EdCs) during zebrafish atrioventricular canal (AVC) and atrioventricular heart valve development. The method involves temporally stopping the heart with a drug in order for accurate photoconversion to take place. Hearts are allowed to resume beating upon removal of the drug and embryonic development continues normally until the heart is stopped again for high-resolution imaging of photoconverted EdCs at a later developmental time point. The second part of the protocol describes an image analysis method to quantify the length of a photoconverted or non-photoconverted region in the AVC in young embryos by mapping the fluorescent signal from the three-dimensional structure onto a two-dimensional map. Together, the two parts of the protocol allows one to examine the origin and behavior of cells that make up the zebrafish AVC and atrioventricular heart valve, and can potentially be applied for studying mutants, morphants, or embryos that have been treated with reagents that disrupt AVC and/or valve development.

This protocol describes a method for the photoconversion of Kaede fluorescent protein in endocardial cells of the living zebrafish embryo that enables the tracking of endocardial cells during atrioventricular canal and atrioventricular heart valve development.

During embryogenesis, cells undergo dynamic changes in cell behavior, and deciphering the cellular logic behind these changes is a fundamental goal in the ...

Probing Cell Mechanics with Bead-Free Optical Tweezers in the Drosophila Embryo

Morphogenesis requires coordination between genetic patterning and mechanical forces to robustly shape the cells and tissues. Hence, a challenge to understand morphogenetic processes is to directly measure cellular forces and mechanical properties in vivo during embryogenesis. Here, we present a setup of optical tweezers coupled to a light sheet microscope, which allows to directly apply forces on cell-cell contacts of the early Drosophila embryo, while imaging at a speed of several frames per second. This technique has the advantage that it does not require the injection of beads into the embryo, usually used as intermediate probes on which optical forces are exerted. We detail step by step the implementation of the setup, and propose tools to extract mechanical information from the experiments. By monitoring the displacements of cell-cell contacts in real time, one can perform tension measurements and investigate cell contacts&#39; rheology.

Probing Cell Mechanics with Bead-Free Optical Tweezers in the Drosophila Embryo

We present a setup of optical tweezers coupled to a light sheet microscope, and its implementation to probe cell mechanics without beads in the Drosophila embryo.

Morphogenesis requires coordination between genetic patterning and mechanical forces to robustly shape the cells and tissues. Hence, a challenge to ...

Microinjection-based System for In Vivo Implantation of Embryonic Cardiomyocytes in the Avian Embryo

Interpreting the relative impact of cell autonomous patterning versus extrinsic microenvironmental influence on cell lineage determination represents a general challenge in developmental biology research. In the embryonic heart, this can be particularly difficult as regional differences in the expression of transcriptional regulators, paracrine/juxtacrine signaling cues, and hemodynamic force are all known to influence cardiomyocyte maturation. A simplified method to alter a developing cardiomyocyte's molecular and biomechanical microenvironment would, therefore, serve as a powerful technique to examine how local conditions influence cell fate and function. To address this, we have optimized a method to physically transplant juvenile cardiomyocytes into ectopic locations in the heart or the surrounding embryonic tissue. This allows us to examine how microenvironmental conditions influence cardiomyocyte fate transitions at single cell resolution within the intact embryo. Here, we describe a protocol in which embryonic myocytes can be isolated from a variety of cardiac sub-domains, dissociated, fluorescently labeled, and microinjected into host embryos with high precision. Cells can then be directly analyzed in situ using a variety of imaging and histological techniques. This protocol is a powerful alternative to traditional grafting experiments that can be prohibitively difficult in a moving tissue such as the heart. The general outline of this method can also be adapted to a variety of donor tissues and host environments, and its ease of use, low cost, and speed make it a potentially useful application for a variety of developmental studies.

In this method, embryonic cardiac tissues are surgically microdissected, dissociated, fluorescently labeled, and implanted into host embryonic tissues. This provides a platform for studying the individual or tissue level developmental organization under ectopic hemodynamic conditions, and/or altered paracrine/juxtacrine environments.

Interpreting the relative impact of cell autonomous patterning versus extrinsic microenvironmental influence on cell lineage determination represents a ...

Minimally Invasive Embryo Transfer and Embryo Vitrification at the Optimal Embryo Stage in Rabbit Model

Podsumowanie

Przeglądaj więcej filmów

Rozdziały w tym wideo

Microbead Implantation in the Zebrafish Embryo

Imaging Subcellular Structures in the Living Zebrafish Embryo

Observing Mitotic Division and Dynamics in a Live Zebrafish Embryo

Embryo Microinjection and Electroporation in the Chordate Ciona intestinalis

Use of Immunolabeling to Analyze Stable, Dynamic, and Nascent Microtubules in the Zebrafish Embryo

AFM and Microrheology in the Zebrafish Embryo Yolk Cell

Effectiveness of the Air Stripping in Two Salmonid Fish, Rainbow Trout (Oncorhynchus Mykiss) and Brown Trout (Salmo Trutta Morpha fario)

Following Endocardial Tissue Movements via Cell Photoconversion in the Zebrafish Embryo

Probing Cell Mechanics with Bead-Free Optical Tweezers in the Drosophila Embryo

Microinjection-based System for In Vivo Implantation of Embryonic Cardiomyocytes in the Avian Embryo