Utero-tubal Embryo Transfer and Vasectomy in the Mouse Model

Summary

Utero-tubal embryo transfer uses the utero-tubal junction as a barrier to prevent the embryo outflow that may occur when performing uterine transfer. Vasectomized males are required to obtain pseudopregnant recipients for embryo transfer. Both techniques are discussed.

Abstract

The transfer of preimplantation embryos to a surrogate female is a required step for the production of genetically modified mice or to study the effects of epigenetic alterations originated during preimplantation development on subsequent fetal development and adult health. The use of an effective and consistent embryo transfer technique is crucial to enhance the generation of genetically modified animals and to determine the effect of different treatments on implantation rates and survival to term. Embryos at the blastocyst stage are usually transferred by uterine transfer, performing a puncture in the uterine wall to introduce the embryo manipulation pipette. The orifice performed in the uterus does not close after the pipette has been withdrawn, and the embryos can outflow to the abdominal cavity due to the positive pressure of the uterus. The puncture can also produce a hemorrhage that impairs implantation, blocks the transfer pipette and may affect embryo development, especially when embryos without zona are transferred. Consequently, this technique often results in very variable and overall low embryo survival rates. Avoiding these negative effects, utero-tubal embryo transfer take advantage of the utero-tubal junction as a natural barrier that impedes embryo outflow and avoid the puncture of the uterine wall. Vasectomized males are required for obtaining pseudopregnant recipients. A technique to perform vasectomy is described as a complement to the utero-tubal embryo transfer.

Introduction

Embryo transfer is probably the most frequent surgical procedure performed in the mouse model. This technique is essential to obtain offspring from embryos subjected to in vitro manipulation techniques and, therefore, constitutes a necessary step for the development of genetically modified models by pronuclear injection, lentiviral transduction, or chimera formation. Besides, the technique allows the study of the developmental effects of diverse insults occurring during preimplantation development. The use of artificial reproduction techniques¹ or the exposure to abnormal concentrations of different substances or metabolites² may affect embryo development resulting in implantation or placentation failures and long term effects in the offspring. A reliable and reproducible embryo transfer technique is crucial to test the possible negative effects of the experimental treatment on implantation and fetal development in a consistent manner.

Murine preimplantation embryos can be transferred to a recipient female either into the oviduct via the ampullae of 0.5 days post coitum (dpc) pseudopregnant recipients (oviduct transfer)^3,4 or into the uterus of 2.5 dpc pseudopregnant recipient (uterine transfer)^5,6 depending on their developmental stage. Embryos at the blastocyst stage, such as those used to generate chimeric mice by injection of embryonic or induced pluripotent stem cells, are usually transferred by uterine transfer. Blastocysts can also be transferred to the oviduct of a 0.5 dpc recipient, but it constitutes a less physiological test for developmental disruptors, because the embryo undergoes diapause and has 2 days to recover from the insult before implantation takes place. Uterine transfer involves puncturing the uterine wall with a narrow needle in order to generate an aperture that allows the access of an embryo manipulation pipette into the uterine lumen. Although this technique can yield good results, the survival to term (i.e. the percentage of embryos transferred that develop to a pup) is often low and unpredictable^7,8.

The puncture of the uterine wall entails some detrimental side effects. First, myometrium is a highly vascularized tissue and its puncture often results in a small hemorrhage. Blood may block the embryo transfer pipette or invade the uterine lumen causing embryonic death and/or implantation failure. This is particularly relevant when embryos without zona are transferred, as the blood cells and debris can attach to the blastomeres. Second, the opening performed does not seal after the embryos have been transferred, so they can flow back through the orifice and be expelled to the abdominal cavity when a too large volume has been introduce into the uterus. The utero-tubal embryo transfer described herein take advantage of the utero-tubal junction to deliver the embryos into the uterus without the need of puncturing the uterine wall and thereby avoiding its adverse consequences⁹.

The pseudopregnant recipient females used for embryo transfer are obtained by natural mating with vasectomized males⁸. The seminal secretions produced by a sterile male are required for the uterus to become receptive to the transferred embryos. To obtain a recipient, a maximum of 2 females of 8 weeks to 6 months of age are placed with a vasectomized male in the afternoon. The following morning, females are checked for the presence of a vaginal copulation plug, a clump of coagulated proteins from the male seminal fluid. As mating usually occurs during midnight, the day of vaginal plug detection is considered to be 0.5 dpc. Although vasectomized males can be purchased from some vendors, the surgical procedure described herein is relatively easy and does not require any additional instruments than required for embryo transfer.

Protocol

All animal experiments were approved by the Beltsville Area Animal Care and Use Comittees (BAACUC 11-015) in accordance with USDA Animal Care and Use Guidelines.

1. Anesthesia and Analgesia (Common for Both Surgical Procedures)

1. Weigh the mouse and load the following anesthetics and analgesic in two 1 ml syringes with 27 G needles:
   1. Ketamine (0.1 mg/g: 0.01 ml/g of a 10 mg/ml solution) and xylazine (0.01 mg/g: 0.005 ml/g of a 2 mg/ml solution).
   2. Buprenorphine (0.1 μg/g: 0.01 ml of a 0.01 mg/ml solution).
2. Immobilize the mouse by picking up the scruff of its neck as close to the jaws as possible with the thumb and index fingers and holding the tail between the little and ring fingers.
3. Inject Ketamine-Xylazine mixture intraperitoneally. In order to avoid puncturing internal organs, hold the mouse with its head slightly below the level of its hips (Figure 1A)
4. Inject Buprenorphine subcutaneously in the scruff of neck hold between the thumb and index fingers (Figure 1B).
5. Leave the mouse in the cage (clean and without any other animal) on a warm stage.
6. Once unconscious, check for the absence of rear foot reflex (checked by toe pinch). Apply eye ointment to avoid dryness of the eye and to check for the absence of palpebral reflex (Figure 1C).
7. This protocol provides a surgical anesthesia plane for a minimum of 30 min, enough to perform the procedures described below (Protocols 2 and 3). If longer times are required, an additional injection of ketamine + xylazine with half of the dosage described in 1.1.1 can be applied after 30 min. A change in the breathing pattern to a faster and irregular one indicates the loss of the proper anesthesia plane.

2. Vasectomy

1. Use a male with a proven mating performance.
2. Sterilize surgical instruments, clean the surfaces where surgery will be performed and wipe them with 70% ethanol.
3. Perform anesthesia as previously detailed (Protocol 1), checking for loss of reflexes.
4. Place the mouse on a warm stage, remove fur with electric clippers from the ventral area between two imaginary transversal lines placed 0.5 cm and 2.5 cm above the penis (Figure 2A).
5. Sanitize the shaved area by sequential wiping with 10% povidone iodine and 70% ethanol.
6. Place the mouse in supine position with its tail towards the surgeon and cover with a sterile towel with a hole exposing the shaved area. Illuminate the surgical area.
7. Perform a 10-15 mm longitudinal skin incision in the medial line of the abdomen, about 1 cm above the penis. Hold the skin with dressing serrated forceps and then cut with scissors (Figures 2A and 2B).
8. Perform a 5-10 mm longitudinal incision in the linea alba. Hold the muscle with microdissecting serrated forceps and cut with scissors (Figure 2C).
9. Grab the testicular adipose pad of one side with micro dissecting serrated forceps and pull it to expose testis, vas deferens and epididymis. Vas deferens is located medial to the testis and it is a clearly distinguishable free tube (not attached to the testis wall like the epididymis) with a blood vessel running along one side (Figure 2D).
10. Holding the vas deferens with a micro dissecting serrated forceps, flame dressing forceps until they turn red (Figure 2E), and then use them to cut and cauterize the vas deferens in two points at once (Figure 2F). The cut should remove a portion of about 5 mm and leave two clearly separated cauterized ends (Figure 2G).
11. Move the testicle, epididymis and vas deferens back to the abdominal cavity.
12. Proceed from step 9 in the other testicle.
13. Suture the muscle with one or two horizontal mattress stitches made with 5/0 absorbable suture (Figure 2H).
14. Suture the skin with one or two wound clippers (Figure 2I).
15. Identify the vasectomised male (ear ring, finger tattoo…), move it the cage placed on a warm stage and observe until it recovers from anesthesia (conscious and maintain sternal recumbency). A 0.5-1 ml subcutaneous injection of warm saline solution improves recovery. Record the possible incidences occurring during vasectomy transfer, add antibiotic to the drinking water.
16. Wound clips can be removed 10 days after vasectomy with a wound clipper remover or a pair of teeth forceps. The vasectomized male will be ready to mate 2 weeks after surgery.
17. Test the infertility of the vasectomized male by mating with fertile females before using it to obtain recipients.

3. Utero-tubal Embryo Transfer

1. Mouse morulae or blastocysts can be transferred by this technique to a pseudopregnant recipient female at 2.5 dpc.
2. Prepare embryo manipulation glass pipette:
   1. Polish the tips of the glass capillaries in order to avoid damaging the pipette holder.
   2. Soften a middle portion of the glass capillary by heating with a fine flame while slightly rotating the capillary with both hands synchronously. Once the capillary section becomes soft and malleable (light red color), withdraw it quickly from the flame and pull both ends to narrow its external diameter to 130-150 μm.
   3. Wait for the glass to cool down and then cut it by lightly scoring the narrow part with a diamond-point pencil, abrasive stone or nail file and pulling from both sides. The break should be clean and perpendicular
3. Polish the tip by very quickly flaming, leaving a 100-130 μm aperture. Pipettes can be stored for later use.
4. Warm embryo manipulation media (CZBH or M2, see discussion).
5. Sterilize surgical instruments, clean the surfaces where surgery will be performed and wipe them with 70% ethanol.
6. Perform anesthesia as previously detailed (Protocol 1), checking for loss of reflexes.
7. Keeping the mouse on a warm stage, remove fur with electric clippers from the dorsal area between the knees and the distal ribs (Figures 3A and 3B).
8. Sanitize the shaved area by sequential wiping with 10% povidone iodine and 70% ethanol.
9. Move the embryos from the incubator to the prewarmed embryo manipulation media.
10. Move the recipient to a warm stage under the stereomicroscope and place it in prone position laterally to the surgeon (with its head looking to the right or left side of the surgeon).
11. Cover the area with a sterile towel with a hole exposing the shaved area and illuminate the surgical area.
12. Perform a 1 cm transversal (vertical) incision in the skin in a spot located on the cranial ⅓ of the line between the last rib and the hips and the dorsal ⅓ of the line between the back and the abdomen (Figures 3A and 3B). Hold the skin with dressing serrated forceps and cut with scissors (Figure 3C).
13. Once the skin has been cut, ovary (red/orange) or the adipose pad surrounding the ovary (white) can be visualized through the body wall. Perform a 0.3-0.5 cm transversal (vertical) incision in the body wall over the ovary or adipose pad in a spot where the incision does not cut any large blood vessel. Hold the muscle with micro dissecting serrated forceps and cut with scissors (Figure 3D).
14. Move the mouse to have its head facing towards the surgeon.
15. Load the embryo manipulation pipette (Figure 3E):
    1. Allow CZBH media to ascend by capillarity through the narrow portion of the manipulation pipette until about 5 mm of the wider part.
    2. Take up a small air bubble (0.2-0.5 mm).
    3. Introduce the embryos (5-10) in a minimal amount of media (2-4 mm).
    4. Take up another small air bubble (0.2-0.5 mm) and a small amount of media (0.5-1 mm).
    5. Leave the glass pipette attached to the mouth aspirator holder or to the hand-operated device, ready for step 3.17.
16. Grab the adipose pad surrounding the ovary with micro dissecting serrated forceps and pull it towards the mouse head to expose the ovary, oviduct, and a small portion of the upper uterus out of the abdominal cavity (Figures 3F and 3G).
17. Holding the aspirator mouth piece in the mouth, ready to be used, grab the adipose pad with micro dissecting serrated forceps to move the oviduct and expose the utero-tubal junction (i.e. where the oviduct meets the uterus).
18. Keeping the utero-tubal junction accessible, take slight curved micro dissection forceps with your left hand (if right handed) and place them just below the utero-tubal junction grabbing the oviduct about 2 mm above that portion.
19. Holding the utero-tubal junction with the slight curved micro dissection forceps, puncture the oviduct section close to the forceps with a 27 G needle (Figure 3H).
20. Insert the embryo manipulation pipette into the orifice performed with the needle and advance to the uterus through the utero-tubal junction (Figures 3I and 3J). Once the pipette has passed the utero-tubal junction (Figure 3K) it slides easily. Do not progress very far into the uterus to prevent endometrial damage (no more than 3 mm) and pipette blocking by the debris.
21. Release the embryos into the uterus by gently blowing (Figure 3L). Both air bubbles must pass through the uterus. Some of the media above the first bubble may also be released into the uterus, but avoid introducing more air, as it may impede implantation.
22. Remove the pipette just after embryos have been released into the uterus.
23. Move the oviduct and ovary back to the abdominal cavity by grabbing the adipose pad.
24. Suture the muscle with a horizontal mattress stitch with 5/0 absorbable suture (Figure 3M).
25. Suture the skin with a wound clipper (Figure 3N).
26. Proceed from step 10 on the other side if required.
27. Identify the recipient (ear ring, finger tattoo…), move it to its cage (placed on a warm stage) and observe until it recovers from anesthesia (conscious and maintain sternal recumbency).
28. Annotate the possible incidences occurring during embryo transfer and add antibiotic to the drinking water. A 0.5-1 ml subcutaneous injection of warm saline solution improves recovery.
29. Wound clips can be removed 10 days after embryo transfer with a wound clipper remover or two pairs of forceps (Figure 3O). The recipient can be weighed on that day to assess pregnancy and estimate the number of pups. Provide nestling material to the recipient 15 days after embryo transfer.

Results

Utero-tubal embryo transfer provides a mean to transfer embryos to the uterus avoiding some of the complications associated to the uterine embryo transfer^2,9,10. In Table 1 we show some representative result we have obtained transferring CD1 blastocysts subjected to different kinds of manipulations to CD1 recipients following the protocol described. The survival to term (% of embryos resulting in a pup) or survival to E15 (in the case of lentivirus exposed) is similar between embryos simply cu...

Discussion

Vasectomy is a relatively straight forward surgical technique that does not involve major difficulties. When sanitizing with povidone iodine and ethanol make sure that the last wash (with ethanol) removes povidone iodine, as it may irritate the peritoneum. The access to vas deferens can also be achieved by the scrotum or performing a transversal incision in the abdomen⁸. Scrotal incision has been recommended to transversal abdominal incision due to the comparatively smaller incision needed and slightl...

Materials

List of materials used in this article

Name	Company	Catalog Number	Comments
Ketamine	VEDCO	Ketaved ANADA 200-257	To be ordered by a licensed veterinarian.
Xylazine	Lloyd Laboratories	Anased NADA #139-236	To be ordered by a licensed veterinarian.
Buprenorphine	Generic	NDC 400-42-010-01	To be ordered by a licensed veterinarian.
Eye ointment	Novartis	Genteal
Antibiotic	Pfizer	Clavamox NADA #55-101.	Added to drinking water (0.3 mg/ml of amoxicillin trihydrate and 0.075 mg/ml of clavulanate potassium). Add 1.5 ml of the reconstituted 15 ml bottle to 250 ml of water.
Dressing serrated forceps	ROBOZ	RS-8120	Any medium size surgical-grade steel straight forceps will work.
Microdissecting serrated forceps	ROBOZ	RS-5137	These ones are curved at a 90º angle. Straight forceps can be used if preferred.
Slight curved micro dissection forceps	ROBOZ	RS-5136	This model is particularly useful to hold the oviduct.
Scissors	ROBOZ	RS-5880	Any regular surgical grade steel small straight scissors will work.
27 G needles	Beckton-Dickinson	305136	Smaller needles (30 G) can be also used. 25 G may be a bit too big.
Clip applier	MiKRon	42763
9 mm Clips	MiKRon	427631
Clip remover	MiKRon	7637	Two pairs of teeth forceps (ROBOZ RS-8160) can be used instead.
Suture needle holder	ROBOZ	RS-7820
Suture	Dowist Gell	5-0 Dexon S 7204-21	Can be substituted for any 4-0 to 6-0 absorbable suture with a narrow curved needle.
Glass capillaries	VWR	100 ul calibrated pipettes 53432-921	It includes a mouth aspirator system that only requires to attach a 0.22 µm filter in the tubing to be ready to use. More information can be obtained in Nagy et al.8
Burner	KISAG AG	Typ 2002	Gas operated burner, can be charged with Kigas (CH-4512, from the same vendor). Alcohol burners may be also used, but gas provides a higher temperature and this burner provides a small and precise flame.
Stereomicroscope	Leica	MZFLIII	This is an expensive estereomicroscope with fluorescence, that can be also used for other purposes. There are cheaper options such as Leica MZ8 or Nikon SMZ-10 or SMZ-2B, to name a few. It is better to use two stereomicroscopes, one for handling the embryos (which does not need to be a very nice one) and another one for the recipient. The one used for the recipient should display a long distance from the stage plate to the objective lense, in order to be able to focus 3-4 cm above the stage plate (where the oviduct will be placed) and still leave some room for the surgeon; most of the stereomicroscopes can do this, but some cannot.
Fiber optics ilumination	Dolan Jenner	Fiber lite	To iluminate the surgical area. There are different systems available.
Warm stages	American scope	http://store.amscope.com/tcs-100.html	These can be placed over the stage plate of the stereomicroscope. Some modifications (inserting a stick to level the stage) may be needed if it is too short for the stereomicroscope. A big warm stage can be used for warming the cage if it is available. If not, a regular heating pad can be used, but temperature must be checked.
Culture dishes for embryo manipulation	Falcon	353001	351008 may be also used; they made narrower drops.