Production of Transgenic and Knockout Pigs by Somatic Cell Nuclear Transfer

  • Angelica M. Giraldo
  • Suyapa Ball
  • Kenneth R. Bondioli
Protocol
Part of the Methods in Molecular Biology book series (MIMB, volume 885)

Abstract

Xenotransplantation is one alternative to transplantation of human organs which has been investigated. It is generally accepted that the pig represents the most logical choice of animals to serve as organ donors for xenotransplantation. Moreover, the implementation of cloning by somatic cell nuclear transfer (SCNT) and transgenic techniques have resulted in the production of numerous transgenic pigs than can be used for xenotransplantation purposes as well as models for human diseases.

Key words

Cloning Somatic cell nuclear transfer Transgenesis Knockout Pig engineering 

1 Introduction

The shortage of human organs for transplantation is well documented, and few practical solutions have been offered. Xenotransplantation is one alternative to transplantation of human organs which has been investigated. It is generally accepted that the pig represents the most logical choice of animals to serve as organ donors for xenotransplantation (1). This consensus has been arrived at because of physiological and genetic similarities between humans and pigs and the long history of safe utilization of porcine tissues and proteins in human medicine. There are many biological barriers to successfully transplanting animal organs into humans, particularly those involving the inflammatory and immune reactions induced by the xenograft. A number of transgenic strategies have been investigated and proposed to overcome these reactions. These include, but are not limited to, elimination of complement-mediated hyperacute rejection and stimulation of cell-mediated immune rejection and are reviewed elsewhere in this volume. Many of these strategies are best accomplished through a gene targeting process resulting in a functional removal of a gene function, “gene knockout,” or a replacement of part of a gene sequence, “gene knock in.” Such gene targeting reactions can be accomplished in cultured cells by homologous recombination and subsequent selection of the targeted cells by a variety of strategies. In mice, these procedures are performed in embryonic stem (ES) cells that readily support homologous recombination and are readily propagated in culture (2). Genetically modified and selected murine ES cells can be combined with normal embryos and have the capability of transferring the modified ES cell genome through the germ line of resulting animals. Unfortunately, numerous attempts to isolate ES cells from pigs have failed to produce cells with this germ line transfer capability. Somatic cell nuclear transfer (SCNT) has provided a means to create live animals capable of germ line transfer from cells genetically modified in culture from non-rodent species such as the pig. Several recent reports have demonstrated the feasibility of these types of genetic manipulations utilizing homologous recombination in somatic cells and nuclear transfer to produce live pigs (3).

Strategies for the design of targeting constructs have been described in the literature for a number of years. There are no indications that the design principles for these constructs are any different for targeting in porcine fetal fibroblasts. The acquisition of isogenic DNA to include as the homologous arms in a targeting construct is not a practical reality in this application and successful targeting with non-isogenic DNA has shown this to be a moot point. A strategy that has been successful in porcine fetal fibroblasts when the gene to be targeted is expressed consists of a single positive selection strategy followed by screening for targeted insertions by PCR. The positive selection strategy consists of a promoter trap design with a promoterless Neomycin resistance gene targeted to be inserted in frame behind or to replace the ATG of the gene to be disrupted. Cloned construct DNA is prepared for transfection in the same manner as DNA prepared for nuclear injection. In this chapter, we describe the procedures for transfection and selection of porcine somatic cells and the production of live animals from these cells by SCNT.

2 Materials

2.1 Collection of Porcine Oocytes

2.1.1 Collection and Transport of Ovaries

  1. 1.

    Disinfectants: 70% alcohol and Nolvasan.

     
  2. 2.

    Clean plastic container with lid and colander.

     
  3. 3.

    Washing solution: 0.9% saline solution (autoclaved) supplemented with 10 μg/mL of gentamicin.

     
  4. 4.

    Optional: Betadine (see Note 1).

     

2.1.2 Aspiration of Follicles

  1. 1.

    5-mL air syringes, 50-mL conical tubes, and 20–22 G needles.

     
  2. 2.

    Optional: Regulated vacuum pump (Cook, Indiana, USA).

     

2.1.3 Collection and In Vitro Maturation of Oocytes

  1. 1.

    Oocyte manipulation supplies: Pasteur pipettes, pulled capillaries of about 400–500 μm in diameter, mouth pipette, suction bulb, or Drummond pipette.

     
  2. 2.

    Culture supplies: Square grid dishes, Petri dishes, four-well dishes.

     
  3. 3.

    Holding medium: M199 with Hank’s salts, l-glutamine and 25 mM HEPES supplemented with 0.1% bovine serum albumin (BSA), 100 U/mL of penicillin, and 100 μg/mL of streptomycin. Filter and store for up 1 month at 4°C. Bring the medium at 38.5°C before use.

     
  4. 4.

    Maturation medium: The medium of choice should be placed in the incubator at least 1 h before use.

     
  5. 5.

    Equipment: Dissection microscope with under-stage illumination, dry bath and CO2 incubator at 38.5°C.

     

2.2 Establishment of Cell Lines and Preparation of Donor Cells

2.2.1 Establishment and Maintenance of a Cell Line

  1. 1.

    Rubbing alcohol.

     
  2. 2.

    Collection and dissection tools: Razors, biopsy punches, scissors, scalpels, blades.

     
  3. 3.

    Culture supplies: 15- or 50-mL conical tubes, culture dishes, Pasteur pipettes, pipettors, tips.

     
  4. 4.

    Washing medium: Dulbecco’s phosphate-buffered saline (PBS, calcium and magnesium free) containing 100 U/mL of penicillin and 100 μg/mL of streptomycin.

     
  5. 5.

    Cell culture medium: Dulbecco’s modified Eagle’s medium (DMEM) high glucose containing 10% of fetal bovine serum, 100 U/mL of penicillin, and 100 μg/mL of streptomycin.

     
  6. 6.

    Trypsin, 0.05% with ethylenediamine tetraacetic acid (EDTA).

     
  7. 7.

    Equipment: Inverted microscope, aspiration pump, CO2 incubator, centrifuge, and hemocytometer.

     

2.2.2 Electroporation and Selection of Donor Cells

  1. 1.

    Cytosalts: 120 mM KCl, 0.15 mM CaCl2⋅2H2O, 10 mM K2HPO4, 5 mM MgCl2⋅5H2O. Adjust pH to 7.6. Sterilize by filtration and store at room temperature (see Note 2).

     
  2. 2.

    Opti-MEM (Invitrogen, Carlsbad, CA, USA).

     
  3. 3.

    Electroporation medium: 75% cytosalts, 25% Opti-MEM mixed fresh daily.

     
  4. 4.

    Electroporation cuvette with 4-mm gap.

     
  5. 5.

    Cell culture supplies: 15-mL conical tubes, cryovials, fine tip transfer pipettes, Pasteur pipettes, pipettors, tips, 25-cm2 tissue culture flasks, 100-mm tissue culture plates, 24- and 48-well tissue culture plates, sterile cloning rings.

     
  6. 6.

    Washing medium: see Subheading 2.2.1.

     
  7. 7.

    Cell culture medium: see Subheading 2.2.1.

     
  8. 8.

    Trypsin, 0.05% with EDTA.

     
  9. 9.

    Geneticin (G418).

     
  10. 10.

    Freezing medium: 10% DMSO in fetal bovine serum.

     
  11. 11.

    Equipment: Cell electroporator.

     

2.2.3 Preparation of Donor Cells for Cloning

  1. 1.

    Culture supplies: 15-mL conical tubes, 1.5 mL vials, cryovials, culture dishes, Pasteur pipettes, pipettors, tips.

     
  2. 2.

    Washing medium: see Subheading 2.2.1.

     
  3. 3.

    Cell culture medium: see Subheading 2.2.1.

     
  4. 4.

    Trypsin, 0.05% with EDTA.

     
  5. 5.

    Equipment: Liquid nitrogen tank, inverted microscope, aspiration pump, centrifuge, hemocytometer, humidified CO2 incubator, and water bath at 38°C.

     

2.3 Production of Cloned Embryos

2.3.1 Removal of Cumulus Cells and Selection of Oocytes

  1. 1.

    Glass pipettes: Pasteur pipettes with polished end, glass capillaries with internal diameter of 180–200 and 250–300 μm.

     
  2. 2.

    Culture supplies: four-well dishes, Petri dishes, and 15-mL conical tubes.

     
  3. 3.

    Holding medium: M199 with Hank’s salts, l-glutamine, and 25 mM HEPES supplemented with 0.4% BSA and 50 μg/mL of gentamicin. Filter and store for up to 1 month at 4°C. Bring the medium at 38.5°C before use.

     
  4. 4.

    Hyaluronidase solution: 0.3 mg/mL of hyaluronidase in holding medium (see Subheading 2.3.1). Bring the solution at 38.5°C before use.

     

2.3.2 Enucleation of Mature Oocytes

  1. 1.

    Mineral oil.

     
  2. 2.

    Depression slide.

     
  3. 3.

    Micromanipulation tools: Holding pipette with an internal diameter of 80–100 μm, and enucleation pipette of 18–20 μm in diameter.

     
  4. 4.

    Holding medium: see Subheading 2.3.1.

     
  5. 5.

    Enucleation medium: 7.5 μg/mL of Hoechst and 7.5 μg/mL of cytochalasin B in holding medium (see Subheading 2.3.1). Bring the solution at 38.5°C before use. It is recommended to prepare 100× stock solutions of Hoechst and cytochalasin B. Cytochalasin B is soluble in DMSO. The stocks can be aliquoted and store at −20°C.

     
  6. 6.

    Equipment: Inverted microscope equipped with UV light, Hoffman or phase-contrast system, stage warmer; and a set microinjectors and micromanipulators.

     

2.3.3 Reconstruction of Embryos

  1. 1.

    See Subheading 2.3.2.

     
  2. 2.

    Four-well dishes.

     
  3. 3.

    Injection pipette: Injection needle of 20–25 μm in diameter.

     
  4. 4.

    Fusion medium: 260 mM mannitol, 0.1 mM MgSO4, 0.001 mM CaCl2, and 0.1% polyvinyl alcohol (PVA) in water. Add the PVA to prewarmed water and stir. Once the PVA dissolves, add the mannitol. Dissolve the MgSO4 and CaCl2 in separate tubes and then add them to the solution. Filter and store at 4°C for up to 6 months.

     
  5. 5.

    Equipment: Electro cell manipulator (BTX Inc., San Diego, CA), fusion/activation chamber with a 3.2-mm gap (BTX Inc.) and dissection microscope with under-stage illumination.

     

2.3.4 Activation of Fused Oocytes

  1. 1.

    See Subheading 2.3.3.

     
  2. 2.

    Activation medium: 260 mM mannitol, 0.1 mM MgSO4, 0.1 mM CaCl2, and 0.1% PVA in water. Add the PVA to prewarmed water and stir. Once the PVA dissolves, add the mannitol. Dissolve the MgSO4 and CaCl2 in separate tubes and then add to the solution. Filter and store at 4°C for up to 6 months.

     

2.4 Embryo Culture

  1. 1.

    Petri dishes and mineral oil.

     
  2. 2.

    Embryo culture medium.

     
  3. 3.

    Equipment: Humidifier incubator.

     

2.5 Synchronization of Recipients

  1. 1.

    Gilts between 280 and 400 lb in good body condition and health.

     
  2. 2.

    Altrenogest Solution 0.22%, 2.2 mg/mL (Matrix®, Millsboro, DE, USA).

     
  3. 3.

    Human chorionic gonadotropin (hCG), 1,000 IU/mL.

     
  4. 4.

    Syringes and needles.

     

2.6 Loading of Embryos in the Transfer Tubing

  1. 1.

    Petri dish and 1-mL air syringe.

     
  2. 2.

    Sterile saline solution.

     
  3. 3.

    Holding medium: see Subheading 2.3.1.

     
  4. 4.

    Transfer tubing: 20-cm long polyethylene tube (PE90 tube, Becton Dickinson, Franklin Lakes, NJ, USA). Immerse the tubing on 70% alcohol and rinse it with holding medium before use.

     

2.7 Surgical Embryo Transfer

  1. 1.

    Recipient animals.

     
  2. 2.

    Flunixin meglumine injection (50 mg/mL), antibiotic and sterile saline solution (0.9%).

     
  3. 3.

    Ethanol and Nolvasan.

     
  4. 4.

    Drape, gauze and catgut chromic suture.

     
  5. 5.

    1-mL syringe.

     
  6. 6.

    Pack of surgical instruments.

     

2.8 Post-Surgical Care

  1. 1.

    Pregnant mare serum gonadotropin (PMSG; 1,000 IU/mL) and hCG (1,000 IU/mL).

     
  2. 2.

    Syringes and needles.

     

2.9 Vaccination and Deworming Protocols Prior to Farrowing

  1. 1.

    Dewormer of choice.

     
  2. 2.

    Vaccine 1—Bordetella bronchiseptica-Erysipelothrix rhusiopathiae-Pasteurella multocida bacterin-toxoid-rhinitis bacterin-toxoid.

     
  3. 3.

    Vaccine 2—Mycoplasma hyopneumoniae bacterin.

     
  4. 4.

    Vaccine 3—Clostridium perfringens type C and enterotoxigenic strains of Escherichia coli.

     
  5. 5.

    Syringes and needles.

     

2.10 Induction of Labor

  1. 1.

    Dexamethasone (2 mg/mL).

     
  2. 2.

    Cloprostenol sodium (263 μg/mL).

     
  3. 3.

    Syringes and needles.

     

2.11 Post-Natal Care of Piglets

  1. 1.

    Vaccines: Parvovirus vaccine-killed virus-Erysipelothrix rhusiopathiae-Leptospira canicola-grippotyphosa-hardjo-icterohaemorrhagiae-pomona bacterin, Bordetella bronchiseptica-Erysipelothrix rhusiopathiae-Pasteurella multocida bacterin-toxoid-rhinitis bacterin-toxoid and Mycoplasma hyopneumoniae bacterin.

     
  2. 2.

    Dewormer of choice.

     
  3. 3.

    Iron dextran complex injection (200 mg/mL), antibiotic and vitamin A and D.

     
  4. 4.

    Iodine solution and Nolvasan solution.

     
  5. 5.

    Scale, Pig tooth nipper, small V ear notcher, tail docker.

     
  6. 6.

    Syringes and needles.

     

3 Methods

3.1 Collection of Porcine Oocytes

3.1.1 Collection and Transport of Ovaries

  1. 1.

    All reusable materials used to collect ovaries and oocytes should be disinfected with 70% alcohol and/or Nolvasan solution.

     
  2. 2.

    Collect the ovaries in a clean container containing warmed washing solution as soon as the tissue is harvested from the carcasses.

     
  3. 3.

    Once enough number of ovaries is collected, discard the liquid from the container and add enough washing solution to cover the ovaries. Stir the ovaries, discard the liquid and wash ovaries with saline solution again. Repeat this process until the washing solution has a clear appearance. Alternatively, place the ovaries in a clean colander and rinse them thoroughly with washing solution until blood and small debris are no longer observed. This method is especially useful when a large amount of ovaries is processed (see Note 1).

     
  4. 4.

    Transport ovaries to the laboratory. Transport temperature and time of ovary collection to oocyte aspiration may influence the quality of oocytes. Thus, ovaries can be held in washing solution between 25 and 39°C, while time between collection and aspiration of ovaries should not exceed 6 h.

     

3.1.2 Aspiration of Follicles

  1. 1.

    Use a 20–22 G needle attached to a 5-mL air syringe to aspirate every follicle in the size range of 3–8 mm (see Note 3). Alternatively, a vacuum pump set at 40–50 mmHg can be used to aspirate the follicles. Direct the bevel of the needle towards the ovary and try to aspirate as many follicles as possible using a single point of entry. Apply only enough suction to empty the follicles and avoid spillage of follicular fluid.

     
  2. 2.

    Deposit the follicular fluid in a 50-mL conical tube. After aspirating all the ovaries, allow the oocytes to settle. Keep the follicular fluid at 38°C.

     

3.1.3 Collection and In Vitro Maturation of Oocytes

  1. 1.

    Using a Pasteur pipette attached to a suction bulb, slowly aspirate a portion of the pellet from the bottom of the 50-mL tube. Be careful not to disturb the pellet.

     
  2. 2.

    Place the pellet in a grid dish containing holding medium. Do not overload the search dish with a large volume of pellet. Otherwise, the medium will turn cloudy and it will impair the identification of good quality oocytes. Instead, use as many grid dishes as needed to search throughout the pellet (see Note 4).

     
  3. 3.

    Collect the oocytes as quick as possible to avoid a drastic drop in temperature. Porcine oocytes are very sensitive to cold or drastic changes in temperature.

     
  4. 4.

    Using a pulled glass capillary mouth pipette attached to suction bulb or a Drummond pipette, select oocytes with at least three layers of cumulus cells, and dark and homogeneous ooplasm.

     
  5. 5.

    Transfer the selected oocytes to a Petri dish containing clean holding medium. Place the dish on a dry bath to keep the oocytes warm.

     
  6. 6.

    Wash the oocytes three additional times with holding medium. At this point the oocytes should be free of debris and possible contaminants.

     
  7. 7.

    Rinse the oocytes in a dish containing equilibrated maturation medium (see Note 5).

     
  8. 8.

    Transfer 50 oocytes to each of the wells of a four-well dish containing 500 μL of maturation medium.

     
  9. 9.

    Incubate the oocytes at 38°C and 5% CO2 in 90% humidity for 38–42 h (see Note 6).

     

3.2 Establishment of Cell Lines and Preparation of Donor Cells

3.2.1 Establishment and Maintenance of Cell Lines

  1. 1.

    Fetuses should be decapitated and eviscerated before processing. If adult animals are used to establish the cell culture, the skin should be shaved and disinfected prior to the collection of the sample. The skin sample, of approximately 10-mm in diameter, can be collected using a biopsy punch or a scalpel. All the instruments used during the procedure should be sterile.

     
  2. 2.

    Place the skin sample or fetus in a sterile tube containing holding medium. The sample can be processed immediately or refrigerated. If kept at 4°C, biopsy samples will survive for at least 24 h and even up to 3–4 days, although the longer the time from collection to culture, the lower the likelihood of establishing a healthy cell line.

     
  3. 3.

    Transfer the sample to a culture dish. Finely chop the tissue using scissors or scalpels to about 1-mm cubes. Rinse the pieces with holding medium.

     
  4. 4.

    Place the pieces onto a tissue culture dish and add just enough culture medium to cover the culture surface of the dish and hold the tissue pieces in place.

     
  5. 5.

    Culture the explants under 5% CO2 in air and 90% humidity at 38°C.

     
  6. 6.

    Check the culture dish 3–5 days later. If the pieces have adhered, add enough culture medium to cover the explants. Cells will start to migrate from the explants 5–10 days after seeding.

     
  7. 7.

    After 5–7 days in culture, the medium should be changed every 3 days. With a sterile Pasteur pipette attached to a vacuum pump, remove the medium from the dish and add fresh medium.

     
  8. 8.

    After the primary culture is established and the cells reach 80–100% of confluence, the cell line can be sub-cultured. Discard the culture medium of the culture dish.

     
  9. 9.

    Add enough PBS to cover the surface of the dish and dilute the remaining culture medium. Gently, agitate the dish in an effort to rinse the cell culture.

     
  10. 10.

    Remove the PBS and the explants that come lose during the process. Repeat this process two more times.

     
  11. 11.

    Add enough prewarmed trypsin to cover the cell culture. Incubate the dish at 38°C for 5 min.

     
  12. 12.

    Check the dish under an inverted microscope to verify that the cells have detached from the bottom of the dish. Cells in primary cultures sometimes require longer incubation time in trypsin. If cells are not completely detached after 10 min, tap the sides of the dish vigorously.

     
  13. 13.

    Transfer the cell suspension into a 15-mL tube containing prewarmed culture medium.

     
  14. 14.

    Centrifuge the cells at 350  ×  g for 5 min and discard the supernatant.

     
  15. 15.

    Add enough culture medium to resuspend the cell pellet. Pipette up and down several times to obtain a cell suspension free of cell aggregates.

     
  16. 16.

    Count cells using a hemocytometer. Seed the desired number of cells into a new culture dish containing fresh medium. Generally, skin fibroblasts can be sub-cultured with a split ratio of 1:5 or 1:10. However, some cell lines may require higher or lower initial seeding densities to obtain a satisfactory growth curve.

     
  17. 17.

    Steps 8–16 can be repeated every time the cells reach 80–100% of confluence.

     

3.2.2 Electroporation and Selection of Donor Cells

  1. 1.

    Aseptic cell culture techniques should be utilized throughout the electroporation and selection procedure.

     
  2. 2.

    Cells from Subheading 3.2.1 can be used directly following removal from culture dish with trypsin. Alternatively, cells from Subheading 3.2.1 which have been cryopreserved at early passage can be used for electroporation immediately after thawing (see Note 7).

     
  3. 3.

    Prepare electroporation medium by mixing 75% cytosalts and 25% Opti-MEM medium.

     
  4. 4.

    Pellet 1–2  ×  106 cultured cells released by trypsin or cryopreserved cells after thawing in a 38°C water bath by centrifugation at 350  ×  g for 5 min.

     
  5. 5.

    Resuspend cells in 5 mL of electroporation medium and centrifuge at 350  ×  g for 5 min.

     
  6. 6.

    Resuspend cells in 800 mL of electroporation medium containing 20 μg of linearized DNA. Let cells incubate at room temperature in electroporation medium containing DNA for approximately 10 min, then load cells into electroporation cuvette.

     
  7. 7.

    Apply an electroporation pulse of 450 V and 350 μF capacitance.

     
  8. 8.

    Remove cells from the electroporation cuvette with a fine-tipped transfer pipette and place in a 25-cm2 tissue culture flask with culture medium. Culture for 48 h without selection to allow expression of the antibiotic resistance gene.

     
  9. 9.

    Release cells with trypsin and centrifuge at 350  ×  g for 5 min. Resuspend in selection medium consisting of culture medium plus 600 μg/mL of G418. Plate cells in 100-mm tissue culture plates at 2  ×  105 cells per plate in selection medium (see Note 8).

     
  10. 10.

    Culture under selection with medium changes every 3–4 days for approximately 10 days until colonies of 100–200 cells appear.

     
  11. 11.

    Isolate individual colonies by trypsinization using an appropriate diameter cloning ring for the size of the colony. Passage isolated colonies in wells of a 48-well plate with selection medium.

     
  12. 12.

    When surviving colonies reach 80–100% confluency, trypsinize and place the cells in a 24-well plate while utilizing approximately 10% of the cells for initial screening by PCR (see Note 9).

     
  13. 13.

    When colonies selected from the initial PCR screen reach 80–100% confluency, passage by trypsin release to a new well of a 24 well plate while cryopreserving approximately half of the cells in small aliquots of 1–2  ×  105 cells in 100 μL of freezing medium. Repeat this step until sufficient cell aliquots for nuclear transfer have been cryopreserved (see Note 10).

     
  14. 14.

    Expand colonies to larger tissue culture plates to obtain enough DNA for final screening by PCR and/or Southern blot if desired.

     

3.2.3 Preparation of Donor Cells for Cloning

  1. 1.

    Cells successfully transfected with the gene(s) of interest can be used fresh or stored in liquid nitrogen. It is recommended to freeze the cells to avoid prolonged time in culture and sequential passages.

     
  2. 2.

    Remove the vial containing the transgenic cells from the liquid nitrogen tank and hold it at room temperature for a couple of seconds.

     
  3. 3.

    Place the vial in a water bath until the medium thaws out.

     
  4. 4.

    Transfer the thawed cell suspension to a 15-mL tube containing cell culture medium.

     
  5. 5.

    Centrifuge the cell suspension at 350  ×  g for 5 min and discard the supernatant.

     
  6. 6.

    Add enough culture medium to resuspend the cell pellet. Pipette up and down several times to obtain a cell suspension free of cell aggregates (see Note 11).

     
  7. 7.

    Seed cells at an appropriate dilution such that the cells reach confluency 24–48 h prior to nuclear transfer.

     
  8. 8.

    Culture the cells until in a humidifier incubator at 5% O2, 5% CO2 balanced with N2.

     
  9. 9.

    Approximately 1–2 days after cells reach confluency, discard the culture medium from the dish.

     
  10. 10.

    Add enough PBS to cover the surface of the dish. Gently, agitate the dish in an effort to rinse the cell culture.

     
  11. 11.

    Remove the PBS from the dish.

     
  12. 12.

    Add enough prewarmed trypsin to cover the cell culture. Incubate the dish at 38°C for 5 min.

     
  13. 13.

    Check the dish under an inverted microscope to verify that the cells have detached from the bottom of the dish and transfer the cell suspension into a 15-mL tube containing prewarmed culture medium.

     
  14. 14.

    Centrifuge the cells at 350  ×  g for 5 min and discard the supernatant.

     
  15. 15.

    Resuspend the cells in 1 mL of holding medium and transfer the suspension to a 1.5-mL vial. The cells can be held at 38°C or room temperature for a couple of hours.

     

3.3 Production of Cloned Embryos

3.3.1 Removal of Cumulus Cells and Selection of Oocytes

  1. 1.

    Using a Pasteur pipette (with polished opening of ∼400 μm), transfer ≤300 expanded cumulus oocytes complexes (COC’s) in each well of a four-well dish containing 500 μL of hyaluronidase solution.

     
  2. 2.

    Pipette the COC’s up and down to remove some of the cumulus cells surrounding the oocytes.

     
  3. 3.

    Transfer the oocytes of each well into a 15-mL tube containing 2–3 mL of hyaluronidase solution.

     
  4. 4.

    Vortex the tube vigorously for 4–5 min. Use 5 mL of holding medium to rinse the walls of the tube forcing the oocytes adhered to the wall to settle in the medium.

     
  5. 5.

    Centrifuge the oocytes at 250  ×  g for 1 min and transfer the pellet to a dish containing holding medium.

     
  6. 6.

    Use a dissection microscope to select oocytes with intact membrane and transfer them to a new dish containing holding medium (see Note 12). Use a glass capillary of 250–300 μm of intern diameter to move the oocytes.

     
  7. 7.

    Wash the oocytes two more times in holding medium.

     
  8. 8.

    Place pools of 50–100 denuded oocytes in 50 μL drops of holding medium covered with mineral oil. Keep the oocytes at 38°C.

     

3.3.2 Enucleation of Mature Oocytes

  1. 1.

    Prepare the micromanipulation chamber by placing ∼100 μL of enucleation medium in the center of a depression slide (see Note 13) and cover the medium with mineral oil.

     
  2. 2.

    Under the microscope, place a pool of oocytes (see Note 14) on the depression slide containing enucleation medium. Incubate the oocytes in this medium for 10 min before exposure to UV light.

     
  3. 3.

    Use the holding pipette to hold an oocyte by applying pressure on the holding system and the aspiration needle to rotate the oocyte and search for the first polar body (PB). Discard all immature oocytes.

     
  4. 4.

    Place the PB at around 4–5 o’clock. Usually, the metaphase II (MII) is in the vicinity of the PB; however, it is recommended to verify the localization of the metaphase plate using UV light.

     
  5. 5.

    With the bevel facing down, insert the enucleation needle throughout the zona pellucida and remove the PB and metaphase plate.

     
  6. 6.

    Verify the removal of the MII by exposing the aspirated karyoplast to UV light. If the metaphase is still present, rotate the oocyte and relocate the MII around 4–5 o’clock. Remove the metaphase.

     
  7. 7.

    Discard the PB and cytoplasm from the pipette.

     
  8. 8.

    Place pools of 50–100 enucleated oocytes in 50 μL drops of holding medium covered with mineral oil.

     

3.3.3 Reconstruction of Embryos

  1. 1.

    Prepare the micromanipulation chamber by placing ∼100 μL of holding medium in the center of a depression slide and cover the medium with mineral oil.

     
  2. 2.

    Place the enucleated oocytes on the left side of the micromanipulation chamber and a pool of donor cells on the right hand corner of the depression slide (see Subheading 3.2). Allow the cells to settle.

     
  3. 3.

    Focus the microscope on the bottom of the chamber to visualize the cells. Aspirate a cell into the injection pipette (see Note 15).

     
  4. 4.

    Select an oocyte and keep it in place using the holding pipette. Insert the reconstruction pipette through the zona pellucida and deposit the donor cell into the perivitelline space. Remove the pipette out of the oocyte.

     
  5. 5.

    Place pools of 50–100 reconstructed oocytes in 50 μL drops of holding medium covered with mineral oil.

     
  6. 6.

    In a four-well dish, place 500 μL of holding medium in well one; 250 μL of holding and 250 μL of fusion medium in well two, 500 μL of fusion medium in well three and 500 μL of holding medium in well four. Place pools of 50–100 oocytes in well one of the fusion dish.

     
  7. 7.

    Transfer 10–15 oocytes to the well two and allow the oocytes to settle. Then, transfer the oocytes to the well three and allow them to settle.

     
  8. 8.

    Place the fusion chamber onto the stage of a dissection microscope and connect the wires to a BTX fusion machine.

     
  9. 9.

    Place the oocytes into the fusion chamber containing fusion medium. Move each oocyte until the plane of contact between the cell and the ooplasm is parallel to the electric bars of the fusion chamber (see Note 16).

     
  10. 10.

    Fusion is induced with an AC pulse of 5 V for 5 s followed by two DC pulses of 1.5 kV/cm for 60 μs (see Note 17).

     
  11. 11.

    Wash the oocytes in the well four of the fusion dish and then transfer the couplets to 50 μL drops of holding medium covered with mineral oil.

     
  12. 12.

    Repeat steps 7–11 for all the couplets.

     
  13. 13.

    Check fusion visually approximately 30 min after the fusion procedure. Discard the unfused oocytes.

     

3.3.4 Activation of Fused Oocytes

  1. 1.

    Fused oocytes can be activated 1–4 h post-fusion.

     
  2. 2.

    In a four-well dish, place 500 μL of holding medium in well one; 250 μL of holding and 250 μL of activation medium in well two, 500 μL of activation medium in well three and 500 μL of holding medium in well four. Place pools of 50–100 oocytes in well one of the activation dish.

     
  3. 3.

    Transfer 20–30 oocytes to the well two and allow the oocytes to settle. Then, transfer the oocytes to the well three and allow them to settle.

     
  4. 4.

    Place the activation chamber onto the stage of a dissection microscope and connect the wires to a BTX fusion machine.

     
  5. 5.

    Place the oocytes into the activation chamber containing 700 μL of activation medium.

     
  6. 6.

    Activate the oocytes with an AC pulse of 5 V for 5 s followed by two DC pulses of 1.25 kV/cm for 60 μs.

     
  7. 7.

    Wash the oocytes in the well four of the activation dish and then transfer the couplets to 50 μL drops of holding medium covered with mineral oil.

     

3.4 Embryo Culture

Activated oocytes should be transferred into a recipient animal as soon as possible. However, the presumptive zygotes can be held overnight or for a couple of days in embryo culture medium.
  1. 1.

    Place pools of 10–15 activated couplets in 25 μL drops of embryo culture medium covered with mineral oil (see Note 18).

     
  2. 2.

    Incubate the embryos at 38°C in a humidifier incubator at 5% O2, 5% CO2, and 90% N2.

     

3.5 Synchronization of Recipients

The recipient animals should be synchronized and show signs of heat the day that cloning is performed, regardless of the day of embryo transfer (ET). If the embryos are to be transferred after some days in culture, the recipient gilts should have shown heat the day that cloning was performed.
  1. 1.

    House the gilts individually in synchronization crates or pens.

     
  2. 2.

    Deworm gilt at the same time as synchronization begins.

     
  3. 3.

    Days 1 through 14, feed recipients with 8 mL of altrenogest solution (18 mg/day) with ration once a day. Draw out with a 12-mL syringe, 8-mL of altrenogest solution and administer directly into the feed once per day. Repeat this process daily for 14 days. Wear protective gloves while handling the altrenogest solution and bottle.

     
  4. 4.

    At day 18, administer 500 IU of hCG intramuscularly in the neck, behind the ear or ham with a 3-mL syringe and 18 G 1 ½-in. needle.

     
  5. 5.

    At day 19, transfer 1-cell embryos.

     

3.6 Loading of the Embryo in the Transfer Tubing

  1. 1.

    After activation, place the reconstructed embryos in a dish containing holding medium.

     
  2. 2.

    Group the activated oocytes very tightly in the center of the dish.

     
  3. 3.

    Attach the transfer tubing to a 1-mL air syringe and aspirate enough holding medium to create a column of liquid of approximately 0.5 cm.

     
  4. 4.

    Then, create a column of about 0.5 cm of air in the tubing.

     
  5. 5.

    Place the tip of the transfer tubing directly on top of the cloned embryos and load them with the minimum possible amount of holding medium.

     
  6. 6.

    After all the embryos have been loaded (see Note 19), create a column of air followed by one of medium.

     
  7. 7.

    Carefully, detach the syringe from the transfer tubing and check that the embryos are still in the tubing.

     
  8. 8.

    The embryos should be held and transported at 38°C.

     

3.7 Surgical Embryo Transfer

  1. 1.

    The pigs are denied access to food or water the morning before surgery. One antibiotic treatment should be performed the day prior to surgery.

     
  2. 2.

    Sedate and anesthetize the recipient gilts. Follow the protocols recommended by the veterinarian in charge.

     
  3. 3.

    Place the pig on a dorsal position and disinfect the abdomen.

     
  4. 4.

    Administer 2 mL of flunixin meglumine and give one more antibiotic treatment prior to surgery.

     
  5. 5.

    Clean the abdominal area to remove any loose dirt and debris. Clip the ventral abdomen and clean and disinfect the skin using gauze soaked in Nolvasan scrub and gauze soaked in ethanol.

     
  6. 6.

    Cover the abdomen with a sterile drape and secure in place with towel clips. Cut an oval shape hole 7-cm wide by 20-cm long with operating scissors. Make a midline incision about 5–10-cm long through the abdominal wall between the last and second to last teats.

     
  7. 7.

    Carefully exteriorize one of the uterine horns. Insert the transfer tubing through the ostium of the infundibulum so that the end of the tubing is in the region of the ampullary-isthmic junction. Embryos are expelled from the tubing using the 1-mL syringe.

     
  8. 8.

    Rinse the uterus with 35°C sterile saline solution before returning it to the abdomen. Gently pour remaining sterile saline solution into the abdominal cavity to prevent possible adhesions.

     
  9. 9.

    Close the abdomen using an appropriate suture material and technique. The gilt is then taken to a recovery room until consciousness is regained.

     
  10. 10.

    Check on recipient gilt 2–4 h after surgery. Transport recipient gilt to gestation housing 12 h after surgery.

     

3.8 Post-Surgical Care

Several laboratories use a combination of PMSG and hCG injections after embryo transfer in an attempt to help to maintain pregnancy.
  1. 1.

    Recipient gilt must have a suture check at 24 and 48 h post-ET. Check for excess bleeding and swelling of abdominal area.

     
  2. 2.

    At day 11 post-ET, administer 1,000 IU of PMSG intramuscularly in the neck, behind the ear or ham.

     
  3. 3.

    At day 14 post-ET, administer 500 IU of hCG intramuscularly in the neck, behind the ear or ham.

     
  4. 4.

    An ultrasound should be performed at day 28 post-ET to confirm pregnancy. Weekly ultrasounds can be scheduled to verify the viability of the fetuses.

     

3.9 Vaccination and Deworming Protocols Prior to Farrowing

  1. 1.

    Five weeks prior to farrowing, administer vaccines 1 and 3 intramuscularly in the neck, behind the ear or ham.

     
  2. 2.

    Two weeks prior to farrowing, administer vaccines 1, 2, and 3 intramuscularly in the neck, behind the ear or ham.

     
  3. 3.

    Deworm accordingly to the product directions.

     
  4. 4.

    The vaccination schedule and type of vaccine used depends on the herd and health status.

     
  5. 5.

    Ten days prior to farrowing, transport recipient gilt to farrowing facility. Recipient gilt must be washed with soap or disinfectant upon arrival.

     

3.10 Induction of Labor

Labor should be induced when pregnancy has reached 120–121 days and no signs of parturition are apparent.
  1. 1.

    At day 1, administer 12 mL of dexamethasone and 2 mL of cloprostenol (see Note 20) intramuscularly in the neck, behind the ear or ham.

     
  2. 2.

    A day later, administer 1 mL of cloprostenol intramuscularly.

     
  3. 3.

    Pig should start showing signs of parturition 24–36 h after the first set on injections. Farrowing should occur no later than 48 h after initial treatment.

     

3.11 Post-Natal Care of Piglets

  1. 1.

    Dip umbilical cord in iodine solution as soon as possible after birth.

     
  2. 2.

    Process newborn litter 24–48 h after birth or when suitable for piglets health.

     
  3. 3.

    Assign litter and pig number at day 1. Notch the right ear with the litter number and the left ear with the individual number. Never repeat a litter/piglet number, it can be very confusing to track specific animals.

     
  4. 4.

    Place the piglet in lateral recumbency between arm and side. Restrain the head with your hand. Locate the long, sharp teeth on both sides of the upper and lower jaw. Cut teeth off slightly above the gum line.

     
  5. 5.

    Hold the piglet by one rear leg so that it is suspended in the air upside down, place tail docker around tail so that approximately 3/4 in. will remain, and with the crimped side facing the piglet, dock the tail. Rinse instruments with Nolvasan solution between piglets.

     
  6. 6.

    With piglet in same position, administer 1 mL of iron dextran complex injection, antibiotic, and vitamin A and D. Deliver these injections intramuscularly in the hams-two injections per side. Withdraw needle slowly and place finger firmly on injection sites for several seconds so as to prevent medications from coming out.

     
  7. 7.

    Place piglet on scale to determine weight and record on sheet.

     
  8. 8.

    At day 21, administer the vaccines intramuscularly into the ham and deworm.

     
  9. 9.

    Wean 1–2 weeks after vaccination to minimize the stress on the piglet.

     
  10. 10.

    All instruments must be cleaned and autoclave prior to their next use.

     

4 Notes

  1. 1.

    Chance of contamination generally increases in abattoirs with poor sterilization techniques or when a large quantity of ovaries is collected. To prevent contamination carried over from the surface of biological tissues, ovaries can be immersed for 3–5 min in washing solution containing 1–2% of Betadine. Rinse the ovaries thoroughly with washing solution to remove any traces of the antiseptic.

     
  2. 2.

    The cytosalts formulation (4) is intended to more closely mimic intracellular ionic conditions rather than extracellular conditions and thus limit loss of viability upon electroporation. Alternatively, electroporation can be carried out in Opti-MEM.

     
  3. 3.

    Substances present in the rubber plunger tips of syringes have been reported to have detrimental effects on oocytes and embryos. Only air syringes should be used during aspiration of oocytes, preparation of culture media, and manipulation of embryos (5, 6).

     
  4. 4.

    Alternatively, remove the supernatant from the 50-mL tube and add 50 mL of Ringer lactate or PBS supplemented with 0.1% of BSA. Invert the tube carefully to resuspend the pellet. Place the content of the tube in an embryo filter with a 75-μm mesh nylon membrane and filter the cell suspension. Rinse the filter with abundant medium to eliminate blood and other small particles. Do not allow the membrane to dry out at any time. Place the cell suspension remaining on top of the membrane on the search dish.

     
  5. 5.

    Successful in vitro maturation of porcine oocytes has been achieved using a variety of culture media types (simple or complex) containing fetal calf serum (7) or follicular fluid (8) and other supplements, such as gonadotropins and growth factors (9). Additionally, serum-free media have been successfully used for the in vitro maturation of pig oocytes (10).

     
  6. 6.

    Some protocols describe a sequential maturation system. In these protocols, the oocytes are cultured in maturation medium containing gonadotropins for the first 20–22 h before being changed to a gonadotropin-free medium for additional 18–20 h.

     
  7. 7.

    Cells can be expanded post thaw prior to electroporation if needed. This is not recommended unless necessary. The overall goal is to have cells for nuclear transfer with a minimum number of population doublings and expansion will only increase the number of population doublings.

     
  8. 8.

    The seeding density of 2  ×  105 is a starting point and may need to be adjusted for different DNA constructs or donor cell lines. The goal is to form single cell colonies that are sufficiently separated to allow isolation. Too low of a seeding density may decrease cell viability.

     
  9. 9.

    One of the direct PCR reagents sold by several molecular biology reagents suppliers that eliminate the need to purify DNA can be beneficial at this step.

     
  10. 10.

    Three or four cryopreserved cell aliquots should be sufficient for any single colony. If the nuclear transfer procedures described here are followed and a pregnancy does not occur after four embryo transfers, the investigator is advised to try a different cell line.

     
  11. 11.

    It is recommended to culture the cells for a couple of days to allow them to recover from the freezing–thawing procedure. However, frozen/thawed cells can be used directly as donors without intermediate culture.

     
  12. 12.

    Remaining cumulus cells after the vortex procedure can be stripped by pipetting the oocytes up and down with a glass capillary of an internal diameter of 180–200 μm.

     
  13. 13.

    Alternatively, a microscope slide surrounded by a thick wall of silicone or a 100-mm dish can be used as micromanipulation chamber.

     
  14. 14.

    Place as many oocytes on the micromanipulation medium as you can enucleate in about 30–60 min. Incubation of the oocytes in Hoechst for extended periods of time may have detrimental effects on embryo development.

     
  15. 15.

    Insert as many cells as possible in the injection pipette without compromising the suction control.

     
  16. 16.

    If the membranes of the ooplasm and the somatic cell are not in contact, you can reduce the osmolarity of the fusion medium by adding water. An osmolarity below 260 mM will induce a rapid increase in the volume of the cytoplasms, better contact between cells and consequently higher fusion rates. However, dramatic reduction in the osmolarity will cause lysis of the oocyte.

     
  17. 17.

    BTX offers fusion/activation chambers with different gap sizes. The voltage settings and the volume of medium should be adjusted accordingly to the gap size. Additionally, the number of oocytes that can be efficiently aligned at once depends of the space between gaps.

     
  18. 18.

    A variety of culture media have been used to successfully culture porcine oocytes. Common culture media include but are not limited to NCSU-23 (11), BECM-3 (12), and PZM-3 (13).

     
  19. 19.

    The appropriate number of embryos that should be transferred per recipient animal depends of factors such as the cell line used as donor, the transgene and the quality of oocytes. Under optimal conditions, transfer of 100–200 cloned embryos into a single recipient should be sufficient to establishing a pregnancy.

     
  20. 20.

    Estrumate tends to work better that Lutalyse for induction of parturition.

     

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Copyright information

© Springer Science+Business Media, LLC 2012

Authors and Affiliations

  • Angelica M. Giraldo
    • 1
    • 2
  • Suyapa Ball
    • 1
  • Kenneth R. Bondioli
    • 3
  1. 1.Revivicor, Inc.BlacksburgUSA
  2. 2.DeSoto Biosciences Inc.SeymourUSA
  3. 3.Louisiana State University Agricultural Center, School of Animal SciencesBaton RougeUSA

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