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Method Article

Co-transplantation of Human Ovarian Tissue with Engineered Endothelial Cells: A Cell-based Strategy Combining Accelerated Perfusion with Direct Paracrine Delivery

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DOI:

10.3791/57472

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May 16th, 2018

 ,  ,  ,  ,  ,  ,  ,  , 
1Center for Reproductive Medicine and Infertility, Weill Cornell Medical College, 2Angiocrine Biosciences, Inc., 3Tri-Institutional Stem Cell Derivation Laboratory, Weill Cornell Medical College, 4Joan and Sanford I. Weill Department of Medicine, Weill Cornell Medical College

In This Article

Summary

For some patients, the only option for fertility preservation is cryopreservation of ovarian tissue. Unfortunately, delayed revascularization undermines follicular viability. Here, we present a protocol to co-transplant human ovarian tissue with endothelial cells for utilization as a cell-based strategy combining accelerated perfusion with a direct paracrine delivery of bioactive molecules.

Abstract

Infertility is a frequent side effect of chemotherapy and/or radiotherapy and for some patients, cryopreservation of oocytes or embryos is not an option. As an alternative, an increasing number of these patients are choosing to cryopreserve ovarian tissue for autograft following recovery and remission. Despite improvements in outcomes among patients undergoing auto-transplantation of cryopreserved ovarian tissue, efficient revascularization of grafted tissue remains a major obstacle. To mitigate ischemia and thus improve outcomes in patients undergoing auto-transplantation, we developed a vascular cell-based strategy for accelerating perfusion of ovarian tissue. We describe a method for co-transplantation of exogenous endothelial cells (ExECs) with cryopreserved ovarian tissue in a mouse xenograft model. We extend this approach to employ ExECs that have been engineered to constitutively express Anti-Mullerian hormone (AMH), thus enabling sustained paracrine signaling input to ovarian grafts. Co-transplantation with ExECs increased follicular volume and improved antral follicle development, and AMH-expressing ExECs promoted retention of quiescent primordial follicles. This combined strategy may be a useful tool for mitigating ischemia and modulating follicular activation in the context of fertility preservation and/or infertility at large.

Introduction

Cancer remains among the leading causes of death in the developed world, yet decades of research have yielded significant progress for most types of cancer, and in some cases nearly doubled survival rates1. Unfortunately, chemotherapeutic agents are often gonadotoxic, depleting the reserve of primordial follicles in ovaries and reducing fertility2. This growing population can benefit from various methods of fertility preservation including oocyte and/or embryo cryopreservation, however, patients requiring prompt initiation of cancer therapy and pre-pubertal patients are ineligible for these options. As an alternative, some patients have chosen to cryopreserve ovarian tissue before undertaking their therapeutic regimen, and upon recovery and remission, auto-transplanting tissue to restore fertility3. Yet, to date, graft survival and follicular output following auto-transplantation remain relatively low4, mainly due to tissue ischemia and hypoxia5,6,7. Despite numerous efforts to improve the viability of ovarian cortical grafts using anti-oxidants8,9, pro-angiogenic cytokines10,11,12,13, or mechanical manipulations14, graft ischemia in a 5 to 7 day window post-transplant undermines the viability and survival of the graft7. To address this, we developed a cell-based strategy to facilitate anastomosis of host and graft vessels and thus hasten reperfusion of ovarian tissue.

In addition to the ischemic insult to grafted ovarian tissue in the post-transplant window, the disruption of inter-follicular signaling may contribute to depletion of the pool15,16. Because exogenous endothelial cells (ExECs) contribute to stable and functioning vessels in the periphery of the graft, they present a unique opportunity to convey a defined molecular input to transplanted tissue. As a proof of principle, ExECs were engineered to express super-physiological levels of Anti-Mullerian hormone (AMH), a member of the transforming growth factor beta (TGFβ) superfamily that has been shown to restrict follicular growth17. Comparison of follicular distribution in grafts co-transplanted with control and AMH-expressing cells verifies the biological activity and potency of engineered exECs.

In summary, by improving graft viability and suppressing premature mobilization of the follicular pool, this approach can increase the productivity of auto-transplanted ovarian tissue in patients undergoing fertility preservation. Moreover, the ExEC-based platform enables experimental interrogation of molecular regulators that have been implicated in follicular development.

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Protocol

All procedures involving animal subjects have been approved by the Institutional Animal Care and Use Committee (IACUC) at Weill Cornell Medical College. All xenotransplantation experiments using ovarian tissue were performed in accordance with relevant guidelines and regulations. Human ovarian tissue was collected from patients scheduled for chemotherapy or radiotherapy for cancer treatment or prior bone marrow transplantation. The institutional review board (IRB) Committee of Weill Cornell Medical College approved the collection of tissue for potential autologous use, and upon the patient's informed consent a donation of up to 10% of their ovarian tissue for research use was performed.

1. Collection of Human Ovarian Tissue

NOTE: When an ovarian tissue is transported from a remote facility transit, time should not exceed 5 h18,19.

  1. Collect the ovarian tissue and rinse with a sterile saline solution.
  2. Place the ovary in a sterile container.
  3. Pour Leibovitz's L-15 medium until the ovary is completely immersed in the medium.
  4. Close and seal the container.
  5. Transport the container on ice.

2. Processing the Procured Ovarian Tissue, Adapted from Schmidt et al.18

  1. Preparations for ovarian processing and freezing
    1. Prepare 100 mL of medium for processing: Leibovitz's L-15 medium 99 mL + 1 mL antibiotic-antimycotic solution. Filter media using 0.2 µm filter. Keep the medium refrigerated.
    2. Prepare 100 mL of freezing solution using DMSO as a cryo-protectant. Add 69.64 mL Leibovitz's L-15 medium, 17.66 mL fetal bovine serum (FBS), 3.42 g of sucrose (to create a final concentration of 0.1 mol/L), 10.65 mL DMSO (to create a final concentration of 1.5 mol/L), and 1 mL antibiotic-antimycotic solution. Filter the medium using a 150 mL filter unit, 0.2 µm. Store the medium at 4 °C.
    3. Sterilize the surgical tools using an autoclave programmed for a wrapped solids sterilization cycle.
  2. Upon arrival of the tissue
    1. Set the sterile surgical tools in the biosafety cabinet: scalpel with blade number 21, sharp fine curved scissors and forceps at varied sizes: long (about 150 mm length) and 2 medium-sized (about 110 mm length).
    2. Wear sterile gloves, open the container within the biosafety cabinet. Take the ovary and place it in a 150 mm Petri dish and pour cold medium prepared in step 2.1.1 on top of the ovary to prevent dehydration of the ovarian tissue.
    3. Isolate the ovary from any residual tissue and rinse it with cold medium until it's devoid of tissue and blood.
    4. Place the ovary in a clean 150 mm Petri dish, add cold medium, prepared in step 2.1.1 until the ovary is half submerged in it.
  3. Dissection of the ovarian tissue
    1. Bisect the ovary and remove the medulla first by sharp dissection, using curved fine scissors. Then scrape the medulla away, using a sterile scalpel with a blade number 21. Precede until the cortical tissue is 1-1.5 mm in thickness.
    2. Cut the cortical tissue into slivers of 2-3 mm in width, the length of the strip will be the entire length of the ovarian piece that was processed.

3. Ovarian Tissue Slow Freezing, Adapted from Newton et al. and Oktay et al.6,20

  1. Preparation for freezing
    1. Label cryovials (1.8 mL).
    2. Add 1.5 mL of the freezing solution prepared in step 2.1.2 to each vial.
    3. Transfer one cortical strip per cryogenic vial containing the freezing solution.
    4. Equilibrate the cortical strip for 20 min on a rotating plate, apply gentle agitation at 4 °C.
  2. Slow freezing of ovarian tissue
    1. Load the cryovials into a programmable planer freezer starting at 0 °C.
    2. Cool at 2 °C/min to -7 °C.
    3. Keep the tissues at this constant temperature for 10 min.
    4. Perform manual seeding for ice crystal nucleation induction, by touching each cryovial with a cotton tip immersed in liquid nitrogen (LN2).
    5. Continue to cool at 0.3 °C/min until sample temperature reaches -40 °C.
    6. Cool at a faster rate of 10 °C/min to -140 °C.
    7. Transfer cryovials to the dewar for storage in LN2 (-196 °C).

4. Preparations for the Surgeries (Bilateral Oophorectomy and Co-transplantation)

  1. Preparation of plates
    1. A day prior to thawing the ovarian tissue for transplantation
      1. Place a piece of a plastic paraffin film at the bottom of a 50 mm Petri dish, spray it with 70% Ethanol until it will be completely covered.
      2. Leave the Petri dishes overnight in the biosafety cabinet. Prepare 2 Petri dishes per mouse.
    2. At the day of surgeries
      1. Aspirate the ethanol from the Petri dish containing the plastic paraffin film within the biosafety cabinet.
      2. Leave the Petri dish lead half open until ethanol evaporates completely.
      3. Close the lid of the Petri dish when the plastic paraffin film is completely dry. Keep it within the biosafety cabinet.
      4. Label wells of a 6 well plate accordingly, 0.1 mol/L Sucrose+ 1 mol/L DMSO, 0.1 mol/L Sucrose+ 0.5 mol/L DMSO, 0.1 mol/L Sucrose, Basic Thawing Solution (BTS), Medium.
  2. Preparation of the solutions
    1. Prepare 100 mL of BTS: Leibovitz's L-15 medium 79 mL + FBS 20 mL + Antibiotic-Antimycotic solution 1 mL. Filter with a 0.22 µm filter system.
    2. Prepare 10 mL of BTS with 0.1 mol/L Sucrose. Scale 0.342 g of Sucrose and add 10 mL of BTS prepared in step 4.2.1, Agitate gently until the Sucrose is completely dissolved. Filter the solution using a syringe filter 0.22 µm.
    3. Prepare the solutions according to Table 1. Cover tubes containing DMSO with aluminum foil. Keep refrigerated until use.

5. Ovarian Tissue Rapid Thawing

  1. Take out of the LN2 dewar a vial containing frozen ovarian tissue and keep it at room temperature for 30 s.
  2. Wipe the vial clean using a tissue paper.
  3. Immerse the vial in a 30 °C water bath for 1-2 min until its' content is thawed.
  4. Open the vial and place the cortical strip in the first well that contains 3ml of the first solution (0.1 mol/L Sucrose+ 1 mol/L DMSO, prepared in step 4.2.3).
    NOTE: All steps involving opening the lid of the plate will be done under laminar flow.
  5. Incubate the 6 well plate on a rotating plate at 4 °C for 5 min. Keep the plate covered with aluminum foil for this step.
  6. Transfer the cortical strip into the second well containing 3 mL of the second solution (0.1 mol/L Sucrose+ 0.5 mol/L DMSO, prepared in step 4.2.3).
  7. Incubate the 6 well plate on a rotating plate for gentle agitation at 4 °C for 5 min. Keep the plate covered with aluminum foil for this step.
  8. Transfer the cortical strip into the third well, containing 4 mL of solution (BTS with 0.1 mol/L Sucrose, prepared in step 4.2.2).
  9. Incubate on a rotating plate for gentle agitation at 4 °C for 5 min.
  10. Transfer the cortical strip into the fourth well containing 4 mL of solution (BTS, prepared in step 4.2.1).
  11. Incubate on a rotating plate for gentle agitation at 4 °C for 5 min.
  12. Transfer the cortical strip into the last well containing 4 mL of cold medium (prepared in step 2.1.1). Keep thawed ovarian tissue on ice until performing the transplantation.

6. Encapsulation of the Ovarian Tissue

  1. Prepare the following solutions
    1. Prepare 25 mL of 20 mmol/L HEPES buffer in 0.9% saline. Filter and store at 4 °C.
    2. Prepare 1 mL of CaCl2 at the concentration of 1 mol/L. Filter and store at 4 °C.
  2. Prepare a Fibrinogen 50 mg/mL stock solution
    1. Add 1 g of Fibrinogen into 20 mL of 20 mmol/L HEPES buffer in 0.9% saline by slowly mixing fibrinogen into the HEPES buffered saline over several hours at 37 °C.
    2. Label 1.7 mL micro-centrifuge tubes.
    3. Filter the solution through a 0.45 µm syringe filter and then through a 0.2 µm syringe filter.
    4. Aliquot the solution into 200 µL into micro-centrifuge tube and store at -20 °C.
  3. Prepare a Thrombin 100 U/mL stock solution
    NOTE: Thrombin solutions adsorb to glass, aliquot the solution in plastic tubes/vials.
    1. Add 2.5 mL of 0.9% sterile saline to 250 U of Thrombin.
    2. Gentle shake until the solution to completely dissolve.
    3. Label 1.7 mL micro-centrifuge tubes.
    4. Filter through a 0.2 µm syringe filter, aliquot 50 µL into micro-centrifuge tubes and store at -80 °C.
  4. Make a Fibrin clot of 70 µL, obtaining a final concentration of 10 mg/mL fibrinogen and 5 U/mL Thrombin
    NOTE: Make sure to work fast, the solution clots in a few seconds. Also, avoid the addition of bubbles to the mixture.
    1. Prepare a Fibrinogen solution in a 1.7 mL micro-centrifuge tube.
      1. Add 360 µL of 20 mmol/L HEPES Buffer in 0.9% saline solution to the aliquoted 200 µL of the Fibrinogen stock, prepared in step 6.2.4.
      2. Mix by gentle pipetting.
      3. Keep it on ice.
    2. Prepare a Thrombin solution in a second micro-centrifuge tube.
      1. Add 148.7 µL of DMEM to the aliquoted 50 µL of the Thrombin stock, prepared in step 6.3.4.
      2. Mix by gentle pipetting.
      3. Add 1.3 µL of 1 mol/L CaCl2.
      4. Mix by gentle pipetting.
      5. Keep it on ice.
    3. Prepare a single cell suspension, In a third micro-centrifuge tube.
      1. Detach the engineered endothelial cells from the plate using an enzyme cell detachment medium.
      2. Spin down the and count the cells using a hemocytometer with a cover glass.
      3. Use 20,000 cells per every 1 mm2 of ovarian tissue. Calculate 20,000x area of ovarian tissue in mm2.
      4. Spin down the engineered endothelial cells and re-suspend it in basic medium (DMEM) to reach a total volume of 16.8 µL.
      5. Keep it on ice.
    4. Place a piece of ovarian tissue on a sterile gauze sponge to dry it for a few seconds.
    5. Place the piece of ovarian tissue on top of the plastic paraffin film within the 50 mm Petri dish.
    6. Add 39.2 µL of the Fibrinogen solution, prepared in step 6.4.1.2, into the cell suspension prepared in step 6.4.3.4, mix by gentle pipetting. Keep it on ice.
    7. Add 14 µL of the Thrombin solution prepared in step 6.4.2.4. Mix gently by pipetting once. Work fast.
    8. Place the mixture on top of the ovarian tissue, pipet it in the form of an elongated droplet.
    9. Place the Petri dish with the clot in the incubator at 37 °C.

7. Bilateral Oophorectomy and Co-transplantation of Human Ovarian Tissue with Engineered Endothelial Cells to NSG Mice

NOTE: Ten to fourteen-week-old female NSG mice21 were used (Jackson Labs).

  1. Prepare all surgical tools, as elaborated in step 2.1.3, make sure you have all sutures, pads, and surgical tapes handy.
  2. Anesthetize the mouse using an anesthesia system with Isoflurane.
    1. Place the mouse in the induction chamber, close lid and open the appropriate stopcock.
    2. Open flowmeter to 1,000 mL/min. Turn on the vaporizer and set it to deliver 2-3.5%.
    3. Observe effects of anesthesia; loss of consciousness, slow and regular breaths.
    4. Transfer to the nose cone to maintain anesthesia. Turn the vaporizer to deliver 1-3%, depending on vital signs.
    5. Verify absence of pedal or tail reflex. If the reflex is present, increase isoflurane until it's absent.
  3. Trim the mouse's back hair, from the tail up to the clavicles line, using an electric hair trimmer.
  4. Place the mouse in a prone position on the surgical platform.
  5. Fix the nose cone to the surgical platform using a surgical tape.
  6. Tape the mouse's limbs gently to the surgical platform using a 1.25 cm-wide surgical paper tape.
  7. Use a sterile lubricant jelly and insert a thermometer PR. Fix the thermometer by taping it to the surgical platform using a 1.25 cm wide perforated plastic surgical tape.
  8. Tape the tail to the surgical platform by using a 2.5 cm wide surgical paper tape.
  9. Heat, using an infrared or a hot water heating pad, and monitor the mouse's body temperature throughout the procedure. Keep body temperature within the range of 35-37 °C.
  10. Clean the trimmed area with a sterile Povidone-Iodine Solution swab stick and wipe using sterile alcohol prep.
  11. Repeat step 6.9 twice more.
  12. Place a drop of sterile ocular lubricant vet ointment over each of the mouse's eyes to protect from dehydration and damage to the cornea.
  13. Cover the mouse using a sterile surgical drape.
  14. Inject Buprenorphine (1 mg/Kg) sub-cutaneous (S/C).
  15. Perform a longitudinal medial dorsal incision using a scalpel (blade #21).
  16. Perform a bilateral oophorectomy, use a dorsal approach.
    1. Free the subcutaneous connective tissue by blunt dissection using the scissors.
    2. Pinch the fascia laterally to the midline make an incision and reach the peritoneal cavity using the sharp scissors.
    3. Grab the ovarian fat pad and the ovary gently with tweezers and pull it out of the abdominal cavity.
    4. Grasp the ovary and the fat pad with a vascular clamp.
    5. Ligate the ovary and the fat pad using a 4/0 monofilament absorbable suture at the base of the ovary, distal to the fallopian tube.
    6. Clip the ovary distal to the tie. Make sure there is no bleeding from the base of the stump.
    7. Place the tissue back into the abdominal cavity and suture the fascia using a 6/0 braided absorbable suture.
    8. Repeat stages 7.15.1-7.15.7 at the contralateral side.
  17. Co-transplant the ovarian tissue.
    1. Perform a horizontal incision in the fascia above the Gluteus Maximus, at the length that fits the clots dimensions. Create a pocket within this virtual space by opening the scissors underneath the fascia.
    2. Pick up one piece of encapsulated cortical tissue, as prepared in step 6.4.8 and place it in this pocket.
    3. Suture the fascia using a 6/0 braided absorbable suture.
  18. Repeat stages 7.16.1-7.16.3 at the contralateral side as well.
  19. Close the dorsal wall using simple interrupted stitches with a 4/0 monofilament absorbable suture.
  20. Inject Lidocaine 0.5% (1.2 mL/Kg) S/C at the incision site.
  21. Use a sterile alcohol prep to clean the skin.
  22. Turn off the isoflurane while monitoring the mouse's temperature.
  23. Provide 1-2 min of O2, without Isoflurane. Keep on warming the mouse until it starts to move and return to consciousness.
  24. Place the mouse in a clean recovery cage and later house the mouse in a separate cage until the complete healing of the surgical wound.
  25. Inject Buprenorphine (1 mg/Kg) S/C as needed every 8-12 h, for 24 h postoperatively.
  26. Keep the mice in separate cages when all food, water, bedding, and cages are autoclaved, until the end point of the experiment. Keep the cages in a pressurized ventilated room. Use personal protective equipment whenever handling the mice.

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Results

To determine whether co-transplantation of ExECs provides a benefit to patients' tissue, thawed ovarian cortical strips were divided into equal sized pieces and engrafted bilaterally into immuno-compromised, NOD scid gamma (NSG), mice. With one side embedded in a fibrin clot alone (no ECs) and the other containing ExECs (Figure 1a), each mouse served as its own control. ExECs were obtained via isolation of primary endothelium from human umbilical cords and su...

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Discussion

Here we demonstrate that co-transplantation of exECs provides a significant benefit to ovarian tissue viability and function following xenograft in mice. Standards for clinical application of ovarian tissue auto-transplantation for fertility preservation have not been set and the optimal parameters (size, transplantation site, duration of graft, etc.)32,33,34 for enhanced recovery of the follicular pool remain undefined...

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Disclosures

Michael Ginsberg is an employee of Angiocrine Biosciences, Inc., San Diego, CA, 92130, United States, that isolated, transfected with E4-ORF- 1 and labeled the endothelial cells we used.

Acknowledgements

Omar Alexander Man for the illustrations.
L.M. was supported by a Pilot Award from the Cornell Clinical and Translational Science Center and an ASRM research grant.
The authors would like to thank James lab members for critical reading of the manuscript.

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Materials

List of materials used in this article
NameCompanyCatalog NumberComments
Leibovitz’s L-15 mediumGibco11415064
Antibiotic-AntimycoticGibco15240062Anti-Anti X100
SucroseSigmaS 1888
FibrinogenSigmaF 8630from bovine plasma
ThrombinSigmaT 1063from human plasma
DMSOSigmaD 2650
DMEMGibco12491015
Enzyme Cell Detachment MediumInvitrogen00-4555-56Accutase
Plastic paraffin filmBemis NAParafilm M
Surgical paper tape 2.5 cm3M1530-1Micropore
Surgical Paper tape 1.25 cm3M1530-0Micropore
Perforated plastic Surgical tape 1.25 cm3M1527-0Transpore
Monofilament Absorbable SutureCovidienUM-203Biosyn
Braided Absorbable SutureCovidienGL-889Polysorb
Povidone-iodine Solution USP 10%Purdue Products67618-153-01Betadine Solution Swab Stick
CryovialesNunc377267CryoTube
sterile ocular lubricantDechra17033-211-38Puralube
1.7 ml micro-centrifuge tubeDenvilleC-2172Eppendorf
Anasthesia systemVetEquipV-1 table top system with scavenging
Endothelial cellsAngiocrine Biosciences, Inc., San Diego, CA, USAIsolated, transfected with E4-ORF- 1 and labeled endothelial cells
Trichrome stainSigmaHT15-1ktTrichrome Stain (Masson) Kit
IsolectinInvitrogenI32450isolectin GS-IB4 From Griffonia simplicifolia, Alexa Fluor™ 647 Conjugate

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Ovarian Tissue Co transplantationEndothelial Cell EngineeringFibrin Clot FormationMouse Xenograft ModelFollicle Development AnalysisParacrine Signaling DeliveryIschemia Mitigation StrategyFollicular Activation ModulationCryopreserved Tissue ViabilityAnti Mullerian Hormone Expression
Co-transplantation of Human Ovarian Tissue with Engineered Endothelial Cells: A Cell-based Strategy Combining Accelerated Perfusion with Direct Paracrine Delivery
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