This study was supported by the Swiss National Science Foundation (project grant no. 310030_192736). We would like to thank Dr. Frauke Seehusen from the Institute of Veterinary Pathology of the University of Zurich for her histopathological support.

All animal experiments were performed following Swiss Federal Animal Regulations and approved by the Veterinary Office of Zurich (n&#176; 225/2019), assuring human care. Female virgin Lewis rats (body weight of 170-200 g) and female virgin Brown Norway rats (170-200 g) were used as uterus donors/recipients, while male Lewis rats (300-320 g) were used for mating. The rats were aged from 12-15 months. The animals were obtained from commercial sources (see Table of Materials) and were housed in controlled conditions and an enriched environment with free access to water and standard food.
1. Uterus retrieval
NOTE: For details on the procedure, please see the previously published reports12,13,15.
<ol>
	<li>Induce anesthesia with isoflurane and oxygen within an enclosed Plexiglas container (14 cm x 25 cm x 13 cm) for 1-2 min (5 vol% isoflurane in O2).
	<ol>
		<li>Administer buprenorphine subcutaneously (0.05 mg/kg) and bupivacaine (0.5%, 8 mg/kg) subcutaneously in the region of the planned abdominal incision 30 min before surgery.</li>
		<li>Shave all the abdominal skin of the rat with an electric shaver.</li>
		<li>Use tapes to keep the animal fixed on a heating plate during surgery. Apply eye ointment to both eyes.</li>
		<li>Maintain anesthesia during the procedure with 2-4 vol% isoflurane in oxygen by continuous administration through a small nose cone.</li>
		<li>Monitor the anesthetic depth by clinical parameters without specialized tools (respiratory rate of ~70-120/min-a slow rate drop of 50% is acceptable during anesthesia; checking anesthetic depth with toe pinch; color of mucous membranes should be pink, not blue or grey)16, and adjust the isoflurane concentration accordingly. 
		NOTE: Optional: frequent respiration monitoring during surgery is feasible with the help of an assistant.</li>
		<li>Confirm anesthetic depth by performing a toe pinch.</li>
		<li>Clean&#160;the abdominal skin in a circular motion with three alternating swabs of an antiseptic solution and 70% alcohol. Allow to dry.</li>
		<li>Place a sterile drape (see Table of Materials) with an abdominal window over the animal.</li>
	</ol>
	</li>
	<li>Perform median laparotomy.
	<ol>
		<li>Open the abdomen via a 6-8 cm midline long incision, starting 0.5 cm below the xiphisternum toward the hypogastrium. Use a no. 10 scalpel for the skin incision and small sharp scissors for the linea alba incision. Do not damage the liver or the bladder.</li>
		<li>Move the intestines outside the abdominal cavity using cotton swabs, cover&#160;them gently with a gauze moistened with sterile saline,&#160;and protect&#160;them with a sterile plastic bag for better insulation.</li>
		<li>Insert retractors or clips (see Table of Materials) at the left and right abdominal wall folders to keep the peritoneal muscle aside and the abdomen open, to obtain optimal access and visibility of the uterus and associated vessels. Fix the clips/retractors with tapes.</li>
		<li>Apply prewarmed saline to keep the surgical area and the intestines moist and avoid drying of the viscera.</li>
	</ol>
	</li>
	<li>Harvest the right uterine horn with the common uterine cavity and cervix plus vascular pedicles, including the right uterine, internal, and common iliac vessels.
	<ol>
		<li>Ligate (4/0 polyglactin; see Table of Materials), cauterize, and sever the left uterine horn adjacent to the branching from the common uterine cavity.</li>
		<li>Remove excess fat surrounding the uterus and vagina. 
		NOTE: Keep the fat around the uterine-vascular system.</li>
		<li>Dissect the bladder at its attachment to the cervix with cauterization of all draining and feeding bladder vessels. During cauterization, maintain an adequate distance between the cervix and the vagina to avoid unncecessary cauterization on these two structures. Otherwise, the risk of graft necrosis increases. 
		NOTE: Most surgical manipulation should affect the bladder. Retract or pull the bladder caudally with a vascular&#160;clamp (see Table of Materials) &#160;to obtain a better view of excavatio vesicouterina.</li>
		<li>Cauterize and sever the descending uterine vessels at the level of the ureter as distal to the cervix as possible. 
		NOTE: Maintain microcirculation around the vagina and cervix as much as possible during the division.</li>
		<li>Separate the cervical/vaginal portion of the future graft from the rectal attachment and the paravaginal and paracervical ligaments. 
		NOTE: Avoid any cauterization on the graft vagina.</li>
		<li>Carefully dissect the vagina via diathermy&#160;around 2-3 mm caudal of the cervix. 
		NOTE: No villi (cervix) is visible inside the vaginal lumen.</li>
		<li>Locate both the uterine artery and vein at their origins. Ligate (8/0 polyamide; see Table of Materials), cauterize, and sever the gluteal vessels and all vessels caudal of the uterine vessels. 
		NOTE: Direct ligation of the common iliac vena caudal to the uterine vena is usually possible.</li>
		<li>By blunt dissection, free&#160;the common iliac vessels from each other, from the bifurcation of the aorta and the vena cava down to the division of the uterine vessels. 
		NOTE: One can gain better surgical access to the area by removing one or two adjacent lymph nodes.</li>
		<li>Excise the right uterine horn 3 mm from the Fallopian tube, after cauterizing the utero-ovarian pedicle at the same level. This enables anastomosis of the graft uterine horn to the upper part of the recipient uterine horn.</li>
		<li>Place ligatures (8/0 polyamide) directly around the right common iliac artery and vein, proximal to the aortic and caval bifurcations. Make a small incision (0.5-1 mm) into the right common iliac artery adjacent to the bifurcation, and insert a bent, blunted 30 G needle or a straight, blunted 25 G needle into the lumen (for flushing). Secure it with a ligature (6/0 polyamide). 
		NOTE: A further option is additional securing with a bulldog clamp to avoid displacement of the needle and/or vessel.</li>
		<li>Dissect the common iliac vein caudally of the ligature at the right common iliac vein to enable outflow during flushing.</li>
	</ol>
	</li>
	<li>Flush the graft following the steps below.
	<ol>
		<li>Flush the uterus manually using 3 mL syringes with approximately 9 mL of cold Ringer solution (RHX: Ringer supplemented with 50 IU/mL heparin and 0.4 mg/mL xylazine) at a flowrate of 6 mL/min. Flush again with 6 mL of organ preservation solution supplemented with heparin (50 IU/mL) and xylazine (0.4 mg/mL) (see Table of Materials). 
		&#8203;NOTE: Avoid high flushing pressure and ensure proper needle placement.</li>
		<li>Remove the transplant when the uterine tissue has turned pale. Cut the common iliac artery caudally of the ligature at the bifurcation of the abdominal aorta.</li>
	</ol>
	</li>
	<li>Place the transplant into chilled organ preservation solution (4 &#176;C) for back table preparation and storage before transplantation.</li>
	<li>After removing the graft, euthanize the animal by first turning the isoflurane setting to maximum and then inducing bilateral pneumothorax followed by exsanguination17.</li>
</ol>
2. Syngeneic uterus transplantation
NOTE: For details on the procedure, please see the previously published reports12,13,15.
<ol>
	<li>Induce anesthesia and prepare the animal as mentioned in step 1.1.
	<ol>
		<li>Administer effective analgesia (as described in step 1.1.1) and 200 IU/kg high molecular weight heparin 30 min before surgery.</li>
	</ol>
	</li>
	<li>Perform median laparotomy.
	<ol>
		<li>Open the abdomen via a 6-8 cm long midline incision starting 0.5 cm below the xiphisternum toward the hypogastrium. Use a no. 10 scalpel for the skin incision and small sharp scissors for the linea alba incision. Do not damage the liver and the bladder.</li>
		<li>Move the small intestines outside the abdominal cavity using cotton swabs, wrap them with a sterile moistened gauze, and cover them with a sterile plastic bag for better insulation.</li>
		<li>Insert retractors or clips at the left and right abdominal wall folders to keep the peritoneal muscle aside and the abdomen open, to obtain optimal access and visibility of the uterus and associated vessels. Fix the clips/retractors with tapes.</li>
		<li>Apply prewarmed saline to keep the surgical area and the intestines moist and avoid drying of the viscera.</li>
	</ol>
	</li>
	<li>Perform a hysterectomy with dissection and mobilization of the upper third of the vagina from the rectum and the bladder.
	<ol>
		<li>Cauterise the microvasculature around the uterus, cervix, and vagina. Cut and separate the uterus from the surrounding structures close to the organ to protect the microcirculation of the uterine sinister.</li>
		<li>Remove fat tissue from the surroundings.</li>
		<li>Amputate the left horn by cauterization. On the right side, preserve a 7-8 mm segment of the upper part of the uterus for later anastomosis to the uterine graft.</li>
	</ol>
	</li>
	<li>Perform uterus transplantation.
	<ol>
		<li>Mobilize and separate the right common iliac vessels, from the origin of the uterine vessels up to the aortic/caval bifurcation.</li>
		<li>Position the graft in the abdominal cavity. Wrap the graft in a gauze soaked in cold organ preservation solution. 
		NOTE: The graft needs to be kept cold during anastomosis.</li>
		<li>Place atraumatic vascular clamps onto the right common iliac vein at each side, framing the future anastomosis site. 
		NOTE: Lower the anesthesia to 1-1.5 vol% isoflurane to adapt to the sudden decrease in cardiac preloading and the resulting hypotension.</li>
		<li>Cut a slit slightly larger than the opening of the graft vein into the common iliac vein.</li>
		<li>Position the graft vein.</li>
		<li>Place one stay suture (10/0 polyamide; see Table of Materials) into each corner of the slit on the right common iliac vein. 
		NOTE: Keep the suture knot at the caudal corner loose for better adjustment and to prevent purse-string effects.</li>
		<li>Regularly flush the anastomosis area with cooled RHX during the procedure to prevent thromboses.</li>
		<li>Anastomose one side of the graft vein to the recipient&#39;s vein with six to eight loops of a continuous suture (Figure 1). 
		NOTE: Start with the cranial stay suture (10/0 polyamide) and first anastomose the inbound part of the vessels.</li>
		<li>Anastomose the other side of the vessel in the same manner, this time starting from the outside.</li>
		<li>Tie a knot at the cranial stay suture, and then one at the caudal stay suture (10/0 polyamide), after finishing the anastomoses at both sides. 
		NOTE: Tighten the continuous sutures only as much as necessary to prevent purse-string effects.</li>
		<li>Place atraumatic vascular clamps onto the right common iliac artery at each side, framing the future anastomosis site.</li>
		<li>Perform the arterial anastomosis (right common iliac arteria [RCIA] via 8-10 loops using interrupted sutures (10/0 polyamide). 
		NOTE: Interrupted sutures are easier to control than continuous ones (optional with the &#34;fish-mouth&#34; technique)18. Constant flushing of the anastomosis area with cooled RHX during the procedure helps to prevent thromboses. When using continuous sutures, perform this step analogous to venous anastomosis.</li>
	</ol>
	</li>
	<li>Perform graft reperfusion.
	<ol>
		<li>When both anastomosis sites appear patent and any bleeding is stopped, release the vascular clamps on the graft vessels (Figure 2).</li>
		<li>Inspect the graft for signs of reperfusion, such as reddening, filling of the vein, or pulsation in the graft artery.</li>
		<li>Connect the vaginal cuff of the transplant to the vaginal vault of the recipient by using six to seven intraluminal (6/0 polyglactin) interrupted sutures. 
		NOTE: Start with a single suture at the 12 o&#39;clock position first, and place the next ones at the 10 and 1 o&#39;clock positions. The two sutures at the 9 and 3 o&#39;clock positions should be tied after the sutures in the front row19,20.</li>
		<li>Anastomose the graft uterine horn end-to-end to the remaining cranial uterine segment of the recipient uterus by using five to seven interrupted sutures (7/0 polyamide). 
		NOTE: Do not stitch through the lumen.</li>
	</ol>
	</li>
	<li>Close the abdomen with a continuous suture. Use 4/0 polyglactin for suturing the muscle layer, and 6/0 polyamide or surgical wound clips for the skin.</li>
	<li>Let the animal recover in a warmed cage once the transplant is completed. Remain with the animal until it has regained sternal recumbency ability, and maintain single housing until its full recovery. Provide psotopertaive analgesia treatment by subcutaneously&#160;administering buprenorphine (0.05 mg/kg) and suitable NSAID, however not before 4-8 hours after the first dose of anesthesia. Provide continuous with&#160;buprenorphine via drinking water (1 mg/kg, oral, 5 mL buprenorphine in 160 mL drinking water (0.3 mg/mL)) for three days after surgery.</li>
	<li>The skin&#160;suture is removed 10-14 days post&#160;suregery.</li>
</ol>

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<li>Jakubauskiene, L., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Relaxin+and+erythropoietin+significantly+reduce+uterine+tissue+damage+during+experimental+ischemia-reperfusion+injury%5BTitle%5D)+AND+%22International+Journal+of+Molecular+Sciences%22%5BJournal%5D)">Relaxin and erythropoietin significantly reduce uterine tissue damage during experimental ischemia-reperfusion injury.</a> International Journal of Molecular Sciences. 23 (13), 7120(2022).</li>
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</ol>

The authors declare that they have no competing interests.

The protocol presented here offers detailed instructions for the surgical approach behind uterus transplantation in rats. The protocol has been optimized to increase the odds of live births following UTx and subsequent mating. The original protocol has been taken over from the Br&#228;nnstr&#246;m group12,13, inspired by the mouse work of Akouri et al.10, and modified based on the authors&#39; experiences over the past years. As such, the modifications were driven by true learning curves reflecting the development from failed transplants to reproducible outcomes.
Key modifications that are likely decisive were: (1) the addition of xylazine to the flushing solution during organ retrieval and anastomosis, which stimulated vasodilation leading to reduced thrombotic risks. (2) The use of interrupted sutures for arterial end-to-side anastomosis. Interrupted sutures not only increase surgical control over patency and avoid purse-string effects, but also enable subsequent widening of the anastomotic area to increase the blood flow through the arteria uterina. (3) The application of interrupted and non-absorbable sutures for the uterus horn anastomosis with stitching only within the layer of the perimetrium; this modification prevents constriction of the uterus wall, which may hinder later fertilization. (4) Using absorbable suture material for vaginal anastomosis reduces the risk of vaginal stenosis and thereby increases the chances of live births. (5) Performing any surgical manipulation as far away from the vagina and cervix as possible is a crucial step to avoid vaginal scarring and to increase the chances of live births. (6) The direct ligation of the right common iliac vein and artery instead of ligating the vena cava, abdominal aorta, and the left common iliac vein artery. Direct ligation caudal to the uterine vena is, in most cases, feasible; the direct ligation simplifies organ procurement and shortens surgical times without negatively impacting transplant outcomes. (7) Lowering the isoflurane concentration to 1%-1.5 vol% right after the clamping of the common iliac vena (step 2.4.3) is likewise crucial, as otherwise cardiac death may occur.
The above points are the main elements that distinguish the present protocol from the perhaps most widely used approach described by the Swedish group10,12,13. These measures benefitted the ultimate endpoint of UTx, the live birth, as evinced by the group comparison (original vs. modified protocol; see Table 1). Clearly, the surgical learning curve over time has likewise added to better outcomes; however, its contribution is difficult to estimate.
In addition to a meticulous surgical technique, postoperative recipient care is likewise crucial for the final outcome. Postoperative treatments follow the respective local protocol; the instructions here recommend 0.05 mg/kg buprenorphine after the recipient wakes up from anesthesia, and during the first postoperative day if presenting with pain as defined by our scoring assessment. The scoring criteria are based on behavior (activity, respiration, coat, posture, wound) and body weight. Immediately after surgery, positive signs of recovery are a return of pink coloring of the extremities and ears, as well as a return of red coloring of the eyes. Given the length of the transplant procedure, regular monitoring during the first 3 hours after transplantation is critical. If assessment criteria are not met during these 3 hours, the animal is euthanized to avoid further suffering. Monitoring is continued three times a day for the first 3 postoperative days. If the animal does not fully recover during the first 2 days, it is euthanized. Depending on its condition, the animal remains in a single housing for 2-3 days post-surgery.
Sterile conditions during surgery are yet another aspect relevant to successful outcomes. A&#160;strict hygiene routine is mandatory to minimize infection risks. When entering the animal facility, hands must be thoroughly washed before getting dressed in a clean overall, a face mask, gloves, and a cap. The entire surgical area is disinfected with a special swab&#160;containing benzyl-C12-18-alkyldimethylammonium chloride, didecyldimethylammonium chloride, and glutaraldehyde. Additional disinfection with 70% ethanol follows. Before each surgical manipulation, gloves are briefly cleaned with 70% ethanol, and the area surrounding the surgical site is covered with a sterile drape. These measures greatly help to minimize infection risks.
One issue with progeny success is the difficulty in monitoring actual pregnancy. Bodyweight increases may be indicative of pregnancy, but can also be due to infanticide and generally do not reliably reflect the state of the female. For example, only modest weight changes overall were apparent for group 1 during monitoring. Animals 3 and 4 displayed similar weight increases over the two mating cycles, yet only animal 4 was pregnant upon relaparotomy (Figure 3 and Figure 4). Moreover, weight loss after the first mating cycle with absent birth may point to pregnancy absorption. Although it remains unclear why tissue absorption results in weight loss, this phenomenon has been repeatedly observed in rat UTx recipients12. Vaginal plug formation post successful mating (Figure 11) likewise was often absent and did not show a correlation to later body weight changes. Given the little value in marking pregnancy, body weight monitoring frequency was reduced for group 2 to minimize animal stress exposure. A further consequence of inconsistent plug presence and weight changes was an unreliable definition of the conception time. Due dates for births have thus been calculated using the 2nd day post male exposure as the conception day. Interestingly, body weight changes in group 2 correlated relatively well with the course of pregnancy, with delivery being marked by a clear drop in weight (Figure 6).
Comparing body weight changes (Figure 6) of animals 1, 2, and 3 (pregnant) with animals 5 and 6 (never pregnant) nicely illustrates the effects of pregnancy and live birth versus unsuccessful mating. The reason behind the different body weight behavior in group 2 compared to group 1 remains unclear. However, weight changes may be more reliable simply due to the higher number of full-term pregnancies in group 2. The low pregnancy rate in group 1 may possibly relate to vaginal stenosis, which was overrepresented relative to group 2 (Table 3) and is a cause of hampered conception. The reduced stenotic incidence in group 2 may be due to the use of non-absorbable instead of absorbable sutures for vaginal anastomosis, likely along with improved surgical manipulation during vaginal dissection in this group.
Unlike the Swedish approach10,12, the protocol described here uses IGL-1 (Institute Georges Lopez) solution for graft storage instead of Ringer or other solutions. While the exact impact of IGL-1 solution on UTx outcomes is unknown, various reports suggest overall benefits in experimental transplantation for this solution22,23,24,25. Lastly, a few protocol steps usually performed for UTx12 did not affect outcomes if skipped. These include the clipping at the left or right uterus horn and the postoperative heparin substitution, which the authors deem optional.
One limitation inherent to experimental UTx is also valid for the presented protocol. The procedure is time-consuming, and organ retrieval followed by transplantation can take up to 6 hours. Therefore, rat UTx requires full focus for several hours in order to perform each step flawlessly. The latter remains the most important aspect of success. Such attention can be accomplished provided the microsurgical skills are fully developed and continuously practiced. Likewise, patience and stamina are key traits to avoid mistakes during the whole procedure. Another consequence of the lengthy procedure is that only one animal can be operated on per day. Careful planning and focus on key questions help to develop a productive research design. In general, it is advised to achieve a graft survival above 70% before committing to the actual experimental series to ensure that findings will be sufficiently robust. Finally, one may wish to reconsider live birth as an endpoint. The high incidence of vaginal stenosis and pregnancy resorption12 markedly increase the number of UTxs required to obtain meaningful results. Pending on the experimental question, other endpoints, such as graft survival, may be more effective.
Despite its increasing application, UTx remains an experimental application also in the clinic. Besides the need for improved surgical approaches and immunosuppressive strategies, the use of grafts from deceased donors, in particular, is an area requiring additional research. Strategies to mitigate ischemic injury are not established, but would be most welcome for the expansion of potential donor pools. Indeed, the impact of ischemia on uteri is ill-researched, with applied knowledge relying on findings from other organs26,27. Rat UTx offers a means to explore ischemic injury in controlled settings while using approaches tailored to the clinic28,29. Of note, live donation is associated with certain risks; donors have a high rate of surgical complications, such as urinary tract and bowel trauma. Accordingly, the demand for dead donations and its research is growing5,30.
Many additional questions exist where rat UTx could be informative. For example, the rat system offers the opportunity to identify and/or validate biological markers that could be used to noninvasively monitor the course of uterus transplantations in the clinic. Recent societal developments may create new questions where the rat model may be of use. UTx is now being advocated also for transgender individuals, with the male anatomy asking for adapted surgical approaches31.
In conclusion, a new protocol of rat UTx based on existing rodent work and modified by hands-on experience is presented. The modified protocol has a high likelihood of rat UTx resulting in live births-provided sufficient microsurgical skills and practice. UTx is perhaps the most complex of the major transplantation procedures. The described instructions shall add to a common protocol, which is currently lacking but needed&#160;to establish the demanding procedure of rat UTx within the research community.

Uterine transplantation (UTx) is a novel treatment for absolute uterine factor infertility (AUFI). AUFI results from an absence (congenital or acquired) or malformation of the uterus and affects 3%-5% of women worldwide1. Ethical, legal, or religious reasons rule out adoption or surrogacy for many women who have a desire for motherhood but suffer from AUFI2. For these women, UTx remains the only option to start their own family. UTx has been applied in the clinic, albeit with mixed success; the procedure is technically challenging and requires steady improvement for its clinical establishment.
In 2014, the first transplantation of a uterus from a live donor (LD)-resulting in successful pregnancy-was performed by the pioneering Swedish group of Br&#228;nnstr&#246;m3. The first birth following UTx from a deceased donor (DD) was reported in 2016 in Brazil4. By 2021, more than 80 UTxs have been performed worldwide, however with a success rate of about 50% and with grafts coming from LD for the majority1.
Although not life-saving, UTx is an increasingly popular procedure to fulfill the desires for own progeny. As such, the demand for grafts is mounting, placing DD donation into a future focus. However, DD donation is complicated due to considerably longer cold (and in the case of cardiac death, also warm) ischemic exposures, elevating the risks of graft dysfunction and rejection5,6. Surgical technique, demanding compatibility matching, and associated immunosuppression remain critical issues regarding UTx outcomes7.
To manage the above risks in the clinic, appropriate animal models for the exploration of ischemia and immunosuppression are needed. The most clinically relevant endpoint for animal models remains successful birth; to date, pregnancies following experimental UTx have been achieved in mice, rats, sheep, rabbits, and cynomolgus monkeys8. While larger animals are predestined for acquiring and optimizing surgical techniques, rodents come with the distinct advantage of short gestation periods. Therefore, rodent models are superior regarding practical, financial, and ethical considerations9. However, the main challenge of UTx in mice is the small anatomy, with the highly demanding surgery tied to the low reproducibility of murine UTx10. By contrast, rats are surgically more accessible and retain the advantages of short gestation times. As such, the rat has become the model of choice for UTx9. Wranning et al. introduced the rat model of orthotopic UTx in 2008, and using this model, the first live birth following UTx and natural mating has been reported11,12,13. Subsequent studies have had critical contributions to the implementation of UTx in humans9.
Nonetheless, UTx remains challenging in rats, and only a few groups as of yet have mastered this surgical technique. One relevant obstacle to the spread of rat UTx among researchers is the lack of a precise description of the individual microsurgical steps, the pitfalls, and the according measures for troubleshooting14. This protocol aims to provide a detailed guide for this highly complex microsurgical procedure to facilitate the implementation of this animal model into future research.

<table><tbody><tr><td>Angled to Side Scissor 5 mm</td><td>F.S.T</td><td>15008-08</td><td></td></tr><tr><td>Big Paper Clip</td><td>No specific</td><td></td><td>Used as retractor</td></tr><tr><td>Blunt Bend Needle G30</td><td>Unimed S.A.</td><td></td><td></td></tr><tr><td>Bupivacain 0.5%</td><td>Sintetica</td><td></td><td></td></tr><tr><td>Buprenorphine 0.3 mg/mL</td><td>Temgesic</td><td></td><td></td></tr><tr><td>Dosiernadel G25</td><td>H.SIGRIST&amp;amp; PARTNER AG</td><td></td><td></td></tr><tr><td>Dumont #5SF Forceps</td><td>F.S.T</td><td>11252-00</td><td></td></tr><tr><td>Ethilon 10/0</td><td>Ethicon</td><td>2810G</td><td></td></tr><tr><td>Ethilon 6/0</td><td>Ethicon</td><td>667H</td><td></td></tr><tr><td>Ethilon 7/0</td><td>Ethicon</td><td>EH7446H</td><td></td></tr><tr><td>Ethilon 8/0</td><td>Ethicon</td><td>2808G</td><td></td></tr><tr><td>Femal Brown Norway Rats (150-170 g)</td><td>Janvier</td><td></td><td></td></tr><tr><td>Femal Lewis Rats (150-170 g)</td><td>Charles River Deutschland</td><td></td><td></td></tr><tr><td>Fine Scissors - Sharp</td><td>F.S.T</td><td>14060-09</td><td>Any other small scissor works too</td></tr><tr><td>Halsey Micro Needle Holder</td><td>F.S.T</td><td>12500-12</td><td>Any other small needholder works too</td></tr><tr><td>Heparin Natrium 25000 I.E./ 5 mL</td><td>B. Braun</td><td></td><td></td></tr><tr><td>Institute Georges Lopez Perfusion Solution (IGL)</td><td>Institute Georges Lopez</td><td></td><td>Organ preservation solution&amp;nbsp;&amp;nbsp;</td></tr><tr><td>Male Lewis Rats (300-320 g)</td><td>Charles River Deutschland</td><td></td><td></td></tr><tr><td>Micro Serrefines 13 mm</td><td>F.S.T</td><td>18055-04&amp;nbsp;&amp;nbsp;</td><td></td></tr><tr><td>Micro Serrefines 16 mm gebogen</td><td>F.S.T</td><td>18055-06</td><td></td></tr><tr><td>Micro-Serrefine Clamp Applicator with Lock&amp;nbsp;&amp;nbsp;</td><td>F.S.T</td><td>18056-14&amp;nbsp;&amp;nbsp;</td><td></td></tr><tr><td>M&amp;ouml;lnlyncke Op Towel</td><td>M&amp;ouml;lnlyncke</td><td>800300</td><td>Sterile drape</td></tr><tr><td>NaCl 0.9%</td><td>B.Braun</td><td></td><td></td></tr><tr><td>Octenisept</td><td>Sch&amp;uuml;lke</td><td></td><td></td></tr><tr><td>Paper Tape</td><td>Tesa</td><td></td><td>For fixing the animal</td></tr><tr><td>Philips Avent Schneller Flaschenw&amp;auml;rmer SCF358/02</td><td>Philips</td><td>12824216</td><td></td></tr><tr><td>Ringerfundin</td><td>B.Braun</td><td></td><td></td></tr><tr><td>Rompun 2%</td><td>Bayer</td><td></td><td>Xylazine</td></tr><tr><td>Round Handled Needle Holders</td><td>F.S.T</td><td>12075-12</td><td></td></tr><tr><td>Round Handled Needle Holders</td><td>F.S.T</td><td>12075-12</td><td></td></tr><tr><td>S&amp;amp;T Vessel Dilating Forceps - Angled 45&amp;deg;</td><td>F.S.T</td><td>00276-13</td><td></td></tr><tr><td>Sacryl Naht</td><td>KRUUSE</td><td>152575</td><td></td></tr><tr><td>Scapel No 10</td><td>Swann Morton</td><td>201</td><td></td></tr><tr><td>Small Histo-Container</td><td></td><td></td><td>Any small histo-container works fine-for coldstorage of the graft</td></tr><tr><td>Small Plastik Bags</td><td></td><td></td><td>Any transparant plastic bags are fine</td></tr><tr><td>Steril Cotton swab</td><td>Lohmann-Rauscher</td><td></td><td>Any steril cotton swab is fine</td></tr><tr><td>Sterile Gauze</td><td>Lohmann-Rauscher</td><td></td><td>Any steril gauze is fine</td></tr><tr><td>Straight Scissor 8mm</td><td>F.S.T</td><td>15024-10</td><td></td></tr><tr><td>Surgical microscope &amp;ndash; SZX9</td><td>Olympus</td><td>OLY-SZX9-B</td><td></td></tr><tr><td>Sutter Non Stick GLISS 0.4 mm</td><td>Sutter</td><td>78 01 69 SLS</td><td></td></tr><tr><td>Suture Tying Forceps&amp;nbsp;</td><td>F.S.T</td><td>00272-13</td><td></td></tr><tr><td>ThermoLux warming mat</td><td>ThermoLux</td><td></td><td></td></tr><tr><td>Tissue Forceps for Skin</td><td></td><td></td><td>Any tissue forceps are fine</td></tr><tr><td>Vesseldilatator Forceps</td><td>F.S.T</td><td>00125-11</td><td></td></tr><tr><td>Vicryl&amp;nbsp; plus 4/0</td><td>Ethicon</td><td>VCP292H</td><td></td></tr></tbody></table>

surgical approach, challenges, and resolutions for uterus transplantation in rats

Uterine transplantation (UTx) is a new approach for treating women with absolute uterine factor infertility (AUFI). An estimated 3%-5% of women suffer from AUFI. These women were deprived of the option to have children until the advent of UTx. The clinical application of UTx was driven by experimental studies in animals, and the first successful UTx was achieved in rats. Given their physiological, immunological, genetic, and reproductive characteristics, rats are a suitable model system for such transplants. In particular, their short gestation period is a clear advantage, as the usual endpoint of experimental UTx is successful pregnancy with live birth. The biggest challenge for rat models remains the small anatomy, which requires advanced microsurgical skills and experience. Although UTx has led to pregnancy in the clinic, the procedure is not established and requires continuous experimental optimization. Here, a detailed protocol is presented, including essential troubleshooting for rat UTx, which is expected to make the entire procedure easier to grasp for those without experience in this type of microsurgery.

Results from two groups of rats are presented. UTx was carried out before (group 1, n = 8) and after (group 2, n = 8) protocol adjustment (Table 1)&#160;to demonstrate the effects of our modifications (please see the Discussion for an explanation of our modifications)12,15,21.
The outcome of rat UTx is associated with three key phases. The first phase is successful recovery from UTx. Usually, recipients should recover within the first 2 postoperative days. The second phase relates to the health status of the graft 2 weeks after surgery, which decides the inclusion into the mating process (Table 2). The third phase involves spontaneous mating followed by successful birth as evidence of fertility.
All animals from both groups had an uneventful recovery from surgery. During the second phase, four animals were excluded from group 1 and two from group 2. The exclusion was due to graft thrombosis and abscess (n = 4 for group 1, n = 2 for group 2), and narrowed/malformed utero-uterine anastomosis (additionally for n = 1, group 1) on examination at relaparotomy (Table 2). Relaparotomy (along the scar of the original laparotomy) was performed for all females 2 weeks post transplantation, as the physical appearance of the animals had little value as an indicator of graft health. Overall, the 2 week graft survival rate was 50% and 75% for groups 1 and 2, respectively (Table 3).
During phase 3, four females from group 1 were matched for mating with Lewis males, around 9 weeks after UTx. Two females showed signs of pregnancy (bodyweight increase, Figure 3; nesting behavior); however, no live birth was observed. After two mating cycles of three females, pup body parts (bone and tissue) were found in one female rat (&#176;1). Histological examination of hematoxylin and eosin stained tissue sections by a veterinary pathologist revealed these pups developed up to parturition (Figures 4 and Figure 5).
Six females from group 2 were mated with Lewis males (see Figure 6 for bodyweight changes). Three of the six rats (two Lewis and one Brown Norway rat) gave birth to pups, while another two showed signs of pregnancy. The first litter of the female Lewis rat (#1) consisted of two pups (Figure 7). Shortly after birth, the Lewis female became pregnant again; however, only two of three pups survived postnatally (Figure 8). One likely explanation for the one death is infanticide, as it occurs even with healthy pups under conditions of postpartum stress. Likewise, the Brown Norway female (#2) gave birth twice, each time to four pups per mating cycle (Figure 9). The largest litter number of group 2 was delivered by another Lewis female (#3), with seven pups after the first mating cycle. All surviving pups displayed normal development (Figure 10).
Overall, the adaptation of the protocol increased 2 week graft survival from 50% to 75%. Five of six females became pregnant, compared to two out of four from group 1. Likewise, three of six of females gave birth to live pups compared to zero out of four of the group 1 females. In conclusion, the adapted protocol improved both direct surgical outcomes and the rate of successful live births following UTx (Table 4 and Figure 6).
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/64757/64757fig01.jpg" /> 
Figure 1: Anastomosis of the graft and the recipient&#39;s vein. The right common iliac vein (RCIV) of the graft is connected to the RCIV of the recipient via end-to-side anastomosis (step 2.4). RCIA = right common iliac artery <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/64757/64757fig01large.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/64757/64757fig02.jpg" /> 
Figure 2: Anastomosis of the graft and the recipient artery. The right common iliac artery (RCIA) is connected to the RCIA of the recipient via end-to-side anastomosis (step 2.4). After opening both vascular clamps (2.5), the artery should be fully perfused in the absence of outer bleeding. Black arrow: graft RCIA; red arrow: graft RCIV. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/64757/64757fig02large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 3" class="xfigimg" src="/files/ftp_upload/64757/64757fig03.jpg" /> 
Figure 3: Bodyweight changes after mating of females in group 1. Body weight monitoring of the four group 1 females who displayed an intact graft upon relaparotomy. One animal (&#176;1) was euthanized during Caesarean sectioning to inspect the uterus. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/64757/64757fig03large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 4" class="xfigimg" src="/files/ftp_upload/64757/64757fig04.jpg" /> 
Figure 4: Histological examination of the uterus of the group 1 rat with dead pups. Body parts of pups were found in rat &#176;1. Examination revealed a vital and dilated uterus, suggesting the pups developed normally but could not be delivered. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/64757/64757fig04large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 5" class="xfigimg" src="/files/ftp_upload/64757/64757fig05.jpg" /> 
Figure 5: Pup body parts inside the uterus of group 1 rat &#176;1. The developmental stage of the bones was consistent with full-term pups. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/64757/64757fig05large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 6" class="xfigimg" src="/files/ftp_upload/64757/64757fig06.jpg" /> 
Figure 6: Body weight monitoring of the six group 2 females who displayed an intact graft upon relaparotomy. Green stars mark individual birth events. Hashtag numbers refer to the individual females. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/64757/64757fig06large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 7" class="xfigimg" src="/files/ftp_upload/64757/64757fig07.jpg" /> 
Figure 7: The first live birth after rat UTx following the modified protocol. Two newborn rats and their mother (head to the right; Lewis female #1, group 2). <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/64757/64757fig07large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 8" class="xfigimg" src="/files/ftp_upload/64757/64757fig08.jpg" /> 
Figure 8: The second litter after the second mating cycle post rat UTx following the modified protocol. Female #1 (group 2) gave birth to three pups, two of which survived. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/64757/64757fig08large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 9" class="xfigimg" src="/files/ftp_upload/64757/64757fig09.jpg" /> 
Figure 9: The first litter of the Brown Norway rat after UTx following the modified protocol. After the first mating cycle, the Brown Norway rat (#2, group 2) gave birth to four pups, followed by another four after the second mating cycle. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/64757/64757fig09large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 10" class="xfigimg" src="/files/ftp_upload/64757/64757fig10.jpg" /> 
Figure 10: Development of pups. All surviving pups displayed normal development at 3 weeks of age. One representative example is shown. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/64757/64757fig10large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 11" class="xfigimg" src="/files/ftp_upload/64757/64757fig11.jpg" /> 
Figure 11: Vaginal plug formation following successful mating. White plugs should form to cover the vagina after fertilization to prevent further mating. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/64757/64757fig11large.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></td>
			<td>Group 1</td>
			<td>Group 2</td>
			<td>Syngeneic UTx in rats</td>
			<td>Allogeneic UTx in rats</td>
		</tr>
		<tr>
			<td></td>
			<td>before modification</td>
			<td>after modification</td>
			<td>see ref. 12</td>
			<td>see ref. 15</td>
		</tr>
		<tr>
			<td>Total number of animals&#160;</td>
			<td>8</td>
			<td>8</td>
			<td>27</td>
			<td>14</td>
		</tr>
		<tr>
			<td>Two-weeks graft survival</td>
			<td>4/8 (50%)</td>
			<td>6/8 (75%)</td>
			<td>19/27 (70%)</td>
			<td>9/14 (64%)</td>
		</tr>
		<tr>
			<td>Number of mated females</td>
			<td>4/4 (100%)</td>
			<td>6/6 (100%)</td>
			<td>17/19 (89%)</td>
			<td>9/9 (100%)</td>
		</tr>
		<tr>
			<td>Full term pregnancy&#160;</td>
			<td>1/4 (25%)</td>
			<td>5/6 (83%)</td>
			<td>11/17(65%)</td>
			<td>5/9 (56%)</td>
		</tr>
		<tr>
			<td>Successfully delivered litter</td>
			<td>0</td>
			<td>5a/td&#62;</td>
			<td>1</td>
			<td>5</td>
		</tr>
		<tr>
			<td>Total number of living pups</td>
			<td>0</td>
			<td>20b</td>
			<td>3c</td>
			<td>25d</td>
		</tr>
		<tr>
			<td>Pregnancy to term w/o live birth</td>
			<td>1</td>
			<td>1e</td>
			<td>10</td>
			<td>2</td>
		</tr>
	</tbody>
</table>
Table 1: Outcome comparison of the modified and unmodified protocol for rat UTx. a: Two pregnancies by the same rat; b: Including the infanticide pup; c: Median; d: Sum of the median/delivery; e: Dead pups in resorption stage after three cycles of mating.
<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>Surgical settings</td>
			<td>Group 1 (n = 8)</td>
			<td>Group 2 (n = 8)</td>
		</tr>
		<tr>
			<td>Project stage</td>
			<td>initial stage&#160;</td>
			<td>late stage&#160;</td>
		</tr>
		<tr>
			<td>Cold storage time</td>
			<td>2-3 h</td>
			<td>2-3 h</td>
		</tr>
		<tr>
			<td>Flushing solution during anastomosis</td>
			<td>RH</td>
			<td>RHX</td>
		</tr>
		<tr>
			<td>Vaginal anastomosis</td>
			<td>6/0 Ethilon</td>
			<td>6/0 Vicryl</td>
		</tr>
		<tr>
			<td>Uterus horn anastomosis</td>
			<td>partially continuous suture</td>
			<td>interrupted suture</td>
		</tr>
		<tr>
			<td>Arteria anastomosis</td>
			<td>continuous suture</td>
			<td>Interrupted and continuous</td>
		</tr>
		<tr>
			<td>Micro-vascularization around vagina and cervix</td>
			<td>cauterization close to the vaginal/cervical tissue</td>
			<td>cauterization more distally&#160;</td>
		</tr>
	</tbody>
</table>
Table 2: Surgical settings group 1 versus group 2.
<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>Exclusion criteria#</td>
		</tr>
		<tr>
			<td>Signs of thrombosis (particularly around anastomoses)</td>
		</tr>
		<tr>
			<td>Major adhesion&#160;</td>
		</tr>
		<tr>
			<td>Constricted uterus</td>
		</tr>
		<tr>
			<td>Signs of infection</td>
		</tr>
		<tr>
			<td>Graft necrosis</td>
		</tr>
	</tbody>
</table>
Table 3: Exclusion criteria for mating. #Applied at relaparotomy 2 weeks post UTx.
<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td></td>
			<td>n (group 1)</td>
			<td>n (group 2)</td>
		</tr>
		<tr>
			<td>Animals&#160;</td>
			<td>8</td>
			<td>8</td>
		</tr>
		<tr>
			<td>Healthy graft after 2 weeks</td>
			<td>4</td>
			<td>6</td>
		</tr>
		<tr>
			<td>Mated</td>
			<td>4</td>
			<td>6</td>
		</tr>
		<tr>
			<td>Full term pregnancy&#160;</td>
			<td>1</td>
			<td>5</td>
		</tr>
		<tr>
			<td>Successfully delivered litter</td>
			<td>0</td>
			<td>5a</td>
		</tr>
		<tr>
			<td>Total number of living pups</td>
			<td>0</td>
			<td>20b</td>
		</tr>
		<tr>
			<td>Pregnancy to term, not live birth</td>
			<td>1</td>
			<td>0</td>
		</tr>
	</tbody>
</table>
Table 4: Outcomes of group 1 versus group 2. a: Two consecutive pregnancies in the same rat; b: Including the pup killed by infanticide.

Watch this Scientific Journal Video about Surgical Approach, Challenges, and Resolutions for Uterus Transplantation in Rats at JoVE.com

Surgical Approach, Challenges, and Resolutions for Uterus Transplantation in Rats

The present protocol describes all the essential steps for successful uterine transplantation (UTx) in rats. The rat model has proven suitable to promote the clinical implementation of UTx; however, rat UTx is a highly complex procedure requiring careful instructions.

Uterine transplantation (UTx) is a new approach for treating women with absolute uterine factor infertility (AUFI). An estimated 3%-5% of women suffer ...

surgical-approach-challenges-resolutions-for-uterus-transplantation

Nanyang Technological University

Research

JoVE Journal

Medicine

1.2K Views.  University Hospital Zürich. Rechter baarmoedertransplantatie behoort tot een van de meest complexe microchirurgische procedures die slechts door weinigen wordt beheerst. Ons protocol verschuift de transplantatie van de rechter baarmoeder van een experimenteel stadium naar een toepasbare methodiek. Ons protocol leidt tot een hoog aantal levendgeborenen na baarmoedertransplantatie.Het zal binnenkort de toepassing van rechter baarmoedertransplantatie voor daadwerkelijk onderzoek bevorderen. Geduld en uithoudingsverm...

Video: Surgical Approach, Challenges, and Resolutions for Uterus Transplantation in Rats

Mouse Model of Surgically-induced Endometriosis by Auto-transplantation of Uterine Tissue

Endometriosis is a chronic, painful disease whose etiology remains unknown. Furthermore, treatment of endometriosis can require laparoscopic removal of lesions, and/or chronic pharmaceutical management of pain and infertility symptoms. The cost associated with endometriosis has been estimated at 22 billion dollars per year in the United States1. To further our understanding of mechanisms underlying this enigmatic disease, animal models have been employed. Primates spontaneously develop endometriosis and therefore primate models most closely resemble the disease in women. Rodent models, however, are more cost effective and readily available2. The model that we describe here involves an autologous transfer of uterine tissue to the intestinal mesentery (Figure 1) and was first developed in the rat3 and later transferred to the mouse4. The goal of the autologous rodent model of surgically-induced endometriosis is to mimic the disease in women. We and others have previously shown that the altered gene expression pattern observed in endometriotic lesions from mice or rats mirrors that observed in women with the disease5,6. One advantage of performing the surgery in the mouse is that the abundance of transgenic mouse strains available can aid researchers in determining the role of specific components important in the establishment and growth of endometriosis. An alternative model in which excised human endometrial fragments are introduced to the peritoneum of immunocompromised mice is also widely used but is limited by the lack of a normal immune system which is thought to be important in endometriosis2,7. Importantly, the mouse model of surgically induced endometriosis is a versatile model that has been used to study how the immune system8, hormones9,10 and environmental factors11,12 affect endometriosis as well as the effects of endometriosis on fertility13 and pain14.

A description of the surgical induction of endometriosis in mice and rats by auto-transplantation of uterine tissue to the arterial cascade of the intestinal mesentery.

Endometriosis is a chronic, painful disease whose etiology remains unknown. Furthermore, treatment of endometriosis can require laparoscopic removal of ...

Orthotopic Liver Transplantation in Rats

Clinical progress in the field of liver transplantation has been largely supported by animal models1,2. Since the publication of the first orthotopic rat liver transplantation in 1979 by Kamada et al.3, this model has remained the gold standard despite various proposed alternative techniques4. Nevertheless, its broader use is limited by its steep learning curve5.
In this video paper, we show a simple and easy-to-establish revision of Kamada's two-cuff technique. The suprahepatic vena cava anastomosis is performed manually with a running suture, and the vena porta and infrahepatic vena cava anastomoses are performed utilizing a quick-linker cuff system6. Manufacturing the quick-linker kit is shown in a separate video paper.

We present an easy-to-establish revision of the classical two-cuff technique for orthotopic liver transplantation in rat.

Clinical progress in the field of liver transplantation has been largely supported by animal models1,2. Since the publication of the first orthotopic rat ...

Surgical Procedures for a Rat Model of Partial Orthotopic Liver Transplantation with Hepatic Arterial Reconstruction

Orthotopic liver transplantation (OLT) in rats using a whole or partial graft is an indispensable experimental model for transplantation research, such as studies on graft preservation and ischemia-reperfusion injury 1,2, immunological responses 3,4, hemodynamics 5,6, and small-for-size syndrome 7. The rat OLT is among the most difficult animal models in experimental surgery and demands advanced microsurgical skills that take a long time to learn. Consequently, the use of this model has been limited. Since the reliability and reproducibility of results are key components of the experiments in which such complex animal models are used, it is essential for surgeons who are involved in rat OLT to be trained in well-standardized and sophisticated procedures for this model.
While various techniques and modifications of OLT in rats have been reported 8 since the first model was described by Lee et al. 9 in 1973, the elimination of the hepatic arterial reconstruction 10 and the introduction of the cuff anastomosis technique by Kamada et al. 11 were a major advancement in this model, because they simplified the reconstruction procedures to a great degree. In the model by Kamada et al., the hepatic rearterialization was also eliminated. Since rats could survive without hepatic arterial flow after liver transplantation, there was considerable controversy over the value of hepatic arterialization. However, the physiological superiority of the arterialized model has been increasingly acknowledged, especially in terms of preserving the bile duct system 8,12 and the liver integrity 8,13,14.
In this article, we present detailed surgical procedures for a rat model of OLT with hepatic arterial reconstruction using a 50% partial graft after ex vivo liver resection. The reconstruction procedures for each vessel and the bile duct are performed by the following methods: a 7-0 polypropylene continuous suture for the supra- and infrahepatic vena cava; a cuff technique for the portal vein; and a stent technique for the hepatic artery and the bile duct.

Orthotopic liver transplantation in rats is an indispensable experimental model for biomedical research. Here we present our surgical procedures for orthotopic rat liver transplantation with hepatic arterial reconstruction using a 50% partial graft.

Orthotopic liver transplantation (OLT) in rats using a whole or partial graft is an indispensable experimental model for transplantation research, such as ...

Ultrasonography in Experimental Reproductive Investigations on Rats

With the development of assisted reproductive technology and the ethical limitations of research on humans, rat animal models have been widely used in reproductive medicine. In the past, the study of reproductive system development in rodents has been based on one-time histological examination of excised tissues. Recently, with the development of high-resolution transabdominal ultrasound, high-quality sonography can now be performed to evaluate the reproductive organs of rats, allowing a new method for studying the reproductive system. Images were obtained using a high-resolution ultrasonographic system. Gynecological ultrasonography was performed on 28 eight-week-old non-pregnant rats and 5 pregnant Sprague-Dawley rats. We describe how to recognize organs of the reproductive system and associated structures in typical views during different phases of the estrus cycle. Color flow Doppler was used to measure uterine artery blood flow and evaluate uterine blood flow pattern changes during different stages of pregnancy. We have demonstrated that ultrasound exploration is a useful method for evaluating changes in internal reproductive organs. Its use raises the possibility of conducting additional experiments, including medical or surgical procedures, and provides the ability to monitor sonographic changes to internal organs without sacrificing animals.

This manuscript describes the utility of ultrasound performed on female rats to design experimental models for reproductive and gynecological investigation. A step-by-step explanation of how to perform ultrasonographic evaluation is shown.

With the development of assisted reproductive technology and the ethical limitations of research on humans, rat animal models have been widely used in ...

A Rat Model of Orthotopic Liver Transplantation Using a Novel Magnetic Anastomosis Technique for Suprahepatic Vena Cava Reconstruction

The rat model of orthotopic liver transplantation (OLT) is essential for transplant research. It is a very sophisticated animal model and requires a steep learning curve. The introduction of the cuff technique for anastomosis of the portal vein (PV) and infrahepatic vena cava (IHVC) has significantly simplified the transplant procedure in rats. However, due to the short anterior wall of the recipients' suprahepatic vena cava (SHVC), the cuff technique is very difficult to use for the reconstruction of the SHVC. Most researchers in this field still use the hand-suture technique for SHVC reconstruction, which makes it the bottleneck step in rat orthotopic liver transplantation. The magnetic anastomosis technique (i.e., magnamosis) is a method of connecting two vessels using the attractive force between two magnets. Our recent study has shown that the magnetic anastomosis technique is superior to the hand-suture technique for SHVC reconstruction in rats. In this article, we show a step-by-step protocol for SHVC reconstruction in rats using the novel magnetic anastomosis technique. In this model, the reconstruction of the PV and IHVC was performed by the standard cuff technique, while the reconstruction of the bile duct (BD) was performed by a stent technique. The hepatic re-arterialization was not performed. The magnetic anastomosis technique made SHVC reconstruction much easier and significantly shortened the anphepatic phase. After a reasonable learning curve, even researchers without advanced microsurgical skills can produce reliable and reproducible results using this rat model of OLT.

The reconstruction of the suprahepatic vena cava (SHVC) remains a difficult step in rat orthotopic liver transplantation. In this article, we show a step-by-step protocol for SHVC reconstruction in rats using a novel magnetic anastomosis technique.

The rat model of orthotopic liver transplantation (OLT) is essential for transplant research. It is a very sophisticated animal model and requires a steep ...

Orthotopic Rat Kidney Transplantation: A Novel and Simplified Surgical Approach

Kidney transplantation offers increased survival rates and improved quality of life for patients with end-stage renal disease, as compared to any type of renal replacement therapy. Over the past few decades, the rat kidney transplantation model has been used to study the immunological phenomena of rejection and tolerance. This model has become an indispensable tool to test new immunomodulatory pharmaceuticals and regimens prior to proceeding with expensive preclinical large animal studies.
This protocol provides a detailed overview of how to reliably perform orthotopic kidney transplantation in rats. This protocol includes three distinctive steps that increase the probability of success: perfusion of the donor kidney by flushing through the portal vein and the use of a cuff system to anastomose the renal veins and ureters, thereby decreasing cold and warm ischemia times. Using this technique, we have achieved survival rates beyond 6 months with normal serum creatinine in animals with syngeneic or tolerant kidney transplants. Depending on the aim of the study, this model can be modified by pre- or posttransplant treatments to study the acute, chronic, cellular, or antibody-mediated rejection. It is a reproducible, reliable, and cost-effective animal model to study different aspects of kidney transplantation.

The purpose of this manuscript and protocol is to explain and demonstrate in detail the surgical procedure of orthotopic kidney transplantation in rats. This method is simplified to achieve the correct perfusion of the donor kidney and shorten the reperfusion time by using the venous and ureteral cuff anastomosis technique.

Kidney transplantation offers increased survival rates and improved quality of life for patients with end-stage renal disease, as compared to any type of ...

A Simplified Model for Heterotopic Heart Valve Transplantation in Rodents

There is an urgent clinical need for heart valve replacements that can grow in children. Heart valve transplantation is proposed as a new type of transplant with the potential to deliver durable heart valves capable of somatic growth with no requirement for anticoagulation. However, the immunobiology of heart valve transplants remains unexplored, highlighting the need for animal models to study this new type of transplant. Previous rat models for heterotopic aortic valve transplantation into the abdominal aorta have been described, though they are technically challenging and costly. For addressing this challenge, a renal subcapsular transplant model was developed in rodents as a practical and more straightforward method for studying heart valve transplant immunobiology. In this model, a single aortic valve leaflet is harvested and inserted into the renal subcapsular space. The kidney is easily accessible, and the transplanted tissue is securely contained in a subcapsular space that is well vascularized and can accommodate a variety of tissue sizes. Furthermore, because a single rat can provide three donor aortic leaflets and a single kidney can provide multiple sites for transplanted tissue, fewer rats are required for a given study. Here, the transplantation technique is described, providing a significant step forward in studying the transplant immunology of heart valve transplantation.

This protocol describes a simple and efficient method for the transplantation of aortic valve leaflets under the renal capsule to allow for the study of alloreactivity of heart valves.

There is an urgent clinical need for heart valve replacements that can grow in children. Heart valve transplantation is proposed as a new type of ...

A Rat Orthotopic Renal Transplantation Model for Renal Allograft Rejection

Renal allograft rejection limits the long-term survival of patients after renal transplantation. Rat orthotopic renal transplantation is an essential model to investigate the mechanism of renal allograft rejection in pre-clinical studies and could aid in the development of novel approaches to improve the long-term survival of renal allografts. Donor kidney implantation in rat orthotopic renal transplantation is commonly performed by end-to-side anastomosis to recipients' aorta and inferior vena cava. In this model, the donor's kidney was implanted using end-to-end anastomosis to the recipients' renal artery and renal vein. The donor's ureter was anastomosed to the recipient's bladder in an end-to-side 'tunnel' method. This model contributes to better healing of ureter-bladder anastomosis and increases the recipients' survival by avoiding interference with blood supply and venous reflux of the lower body. This model can be used to investigate the mechanisms of acute and chronic immune and pathologic rejection of renal allografts. Here, the study describes the detailed protocols of this orthotopic renal transplantation between rats.

The rat orthotopic renal transplantation model contributes to investigating the mechanism of renal allograft rejection. The current model increases the recipients' survival without interference with blood supply and venous reflux of the lower body using an end-to-end anastomosis of kidney implantation and an end-to-side "tunnel" method of ureter-bladder anastomosis.

Renal allograft rejection limits the long-term survival of patients after renal transplantation. Rat orthotopic renal transplantation is an essential ...

In Vivo Heart Transplantation to Study Endothelial-to-Mesenchymal Transition

A Neonatal Heterotopic Rat Heart Transplantation Model for the Study of Endothelial-to-Mesenchymal Transition

Endocardial fibroelastosis (EFE), defined by subendocardial tissue accumulation, has major impacts on the development of the left ventricle (LV) and precludes patients with congenital critical aortic stenosis and hypoplastic left heart syndrome (HLHS) from curative anatomical biventricular surgical repair. Surgical resection is currently the only available therapeutic option, but EFE often recurs, sometimes with an even more infiltrative growth pattern into the adjacent myocardium.
To better understand the underlying mechanisms of EFE and to explore therapeutic strategies, an animal model suitable for preclinical testing was developed. The animal model takes into consideration that EFE is a disease of the immature heart and is associated with flow disturbances, as supported by clinical observations. Thus, the heterotopic heart transplantation of neonatal rat donor hearts is the basis for this model.
A neonatal rat heart is transplanted into an adolescent rat&#39;s abdomen and connected to the recipient&#39;s infrarenal aorta and inferior vena cava. While perfusion of the coronary arteries preserves the viability of the donor heart, flow stagnation within the LV induces EFE growth in the very immature heart. The underlying mechanism of EFE formation is the transition of endocardial endothelial cells to mesenchymal cells (EndMT), which is a well-described mechanism of early embryonic development of the valves and septa but also the leading cause of fibrosis in heart failure. EFE formation can be macroscopically observed within days after transplantation. Transabdominal echocardiography is used to monitor the graft viability, contractility, and the patency of the anastomoses. Following euthanasia, the EFE tissue is harvested, and it shows the same histopathological characteristics as human EFE tissue from HLHS patients.
This in vivo model allows for studying the mechanisms of EFE development in the heart and testing treatment options to prevent this pathological tissue formation and provides the opportunity for a more generalized examination of EndMT-induced fibrosis.

Author Spotlight: A Neonatal Heterotopic Rat Heart Transplantation Model for the Study of Endothelial-to-Mesenchymal Transition  

This work presents an animal model of endothelial-to-mesenchymal transition-induced fibrosis, as seen in congenital cardiac defects such as critical aortic stenosis or hypoplastic left heart syndrome, which allows for detailed histological tissue evaluation, the identification of regulatory signaling pathways, and the testing of treatment options.

Endocardial fibroelastosis (EFE), defined by subendocardial tissue accumulation, has major impacts on the development of the left ventricle (LV) and ...

Establishing a Swine Model to Study Uterus Dynamic Preservation and Transplantation

To date, uterus transplantation is the only option for women with absolute uterine infertility, such as those with Rokitansky syndrome, to experience pregnancy and give birth. Despite the growing interest in uterus&#160;transplantation in recent years, several issues still require further research, including ischemia-reperfusion injury and its impact on graft quality and rejection. Recent literature has highlighted a thrombotic complication rate of up to 20% following uterus&#160;transplantation. This type of complication may result from hypoxia-induced endothelial cell damage, often leading to uterine graft rejection. Hypoxia is induced during static cold storage, which remains the gold standard for graft preservation in solid organ transplantation. Recently, dynamic preservation using machine perfusion has been shown to improve the long-term storage of conventional and marginal organs by reducing ischemic and hypoxic injury. In this protocol, we aim to describe every surgical step involved in porcine uterus procurement and dynamic preservation, based on both uterine pedicles, to enable the connection and initiation of the machine perfusion protocol.

A detailed and reproducible swine uterus model is described, from surgical procurement to the initiation of machine perfusion, allowing for the study of uterus preservation in transplantation.

To date, uterus transplantation is the only option for women with absolute uterine infertility, such as those with Rokitansky syndrome, to experience ...

Surgical Approach, Challenges, and Resolutions for Uterus Transplantation in Rats

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