Materials and Reagents

Personal protective materials–hair net, gloves
4-0 black silk sutures (Kent Scientific, catalog number: INS701073 )
9 mm wound clips (BD, catalog number: 427631 )
Kimwipes (KCWW, Kimberly-Clark, catalog number: 34155 )
Insulin syringes (BD, catalog number: 329412 )
Iodine swabs (PDI Healthcare, catalog number: S41350 )
(Optional) Rodent mask diaphragms (Smiths Medical, Surgivet, catalog number: 32247B1 )
Sterile surgical drape
500 ml filter system
Fertilized one cell Sprague Dawley embryos (see Notes)
8-week old female Sprague Dawley recipient rats
Vasectomized Sprague Dawley male rats
Luteinizing hormone releasing hormone agonist (LHRHa) (Sigma-Aldrich, catalog number: L4513 )
70% ethanol
Buprenorphine (Southern Anesthesia & Surgical, catalog number: 12496075705 )
Carprofen (Zoetis Services, catalog number: 060062 )
Ampicillin (Fisher Scientific, catalog number: BP1760-25 )
0.1% epinephrine (Acros Organics, catalog number: 204400010 )
Sterile, nonmedicated ophthalmic ointment (Rugby Laboratories, catalog number: 301905 )
Embryo tested water (Sigma-Aldrich, catalog number: W1503 )
Sodium chloride (NaCl) (Sigma-Aldrich, catalog number: S5886 )
Potassium chloride (KCl) (Sigma-Aldrich, catalog number: P9333 )
Potassium phosphate monobasic (KH₂PO₄) (Sigma-Aldrich, catalog number: P9791 )
Magnesium sulfate heptahydrate (MgSO₄·7H₂O) (Sigma-Aldrich, catalog number: M1880 )
Glucose (Sigma-Aldrich, catalog number: 158968 )
Penicillin (Sigma-Aldrich, catalog number: P7794 )
Streptomycin (Sigma-Aldrich, catalog number: S1277 )
Sodium bicarbonate (NaHCO₃) (Sigma-Aldrich, catalog number: S5761 )
Sodium pyruvate (Sigma-Aldrich, catalog number: P4562 )
EDTA (Sigma-Aldrich, catalog number: 03609 )
L-Glutamine (Sigma-Aldrich, catalog number: G8540 )
Sodium lactate (Sigma-Aldrich, catalog number: L7900 )
Calcium chloride dihydrate (CaCl₂·2H₂O) (Sigma-Aldrich, catalog number: C7902 )
Bovine serum albumin (BSA) (Sigma-Aldrich, catalog number: A7906 )
Phenol red (Sigma-Aldrich, catalog number: P0290 )
Isoflurane (AMERISOURCE BERGEN, catalog number: 10103618 )
LHRHa solution (see Recipes)
KSOM medium (see Recipes) (Cold Spring Harbor, 2006)

Equipment

Biosafety cabinet
pH meter
Mouth pipet (Fisher Scientific, catalog number: NC9048719 )
Manufacturer: BIOTECH, model: MP001Y.
Glass pipettes (Fisher Scientific, catalog number: 13-678-20C )
Flame source to pull pipettes
Personal protective equipment–clean lab coat
9 mm wound clip applier (BD, catalog number: 427630 )
Microscope (Leica Microsystems, model: Leica S8 APO )
Fine forceps (Fine Science Tools, catalog number: 11251-10 )
Spring scissors (Roboz Surgical Instrument, catalog number: RS-5650 )
Large scissors (Roboz Surgical Instrument, catalog number: RS-5910 )
Grip forceps (Roboz Surgical Instrument, catalog number: RS-8100 )
Micro clip (Roboz Surgical Instrument, catalog number: RS-5420 )
Versi-Dry surface protectors (Thermo Fisher Scientific, Thermo Scientific^TM, catalog number: 74000-00 )
37 °C warming plate (C & A Scientific, Premiere, catalog number: XH-2002 )
Animal clippers (Oster, catalog number: 078005-301-003 )
Bead sterilizer (CellPoint Scientific, catalog number: 5-1450 )
Anesthesia machine (Smiths Medical, Surgivet, catalog number: WWV9000 )
Rodent anesthesia circuit set (Smiths Medical, Surgivet, catalog number: V7103 )
Large rubber bands

Procedure

Between 11:00 AM and 1:00 PM five days before surgery, 0.2 ml of 200 μg/ml LHRHa solution (see Recipes) is injected into the intraperitoneal cavity of the female recipient rats to synchronize estrus.
At noon one day prior to surgery the female recipient rats are mated with vasectomized males.
The following morning females are checked for the presence of a vaginal copulation plug as proof of mating. Females with vaginal plugs are used in the procedure and negative females are placed back into the colony for two weeks before reuse.
The surgeon should wear a clean lab coat, hair net and gloves. A disinfected area is prepared by wiping the base of the microscope with 70% ethanol and placement of a clean surface protector. Embryos to be transferred are produced and collected as previously described (Filipiak and Saunders, 2006) and are incubated in KSOM (see Recipes) at 37 °C up to 24 h before the transfer.
Instruments are autoclaved before the first surgery and sterilized in between up to four additional surgeries. Between surgeries, wipe instruments with sterile saline then insert the tip of each instrument into a 250 °C bead sterilizer for 15 sec. Allow to cool a minimum of one minute on a sterile Kimwipe before use.
Anesthesia is induced in the recipient rat typically within 3-5 min of placement in a chamber with 1.0-1.5 liters per minute (L/min) of 5% isoflurane with oxygen as a carrier gas. Anesthesia is maintained during surgery with 3% isoflurane by placement of the nose of the preanesthetized rat in a nose cone (Figure 1). A diaphragm can be created using a glove and rubber band and cutting a small hole to fit snugly around the nose of the anesthetized rat (see Notes).

Figure 1. Anesthetized rat receiving isoflurane through tight fitting nose cone
Apply ophthalmic ointment to the eyes by squeezing a thin layer of ointment onto the eye without contaminating the tube by touching the eye directly.
Separate subcutaneous injections of buprenorphine (0.05 mg/kg), carprofen (5 mg/kg) and 0.5 ml of ampicillin (100 mg/ml) also take place immediately prior to surgery.
Surgical depth of anesthesia is ensured first by observation of a depressed respiratory rate, then by repeated absence of tail- or foot-pinch reflex.
The lower back of the recipient rat is shaved above the left uterine horn, and placed on a sterile tissue on the stage of the microscope.
The shaved area is blotted with iodine and wiped with 70% ethanol-soaked tissue to prepare the surgical field and remove excess hair particles caused by shaving (Figure 2). This is repeated twice.

Figure 2. Rat fully prepared for surgery after shaving and sterilization. Incision location indicated by black line.
A sterile drape is placed over the animal.
A single incision about 1 cm in length in a dorsal to ventral direction is made in the skin using grip forceps and large scissors at the level of the last rib (Figure 3).

Figure 3. Initial incision through skin and subcutaneous fat. D: dorsal; V: ventral; SC: subcutaneous fat; BW: body wall.
The body wall is picked up with fine forceps and a small incision is made with spring scissors (avoiding blood vessels) just over the left ovarian fat pad (Figure 4).

Figure 4. Incision through body wall. D: dorsal; V: ventral; BW: body wall; OFP: ovarian fat pad.
The ovarian fat pad is grasped with fine forceps, and the distal portion of the reproductive tract is gently retracted from the abdominal cavity. The fat pad is held in place by an attached micro clip with a sterile piece of tightly rolled tissue under the uterus (Figure 5).

Figure 5. Externalized ovary and fat pad. D: dorsal; V: ventral; OFP: ovarian fat pad; OB: ovarian bursa; OV: oviduct; U: uterus.
A minimally vascularized area of the ovarian bursa is manually torn with fine forceps, providing a route to the ostium of the oviduct (Figure 6).

Figure 6. Location of minimally vascularized area of ovarian bursa. D: dorsal; V: ventral; OFP: ovarian fat pad; OB: ovarian bursa; OV: oviduct; U: uterus.
1-2 drops of epinephrine (approximately 20 μl) are dripped onto the bursa from a syringe until bleeding ceases. This decreases the length of the surgery and expedites location of the ostium.
The tip of the transfer mouth pipet is loaded with 15-20 embryos for implantation and inserted into the ostium of the oviduct after stabilization with fine forceps. The embryos are injected into the oviduct by gentle pressure through the pipette (Figure 7).

Figure 7. Placement of embryos into the reproductive tract. D: dorsal; V: ventral; OFP: ovarian fat pad; OB: ovarian bursa; OV: oviduct; U: uterus; O: ostium.
The reproductive tract is carefully replaced in the abdomen, and the abdomen is closed by 2 sutures in the body wall (Figure 8) followed by skin closure with 2 wound clips.

Figure 8. Location of sutures in peritoneal wall. D: dorsal; V: ventral; SC: subcutaneous fat; BW: body wall.
Following surgery, the recovering rat is placed inside a rat cage resting on a 37 °C warming plate until the animal is fully awake and upright, typically five to ten minutes (see Notes).
The cage is returned to a cage rack in a rat room and provided subcutaneous injection 0.2 ml carprofen (5 mg/ml) after 24 h as post-surgical analgesia. Up to three females can be housed together.
The rats are monitored daily to verify recovery from the procedure. Females are separated 20 days after the procedure to give birth in individual cages.

Data analysis

The research paper detailing generation of genetically modified rats using oviduct transfer surgery is available online (Lambert et al., 2016).

Notes

Common strains of Rattus norvegicus include Sprague Dawley, Wistar, Dahl SS and F344. No sex determination of the embryos is necessary. Taconic and Charles River are recommended vendors.
Two-cell embryos can also be transferred by this method.
Between 5 and 25 embryos can be transferred to each female.
An additional suture and/or wound clip can be used if the incision is larger than 1 centimeter.
Rats have an excessive amount of fat compared to mice. Be careful to handle the fat pad without puncturing or tearing the tissue.
Diaphragms can also be purchased with or without openings and cut to the desired size (Surgivet).
Indications of a failed operation include depressed respiratory rate, labored breathing and/or gasping and lethargy.

Recipes

LHRHa solution
Dissolve 1 mg of LHRHa powder in 5 ml sterile water for 200 μg/ml stock
Store at -20 °C in 1 ml aliquots
KSOM (Cold Spring Harbor, 2006)
500 ml embryo tested water
2.775 g NaCI
0.095 g KCl
0.025 g KH₂PO₄
0.025 g MgSO₄·7H₂O
0.02 g glucose
0.03 g penicillin
0.025 g streptomycin
1.05 g NaHCO₃
0.01 g sodium pyruvate
0.002 g EDTA
0.073 g L-glutamine
0.935 g sodium lactate
0.125 g CaCl₂·2H₂O
0.5 g bovine serum albumin
50 μl phenol red
Combine reagents in 175 ml embryo tested water. Adjust volume to 500 ml. Using a 500 ml filter system, filter the solution in a biosafety cabinet. Check the pH and store remaining solution at 4 °C. Check the pH after incubation (pH should drop); ideal pH is 7.1-7.7

Acknowledgments

The University of Alabama at Birmingham Transgenic & Genetically Engineered Models facility (RAK) is supported by the National Institutes of Health [grant numbers P30 CA13148, P30 AR048311, P30 DK074038, P30 DK05336 and P60 DK079626] and by the UAB Cystic Fibrosis Research Center. All procedures were conducted with the approval of the IACUC and the UAB Animal Resources Program (ARP) and are in compliance with guidelines for the care and use of laboratory animals and rodent survival surgery (Bernal et al., 2009; Albus, 2012; Cunliff-Beamer, 1993; Waynforth, 1992). We acknowledge the work of Filipiak and Saunders which served as the baseline for adaptation.

References

Albus, U. (2012). Guide for the care and use of laboratory animals (8^th edition). Laboratory Animals 46(3): 267-268.
Bernal, J., Baldwin, M., Gleason, T., Kuhlman, S., Moore, G. and Talcott, M. (2009). Guidelines for rodent survival surgery. J Invest Surg 22(6): 445-451.
Cold Spring Harbor. KSOM. (2006). Cold Spring Harb Protoc.
Cunliff-Beamer, T. L. (1993). Applying principles of aseptic surgery to rodents. AWIC Newsletter 4(2): 3-6.
Filipiak, W. E. and Saunders, T. L. (2006). Advances in transgenic rat production. Transgenic Res 15(6): 673-686.
Hankenson, F. C. (2013). Critical care management for laboratory mice and rats. CRC Press.
Krinke, G. J., Bullock, G. R. and Bunton, T. (2000). The laboratory rat. Elsevier Science.
Lambert, L. J., Challa, A. K., Niu, A., Zhou, L., Tucholski, J., Johnson, M. S., Nagy, T. R., Eberhardt, A. W., Estep, P. N., Kesterson, R. A. and Grams, J. M. (2016). Increased trabecular bone and improved biomechanics in an osteocalcin-null rat model created by CRISPR/Cas9 technology. Dis Model Mech 9(10): 1169-1179.
Smith, J. C., Corbin, T. J., McCabe, J. G. and Bolon, B. (2004). Isoflurane with morphine is a suitable anaesthetic regimen for embryo transfer in the production of transgenic rats. Lab Anim 38(1): 38-43.
Waller, S. J., Ho, M. Y. and Murphy, D. (1995). Production of transgenic rodents by microinjection of cloned DNA in fertilized one-cell eggs. In: Glover, D. M. and Hames, B. D. (Eds). DNA cloning. Vol. 4. Oxford University Press pp: 184-229.
Waynforth, H. B. and Flecknell, P. A. (1992). Experimental and surgical technique in the rat. 2nd edition. Academic Press.