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 methods initially employed to transfer embryos into the maternal uterus have gradually been replaced by less invasive methods in the vast majority of species13,14,15,21,27,29,30. However, in rabbits, intraoviductal ET in early embryo stages of development and in vitro produced embryos becomes unavoidable to ensure a similar result to natural conditions. In rabbits, intraoviductal mucin coat is a crucial factor allowing embryo implantation, as it takes place after the remodelling of the embryonic coatings during blastocyst expansion in the uterine horns. However, mucin coat deposition is limited to the oviduct for 3 days following ovulation, and the molecular mechanisms of coat material deposition are largely unknown31. For these reasons, it is known that in vitro-developed blastocysts did not survive when transferred to the uterus32,33,34, and embryos with a damaged mucin coat have a lower survival rate35. Likewise, groups that reported a transcervical embryo transfer in rabbits resulted in very low live born rates11,26. Here, we present a minimally invasive technique, adapted from Besenfelder and Brem18, to transfer embryos with successful birth rates. According to the results in Table 1, the morula stage in rabbit embryos was the best embryonic stage to achieve a high survival rate at birth. One possible explanation is the greater sensitivity to manipulation of the earliest stages. Interestingly, the success rate increases as the embryonic stage progresses, possibly due to the greater exposure of the embryo to oviductal secretions prior to its recovery. But when embryos reach the blastocyst stage and are place-concordant transferred to the uterus, the values decrease drastically. Not excluding what has been said, a possible explanation could be that the embryos transferred into the oviduct can restore the possible damage generated in the mucin layer during embryo manipulation. Therefore, blastocysts transferred into the uterus would be deprived of this mechanism, which could compromise their implantation capacity.
The technique is performed using a single port instrument (5 mm endoscope trocar), with slight, brief manipulation. Therefore, the5-mm endoscope trocar incision does not require closure.Laparoscopic technique benefits include decreased postoperative pain, quicker return to normal activity, and fewer postoperative complications. In addition, endoscopic procedures induce fewer abdominal adhesions and allow a better immune response by the recipient compared with open surgery21,36,37. Accumulating evidence from our lab has demonstrated the effectiveness of this ET procedure in the rabbit model. Thus, in the last five years a total of 3,909 embryos (1,335 fresh and 2,574 vitrified embryos) were transferred through the procedure described in the present manuscript. As a result of this technique, the offspring rates of fresh and vitrified transfer embryos were 62.9% and 42.5%, respectively38,39,40,41,42,43,44,45,46,47. Many studies are all based on this technique: Marco-Jim&#233;nez et al.38,39,40,41, Vicente et al.42, Viudes-de-Castro et al.43, Saenz-de-Juano et al.44,45,47, Lavara et al.46.
Practical recommendations for carrying out this technique are described below. In embryo culture experiments, it is also advisable to use a new catheter for embryo transfer instead of the one used to move the embryos between the culture media and manipulation media. This avoids transfer of mineral oil and ensures an optimal flow. During ET it is important to minimize handling of the reproductive tract, as excessive manipulation of the oviduct could result in adhesions. If the oviduct is twisted, employ the epidural syringe to try to position it correctly, not the catheter, as it contains the embryos and the mechanical manipulation could cause their loss. Once the catheter passes through the oviduct, it slides easily. If it does not, the catheter may have deviated. Once inside the oviduct, if the media does not flow, move the catheter out slightly and try to reinsert it again. If it still does not flow, the catheter is clogged. Remove it from the oviduct and release the content into a dish with a clean medium. Then, reload the embryos into another catheter and try to reinsert it into the oviduct again. Delivery usually takes places 28-30 days after morula transfer.
In addition, there is evidence indicating that the embryo developmental stage can be more advanced than the uterine environment in pseudopregnant females, but not the opposite. Specifically, embryos have the ability to wait for the favorable womb environment, but the womb environment cannot wait for the embryos at the right stage for implantation10. With regard to vitrified embryos, after a short/Long-term storage it is possible to synchronize the developmental stage of the embryo with the corresponding favorable womb environment. Furthermore, if the embryo donor is also the embryo recipient, the detrimental effects of superovulation on the endometrium can be bypassed by using the vitrification technique and transferring the embryos in a subsequent cycle48. In rabbits, vitrified embryos transferred into oviducts of recipients induced to ovulate 60-62 h beforehand (asynchrony) is a highly efficient technique44,49. Related with this, it has been suggested that the oviductal embryo transition during 10-12 h could explain the beneficial effects in the restoration of cell physiology and replacement of dead cells, and probably repair the damage induced in mucin coat during embryo manipulation. Besides, vitrified embryos present a delay in development, as they have been metabolically suspended during the storage. Therefore, transfer of cryopreserved embryos into asynchronous recipients allows the embryo to reactivate its metabolic activity and thus the embryo stage of development is synchronized with the womb environment. Instead, if cryopreserved embryos are transferred into synchronic receptors, the cross-talk between the mother and the embryo hinders the onset of a successful pregnancy. In rabbit, the highest survival rate has been obtained after intraoviductal transfer of cryopreserved morulae49. Our data are consistent with this report, although the morula stage exhibits different survival rates following cryopreservation depending on their degree of compaction at 70-72 h (Table 2). Here, compacted morulae showed higher survival rates at birth in comparison to non-compacted morulae, which was in concordance with previous reports showing that every stage of development had its own mechanism relative to the permeation of cryoprotectants and the extent of dehydration during the addition of the cryopreservation solution50. Underlying these techniques, we have demonstrated that a combination of vitrification and intraoviductal embryo transfer is a successful strategy to re-establish rabbit populations after 15 years of storage in liquid nitrogen, without adverse effect on their post-thaw survival and live birth51.
The following details should be taken into account to successfully perform this technique. It is important to bear in mind that the increasing density of the consecutive mediums used for vitrification (DPBS, equilibration solution, vitrification solution) could induce embryo contraction due to progressive embryo dehydration. However, its normal appearance is recovered when the embryo is equilibrated with the medium. Furthermore, when the embryo is moved between increasing density media, it tends to move to the surface of the media due to density movements. To avoid embryo loss and ensure the time of vitrification, it is recommendable to perform the vitrification in small drops of the media that will keep the embryo in place.
In conclusion, here we describe both an ET technique and an embryo vitrification method that facilitate future studies which use rabbits as a model. Based on the close phylogenetic distance between rabbits and humans, the use of this model could provide results easily transferable to human clinical medicine. In addition, our method offers some hygienic and economic advantages, conforming to the concept of the 3 Rs of animal welfare (replacement, reduction and refinement), while maintaining the goal of improving humane treatment of experimental animals.

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&amp;iacute;</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&amp;iacute;</td><td>0265DCCJ500B</td><td></td></tr><tr><td>Clorhexidine digluconate solution</td><td>Alvet Escart&amp;iacute;</td><td>0265DCCA500B</td><td></td></tr><tr><td>CO&lt;sub&gt;2&lt;/sub&gt;</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&amp;rsquo;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&amp;eacute;rica S.A.</td><td>30114GK</td><td>Lightweight trocar model.</td></tr><tr><td>Enrofloxacin</td><td>Alvet Escart&amp;iacute;</td><td>9993046</td><td>To be ordered by a licensed veterinarian.</td></tr><tr><td>Epicraneal needle 23G</td><td>Alvet Escart&amp;iacute;</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&amp;iacute;</td><td>5273</td><td></td></tr><tr><td>Ketamine hydrochloride</td><td>Alvet Escart&amp;iacute;</td><td>184</td><td>To be ordered by a licensed veterinarian.</td></tr><tr><td>Laparoscopy equipment</td><td>Karl Storz Endoscopia Ib&amp;eacute;rica S.A.</td><td>26003 AA</td><td>Hopkins&amp;reg; Laparoscope, 0&amp;ordm;-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&amp;eacute;rica S.A.</td><td>Endoflator&amp;reg;</td><td></td></tr><tr><td>Meloxicam</td><td>Alvet Escart&amp;iacute;</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&amp;iacute;</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&amp;eacute;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&amp;iacute;</td><td>087GL010075</td><td></td></tr><tr><td>Sterile gown</td><td>Alvet Escart&amp;iacute;</td><td>12261501</td><td></td></tr><tr><td>Sterile mask</td><td>Alvet Escart&amp;iacute;</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&amp;iacute;</td><td>525225</td><td>To be ordered by a licensed veterinarian.</td></tr><tr><td>Rabbits</td><td>Universitat Polit&amp;egrave;cnica de Val&amp;egrave;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 performed with embryos in the morula stage, either early or compact morulae. Based on these results, we performed a second experiment to demonstrate the survival rate after vitrification of these embryos. Thus, in Table 2 we show the results obtained after transferring vitrified rabbit morulae recovered at the same time, differentiating between those embryos that had reached a good degree of compaction or not. The survival rate at birth was different between the different embryo stages, being higher in compacted morulae. Therefore, laparoscopic embryo transfer is a reliable technique to transfer fresh and vitrified embryos in rabbits
<img alt="Figure 4" class="xfigimg" src="/files/ftp_upload/58055/58055fig4.jpg" /> 
Figure 4: Rabbit embryos. A) Pronuclear. B) Eight cells. C) Early morula. D) Compact morula. E) Blastocyst. Asterisk indicates the two pronuclei. Black arrows indicate the zona pellucida. White arrows indicate the mucin coat, which normally varies between embryos. ICM: Inner Cell Mass. TE: Trophectoderm. Scale bar: 50 &#181;m. <a href="https://www.jove.com/files/ftp_upload/58055/58055fig4large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>Developmental stage1</td>
			<td>Embryos</td>
			<td>Recipients</td>
			<td>Place of transfer</td>
			<td>Pregnancy rate (%)</td>
			<td>Implantation rate (%)</td>
			<td>Survival rate at birth (%)2</td>
		</tr>
		<tr>
			<td>Pronuclear</td>
			<td>78</td>
			<td>7</td>
			<td>Oviduct</td>
			<td>7 (100)</td>
			<td>50 (64.0)b</td>
			<td>34 (43.6)b</td>
		</tr>
		<tr>
			<td>8 cells</td>
			<td>81</td>
			<td>7</td>
			<td>Oviduct</td>
			<td>7 (100)</td>
			<td>60 (74.1)b</td>
			<td>53 (65.4)a</td>
		</tr>
		<tr>
			<td>Early morula</td>
			<td>81</td>
			<td>7</td>
			<td>Oviduct</td>
			<td>7 (100)</td>
			<td>80 (98.8)a</td>
			<td>60 (74.1)a</td>
		</tr>
		<tr>
			<td>Compact morula</td>
			<td>80</td>
			<td>7</td>
			<td>Oviduct</td>
			<td>7 (100)</td>
			<td>80 (100)a</td>
			<td>58 (72.5)a</td>
		</tr>
		<tr>
			<td>Blastocyst</td>
			<td>80</td>
			<td>7</td>
			<td>Uterus</td>
			<td>7 (100)</td>
			<td>73 (91.3)a</td>
			<td>38 (47.5)b</td>
		</tr>
	</tbody>
</table>
Table 1. Efficiency of fresh rabbit embryo transfer (in vivo derived) by laparoscopy. 1Different embryos were recovered at 18-20h (pronuclear), 36-38h (8 cells), 60-62 h (early morula), 70-72 h (compact morula) and 80-82 h (blastocyst) after mating. Compact (&#62;32 cells) and non-compact morulae (&#8776;32 cells) can be founded at 70-72 h, but only compact morulae were transferred. 2Survival rate at birth from recipient pregnant does. a,bValues with different superscripts are statistically different (P&#60;0.001).
<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>Developmental stage</td>
			<td>Transferred embryos</td>
			<td>Recipients</td>
			<td>Pregnancy rate (%)</td>
			<td>Survival rate at birth (%)1</td>
		</tr>
		<tr>
			<td>Non- compacted</td>
			<td>135</td>
			<td>10</td>
			<td>9 (90)</td>
			<td>62 (45.9)b</td>
		</tr>
		<tr>
			<td>Compacted</td>
			<td>150</td>
			<td>10</td>
			<td>10 (100.0)</td>
			<td>98 (65.3)a</td>
		</tr>
		<tr>
			<td>TOTAL</td>
			<td>285</td>
			<td>20</td>
			<td>19 (95)</td>
			<td>160 (56.1)</td>
		</tr>
	</tbody>
</table>
Table 2. Viability of non-compacted vs compact vitrified morula. a,bValues with different superscripts are statistically different (P&#60;0.001). 1Survival rate at birth from recipient pregnant does. Embryos were recovered at the same time (70-72 h) and were distinguished into compact (&#62;32 cells) and non-compact morulae (&#8776;32 cells).

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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

Nanyang Technological University

Research

JoVE Journal

Developmental Biology

12.6K Views.  Universitat Politècnica de València (UPV). Het belangrijkste doel van dit experiment is om een laparoscopische embryo overdracht procedure bij konijn te beschrijven als een model. Het belangrijkste voordeel van deze techniek is dat deze werkt om embryo's over te brengen met behulp van een eenvoudige, minimaal invasieve techniek. Bovendien werkt het effectieve hier beschreven protocol voor embryo cryopreservatie ook tot de langdurige opslag van de embryo's van het konijn, waardoor tijdflexibele logistieke capaciteiten en de mogelijkhei...

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 ...

Modified MicroSecure Vitrification: A Safe, Simple and Highly Effective Cryopreservation Procedure for Human Blastocysts

Clinical embryo vitrification evolved with the development of unique vitrification devices in the 21st century and with the misconception that ultra-rapid cooling in an &#34;open&#34; system (i.e., direct LN2 contact) was a necessity to optimize vitrification success. The dogma surrounding the importance of cooling rates led to unsafe practices subject to technical variation and to the creation of vitrification devices that disregarded important quality-control factors (e.g., ease of use, repeatability, reliability, labeling security, and storage safety). Understanding the quality-control flaws of other devices allowed for the development of a safe, secure, repeatable, and reliable &#181;S-VTF method aimed to minimize intra- and inter-technician variation. Equally important, it combined the availability of two existing FDA-compliant devices: 1) a 0.3-mL ionomeric resin embryo straw with internalized, dual-colored, tamper-proof labeling with repeatable weld seal potential; and 2) shortened, commonly-used, 300-&#181;m ID sterile flexipettes to directly load the embryo(s) in order to create a highly-effective global vitrification device. Like other aseptic, closed vitrification systems (e.g., High Security Vitrification (HSV), Rapid-i, and VitriSafe) effectively used in reproductive medicine, microSecure Vitrification (&#181;S-VTF) has proven that it can achieve high post-warming survival and pregnancy outcomes with its attention to simplicity, and reduced technical variation. Although the 0.3-mL embryo straw containing an internal hydrophobic plug was commercially replaced with a standard semen straw possessing cotton-polyvinyl pyrrolidone (PVP) plugs, it maintained its ionomeric resin composition to ensure weld sealing. However, the cotton plugs can wick out the fluid-embryo contents of the flexipettes upon contact. A modified &#181;S-VTF method was adapted to include an additional internal weld seal before the plug on the device loading side. The added technical step to the &#181;S-VTF procedure has not affected its successful application, as high survival rates (&#62; 95%) and pregnancy rates continue today.

MicroSecure Vitrification was developed as a non-commercial, aseptic closed vitrification device system that is compliant with FDA good manufacturing and tissue-handling practices. Due to the withdrawal of hydrophobic plugged embryo straws from the industry, the vitrification procedure was modified to include an inner seal before the standard internal cotton plug.

Clinical embryo vitrification evolved with the development of unique vitrification devices in the 21st century and with the misconception that ultra-rapid ...

Production of Apolipoprotein C-III Knockout Rabbits using Zinc Finger Nucleases

Apolipoprotein (Apo) C-III (ApoCIII) resides on the surface of plasma chylomicron (CM), very low density lipoprotein (VLDL) and high density lipoproteins (HDL). It has been recognized that high levels of plasma ApoCIII constitutea risk factor for cardiovascular diseases (CVD). Elevated plasma ApoCIII level often correlates with insulin resistance, obesity, and hypertriglyceridemia. Invaluable knowledge on the roles of ApoCIIIin lipid metabolisms and CVD has been obtained from transgenic mouse models including ApoCIII knockout (KO) mice; however, it is noted that the metabolism of lipoprotein in mice is different from that of humans in many aspects. It is not known until now whether elevated plasma ApoCIII is directly atherogenic. We worked to develop ApoCIII KO rabbits in the present study based on the hypothesis that rabbits can serve as a reasonablemodelfor studying human lipid metabolism and atherosclerosis. Zinc finger nuclease (ZFN) sets targeting rabbit ApoCIIIgene were subjected to in vitro validation prior to embryo microinjection. The mRNA was injected to the cytoplasm of 35 rabbit pronuclear stage embryos, and evaluated the mutation rates at the blastocyst state. Of sixteen blastocysts that were assayed, a satisfactory 50% mutation rate (8/16) at the targeting site was achieved, supporting the use of Set 1 for in vivo experiments. Next, we microinjected 145 embryos with Set 1 mRNA, and transferred these embryos to 7 recipient rabbits. After 30 days gestation, 21 kits were born, out of which five were confirmed as ApoCIII KO rabbits after PCR sequencing assays. The KO animal rate (#KO kits/total born) was 23.8%. The overall production efficiency is 3.4% (5 kits/145 embryos transferred). The present work demonstrated that ZFN is a highly efficient method to produce KO rabbits. These ApoCIII KO rabbits are novel resources to study the roles of ApoCIII in lipid metabolisms.

Recent development in gene targeting tools makes production of knockout (KO) rabbits possible. In the present work, we generated five Apolipoprotein (Apo) C-III KO rabbits using Zinc Finger Nucleases (ZFN). This work demonstrated that ZFN is a highly efficient method to produce KO rabbits.

Apolipoprotein (Apo) C-III (ApoCIII) resides on the surface of plasma chylomicron (CM), very low density lipoprotein (VLDL) and high density lipoproteins ...

Medicine

Utero-tubal Embryo Transfer and Vasectomy in the Mouse Model

The transfer of preimplantation embryos to a surrogate female is a required step for the production of genetically modified mice or to study the effects of epigenetic alterations originated during preimplantation development on subsequent fetal development and adult health. The use of an effective and consistent embryo transfer technique is crucial to enhance the generation of genetically modified animals and to determine the effect of different treatments on implantation rates and survival to term. Embryos at the blastocyst stage are usually transferred by uterine transfer, performing a puncture in the uterine wall to introduce the embryo manipulation pipette. The orifice performed in the uterus does not close after the pipette has been withdrawn, and the embryos can outflow to the abdominal cavity due to the positive pressure of the uterus. The puncture can also produce a hemorrhage that impairs implantation, blocks the transfer pipette and may affect embryo development, especially when embryos without zona are transferred. Consequently, this technique often results in very variable and overall low embryo survival rates. Avoiding these negative effects, utero-tubal embryo transfer take advantage of the utero-tubal junction as a natural barrier that impedes embryo outflow and avoid the puncture of the uterine wall. Vasectomized males are required for obtaining pseudopregnant recipients. A technique to perform vasectomy is described as a complement to the utero-tubal embryo transfer.

Utero-tubal embryo transfer uses the utero-tubal junction as a barrier to prevent the embryo outflow that may occur when performing uterine transfer. Vasectomized males are required to obtain pseudopregnant recipients for embryo transfer. Both techniques are discussed.

The transfer of preimplantation embryos to a surrogate female is a required step for the production of genetically modified mice or to study the effects ...

Biology

Pregnancy and Nursing Management for Embryo-Transferred and Genetically Modified Rabbits

With the advancement of scientific research, the demand for gene-edited rabbit models is increasing. However, there are limited pregnancy and feeding management systems for gene-edited rabbits, leading to low survival rates among gene-edited rabbits prepared by many inexperienced researchers. Therefore, proper guidance is essential. This article summarizes the pregnancy and feeding practices for genetically modified rabbits developed in the author's laboratory and outlines a set of fundamental processes. These include pregnancy diagnosis, antenatal care, midwifery, assisted breastfeeding, weaning, and other procedures, along with the rescue and care of weak newborn rabbits. Compared to the traditional natural childbirth and nurturing methods used in rabbit farms, this approach involves more refined management, requiring additional time and effort but significantly increasing the survival rate of suckling rabbits. The methods described in this article are suitable for most laboratory breeding scenarios involving gene-edited or embryo-transferred rabbits and provide a straightforward and effective reference for other researchers.

This protocol describes a pregnancy and feeding management technique for embryo-transferred and genetically modified rabbits, aimed at reducing newborn rabbit mortality and enhancing the preparation efficiency of gene-edited rabbits.

With the advancement of scientific research, the demand for gene-edited rabbit models is increasing. However, there are limited pregnancy and feeding ...

Rabbit Embryo Thawing: A Warming Procedure to Recover Cryopreserved Rabbit Embryos for Embryo Transfer

Source: Garcia-Dominguez, X. et al., <a href="https://www.jove.com/v/58055">Minimally Invasive Embryo Transfer and Embryo Vitrification at the Optimal Embryo Stage in Rabbit Model.</a> J. Vis. Exp. (2019)
This video demonstrates the procedure for thawing vitrified rabbit embryos and the precautionary measurement steps for optimal embryo recovery.

Source: Garcia-Dominguez, X. et al., Minimally Invasive Embryo Transfer and Embryo Vitrification at the Optimal Embryo Stage in Rabbit Model. J. Vis. Exp. ...

JoVE Encyclopedia of Experiments

Large Animal Models

Leporine

Embryo Transfer Into Rabbit: A Procedure for Laparoscopy-Guided Transfer of Embryos Into Oviduct of Ovulating Recipient Female Rabbit Model

Source: Garcia-Dominguez, X. et al., <a href="https://www.jove.com/v/58055">Minimally Invasive Embryo Transfer and Embryo Vitrification at the Optimal Embryo Stage in Rabbit Model.</a> J. Vis. Exp. (2019)
This video describes the technique of transferring embryos into the oviduct of a recipient female rabbit via Laparoscopy. It allows embryo remodeling in rabbits during its migration through the oviduct, eventually helping in successful implantation. This technique can also be used for stored embryos with good efficiency.

Rabbit Embryo Vitrification Using Cryogenic Straw:  An Ultra-Rapid Cooling Technique to Preserve Rabbit Embryos for Long-term Storage

Source: Garcia-Dominguez, X. et al. <a href="https://www.jove.com/v/58055">Minimally Invasive Embryo Transfer and Embryo Vitrification at the Optimal Embryo Stage in Rabbit Model.</a> J. Vis. Exp. (2019)
In this video, we describe the procedure for vitrification of harvested rabbit embryos at desired development stage using ethylene glycol and dimethyl sulfoxide as cryoprotectants and cryogenic straw as the carrier system. The vitrified embryos have high survival rate and can be stored for an extended period until use.

Source: Garcia-Dominguez, X. et al. Minimally Invasive Embryo Transfer and Embryo Vitrification at the Optimal Embryo Stage in Rabbit Model. J. Vis. Exp. ...

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

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