2 users have reported that they have successfully carried out the experiment using this protocol.
Improved Oviduct Transfer Surgery for Genetically Modified Rat Production

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Disease Models & Mechanisms
14-Oct 2013


Rat embryo transfer surgeries are becoming more common with targeted nucleases increasing the demand for rat models. This protocol details pre-surgical preparation, improved surgical techniques for placing embryos into the oviduct, and post-surgical care of rats to parturition. Direct application of mouse oviduct transfer protocols results in limited success in the rat. By combining techniques from several widely used protocols in the field, increased yield of live pups born to healthy dams is possible. This protocol is distinct from previously published protocols by the use of a modified anesthesia protocol (Smith et al., 2004), use of analgesia, the addition of peritoneal sutures (Filipiak and Saunders, 2006), incision location and number of transfers per animal (Krinke et al., 2000).

Keywords: Rat (大鼠), Transgenics (转基因), Embryos (胚胎), Surgery (手术), Oviduct (输卵管)


The ability to reliably produce healthy pups after microinjection and embryo transfer surgery is critical to model creation and, in particular, the increased likelihood of creating multiple founder animals gives confidence in the phenotypes observed. Therefore, as birth rates were low relative to reported rates even with varied concentrations of injection solution, modifications were systematically made to the existing mouse embryo transfer protocol to better suit the rat.

Multiple publications describe transferring embryos to the oviducts of both horns of the bipartite uterus; however, this increases the length of time the animal is under anesthesia and requires either a midline incision and traversing the peritoneal cavity to reach the lateral reproductive tract, or creating two separate incisions (Krinke et al., 2000). These options are less than ideal since either will increase stress of the animal and thereby the likelihood that the pregnancy will be aborted. By creating a single lateral incision and administering analgesia both preoperatively and postoperatively the stress of the animal is minimized (Smith et al., 2004). The use of isoflurane over injectable anesthetic agents minimizes risk of toxicity (such as seen with tribromoethanol), injury from IP injection, and repeated dosing, all of which are associated with higher mortality rates following rodent surgery (Bernal et al., 2009).

The greatest improvements in litter number and size followed the addition of ampicillin and epinephrine to the procedure [62 born/298 transferred (20.8%) versus 91 born/248 transferred (36.7%) post addition of ampicillin and epinephrine; all projects]. Although the surgery is performed aseptically, ampicillin was shown as early as 1995 to optimize the number of pups born to rats (Waller, 1995) and use of epinephrine on the ovarian bursa reduced bleeding and thereby trauma to the animal, as well as reduced the length of time required to find the infundibulum. These modifications have been used individually in multiple reports; however, this is the first protocol to combine the most advantageous aspects of each protocol while refining procedures that may be detrimental (Krinke et al., 2000; Smith et al., 2004; Filipiak and Saunders, 2006).

Materials and Reagents

  1. Personal protective materials–hair net, gloves
  2. 4-0 black silk sutures (Kent Scientific, catalog number: INS701073 )
  3. 9 mm wound clips (BD, catalog number: 427631 )
  4. Kimwipes (KCWW, Kimberly-Clark, catalog number: 34155 )
  5. Insulin syringes (BD, catalog number: 329412 )
  6. Iodine swabs (PDI Healthcare, catalog number: S41350 )
  7. (Optional) Rodent mask diaphragms (Smiths Medical, Surgivet, catalog number: 32247B1 )
  8. Sterile surgical drape
  9. 500 ml filter system
  10. Fertilized one cell Sprague Dawley embryos (see Notes)
  11. 8-week old female Sprague Dawley recipient rats
  12. Vasectomized Sprague Dawley male rats
  13. Luteinizing hormone releasing hormone agonist (LHRHa) (Sigma-Aldrich, catalog number: L4513 )
  14. 70% ethanol
  15. Buprenorphine (Southern Anesthesia & Surgical, catalog number: 12496075705 )
  16. Carprofen (Zoetis Services, catalog number: 060062 )
  17. Ampicillin (Fisher Scientific, catalog number: BP1760-25 )
  18. 0.1% epinephrine (Acros Organics, catalog number: 204400010 )
  19. Sterile, nonmedicated ophthalmic ointment (Rugby Laboratories, catalog number: 301905 )
  20. Embryo tested water (Sigma-Aldrich, catalog number: W1503 )
  21. Sodium chloride (NaCl) (Sigma-Aldrich, catalog number: S5886 )
  22. Potassium chloride (KCl) (Sigma-Aldrich, catalog number: P9333 )
  23. Potassium phosphate monobasic (KH2PO4) (Sigma-Aldrich, catalog number: P9791 )
  24. Magnesium sulfate heptahydrate (MgSO4·7H2O) (Sigma-Aldrich, catalog number: M1880 )
  25. Glucose (Sigma-Aldrich, catalog number: 158968 )
  26. Penicillin (Sigma-Aldrich, catalog number: P7794 )
  27. Streptomycin (Sigma-Aldrich, catalog number: S1277 )
  28. Sodium bicarbonate (NaHCO3) (Sigma-Aldrich, catalog number: S5761 )
  29. Sodium pyruvate (Sigma-Aldrich, catalog number: P4562 )
  30. EDTA (Sigma-Aldrich, catalog number: 03609 )
  31. L-Glutamine (Sigma-Aldrich, catalog number: G8540 )
  32. Sodium lactate (Sigma-Aldrich, catalog number: L7900 )
  33. Calcium chloride dihydrate (CaCl2·2H2O) (Sigma-Aldrich, catalog number: C7902 )
  34. Bovine serum albumin (BSA) (Sigma-Aldrich, catalog number: A7906 )
  35. Phenol red (Sigma-Aldrich, catalog number: P0290 )
  36. Isoflurane (AMERISOURCE BERGEN, catalog number: 10103618 )
  37. LHRHa solution (see Recipes)
  38. KSOM medium (see Recipes) (Cold Spring Harbor, 2006)


  1. Biosafety cabinet
  2. pH meter
  3. Mouth pipet (Fisher Scientific, catalog number: NC9048719 )
    Manufacturer: BIOTECH, model: MP001Y.
  4. Glass pipettes (Fisher Scientific, catalog number: 13-678-20C )
  5. Flame source to pull pipettes
  6. Personal protective equipment–clean lab coat
  7. 9 mm wound clip applier (BD, catalog number: 427630 )
  8. Microscope (Leica Microsystems, model: Leica S8 APO )
  9. Fine forceps (Fine Science Tools, catalog number: 11251-10 )
  10. Spring scissors (Roboz Surgical Instrument, catalog number: RS-5650 )
  11. Large scissors (Roboz Surgical Instrument, catalog number: RS-5910 )
  12. Grip forceps (Roboz Surgical Instrument, catalog number: RS-8100 )
  13. Micro clip (Roboz Surgical Instrument, catalog number: RS-5420 )
  14. Versi-Dry surface protectors (Thermo Fisher Scientific, Thermo ScientificTM, catalog number: 74000-00 )
  15. 37 °C warming plate (C & A Scientific, Premiere, catalog number: XH-2002 )
  16. Animal clippers (Oster, catalog number: 078005-301-003 )
  17. Bead sterilizer (CellPoint Scientific, catalog number: 5-1450 )
  18. Anesthesia machine (Smiths Medical, Surgivet, catalog number: WWV9000 )
  19. Rodent anesthesia circuit set (Smiths Medical, Surgivet, catalog number: V7103 )
  20. Large rubber bands


  1. 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.
  2. At noon one day prior to surgery the female recipient rats are mated with vasectomized males.
  3. 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.
  4. 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.
  5. 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.
  6. 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

  7. 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.
  8. 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.
  9. Surgical depth of anesthesia is ensured first by observation of a depressed respiratory rate, then by repeated absence of tail- or foot-pinch reflex.
  10. 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.
  11. 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.

  12. A sterile drape is placed over the animal.
  13. 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.

  14. 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.

  15. 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.

  16. 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.

  17. 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.
  18. 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.

  19. 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.

  20. 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).
  21. 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.
  22. 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).


  1. 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.
  2. Two-cell embryos can also be transferred by this method.
  3. Between 5 and 25 embryos can be transferred to each female.
  4. An additional suture and/or wound clip can be used if the incision is larger than 1 centimeter.
  5. Rats have an excessive amount of fat compared to mice. Be careful to handle the fat pad without puncturing or tearing the tissue.
  6. Diaphragms can also be purchased with or without openings and cut to the desired size (Surgivet).
  7. Indications of a failed operation include depressed respiratory rate, labored breathing and/or gasping and lethargy.


  1. 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
  2. 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


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.


  1. Albus, U. (2012). Guide for the care and use of laboratory animals (8th edition). Laboratory Animals 46(3): 267-268.
  2. 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.
  3. Cold Spring Harbor. KSOM. (2006). Cold Spring Harb Protoc.
  4. Cunliff-Beamer, T. L. (1993). Applying principles of aseptic surgery to rodents. AWIC Newsletter 4(2): 3-6.
  5. Filipiak, W. E. and Saunders, T. L. (2006). Advances in transgenic rat production. Transgenic Res 15(6): 673-686.
  6. Hankenson, F. C. (2013). Critical care management for laboratory mice and rats. CRC Press.
  7. Krinke, G. J., Bullock, G. R. and Bunton, T. (2000). The laboratory rat. Elsevier Science.
  8. 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.
  9. 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.
  10. 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.
  11. Waynforth, H. B. and Flecknell, P. A. (1992). Experimental and surgical technique in the rat. 2nd edition. Academic Press.


大鼠胚胎移植手术正变得越来越普遍,靶向核酸酶增加对大鼠模型的需求。 该协议详细介绍了手术前准备,将胚胎置入输卵管的改进的外科技术以及大鼠分娩后的手术后护理。 直接应用小鼠输卵管转移方案在大鼠中的成功有限。 通过组合来自现场广泛使用的几种方案的技术,可以提高健康水坝出生的活的小狗的产量。 该方案与以前公开的方案不同,通过使用改进的麻醉方案(Smith等人,2004),使用止痛剂,添加腹膜缝线(Filipiak和Saunders,2006),切割位置和每只动物的转移数 (Krinke等,2000)。
   多个出版物描述了将胚胎转移到两侧子宫的两个角的输卵管;然而,这增加了动物在麻醉下的时间长度,并且需要中线切口并穿过腹膜腔以到达侧生殖道,或者产生两个单独的切口(Krinke等人,2000)。这些选择不太理想,因为这两个选项都会增加动物的压力,从而增加妊娠中止的可能性。通过在术前和术后创建单侧切口并施用镇痛,动物的压力被最小化(Smith等人,2004)。使用异氟烷可注射麻醉剂可以最大程度地降低毒性风险(如三溴乙醇所见),IP注射损伤和重复给药,所有这些都与啮齿动物手术后的死亡率相关(Bernal et al。,2009)。
   随着氨苄西林和肾上腺素的添加,氨苄青霉素和肾上腺素加入后,最大的改善就是将氨苄青霉素和肾上腺素加入[62例/ 298次转移(20.8%),而加入氨苄青霉素和肾上腺素后,转出91例/ 248例(36.7%);所有项目]。虽然手术是无菌进行的,但早在1995年就显示了氨苄青霉素来优化大鼠出生的幼仔数量(Waller,1995),并且使用肾上腺素对卵巢囊来减少出血,从而减少了对动物的创伤寻找漏斗所需的时间长短。这些修改已经在多个报告中单独使用然而,这是第一个将每个协议的最有利方面结合在一起的方案,同时可以对可能是有害的程序进行优化(Krinke等,2000; Smith等人,2004; Filipiak和Saunders,2006)。

关键字:大鼠, 转基因, 胚胎, 手术, 输卵管


  1. 个人防护用品 - 头发网,手套
  2. 4-0黑丝线(Kent Scientific,目录号:INS701073)
  3. 9毫米伤口夹(BD,目录号:427631)
  4. Kimwipes(KCWW,Kimberly-Clark,目录号:34155)
  5. 胰岛素注射器(BD,目录号:329412)
  6. 碘片(PDI Healthcare,目录号:S41350)
  7. (可选)啮齿动物面膜隔膜(Smiths Medical,Surgivet,目录号:32247B1)
  8. 无菌外科覆盖物
  9. 500 ml过滤系统
  10. 受精的一个细胞Sprague Dawley胚胎(见注释)
  11. 8周龄女性Sprague Dawley受体大鼠
  12. Vasectomized Sprague Dawley雄性大鼠
  13. 促黄体激素释放激素激动剂(LHRHa)(Sigma-Aldrich,目录号:L4513)
  14. 70%乙醇
  15. 丁丙诺啡(Southern Anesthesia& Surgical,目录号:12496075705)
  16. Carprofen(Zoeis Services,目录号:060062)
  17. 氨苄青霉素(Fisher Scientific,目录号:BP1760-25)
  18. 0.1%肾上腺素(Acros Organics,目录号:204400010)
  19. 无菌,非药物眼用软膏(Rugby Laboratories,目录号:301905)
  20. 胚胎测试水(Sigma-Aldrich,目录号:W1503)
  21. 氯化钠(NaCl)(Sigma-Aldrich,目录号:S5886)
  22. 氯化钾(KCl)(Sigma-Aldrich,目录号:P9333)
  23. 磷酸二氢钾(KH 2 PO 4)(Sigma-Aldrich,目录号:P9791)
  24. 硫酸镁七水合物(MgSO 4·7H 2 O)(Sigma-Aldrich,目录号:M1880)
  25. 葡萄糖(Sigma-Aldrich,目录号:158968)
  26. 青霉素(Sigma-Aldrich,目录号:P7794)
  27. 链霉素(Sigma-Aldrich,目录号:S1277)
  28. 碳酸氢钠(NaHCO 3)(Sigma-Aldrich,目录号:S5761)
  29. 丙酮酸钠(Sigma-Aldrich,目录号:P4562)
  30. EDTA(Sigma-Aldrich,目录号:03609)
  31. L-谷氨酰胺(Sigma-Aldrich,目录号:G8540)
  32. 乳酸钠(Sigma-Aldrich,目录号:L7900)
  33. 氯化钙二水合物(CaCl 2·2H 2 O)(Sigma-Aldrich,目录号:C7902)
  34. 牛血清白蛋白(BSA)(Sigma-Aldrich,目录号:A7906)
  35. 苯酚红(Sigma-Aldrich,目录号:P0290)
  36. 异氟烷(AMERISOURCE BERGEN,目录号:10103618)
  37. LHRHa溶液(参见食谱)
  38. KSOM培养基(见食谱)(Cold Spring Harbor,2006)


  1. 生物安全柜
  2. pH计
  3. 口吸管(Fisher Scientific,目录号:NC9048719)
  4. 玻璃移液器(Fisher Scientific,目录号:13-678-20C)
  5. 火焰源拉吸液管
  6. 个人防护装备 - 清洁实验室外套
  7. 9毫米伤口夹具(BD,目录号:427630)
  8. 显微镜(Leica Microsystems,型号:Leica S8 APO)
  9. 精镊子(精细科学工具,目录号:11251-10)
  10. 弹簧剪刀(Roboz Surgical Instrument,目录号:RS-5650)
  11. 大型剪刀(Roboz Surgical Instrument,目录号:RS-5910)
  12. 手镊(Roboz手术器械,目录号:RS-8100)
  13. 微型夹(Roboz Surgical Instrument,目录号:RS-5420)
  14. Versi-Dry表面保护膜(Thermo Fisher Scientific,Thermo Scientific TM ,目录号:74000-00)
  15. 37℃加热板(C& A Scientific,Premiere,目录号:XH-2002)
  16. 动物剪刀(Oster,目录号:078005-301-003)
  17. 珠粒灭菌器(CellPoint Scientific,目录号:5-1450)
  18. 麻醉机(Smiths Medical,Surgivet,catalog number:WWV9000)
  19. 啮齿动物麻醉电路组(Smiths Medical,Surgivet,目录号:V7103)
  20. 大橡皮筋


  1. 手术前五天上午11:00至下午1:00,将0.2ml200μg/ ml LHRHa溶液(见食谱)注射到雌性受体大鼠的腹腔内,以同步发情。
  2. 手术前一天中午,雌性受体大鼠与输精管切除的男性配对。
  3. 检查第二天早上女性是否存在阴道交配插塞作为交配证据。在手术中使用阴道插头女性,将阴性女性放回殖民地两周,然后重新使用。
  4. 外科医生应穿干净的实验衣,头发网和手套。通过用70%乙醇擦拭显微镜的底部并放置干净的表面保护剂来制备消毒区域。转移的胚胎如前所述(Filipiak和Saunders,2006)生产和收集,并在转移前至少24小时在37℃下在KSOM(见食谱)中孵育。
  5. 仪器在第一次手术前进行高压灭菌,并在多达四次额外的手术中进行灭菌。在手术之间,用无菌生理盐水擦拭仪器,然后将每个仪器的尖端插入250°C珠粒灭菌器15秒。在使用前,请勿在无菌Kimwipe上冷却至少1分钟。
  6. 通常在接种大鼠的3-5分钟内诱导麻醉,在室内以1.0-1.5升/分钟(L / min)的5%异氟醚与氧气作为载气。通过将预先麻醉的大鼠的鼻子放置在鼻锥中,通过3%异氟烷在手术期间维持麻醉(图1)。可以使用手套和橡皮筋创建膈肌,并切割一个小孔,紧贴麻醉大鼠的鼻子(见注释)。


  7. 通过将薄薄的一层软膏压在眼睛上,直接触摸眼睛而不污染管,将眼药膏涂抹在眼睛上。
  8. 皮下注射丁丙诺啡(0.05mg / kg),卡洛芬(5mg / kg)和0.5ml氨苄青霉素(100mg / ml)也在手术之前立即进行。
  9. 首先通过观察呼吸频率降低,然后重复不再出现尾部或脚部反射来确保手术深度的麻醉。
  10. 将受体大鼠的下背部剃光在左子宫角上方,并放置在显微镜台上的无菌组织上。
  11. 剃光区域用碘吸印并用70%乙醇浸泡的组织擦拭以制备手术区域并除去由剃刮引起的多余的毛发颗粒(图2)。这是重复两次。


  12. 无菌悬垂布置在动物身上。
  13. 在背部至腹侧方向上长约1厘米的单个切口在皮肤上使用夹钳和大剪刀在最后一个肋的水平(图3)进行。

    图3.初始切口皮肤和皮下脂肪 D:背部; V:腹侧SC:皮下脂肪; BW:体壁。

  14. 使用精细的镊子取出体壁,用左侧卵巢脂肪垫上的弹簧剪刀(避开血管)制成小切口(图4)。

    图4.通过体壁切口。 D:背侧; V:腹侧BW:体壁; OFP:卵巢脂肪垫。

  15. 用精细的镊子抓住卵巢脂肪垫,并将生殖道的远端部分从腹腔轻轻地收回。通过附着的微型夹子将脂肪垫保持在适当的位置,在子宫下方有一个紧密卷绕的组织的无菌片(图5)。

    图5.外部卵巢和脂肪垫。 D:背侧; V:腹侧OFP:卵巢脂肪垫; OB:卵巢囊OV:输卵管U:子宫。

  16. 用精细的镊子手动撕裂卵巢囊的最小血管化区域,提供通向输卵管口的途径(图6)。

    图6.卵巢囊的最小血管化面积的位置。D:背侧; V:腹侧OFP:卵巢脂肪垫; OB:卵巢囊OV:输卵管U:子宫。

  17. 将1-2滴肾上腺素(约20μl)从注射器滴到粘液囊上,直到出血停止。这减少了手术的长度,并加快了口位置。
  18. 转移口吸管的尖端装载有15-20个胚胎用于植入,并且在用细镊子稳定之后插入输卵管的口中。通过移液管通过轻轻的压力将胚胎注入输卵管(图7)

    图7.胚胎进入生殖道的位置。D:背部; V:腹侧OFP:卵巢脂肪垫; OB:卵巢囊OV:输卵管U:子宫O:口。

  19. 腹部仔细更换生殖道,并通过身体壁上的2条缝线闭合腹部(图8),然后用2个伤口夹将皮肤闭合。

    图8.缝线在腹壁中的位置。 D:背部V:腹侧SC:皮下脂肪; BW:体壁。

  20. 手术后,将恢复的大鼠置于置于37°C加温板上的大鼠笼内,直到动物完全清醒和垂直,通常为5至10分钟(见注释)。
  21. 将笼子返回到大鼠室中的笼架,并在24小时后皮下注射0.2ml卡洛芬(5mg / ml)作为手术后镇痛。最多可以将三名女性安置在一起。
  22. 每天监测大鼠,以验证该程序的恢复。女性在手术后20天分开,以分娩出笼。




  1. 普通的黑曲霉菌株包括Sprague Dawley,Wistar,Dahl SS和F344。胚胎没有性别决定是必要的。 Taconic和Charles River是推荐的供应商。
  2. 双细胞胚胎也可以通过这种方法转移
  3. 5至25个胚胎可以转移到每个女性。
  4. 如果切口大于1厘米,可以使用另外的缝线和/或伤口夹
  5. 与老鼠相比,大鼠的脂肪过多。小心处理脂肪垫而不刺穿或撕裂纸巾。
  6. 也可以在有或没有开口的情况下购买隔膜,并切割成所需的尺寸(Surgivet)。
  7. 手术失败的症状包括呼吸频率低下,呼吸困难和/或喘气和嗜睡。


  1. LHRHa解决方案
    将1mg的LHRHa粉末溶解在5ml无菌水中200μg/ ml的储备液 在-20°C储存1 ml等分试样
  2. KSOM(Cold Spring Harbor,2006)
    2.775克NaCl 0.095克KCl
    0.025g KH 2 PO 4
    0.025g MgSO 4·7H 2 O→/ / 0.02 g葡萄糖 0.03克青霉素
    1.05g NaHCO 3
    0.002g EDTA
    0.073g L-谷氨酰胺
    0.125g CaCl 2·2H 2 O
    0.5克牛血清白蛋白 50微升酚红
    将试剂在175 ml胚胎试验水中合并。调节体积至500 ml。使用500 ml过滤系统,将溶液过滤到生物安全柜中。检查pH值并将剩余溶液储存在4°C。孵育后检查pH(pH值应该下降);理想pH值为7.1-7.7


阿拉巴马大学伯明翰大学Transgenic&遗传工程模型设施(RAK)由美国国立卫生研究院支持[授权号P30 CA13148,P30 AR048311,P30 DK074038,P30 DK05336和P60 DK079626]和UAB囊性纤维化研究中心。所有程序均经IACUC和UAB动物资源计划(ARP)批准进行,并符合实验动物和啮齿类动物生存手术的护理和使用指南(Bernal等人, 2009; Albus,2012; Cunliff-Beamer,1993; Waynforth,1992)。我们承认作为适应基准的菲律宾和桑德斯的工作。


  1. Albus,U.(2012)。 护理和使用实验动物(8 th 版)。实验动物 46(3):267-268。
  2. Bernal,J.,Baldwin,M.,Gleason,T.,Kuhlman,S.,Moore,G.and Talcott,M。(2009)。 啮齿动物生存手术指南。投资手册 22(6):445-451。
  3. 冷泉港。 KSOM。(2006)。冷泉Harb Protoc 。
  4. Cunliff-Beamer,TL(1993)。 应用无菌手术的原则到啮齿动物。 AWIC通讯 4(2):3-6。
  5. Filipiak,WE and Saunders,TL(2006)。 Advances转基因大鼠生产中。转基因 Res 15(6):673-686。
  6. Hankenson,FC(2013)。 实验室小鼠和老鼠的重要护理管理 CRC Press 。
  7. Krinke,GJ,Bullock,GR和Bunton,T。(2000)。 实验室大鼠。 Elsevier Science 。
  8. Lambert,LJ,Challa,AK,Niu,A.,Zhou,L.,Tucholski,J.,Johnson,MS,Nagy,TR,Eberhardt,AW,Estep,PN,Kesterson,RA and Grams,JM(2016)。 增加骨小梁骨和改善生物力学在骨钙素无效大鼠由CRISPR / Cas9技术创建的模型。 Dis Model Mech 9(10):1169-1179。
  9. Smith,JC,Corbin,TJ,McCabe,JG和Bolon,B。(2004)。异氟烷与吗啡是转基因大鼠生产中胚胎移植的合适麻醉方案。实验动物 38(1):38-43。 />
  10. Waller,SJ,Ho,MY and Murphy,D。(1995)。 通过在受精的单细胞卵中显微注射克隆的DNA来生产转基因啮齿类动物在:Glover,DM和Hames,BD(Eds)。 DNA克隆卷。牛津大学出版社 pp:184-229。
  11. Waynforth,HB和Flecknell,PA(1992)。 大鼠实验和手术技术。第二版 学术出版社。
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引用:Lambert, L. J., Johnson, L. W., Kennedy, D., Cadillac, J. and Kesterson, R. A. (2017). Improved Oviduct Transfer Surgery for Genetically Modified Rat Production. Bio-protocol 7(16): e2503. DOI: 10.21769/BioProtoc.2503.