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

Podsumowanie

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.

Streszczenie

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.

Wprowadzenie

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 Brännström³. 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 rejection^5,6. 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 reported^11,12,13. 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.

Protokół

All animal experiments were performed following Swiss Federal Animal Regulations and approved by the Veterinary Office of Zurich (n° 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 reports^12,13,15.

1. 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).
   1. 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.
   2. Shave all the abdominal skin of the rat with an electric shaver.
   3. Use tapes to keep the animal fixed on a heating plate during surgery. Apply eye ointment to both eyes.
   4. Maintain anesthesia during the procedure with 2-4 vol% isoflurane in oxygen by continuous administration through a small nose cone.
   5. 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.
   6. Confirm anesthetic depth by performing a toe pinch.
   7. Clean the abdominal skin in a circular motion with three alternating swabs of an antiseptic solution and 70% alcohol. Allow to dry.
   8. Place a sterile drape (see Table of Materials) with an abdominal window over the animal.
2. Perform median laparotomy.
   1. 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.
   2. Move the intestines outside the abdominal cavity using cotton swabs, cover them gently with a gauze moistened with sterile saline, and protect them with a sterile plastic bag for better insulation.
   3. 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.
   4. Apply prewarmed saline to keep the surgical area and the intestines moist and avoid drying of the viscera.
3. Harvest the right uterine horn with the common uterine cavity and cervix plus vascular pedicles, including the right uterine, internal, and common iliac vessels.
   1. Ligate (4/0 polyglactin; see Table of Materials), cauterize, and sever the left uterine horn adjacent to the branching from the common uterine cavity.
   2. Remove excess fat surrounding the uterus and vagina.
      NOTE: Keep the fat around the uterine-vascular system.
   3. 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 clamp (see Table of Materials)  to obtain a better view of excavatio vesicouterina.
   4. 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.
   5. 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.
   6. Carefully dissect the vagina via diathermy around 2-3 mm caudal of the cervix.
      NOTE: No villi (cervix) is visible inside the vaginal lumen.
   7. 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.
   8. By blunt dissection, free 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.
   9. 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.
   10. 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.
   11. Dissect the common iliac vein caudally of the ligature at the right common iliac vein to enable outflow during flushing.
4. Flush the graft following the steps below.
   1. 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).
      ​NOTE: Avoid high flushing pressure and ensure proper needle placement.
   2. 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.
5. Place the transplant into chilled organ preservation solution (4 °C) for back table preparation and storage before transplantation.
6. After removing the graft, euthanize the animal by first turning the isoflurane setting to maximum and then inducing bilateral pneumothorax followed by exsanguination¹⁷.

2. Syngeneic uterus transplantation

NOTE: For details on the procedure, please see the previously published reports^12,13,15.

1. Induce anesthesia and prepare the animal as mentioned in step 1.1.
   1. Administer effective analgesia (as described in step 1.1.1) and 200 IU/kg high molecular weight heparin 30 min before surgery.
2. Perform median laparotomy.
   1. 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.
   2. 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.
   3. 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.
   4. Apply prewarmed saline to keep the surgical area and the intestines moist and avoid drying of the viscera.
3. Perform a hysterectomy with dissection and mobilization of the upper third of the vagina from the rectum and the bladder.
   1. 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.
   2. Remove fat tissue from the surroundings.
   3. 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.
4. Perform uterus transplantation.
   1. Mobilize and separate the right common iliac vessels, from the origin of the uterine vessels up to the aortic/caval bifurcation.
   2. 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.
   3. 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.
   4. Cut a slit slightly larger than the opening of the graft vein into the common iliac vein.
   5. Position the graft vein.
   6. 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.
   7. Regularly flush the anastomosis area with cooled RHX during the procedure to prevent thromboses.
   8. Anastomose one side of the graft vein to the recipient'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.
   9. Anastomose the other side of the vessel in the same manner, this time starting from the outside.
   10. 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.
   11. Place atraumatic vascular clamps onto the right common iliac artery at each side, framing the future anastomosis site.
   12. 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 "fish-mouth" 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.
5. Perform graft reperfusion.
   1. When both anastomosis sites appear patent and any bleeding is stopped, release the vascular clamps on the graft vessels (Figure 2).
   2. Inspect the graft for signs of reperfusion, such as reddening, filling of the vein, or pulsation in the graft artery.
   3. 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'clock position first, and place the next ones at the 10 and 1 o'clock positions. The two sutures at the 9 and 3 o'clock positions should be tied after the sutures in the front row^19,20.
   4. 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.
6. 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.
7. 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 administering buprenorphine (0.05 mg/kg) and suitable NSAID, however not before 4-8 hours after the first dose of anesthesia. Provide continuous with 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.
8. The skin suture is removed 10-14 days post suregery.

Wyniki

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) 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...

Dyskusje

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 Brännström group^12,13, inspired by the mouse work of Akouri et al.¹⁰, and modified based on the authors' experiences over the past years. As such, the ...

Materiały

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