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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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 ...

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 border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<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&#38; 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&#160;&#160;</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&#160;&#160;</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&#160;&#160;</td>
			<td>F.S.T</td>
			<td>18056-14&#160;&#160;</td>
			<td></td>
		</tr>
		<tr>
			<td>M&#246;lnlyncke Op Towel</td>
			<td>M&#246;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&#252;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&#228;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&#38;T Vessel Dilating Forceps - Angled 45&#176;</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 &#8211; 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&#160;</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&#160; 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 re...

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

Research

JoVE Journal

Medicine

1.1K Views.  University Hospital Zürich. 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. 

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

Orthotopic Small Bowel Transplantation in Rats

Small bowel transplantation has become an accepted clinical option for patients with short gut syndrome and failure of parenteral nutrition (irreversible intestinal failure). In specialized centers improved operative and managing strategies have led to excellent short- and intermediate term patient and graft survival while providing high quality of life 1,3. Unlike in the more common transplantation of other solid organs (i.e. heart, liver) many underlying mechanisms of graft function and immunologic alterations induced by intestinal transplantation are not entirely known6,7. Episodes of acute rejection, sepsis and chronic graft failure are the main obstacles still contributing to less favorable long term outcome and hindering a more widespread employment of the procedure despite a growing number of patients on home parenteral nutrition who would potentially benefit from such a transplant. The small intestine contains a large number of passenger leucocytes commonly referred to as part of the gut associated lymphoid system (GALT) this being part of the reason for the high immunogenity of the intestinal graft. The presence and close proximity of many commensals and pathogens in the gut explains the severity of sepsis episodes once graft mucosal integrity is compromised (for example by rejection). To advance the field of intestinal- and multiorgan transplantation more data generated from reliable and feasible animal models is needed. The model provided herein combines both reliability and feasibility once established in a standardized manner and can provide valuable insight in the underlying complex molecular, cellular and functional mechanisms that are triggered by intestinal transplantation. We have successfully used and refined the described procedure over more than 5 years in our laboratory 8-11. The JoVE video-based format is especially useful to demonstrate the complex procedure and avoid initial pitfalls for groups planning to establish an orthotopic rodent model investigating intestinal transplantation.

Small bowel transplantation has become an accepted treatment option for patients with irreversible intestinal failure. Our experimental model of orthotopic small bowel transplantation in rats serves as a reliable tool to address underlying immunologic and inflammatory processes that complicate intestinal transplantation.

Small  bowel transplantation has become an accepted clinical option for patients with  short gut syndrome and failure of parenteral nutrition ...

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 ...

A Novel Surgical Approach for Intratracheal Administration of Bioactive Agents in a Fetal Mouse Model

Prenatal pulmonary delivery of cells, genes or pharmacologic agents could provide the basis for new therapeutic strategies for a variety of genetic and acquired diseases. Apart from congenital or inherited abnormalities with the requirement for long-term expression of the delivered gene, several non-inherited perinatal conditions, where short-term gene expression or pharmacological intervention is sufficient to achieve therapeutic effects, are considered as potential future indications for this kind of approach. Candidate diseases for the application of short-term prenatal therapy could be the transient neonatal deficiency of surfactant protein B causing neonatal respiratory distress syndrome1,2 or hyperoxic injuries of the neonatal lung3. Candidate diseases for permanent therapeutic correction are Cystic Fibrosis (CF)4, genetic variants of surfactant deficiencies5 and &alpha;1-antitrypsin deficiency6.
Generally, an important advantage of prenatal gene therapy is the ability to start therapeutic intervention early in development, at or even prior to clinical manifestations in the patient, thus preventing irreparable damage to the individual. In addition, fetal organs have an increased cell proliferation rate as compared to adult organs, which could allow a more efficient gene or stem cell transfer into the fetus. Furthermore, in utero gene delivery is performed when the individual's immune system is not completely mature. Therefore, transplantation of heterologous cells or supplementation of a non-functional or absent protein with a correct version should not cause immune sensitization to the cell, vector or transgene product, which has recently been proven to be the case with both cellular and genetic therapies7.
In the present study, we investigated the potential to directly target the fetal trachea in a mouse model. This procedure is in use in larger animal models such as rabbits and sheep8, and even in a clinical setting9, but has to date not been performed before in a mouse model. When studying the potential of fetal gene therapy for genetic diseases such as CF, the mouse model is very useful as a first proof-of-concept because of the wide availability of different transgenic mouse strains, the well documented embryogenesis and fetal development, less stringent ethical regulations, short gestation and the large litter size.
Different access routes have been described to target the fetal rodent lung, including intra-amniotic injection10-12, (ultrasound-guided) intrapulmonary injection13,14 and intravenous administration into the yolk sac vessels15,16 or umbilical vein17. Our novel surgical procedure enables researchers to inject the agent of choice directly into the fetal mouse trachea which allows for a more efficient delivery to the airways than existing techniques18.

We developed a novel surgical approach for intratracheal administration of bioactive agents into the mouse fetus. The delivery route is more efficient in targeting the fetal mouse lungs than the commonly used intra-amniotic injection. This procedure has to date not been described in a mouse model.

Prenatal pulmonary delivery of cells, genes or pharmacologic agents could provide the basis for new therapeutic strategies for a variety of genetic and ...

Transplantation of Pulmonary Valve Using a Mouse Model of Heterotopic Heart Transplantation

Tissue engineered heart valves, especially decellularized valves, are starting to gain momentum in clinical use of reconstructive surgery with mixed results. However, the cellular and molecular mechanisms of the neotissue development, valve thickening, and stenosis development are not researched extensively. To answer the above questions, we developed a murine heterotopic heart valve transplantation model. A heart valve was harvested from a valve donor mouse and transplanted to a heart donor mouse. The heart with a new valve was transplanted heterotopically to a recipient mouse. The transplanted heart showed its own heartbeat, independent of the recipient&#8217;s heartbeat. The blood flow was quantified using a high frequency ultrasound system with a pulsed wave Doppler. The flow through the implanted pulmonary valve showed forward flow with minimal regurgitation and the peak flow was close to 100 mm/sec. This murine model of heart valve transplantation is highly versatile, so it can be modified and adapted to provide different hemodynamic environments and/or can be used with various transgenic mice to study neotissue development in a tissue engineered heart valve.

In order to understand the cellular and molecular mechanisms underlying neotissue formation and stenosis development in tissue engineered heart valves, a murine model of heterotopic heart valve transplantation was developed. A pulmonary heart valve was transplanted to recipient using the heterotopic heart transplantation technique.

Tissue engineered heart valves, especially decellularized valves, are starting to gain momentum in clinical use of reconstructive surgery with mixed ...

Ex Situ Normothermic Machine Perfusion of Donor Livers

In contrast to conventional static cold preservation (0-4 &#176;C), ex situ machine perfusion may provide better preservation of donor livers. Continuous perfusion of organs provides the opportunity to improve organ quality and allows ex situ viability assessment of donor livers prior to transplantation. This video article provides a step by step protocol for ex situ normothermic machine perfusion (37 &#176;C) of human donor livers using a device that provides a pressure and temperature controlled pulsatile perfusion of the hepatic artery and continuous perfusion of the portal vein. The perfusion fluid is oxygenated by two hollow fiber membrane oxygenators and the temperature can be regulated between 10 &#176;C and 37 &#176;C. During perfusion, the metabolic activity of the liver as well as the degree of injury can be assessed by biochemical analysis of samples taken from the perfusion fluid. Machine perfusion is a very promising tool to increase the number of livers that are suitable for transplantation.

Ex Situ Normothermic Machine Perfusion of Donor Livers

Here we present a protocol describing oxygenated ex situ machine perfusion of donor liver grafts. This article contains a step by step protocol to procure and prepare the liver graft for machine perfusion, prepare the perfusion fluid, prime the perfusion machine and perform oxygenated normothermic machine perfusion of the liver graft.

In contrast to conventional static cold preservation (0-4 &#176;C), ex situ machine perfusion may provide better preservation of donor livers. Continuous ...

The Mouse Round-window Approach for Ototoxic Agent Delivery: A Rapid and Reliable Technique for Inducing Cochlear Cell Degeneration

Investigators have utilized a wide array of animal models and investigative techniques to study the mammalian auditory system. Much of the basic research involving the cochlea and its associated neural pathways entails exposure of model cochleae to a variety of ototoxic agents. This allows investigators to study the effects of targeted damage to cochlear structures, and in some cases, the self-repair or regeneration of those structures. Various techniques exist for delivery of ototoxic agents to the cochlea. When selecting a particular technique, investigators must consider a number of factors, including the induction of inadvertent systemic toxicity, the amount of cochlear damage produced by the surgical procedure itself, the type of lesion desired, animal survivability, and reproducibility/reliability of results. Currently established techniques include parenteral injection, intra-peritoneal injection, trans-tympanic injection, endolymphatic sac injection, and cochleostomy with perilymphatic perfusion. Each of these methods has been successfully utilized and is well described in the literature; yet, each has various shortcomings. Here, we present a technique for topical application of ototoxic agents directly to the round window niche. This technique is non-invasive to inner ear structures, produces rapid onset of reliably targeted lesions, avoids systemic toxicity, and allows for an intra-animal control (the contra-lateral ear). Results stemming from this approach have helped deeper understanding of auditory pathophysiology, cochlear cell degeneration, and regenerative capacity in response to an acute injury. Future investigations may use this method to conduct interventional studies involving gene therapy and stem cell transplantation to combat hearing loss.

Various methods exist for introducing ototoxic agents to the cochleae of animal models. Presented is a surgical protocol for delivery of ototoxic agents to the round window niche. The procedure is reliable, creates targeted intra-cochlear lesions, and avoids mechanical damage to the microarchitecture. Examination of cochlear self-repair/regeneration is possible.

Investigators have utilized a wide array of animal models and investigative techniques to study the mammalian auditory system. Much of the basic research ...

Surgical Approach for Middle Cerebral Artery Occlusion and Reperfusion Induced Stroke in Mice

Stroke is a leading cause of death worldwide and continues to be one of the major causes of long-term adult disabilities. About 87% of strokes are ischemic in origin and occur in the territory of the middle cerebral artery (MCA). Currently the only Food and Drug Administration (FDA) approved drug for the treatment of this devastating disease is tissue plasminogen activator (tPA). However, tPA has a small therapeutic window for administration (3 - 6 hr), and is only effective in 4% of the patients who actually receive it. Current research focuses on understanding the pathophysiology of stroke in order to find potential therapeutic targets. Thus, reliable models are crucial, and the MCA occlusion (MCAo) model (also termed the intraluminal filament or suture model) is deemed to be the most clinically relevant surgical model of ischemic stroke, and is fairly non-invasive and easily reproducible. Typically the MCAo model is used with rodents, especially with mice due to all the genetic variations available for this species. Here we describe (and present in the video) how to successfully perform the MCAo model (with reperfusion) in mice to generate reliable and reproducible data.

In order to understand the pathophysiology of stroke, it is important to use reliable models. This paper will describe one of the most frequently used stroke models in mice, termed the middle cerebral artery occlusion (MCAo) model (also termed the intraluminal filament or suture model) with reperfusion.

Stroke is a leading cause of death worldwide and continues to be one of the major causes of long-term adult disabilities. About 87% of strokes are ...

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 ...

Designing CAD/CAM Surgical Guides for Maxillary Reconstruction Using an In-house Approach

Computer-aided design/computer-assisted manufacturing (CAD/CAM) is now being evaluated as a preparative technique for maxillofacial surgery. Because this technique is expensive and available in only limited areas of the world, we developed a novel CAD/CAM surgical guide using an in-house approach. By using the CAD software, the maxillary resection area and cutting planes and the fibular cutting planes and angles are determined. Once the resection area is decided, the necessary faces are extracted using a Boolean modifier. These superficial faces are united to fit the surface of the bones and thickened to stabilize the solids. Not only the cutting guides for the fibula and maxilla but also the location arrangement of the transferred bone segments is defined by thickening the superficial faces. The CAD design is recorded as .stl files and three-dimensionally (3-D) printed as actual surgical guides. To check the accuracy of the guides, model surgery using 3-D-printed facial and fibular models is performed. These methods may be used to assist surgeons where commercial guides are not available.

Methods for designing a computer-aided design/computer-aided manufacturing (CAD/CAM) surgical guide are shown. Cutting planes are separated, united, and thickened to easily visualize the necessary bone transfer. These designs can be three-dimensional printed and checked for accuracy.

Computer-aided design/computer-assisted manufacturing (CAD/CAM) is now being evaluated as a preparative technique for maxillofacial surgery. Because this ...

Lentiviral Vector-mediated Gene Therapy of Hepatocytes Ex Vivo for Autologous Transplantation in Swine

Gene therapy is an ideal choice to cure many inborn errors of metabolism of the liver. Ex-vivo, lentiviral vectors have been used successfully in the treatment of many hematopoietic diseases in humans, as their use offers stable transgene expression due to the vector&#39;s ability to integrate into the host genome. This method demonstrates the application of ex vivo gene therapy of hepatocytes to a large animal model of hereditary tyrosinemia type I. This process consists of 1) isolation of primary hepatocytes from the autologous donor/recipient animal, 2) ex vivo gene delivery via hepatocyte transduction with a lentiviral vector, and 3) autologous transplant of corrected hepatocytes via portal vein injection. Success of the method generally relies upon efficient and sterile removal of the liver resection, careful handling of the excised specimen for isolation of viable hepatocytes sufficient for re-engrafting, high-percentage transduction of the isolated cells, and aseptic surgical procedures throughout to prevent infection. Technical failure at any of these steps will result in low yield of viable transduced hepatocytes for autologous transplant or infection of the donor/recipient animal. The pig model of human type 1 hereditary tyrosinemia (HT-1) chosen for this approach is uniquely amenable to such a method, as even a small percentage of engraftment of corrected cells will lead to repopulation of the liver with healthy cells based on a powerful selective advantage over native-diseased hepatocytes. Although this growth selection will not be true for all indications, this approach is a foundation for expansion into other indications and allows for manipulation of this environment to address additional diseases, both within the liver and beyond, while controlling for exposure to viral vector and opportunity for off-target toxicity and tumorigenicity.

Lentiviral Vector-mediated Gene Therapy of Hepatocytes Ex Vivo for Autologous Transplantation in Swine

This protocol is intended to describe porcine hepatocyte isolation and ex vivo gene delivery to cure models of metabolic diseases via autologous cell transplantation. Although this particular model enjoys unique advantages that favor successful therapy, the application is a relevant foundation to address additional diseases and indications.

Gene therapy is an ideal choice to cure many inborn errors of metabolism of the liver. Ex-vivo, lentiviral vectors have been used successfully in the ...