Allogeneic Uterus Transplantation in Baboons

Transplantation of the uterus (UTx) is seen as the treatment option for women with absolute uterine factor infertility. The UTx procedure has been performed successfully in animal models (1–16), and a few clinical UTx attempts have also been reported (17, 18). Because the uterine graft is life-enhancing and not life-sustaining, the development of techniques for its safe performance is of paramount significance. Nonhuman primates (NHP) have been previously used for transplant studies (19–21) and also in UTx experiments from a living donor (8, 12, 13). Our study aimed to ascertain the feasibility and safety of UTx using a deceased donor model. Animals Twelve female Papio hamadryas baboons (11–19 kg; age, 5–17 years) were included in the experiments that took place at the Mannheimer Foundation, Homestead, FL. Approval for the experiments was obtained by the institutional IACUCs (Table 1A).TABLE 1A: Pre-transplant characteristics of the study subjectsFollow-Up Period The primary endpoint of the experiment was the safety of the transplant procedure: Animals were initially followed up per protocol for 2 weeks to 2 months after transplantation. The follow-up period was amended later on to include longer survival. Immunosuppression As induction treatment, the animals received antithymocyte globulin (ATG) (10 mg/kg of body weight intravenous; Thymoglobulin, Genzyme, Cambridge, MA) together with intramuscular (IM) methylprednisolone (Solumedrol; Pharmacia Upjohn Co., NY, NY) on day −2 and −1 (day 0 defined as day of surgery). Maintenance treatment consisted of tacrolimus (TAC) (Prograf; Astellas, Northbrook, IL) and methylprednisolone administered daily. Tacrolimus was initially administered IM with target 24-hr trough levels of 10 to 15 ng/mL and converted to oral administration after the first week. After severe rejections and inability to obtain therapeutic levels in the first two transplants, the protocol was amended to include TAC IM with target 24-hr trough levels of 20 to 50 ng/mL. Immunosuppression was spaced gradually after the first 3 months posttransplant. Mycophenolate mofetil, 50 mg/kg BW, (Cellcept Genentech, San Francisco, CA) was given orally but was discontinued after the first four transplants, because therapeutic levels could not be obtained. Anesthesia and nonimmunosuppressive medications were administered as previously described (12, 22). Donor Surgery The vault of the vagina, the uterus, the fallopian tubes, and the ovaries were recovered en bloc with their arterial inflows and venous outflows, including the abdominal aorta, inferior vena cava (IVC) and the iliac vessels. The peritoneal cavity was entered through a wide cruciate incision. The intestinal loops were mobilized and retracted to expose the infra-renal abdominal aorta and the IVC. The inferior mesenteric artery was ligated and transected. After the right and left ureters were dissected and transected, both right and left ovarian arteries and veins were identified. A segment of the bladder was included in the specimen to facilitate suspension of the graft. The external iliac vessels were tied and transected bilaterally at the level of the inferior epigastric vessels, whereafter the uterine and inferior iliac vessels were mobilized. Superior vesical, pudendal, gluteal, and sacral branches of the internal iliac arteries and veins were ligated and transected along with all lumbar branches of the abdominal aorta and inferior vena cava, up to the level of the renal vessels. The graft was mobilized further and transected at the level of the vagina. At this point, it was only attached by its vascular pedicles of the uterine vessels through paracervical or metrum and the ovarian vessels through the infundibulopelvic ligaments (Fig. 1A). Anticoagulant (heparin) was administered, a large vascular catheter was inserted into the abdominal aorta and the graft was perfused with cold heparinized University of Wisconsin solution. The abdominal aorta and vena cava were transected above the renal vessels, and the graft was removed from the abdomen (Fig. 1B and C).FIGURE 1: Uterine graft retrieval and transplantation into the recipient. Donor surgery: (A) mobilization of the graft that has been transected at the level of the vagina and attached only to its vascular pedicles. (B and C) The abdominal aorta (red arrow) and vena cava (blue arrow) were transected above the renal vessels and the uterine graft was removed from the abdomen en bloc with the vault of the vagina, the uterus, the Fallopian tubes and the ovaries. Recipient surgery: the graft was brought to the field, and an end-to-side running anastomosis was performed on the aorta and the inferior vena cava of the recipient (D). After completion of the two anastomoses, the vascular clamps were released, and the graft was reperfused (E). The graft was fixed to its orthotopic position through suturing the round ligaments and the bladder peritoneums as well as vaginal anastomosis (F). (A) Uterus graft; (B) donor vascular pedicle; (C) donor fallopian tubes or ovaries; (D) donor vault of the vagina; (E) recipient ovary.Recipient Surgery The peritoneal cavity was entered through a midline incision, and the intestinal loops were mobilized and retracted to expose the infrarenal abdominal aorta and IVC. A standard total hysterectomy was performed. The vaginal vault was left open with marking sutures of its corners. The infrarenal abdominal aorta was partially cross-clamped, and a longitudinal arteriotomy was made. The graft was brought to the field and an end-to-side running anastomosis was performed on the aorta. After the IVC was cross-clamped, a longitudinal venotomy was made, and the venous anastomosis was performed (Fig. 1D). Then, the vascular clamps were released, and the graft was reperfused (Fig. 1E). The graft and anastomoses were adjusted for optimal positioning and hemostasis. The upper part of the donor vagina attached to the graft was subsequently anastomosed with the vaginal vault of the recipient (Fig. 1F). The abdominal wall and skin were closed separately with running sutures. Postoperative Assessment Labs were collected twice weekly immediately postoperatively and weekly thereafter, concurrently with graft biopsies, transabdominal ultrasound of the graft, and clinical examination. Biopsies were obtained transvaginally using a hysteroscope (Gyrus ACMI IDH-4 flextip; Gyrus ACMI, Southborough, MA). Rejection was scored as previously reported (12). Six transplantations were performed. All transplant surgeries were uneventful without any need for blood transfusion. All animals recovered from anesthesia (Table 1B, Fig. 2).TABLE 1B: Transplant surgery dataFIGURE 2: Intraoperative parameters during recipient transplant surgery and anesthesia.Animals #1, #2, presented biopsy proven severe rejection on week 2 after transplantation. Both presented increased lymphocyte counts, low tacrolimus levels (Fig. 3B and C) and showed notable graft enlargement. Rejection was treated with ATG and conversion of TAC from oral to IM. Animal #1 did not respond to treatment and was euthanized on postoperative day (POD) 31. The graft was surrounded by extensive adhesions. The vascular anastomosis were patent, with partial arterial and venous thrombosis at the vascular pedicle distal to the anastomosis. At histopathology, some areas of the graft were necrotic, whereas other parts presented only serosal inflammation (Table 1C). Animal #2 responded to treatment. At 4 months after transplantation, the animal presented estrous, hysteroscopy showed a viable graft (Fig. 4A and B), and ultrasound evaluation after intrauterine infusion of opaque medium showed normal uterus morphology. The animal was alive and well 13 months after transplantation, housed in a pen with a male partner. Repeat endoscopy at 14 months showed an atrophic graft, which was confirmed by laparotomy, and the animal was euthanized. Histopathology showed an atrophic graft with superimposed mild rejection.FIGURE 3: Peripheral blood absolute lymphocyte count, tacrolimus 24 hr trough levels and serum creatinine.TABLE 1C: Posttransplant outcomes of the study subjectsFIGURE 4: Images of hysteroscopic views and histopathology. Figure 4A, B: Hysteroscopy of graft of animal #2, 4 months after transplantation. Normal endometrium (A) and bleeding from the site of the biopsy (B) are seen. (C) Explanted graft of animal #4: acute and chronic rejection, moderate-severe. (D) Explanted graft animal #5: normal graft.Animal #3 presented with increased cytomegalovirus (CMV) viral load in the peripheral blood on POD 16 that did not respond to treatment, BW loss greater than 25% and was euthanized on POD 34. Macroscopically, the graft looked viable with patent anastomosis. Histopathology demonstrated arterial thrombi in peripheral graft arteries. The graft presented with mild inflammation, no significant epithelial injury and normal stroma and myometrium. Animal #4 presented increased CMV viral load at 5 weeks after transplantation that responded to treatment and severe rejection on POD 57, at the time its immunosuppression (ISP) was spaced, however without graft enlargement as in animals 1 and 2. Rejection was reversed with an increase of ISP, without need for ATG administration. The animal presented with BW loss greater than 25% and was euthanized on POD 112. Macroscopically the graft looked viable with patent anastomosis. Histopathology showed partially organized thrombus in the graft vascular pedicle and changes suggestive of acute and chronic rejection, moderate-severe (Fig. 4C). Histopathology of the kidneys showed mild acute tubular necrosis, possibly attributable to TAC toxicity. Animal #5 presented with acute tubular necrosis and renal failure and expired a week after transplantation (Fig. 3D). Macroscopically, there was fluid present in the abdominal cavity and serosal inflammation in the intraabdominal native organs. The graft looked viable, without any inflammation. Pathology showed mucosal necrosis and serositis of the stomach and the intestine, acute tubular necrosis of the kidney, and serositis of the native vaginal stump. The vascular anastomosis was patent. The graft was normal: no rejection, no presence of intravascular thrombi (Fig. 4D). Animal #6 presented with increased CMV viral load and weight loss and was sacrificed on POD 42. Histology revealed a normal graft with no rejection and patent vascular anastomosis. There was a mural, nonoccluding arterial thrombus distally, without ischemic injury findings. Our study aimed to ascertain the safety of UTx in the primate model using techniques applicable from a deceased donor. Deceased donor organ advantages include primarily that the risk subjected to an otherwise healthy third-party subject is avoided, and secondarily that we can retrieve the large intraabdominal vessels (aorta and vena cava) and use them in the anastomoses, possibly shortening the transplantation procedure and also decreasing the risk of vessel constriction and the associated postoperative thrombosis risk. In our experiment, all donors and recipients remained stable during the organ recovery and transplantation procedures, and the uterine grafts appeared to be normal after reperfusion in blood flow and macroscopic appearance. The initial follow-up period was 2 weeks to 2 months, to evaluate the short-term outcomes after the transplant surgical procedure. For two of the study subjects (#2 and #4) that presented with a good clinical picture, the experiment was extended to allow additional observations. Animal #2 was able to resume apparently normal physical activities, estrus, and sexual activities when it was paired with a male partner. Its survival was 13.8 months. The graft of this animal was in excellent condition confirmed by hysteroscopy at 4 months after transplantation, but subsequently failed most likely because of a drastic reduction (50%) and spacing of ISP. This spacing was done because it was judged to still be efficacious and to avoid daily intramuscular injections and the associated animal discomfort. Delivery of adequate ISP was one of the most significant problems encountered: it resulted in severe rejections in two animals ( #1, #2) and forced us to use the IM route to achieve levels that effectively controlled rejection (Fig. 3C). A second problem was the great difficulty in maintaining the animals’ BW. Weight loss was most accentuated during the first 2 weeks after transplantation. The severity of the operation and frequent anesthesia or sedations, the resulting limitation of the activities, as well as the relatively high blood levels of TAC, may have been contributing factors. Tacrolimus has been known to exhibit species-related toxicity and species difference of lymphocyte sensitivity to its effects (19, 20). In animals #3 and #6, there was also coexistent CMV viremia which may have also contributed. Overall, 50% of the animals developed CMV viremia. No invasive CMV infection was detected on any of them. Immunosuppression Toxicity All animals but one (#5) presented with adequate urinary production and creatinine levels within reference values throughout the study (Fig. 3D). We suspect that the rapid demise of animal #5 was caused by TAC toxicity which caused acute renal failure and seemed to affect all native tissues. Interestingly, the uterine graft of that animal was not affected. Another animal (#4) presented TAC-related changes in kidney biopsies performed at euthanasia, but did not present any clinical signs of TAC toxicity, and its renal function tests were normal. Mycophenolate mofetil was initially used in the immunosuppression protocol; however, in a future study where pregnancy would be the main outcome, teratogenic immunosuppressants, such as mycophenolate mofetil, would not be included. Vascular Thrombosis Four animals developed mural thrombi in the arterial conduits although the aortoaortic anastomosis was patent and without apparent technical imperfections. This could be because of the length of the arterial conduit, low flow, dehydration or hypercoagulable (maybe hypotension) state associated with the animals’ weight loss. In three of the animals, the thrombi were not occlusive, and their graft was found to be in excellent condition. This complication could possibly be avoided with better nutrition, hydration, shorter arterial conduits and use of antiplatelet agents and more intense anticoagulation prophylaxis. Species-Specific Challenges Nonhuman primates presented several challenges, which are distinctively different from humans. Oral administration of any medication, particularly ISP, was challenging and necessitated parenteral administration, which would be difficult to sustain for the long duration of complete UTx experiments. Cytomegalovirus was a potentially lethal threat, and weight loss of more than 25%, an endpoint of the experiment in most study settings, occurred frequently. These challenges are unique to the NHP models and are not anticipated in human UTx. Future studies of UTx with NHPs should take these facts into consideration. Panagiotis Tryphonopoulos 1 Andreas G. Tzakis2 Akin Tekin1 Liza Johannesson3 Krishna Rivas4 Pablo R. Morales4 Joseph Wagner4 Johan Mölne5 Anders Enskog6 Cesar Diaz-Garcia7 Pernilla Dahm-Kähler3 Mariana Berho8 Stephen Zimberg9 Tommaso Falcone10 Philip Ruiz1 Michael Olausson11 Mats Brännström3 1 Division of Liver and Intestinal Transplantation Department of Surgery University of Miami School of Medicine Miami, FL 2 Department of Surgery Cleveland Clinic Weston, FL 3 Department of Obstetrics and Gynecology Sahlgrenska Academy University of Gothenburg Göteborg, Sweden 4 The Mannheimer Foundation, Inc. Homestead, FL 5 Department of Pathology Sahlgrenska Academy University of Gothenburg Göteborg, Sweden 6 Department of Anesthesia and Intensive Care Sahlgrenska Academy University of Gothenburg Göteborg, Sweden 7 Department of Gynecology and Obstetrics La Fe University Hospital University of Valencia Valencia, Spain 8 Department of Pathology Cleveland Clinic Weston, FL 9 Department of Gynecology Cleveland Clinic Weston, FL 10 Department of Gynecology Cleveland Clinic Weston, FL 11 Sahlgrenska Transplant Institute, Department of Transplantation, Sahlgrenska Academy University of Gothenburg Göteborg, Sweden