A Mouse Model of Postoperative Pain

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The Journal of Neuroscience
Jun 2018



Postoperative pain is highly debilitating and hinders recovery. Opioids are the main pain medication used for acute postoperative pain. Given the devastating opioid addiction and overdose epidemic across the US, non-opioid pain therapeutics are desperately needed. In order to develop novel, non-opioid therapies for the treatment of postoperative pain and identify the mechanisms underlying this pain, rodent models of incisional pain have been established. The protocol herein describes in detail how to create a mouse model of postoperative pain that was adapted from established protocols. This model of postoperative pain is frequently-used, highly reproducible, and results in peripheral and central nervous system alterations.

Keywords: Postoperative pain (术后疼痛), Inflammatory pain (炎症性疼痛), Hypersensitivity (超敏反应), Plantar incision (足底切口), Mouse (小鼠), Skin and muscle incision (皮肤和肌肉切口)


Postoperative pain is a significant, worldwide problem. Approximately 234.2 million people undergo major surgeries each year (Weiser et al., 2008) and about 80% of patients experience acute postoperative pain (Gan, 2017). Of these, between 10% and 50% of patients, develop chronic pain that continues to severely impact their quality of life (Chapman and Vierck, 2017). One of the factors that are associated with the development chronic postoperative pain, but unlikely the cause, is the severity of acute pain experienced during the first postoperative week (Fletcher et al., 2015; Chapman and Vierck, 2017). Opioids are the main pain medication used for acute postoperative pain (Sen and Bathini, 2015; Tan et al., 2018). Given the opioid epidemic, non-opioid pain therapeutics are needed. Therefore, identifying the mechanisms that underlie acute postoperative pain is necessary for the development of optimal therapies for postoperative pain that may ultimately decrease the severity and/or incidence of chronic postoperative pain. Both rat (Brennan et al., 1996) and mouse (Pogatzki and Raja, 2003) models of acute incisional pain have been developed as preclinical models to identify the molecular, cellular and physiological mechanisms that underlie postoperative pain. However, a detailed description of the mouse model of postoperative pain is lacking. Here we describe in detail a mouse model of postoperative pain that requires incision of both the skin and muscle. Incision of both skin and muscle best mimics invasive surgery that causes intense acute pain and leads to chronic pain (Brennan, 2011; Chapman and Vierck, 2017). Furthermore, incision of skin and muscle (~6 days) creates hypersensitivity that lasts substantially longer than the skin-only (~3 days) incision model (Xu and Brennan, 2010). In this protocol, we provide detailed, step-by-step methods adapted from previous protocols (Brennan et al., 1996; Pogatzki and Raja, 2003) for development of a mouse model of postoperative pain.

Materials and Reagents

  1. Stainless steel sterile No. 11 surgical blade (World Precision Instruments, Feather Safety Razor Co. Ltd., catalog number: 504170)
  2. Sterile 5-0 nylon surgical sutures (AD Surgical, Unify, catalog number: S-N518R13)
  3. Surgical tape (3M, Transpore, catalog number: 1527-0)
  4. Cotton swab (VWR, Critical Swab, catalog number: 89031-270)
  5. Glad Press’n Seal (SAI Infusion Technologies, Glad, catalog number: PSS-70)
  6. Sterile nitrile gloves (Kimberly-Clark Professional, Kimtech Pure, catalog number: HC61170)
  7. Petri dishes [VWR, 14.5 and 9 663161, Greiner Bio-One, catalog numbers: 82050-912 (small) and 82050-600 (large)]
  8. Sterile gauze (Allied Medical, Ardes, catalog number: GA441221)
  9. Sharpie extra fine point permanent marker (Staples, Sharpie, catalog number: 37001) 
  10. 8-16 week old C57BL/6J mice (JAX, catalog number: 000664)
  11. Bacitracin zinc ointment (Fougera Pharmaceuticals Inc, catalog number: 0168-0011-04)
  12. Isoflurane (Clipper distributing company LLC., Phoenix, catalog number: 0010250)
  13. 75% ethanol (Fisher Scientific, Decon Laboratories, Inc., catalog number: 22-281-562)
  14. Surgical scrub 7.5% povidone-iodine (Betadine, Veterinary, catalog number: 67618-154-01)
  15. Eye lube (Patterson Veterinary, Optixcare Ophthalmic, catalog number: 07-893-2779)


  1. 1,000 ml beaker (VWR, PYREX, catalog number: 13912-284)
  2. #55 Dumostar Forceps (Fine Science Tools, Dumont, catalog number: 11295-51)
  3. Scalpel handle (Fine Science Tools, catalog number: 10003-12)
  4. Iris Forceps, 10 cm, Curved, Serrated (World Precision Instruments, catalog number: 15915)
  5. Halsted Mosquito Hemostatic Forceps, 12.5 cm, Straight (World Precision Instruments, catalog number: 15920-G)
  6. Vannas Scissors, 8 cm, Curved (World Precision Instruments, catalog number: 14122)
  7. Small animal surgery board (Braintree Scientific, Inc., CD+, catalog number: ACD 014)
  8. Isoflurane dispenser (Highland Medical Equipment, Drager, catalog number: 16-7001)
  9. Sliding top isoflurane induction chamber (Kent Scientific Corporation, catalog number: VetFlo-0530LG)
  10. Heat Therapy Pump with Pad (Adroit Medical Systems, catalog number: HTP-1500)
  11. Isothermal pad (Braintree Scientific, Inc., Deltaphase, catalog number: DPIP)
  12. Digital calipers (VWR, catalog number: 62379-531)
  13. Steri 250 Bead Sterilizer Bead Bath (Lab Unlimited, Simon Keller Ltd., catalog number: 4AJ-6286283)
  14. Microwave (Emerson, 1,000 W, catalog number: B007Q45CIS)
  15. Home cage containing Aspen Sani Chips® (P.J. Murphy Forest and Products, Sani Chips®)


  1. GraphPad Prism 7


  1. Sterilize surgical area with ethanol and sterilize tools in bead bath of at least 325 °C for 30 s. Rest sterile tools, sutures, and blade on sterile gauze.
  2. Warm Heat Therapy Pump pad to 41 °C and warm the Isothermal pad in microwave for 1 min. Place a new, clean home cage on the Heat Therapy Pump pad. Place a laboratory paper towel over the warm Isothermal pad and cover with sterile Press’n Seal and surgery board.
  3. To induce anesthesia, put the mouse in the beaker chamber with paper towel on lid containing 1 ml of isoflurane (Figure 1) and wait for approximately 30 s or until the mouse can no longer right itself. Alternatively, an isoflurane induction chamber that is included with the isoflurane dispenser can be used to induce anesthesia.

    Figure 1. Brief inhalational anesthesia. The mouse was briefly anesthetized by inhalation in chamber with 1 ml isoflurane.

  4. Quickly remove the mouse from the isoflurane chamber and apply eye lube using a cotton swab to each eye. Further anesthetize the mouse with 1.5%-2% inhaled isoflurane by placing the head of the mouse into the nose cone attached to the isoflurane dispenser and cover the mouse (except the hindpaw designated for operation) with sterile Press’n Seal (Figure 2).

    Figure 2. Continuous inhalational anesthesia. The mouse was continuously anesthetized through a nose cone with 1.5%-2% isoflurane.

    For Steps 5-13 also see Video 1.
    Video 1. Plantar incision surgery. This video shows how the model of postoperative pain is made by making an incision through the plantar skin and flexor digitorum brevis muscle (This video was made at the Medical College of Wisconsin and was performed according to guidelines on Animal Care and approved by the Animal Research Ethics Board of the Medical College of Wisconsin under protocol #0383).

  5. Adhere the hindpaw to the surface by taping the toes down with surgical tape (Figure 3).

    Figure 3. Securing the hindpaw to surgical surface. The hindpaw was secured to the surface using surgical tape.

  6. Once the hindpaw is secured, apply a cotton swab of 75% ethanol followed by a new cotton swab of betadine. Repeat for a total of 3 applications each. 
  7. Measure 2 mm from the proximal edge of heel using a digital caliper and place a dot with a permanent marker at this location in the middle of the hindpaw. From the first dot, measure 5 mm towards the toes down the center of the hindpaw and place a second dot (Figure 4).

    Figure 4. Measurement of incision. Two dots were placed in the middle of the hindpaw, one 2 mm from the heel and the second 5 mm from the first dot.

  8. Check that the mouse is fully anesthetized by lightly pinching the most medial toe (most likely this toe could not be secured by the surgical tape) with the forceps. If the mouse flinches, wait until the mouse no longer reacts to the toe pinch before proceeding to Step 9. 
  9. Stabilize the hindpaw by placing the forceps on each side of the heel and make a longitudinal incision through the skin and fascia from the first dot to the second dot (Figure 5).

    Figure 5. Cutaneous incision. A 5 mm longitudinal incision was made with a No. 11 scalpel.

  10. Spread the skin away from the flexor digitorum brevis muscle with the forceps. Elevate the flexor digitorum brevis muscle by inserting one end of the curved forceps underneath the lateral edge of the flexor digitorum brevis muscle and pushing the forceps through to the medial side of the muscle (Figure 6).

    Figure 6. Elevation of flexor digitorum brevis muscle. Curved forceps were inserted under the flexor digitorum brevis muscle to elevate the muscle.

  11. Make a longitudinal incision with the scalpel through the entire belly of the muscle from the origin and insertion taking care not to sever the muscle completely from the origin and insertion, making sure to cut the belly of the muscle into two halves (Figure 7).

    Figure 7. Muscle incision. A longitudinal incision was made through the muscle belly of the elevated flexor digitorum brevis muscle from proximal to distal ends of the cutaneous incision.

  12. To suture the wound, remove the curved forceps from underneath the muscle and elevate the edges of the skin surrounding the wound with forceps. Close the wound by putting two sutures in the skin (but not muscle) approximately 2 mm apart using 5-0 nylon sutures and a hemostat (Figure 8).

    Figure 8. Cutaneous suturing. The skin was closed with two 5-0 nylon sutures.

  13. Apply a generous amount of bacitracin ointment to the wound using a cotton swab and place the mouse in the new cage located on the Heat Therapy Pump pad from Step 2.

Data analysis

Statistical significance was determined with GraphPad Prism 7 Software and graphs are shown as mean ± SEM. A two-way ANOVA with a Sidak post-hoc was used to determine statistical significance for mechanical and heat thresholds. A complete description of statistics used for analyzing mechanical and heat threshold behavioral data is provided in Cowie et al. (2018).


  1. It is important to perform the operation as quickly as possible to reduce the need for repeated anesthesia and any side effects from anesthesia. In our laboratory, it takes a trained surgeon approximately 5 min to perform this procedure.
  2. For a control, a sham surgery is performed by anesthetizing the mouse, sterilizing the hindpaw as in Step 5, and applying bacitracin ointment to the plantar hindpaw.
  3. If bleeding occurs during the procedure, apply pressure to the incision site with a cotton swab until the bleeding stops.
  4. Mice are housed together.
  5. No care is required after surgery except for monitoring of sutures. 
  6. If using mice past postoperative day 3, remove sutures on postoperative day 4. Mice that pulled out sutures before postoperative day 2 must be removed from the study due to poor wound closure.
  7. For consistent behavioral results, apply mechanical and heat stimuli to the medial-posterior aspect of the plantar hindpaw (Figure 9). This area is the least sensitive at baseline because the heel is weight bearing whereas other areas near the pads are more sensitive and variable in withdrawal threshold. Therefore, using the heel area that provides a consistently high baseline withdrawal threshold allows for the best detection of change due to incision (Brennan et al., 1996). Mice were acclimated for 1 h in Plexiglass chambers placed on either a mesh platform (mechanical threshold) or glass platform (heat threshold). Calibrated von Frey monofilaments (0.09 to 19.6 mN) were applied to the plantar hindpaw and the withdrawal threshold for each animal was calculated using the up-down method (Dixon, 1980; Chaplan et al., 1994). The Hargreaves assay was used to measure heat sensitivity as previously established (Hargreaves et al., 1988; Jackson et al., 1995; Barabas and Stucky, 2013; Cowie et al., 2018). Withdrawal latencies to a focused radiant heat source (IITC, Life Sciences Instruments) underneath the glass platform were measured 3 times and averaged for each mouse. A cutoff of 20 s was used to avoid injury. Examples of mechanical and heat hypersensitivity following incision are shown in Figure 10.

    Figure 9. Application of von Frey monofilament. An orange von Frey monofilament was applied to most sensitive location following incision.

    Figure 10. Mechanical and heat thresholds following incision. A. von Frey monofilaments was applied to most sensitive location following incision and the Dixon up-down method (Dixon, 1980) was used to determine mechanical threshold. B. The Hargreaves assay (Hargreaves et al., 1988) was used to measure the withdrawal threshold in response to a radiant heat source that was applied to the most sensitive location following incision. These data were modified from Cowie et al. (2018). Data shown as mean ± SEM, repeated-measures two-way ANOVA and Sidak post-hoc analysis, **P < 0.01 and ****P < 0.0001 sham versus incision. For (A) and (B), n = 8 male mice per group.


This protocol was adapted from established published procedures (Brennan et al., 1996; Pogatzki and Raja, 2003). This work was supported by the National Institute of Neurological Disorders and Stroke grants NS040538 and NS070711 to C.L.S and F31GM123778 to A.M.C. The Research and Education Component of the Advancing a Healthier Wisconsin Endowment at the Medical College of Wisconsin provided partial support. The authors thank Timothy J Brennan, Ph.D., MD for his review of the manuscript.

Competing interests

The authors declare no competing financial or non-financial interests.


All animal procedures were carried out in accordance with the National Institute of Health guidelines and approved by the Institutional Animal Care and Use Committee of the Medical College of Wisconsin (AUA #0383).


  1. Barabas, M. E. and Stucky, C. L. (2013). TRPV1, but not TRPA1, in primary sensory neurons contributes to cutaneous incision-mediated hypersensitivity. Mol Pain 9: 9.
  2. Brennan, T. J. (2011). Pathophysiology of postoperative pain. Pain 152(3 Suppl): S33-40.
  3. Brennan, T. J., Vandermeulen, E. P. and Gebhart, G. F. (1996). Characterization of a rat model of incisional pain. Pain 64(3): 493-501.
  4. Chaplan, S. R., Bach, F. W., Pogrel, J. W., Chung, J. M. and Yaksh, T. L. (1994). Quantitative assessment of tactile allodynia in the rat paw. J Neurosci Methods 53(1): 55-63.
  5. Chapman, C. R. and Vierck, C. J. (2017). The transition of acute postoperative pain to chronic pain: An integrative overview of research on mechanisms. J Pain 18(4): 359 e1-359 e38.
  6. Cowie, A. M., Moehring, F., O'Hara, C. and Stucky, C. L. (2018). Optogenetic inhibition of CGRPα sensory neurons reveals their distinct roles in neuropathic and incisional pain. J Neurosci 38(25): 5807-5825.
  7. Dixon, W. J. (1980). Efficient analysis of experimental observations. Annu Rev Pharmacol Toxicol 20: 441-462.
  8. Fletcher, D., Stamer, U. M., Pogatzki-Zahn, E., Zaslansky, R., Tanase, N. V., Perruchoud, C., Kranke, P., Komann, M., Lehman, T. and Meissner, W. (2015). Chronic postsurgical pain in Europe: An observational study. Eur J Anaesthesiol 32(10): 725-734.
  9. Gan, T. J. (2017). Poorly controlled postoperative pain: prevalence, consequences, and prevention. J Pain Res 10: 2287-2298.
  10. Hargreaves, K., Dubner, R., Brown, F., Flores, C. and Joris, J. (1988). A new and sensitive method for measuring thermal nociception in cutaneous hyperalgesia. Pain 32(1): 77-88.
  11. Jackson, D. L., Graff, C. B., Richardson, J. D. and Hargreaves, K. M. (1995). Glutamate participates in the peripheral modulation of thermal hyperalgesia in rats. Eur J Pharmacol 284(3): 321-325.
  12. Pogatzki, E. M. and Raja, S. N. (2003). A mouse model of incisional pain. Anesthesiology 99(4): 1023-1027.
  13. Sen, S. and Bathini, P. (2015). Auditing analgesic use in post-operative setting in a teaching hospital. J Clin Diagn Res 9(4): FC01-04.
  14. Tan, W. H., Yu, J., Feaman, S., McAllister, J. M., Kahan, L. G., Quasebarth, M. A., Blatnik, J. A., Eagon, J. C., Awad, M. M. and Brunt, L. M. (2018). Opioid medication use in the surgical patient: An assessment of prescribing patterns and use. J Am Coll Surg 227(2): 203-211.
  15. Weiser, T. G., Regenbogen, S. E., Thompson, K. D., Haynes, A. B., Lipsitz, S. R., Berry, W. R. and Gawande, A. A. (2008). An estimation of the global volume of surgery: a modelling strategy based on available data. Lancet 372(9633): 139-144.
  16. Xu, J. and Brennan, T. J. (2010). Guarding pain and spontaneous activity of nociceptors after skin versus skin plus deep tissue incision. Anesthesiology 112(1): 153-164.


术后疼痛非常虚弱,阻碍了恢复。 阿片类药物是用于急性术后疼痛的主要止痛药。 鉴于美国各地的阿片类药物成瘾和过量流行,非急需非阿片类药物疼痛治疗。 为了开发用于治疗术后疼痛的新型非阿片类药物疗法并确定这种疼痛的机制,已经建立了切口疼痛的啮齿动物模型。 本文的方案详细描述了如何创建根据已建立的方案改编的术后疼痛的小鼠模型。 这种术后疼痛模型经常使用,高度可重复,并导致外周和中枢神经系统改变。

【背景】术后疼痛是一个重要的全球性问题。每年大约有2.342亿人接受大手术(Weiser et al。,2008),大约80%的患者出现急性术后疼痛(Gan,2017)。其中,10%至50%的患者出现慢性疼痛,继续严重影响其生活质量(Chapman和Vierck,2017)。与发生慢性术后疼痛相关的因素之一,但不太可能的原因,是术后第一周出现急性疼痛的严重程度(Fletcher et al。,2015; Chapman和Vierck, 2017年)。阿片类药物是用于急性术后疼痛的主要止痛药(Sen和Bathini,2015; Tan et al。,2018)。鉴于阿片类药物流行,需要非阿片类药物疼痛治疗。因此,确定导致急性术后疼痛的机制对于开发术后疼痛的最佳疗法是必要的,这可能最终降低慢性术后疼痛的严重性和/或发生率。大鼠(Brennan et al。,1996)和小鼠(Pogatzki和Raja,2003)急性切口疼痛模型已被开发为临床前模型,以确定术后疼痛的分子,细胞和生理机制。 。然而,缺乏对术后疼痛的小鼠模型的详细描述。在这里,我们详细描述了需要切开皮肤和肌肉的术后疼痛的小鼠模型。皮肤和肌肉的切口最能模仿侵入性手术,导致剧烈的急性疼痛并导致慢性疼痛(Brennan,2011; Chapman和Vierck,2017)。此外,皮肤和肌肉的切口(约6天)会产生超敏反应,其持续时间比仅皮肤(~3天)切口模型长得多(Xu和Brennan,2010)。在该协议中,我们提供了从先前的方案(Brennan 等人,<1996; Pogatzki和Raja,2003)改编的详细的逐步方法,用于开发术后疼痛的小鼠模型。

关键字:术后疼痛, 炎症性疼痛, 超敏反应, 足底切口, 小鼠, 皮肤和肌肉切口


  1. 不锈钢无菌11号手术刀片(World Precision Instruments,Feather Safety Razor Co. Ltd.,目录号:504170)
  2. 无菌5-0尼龙手术缝合线(AD Surgical,Unify,目录号:S-N518R13)
  3. 手术胶带(3M,Transpore,目录号:1527-0)
  4. 棉签(VWR,Critical Swab,目录号:89031-270)
  5. Glad Press'n Seal(SAI Infusion Technologies,Glad,产品目录号:PSS-70)
  6. 无菌丁腈手套(Kimberly-Clark Professional,Kimtech Pure,目录号:HC61170)
  7. 培养皿[VWR,14.5和9 663161,Greiner Bio-One,目录号:82050-912(小)和82050-600(大)]
  8. 无菌纱布(Allied Medical,Ardes,目录号:GA441221)
  9. Sharpie超细点永久性标记(Staples,Sharpie,目录号:37001)&nbsp;
  10. 8-16周龄C57BL / 6J小鼠(JAX,目录号:000664)
  11. 杆菌肽锌软膏(Fougera Pharmaceuticals Inc,目录号:0168-0011-04)
  12. Isoflurane(Clipper distribution company LLC。,Phoenix,目录号:0010250)
  13. 75%乙醇(Fisher Scientific,Decon Laboratories,Inc。,目录号:22-281-562)
  14. 手术磨砂7.5%聚维酮碘(Betadine,兽医,目录号:67618-154-01)
  15. 眼润滑剂(Patterson Veterinary,Optixcare Ophthalmic,目录号:07-893-2779)


  1. 1,000毫升烧杯(VWR,PYREX,目录号:13912-284)
  2. #55 Dumostar Forceps(精细科学工具,Dumont,目录号:11295-51)
  3. 手术刀手柄(精细科学工具,目录号:10003-12)
  4. 虹膜镊子,10厘米,弯曲,锯齿(世界精密仪器,目录号:15915)
  5. Halsted Mosquito Hemostatic Forceps,12.5 cm,Straight(World Precision Instruments,目录号:15920-G)
  6. Vannas剪刀,8厘米,弯曲(World Precision Instruments,目录号:14122)
  7. 小动物手术板(Braintree Scientific,Inc.,CD +,目录号:ACD 014)
  8. 异氟醚分配器(Highland Medical Equipment,Drager,目录号:16-7001)
  9. 滑动顶部异氟烷诱导室(Kent Scientific Corporation,目录号:VetFlo-0530LG)
  10. 带垫的热疗泵(Adroit Medical Systems,目录号:HTP-1500)
  11. 等温垫(Braintree Scientific,Inc.,Deltaphase,目录号:DPIP)
  12. 数显卡尺(VWR,目录号:62379-531)
  13. Steri 250 Bead Sterilizer Bead Bath(Lab Unlimited,Simon Keller Ltd.,目录号:4AJ-6286283)
  14. 微波炉(艾默生,1,000 W,目录号:B007Q45CIS)
  15. 包含Aspen Sani芯片®的主笼(P.J.Murphy Forest and Products,Sani Chips ®)


  1. GraphPad Prism 7


  1. 用乙醇对手术区域进行消毒,并在至少325°C的珠浴中对工具进行消毒30秒。将无菌工具,缝合线和刀片放在无菌纱布上。
  2. 温热治疗泵垫至41°C并在微波炉中加热等温垫1分钟。在热疗泵垫上放置一个干净的新笼子。将实验室纸巾放在温暖的Isothermal垫上,并用无菌Press'n Seal和手术板覆盖。
  3. 为了诱导麻醉,将小鼠放入烧杯室,用纸巾盖上含有1毫升异氟醚的盖子(图1),然后等待大约30秒或直到老鼠不能再对自己。或者,异氟醚分配器中包含的异氟醚诱导室可用于诱导麻醉。

    图1.简单的吸入麻醉。在室内吸入1 ml异氟醚,将小鼠短暂麻醉。

  4. 从异氟醚腔室中快速取出鼠标,并使用棉签将眼润肤液涂抹在每只眼睛上。用1.5%-2%吸入异氟醚进一步麻醉小鼠,将小鼠头部放入与异氟醚分配器相连的鼻锥中,用无菌Press'n Seal覆盖小鼠(指定操作的后爪除外)(图2) 。


    对于步骤5-13 ,请参阅视频1.

  5. 用手术胶带将脚趾贴在脚趾上,将后爪粘在表面上(图3)。

    图3.将后爪固定到手术表面。 使用手术胶带将后爪固定在表面上。

  6. 一旦后爪被固定,使用75%乙醇的棉签,然后使用新的棉签棉签。重复每次共3次申请。&nbsp;
  7. 使用数字卡尺从脚跟的近端边缘测量2mm,并在后爪中间的该位置处放置带有永久性标记的点。从第一个点开始,沿着后爪中心向下测量5毫米,并放置第二个点(图4)。

    图4.切口的测量。 在后爪中间放置两个点,一个距离脚跟2毫米,第二个点距离第一个点5毫米。

  8. 用镊子轻轻夹住最内侧的脚趾(很可能这个脚趾不能用手术带固定),检查鼠标是否完全麻醉。如果鼠标退缩,请等到鼠标不再对脚趾捏反应,然后再进行步骤9.&nbsp;
  9. 通过将镊子放在脚后跟的两侧来稳定后爪,并通过皮肤和从第一个点到第二个点的筋膜进行纵向切口(图5)。

    图5.皮肤切口。 用11号手术刀做5毫米纵切口。

  10. 用镊子将皮肤远离屈肌腱屈肌。通过插入趾短屈肌肌肉的侧向边缘下面的弯钳的一端,并通过与肌肉的内侧推动钳子(图6)提升所述趾短屈肌肌肉。


  11. 用手术刀从原点开始纵向切开肌肉的整个腹部并插入,注意不要将肌肉完全切断原点和插入,确保将肌肉的腹部切成两半(图7)。

    图7.肌肉切口。 从皮肤切口的近端到远端通过屈肌腱短肌的肌腹进行纵向切口。

  12. 要缝合伤口,从肌肉下方取下弯曲的镊子,用镊子抬高伤口周围的皮肤边缘。使用5-0尼龙缝线和止血钳将两条缝合线分开约2毫米(图8),将两条缝合线缝合在皮肤上(但不是肌肉)。

    图8.皮肤缝合。 用两根5-0尼龙缝线封闭皮肤。

  13. 使用棉签将大量的杆菌肽软膏涂抹在伤口上,然后将鼠标放在步骤2的热疗泵垫上的新笼子中。


用GraphPad Prism 7软件测定统计学显着性,图表显示为平均值±SEM。使用具有Sidak事后的双向ANOVA来确定机械和热阈值的统计显着性。 Cowie 等人(2018)提供了用于分析机械和热阈值行为数据的统计数据的完整描述。


  1. 重要的是尽可能快地进行手术,以减少麻醉重复麻醉和任何副作用的需要。在我们的实验室中,需要训练有素的外科医生大约5分钟才能执行此程序。
  2. 对于对照,假手术通过麻醉小鼠,如步骤5中对后爪消毒,并将杆菌肽软膏施用于足底后爪来进行。
  3. 如果在手术过程中出现出血,请用棉签对切口部位施加压力,直至出血停止。
  4. 将小鼠放在一起。
  5. 除了监测缝线外,手术后无需护理。&nbsp;
  6. 如果在术后第3天使用小鼠,则在术后第4天取出缝线。由于伤口闭合不良,必须在术后第2天拔除缝合线的小鼠。
  7. 为了获得一致的行为结果,将机械和热刺激应用于足底后爪的内侧 - 后侧(图9)。该区域在基线处是最不敏感的,因为脚跟是承重的,而垫附近的其他区域在退出阈值中更敏感和可变。因此,使用提供始终如一的高基线退缩阈值的足跟区域可以最好地检测切口引起的变化(Brennan et al。,1996)。将小鼠在放置在网状平台(机械阈值)或玻璃平台(热阈值)上的有机玻璃室中适应1小时。将校准的von Frey单丝(0.09至19.6mN)施用于足底后爪,并使用上下方法计算每只动物的退缩阈值(Dixon,1980; Chaplan 等人,,1994)。 。 Hargreaves测定用于测量热敏感度,如先前所建立的(Hargreaves et al。,1988; Jackson et al。,1995; Barabas and Stucky,2013; Cowie 等人,2018)。对玻璃平台下方的聚焦辐射热源(IITC,Life Sciences Instruments)的退出延迟进行3次测量并对每只小鼠取平均值。截止20秒用于避免伤害。切口后的机械和热超敏反应的例子如图10所示。

    图9. von Frey单丝的应用。 在切口后最敏感的位置应用橙色von Frey单丝。

    图10.切口后的机械和热阈值。 :一种。将von Frey单丝应用于切口后最敏感的位置,并使用Dixon上下方法(Dixon,1980)确定机械阈值。 B. Hargreaves测定法(Hargreaves et al。,1988)用于测量响应于切口后最敏感位置的辐射热源的退缩阈值。这些数据来自Cowie 等人(2018)。数据显示为平均值±SEM,重复测量双向ANOVA和Sidak事后分析,** P &lt; 0.01和**** P &lt; 0.0001假与切口相比。对于(A)和(B),每组n = 8只雄性小鼠。


该方案改编自已建立的公布程序(Brennan et al。,1996; Pogatzki和Raja,2003)。这项工作得到国家神经疾病和中风研究所的支持NS040538和NS070711对C.L.S和F31GM123778的支持给A.M.C.在威斯康星医学院推进更健康的威斯康星州捐赠的研究和教育部分提供了部分支持。作者感谢Timothy J Brennan,博士,医学博士对手稿的评论。






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Copyright: © 2019 The Authors; exclusive licensee Bio-protocol LLC.
引用: Readers should cite both the Bio-protocol article and the original research article where this protocol was used:
  1. Cowie, A. M. and Stucky, C. L. (2019). A Mouse Model of Postoperative Pain. Bio-protocol 9(2): e3140. DOI: 10.21769/BioProtoc.3140.
  2. Cowie, A. M., Moehring, F., O'Hara, C. and Stucky, C. L. (2018). Optogenetic inhibition of CGRPα sensory neurons reveals their distinct roles in neuropathic and incisional pain. J Neurosci 38(25): 5807-5825.