Sacral Spinal Cord Transection and Isolated Sacral Cord Preparation to Study Chronic Spinal Cord Injury in Adult Mice

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Feb 2017


Spinal cord injury (SCI) is characterized by multiple sensory/motor impairments that arise from different underlying neural mechanisms. Linking specific sensory/motor impairments to neural mechanism is limited by a lack of direct experimental access to these neural circuits. Here, we describe an experimental model which addresses this shortcoming. We generated a mouse model of chronic spinal cord injury that reliably reproduces spasticity observed after SCI, while at the same time allows study of motor impairments in vivo and in an in vitro preparation of the spinal cord. The model allows for the combination of mouse genetics in in vitro and in vivo conditions with advanced imaging, behavioral analysis, and detailed electrophysiology, techniques which are not easily applied in conventional SCI models.

Keywords: Spinal cord injury (脊髓损伤), Complete transection (完全横断), in vitro preparation (体外制备), Sacral spinal cord (骶髓), Spasticity (痉挛状态)


Spinal cord injury results in devastating sensory-motor disabilities. Extensive work in animal models has investigated the pathophysiological state of spinal circuits after SCI. Most studies in spinal cord injury are carried out in animal models relevant for clinical evaluation of motor impairment and recovery after SCI (Sharif-Alhoseini et al., 2017). One of the main limitations of these models is the difficulty to relate clinical features of sensory-motor dysfunction to specific cellular mechanism(s). In the last decade, new insights into possible cellular mechanisms underlying motor impairments after SCI have come from transection models of SCI. Here, the sacral spinal cord is surgically transected resulting in paralysis only of the tail muscles. This model was first introduced in cats (Ritz et al., 1992), and later in rats (Bennett et al., 1999), and has provided insights into cellular mechanisms underlying sensory-motor dysfunction after injury (Bennett et al., 2001). Nevertheless, the genetic tools for manipulating neuronal activity in cats and rats are very limited. In contrast, mice allow electrophysiology to be combined with genetics for identification and manipulation of the activity of specific neurons in the spinal cord (Jiang and Heckman, 2006; Kiehn, 2016). In recent efforts, these advantages have allowed optical approaches (e.g., optogenetics, calcium imaging of defined neuronal subtypes) to be used for dissection of the neural mechanisms underlying muscle spasms after SCI (Bellardita et al., 2017). In this protocol, we describe the technical aspects of sacral spinal cord transection in adult mice, and the subsequent use of in vitro sacral spinal cord preparations for direct examination of the neural mechanisms which cause spasm in chronic spinal cord injury.

Materials and Reagents

  1. Glass micropipettes (Harvard Apparatus, catalog number: GC150F-10 )
  2. Wood stick cotton tip swabs (Medline Scientific, catalog number: 300230S )
  3. Non-sterile gauze swabs (Kruuse, catalog number: 160120 )
  4. Surgery cover 60 x 90 cm (Kruuse, catalog number: 141770 )
  5. Absorbent swabs (Kettenbach, catalog number: 001911 )
  6. Suture, straight cutting needles, non-absorbable (eSutures, Ethicon, catalog number: K889H )
  7. Hypodermic needles 26 G brown 16 mm (BD, Microlance, catalog number: 304300 )
  8. 23-Gauge needle (PrecisionGlide IM; BD, catalog number: 305145 )
  9. Vetbond tissue adhesive (3M, catalog number: 1469C )
  10. Surgical glue (3M, Vetbond, catalog number: 372146 )
  11. Facial Tissue (VWR, catalog number: 115-0600 )
  12. Bench liner paper (ScienceWare, VWR, catalog number: 470145-292 )
  13. 8 weeks old mice (JAX Mice Strain; THE JACKSON LABORATORY, catalog number: 000664 )
    Note: All experiments should be performed in accordance with relevant guidelines and regulations. The local Swedish and Danish ethical committees approved all procedures described here.
  14. 100% oxygen (O2)
  15. Isoflurane (Baxter, catalog number: 1001936060 )
  16. Lidocaine Hydrochloride (Sigma-Aldrich, catalog number: BP214 )
  17. Betadine® Surgical Scrub (povidone-iodine, 7.5%)
  18. Xilocaine (1%)
  19. Buprenorphine hydrochloride (Reckitt Benckiser Healthcare, 0.3 mg/ml)
  20. Carprofen (Canidryl, catalog number: 122964 )
  21. Physiologic solution (0.9% sodium chloride) (Grovet, Braun Ecotainer, catalog number: 99512 )
  22. Viscotears liquid gel for dry eyes (Novartis, catalog number: 2082642 )
  23. 70% ethanol
  24. Sylgard (Sigma-Aldrich, catalog number: 761028-5EA )
  25. Sodium chloride (NaCl, Sigma-Aldrich, catalog number: 433209 )
    Note: This product has been discontinued.
  26. Potassium chloride (KCl, Sigma-Aldrich, catalog number: P9541 )
  27. Glucose (Sigma-Aldrich, catalog number: G8270 )
  28. Sodium bicarbonate (NaHCO3, Sigma-Aldrich, catalog number: S5761 )
  29. Magnesium sulfate heptahydrate (MgSO4·7H2O, Sigma-Aldrich, catalog number: 63138 )
  30. Potassium phosphate monobasic (KH2PO4, Sigma-Aldrich, catalog number: 92214 )
  31. Calcium chloride (CaCl2, Sigma-Aldrich, catalog number: 449709 )
  32. Magnesium chloride (MgCl2, Sigma-Aldrich, catalog number: V000149 )
  33. HEPES (Sigma-Aldrich, catalog number: H3375 )
  34. Ringer’s solution (see Recipes)
  35. Oxygenated modified artificial cerebrospinal fluid (mACSF, see Recipes)


  1. Forceps (Fine Science Tools, catalog number: 11251-10 )
  2. Toothed forceps (Fine Science Tools, catalog number: 11154-10 )
  3. Vannas Spring scissors–Micro-serrated (Fine Science Tools, catalog number: 15007-08 )
  4. Dumont No. 2 laminectomy forceps (Fine Science Tools, catalog number: 11223-20 )
  5. Fine Scissors–Tungsten Carbide (Fine Science Tools, catalog number: 14568-09 )
  6. Fine Scissors–Tungsten Carbide & ToughCut (Fine Science Tool, catalog number: 14558-11 )
  7. Rechargeable animal clipper (Wahl Arco)
  8. Scalpel (Fine Science Tools, catalog number: 10020-00 )
  9. Temperature-controlled variable heat pad (K&H Manufacturing, model: 1009 )
  10. Diaphragm vacuum pump-lubricated-single-stage (Environmental Express, catalog number: EE0753280 )
  11. Isofluorane vaporizer (Soarmed, model: MSS-3 )


  1. Complete lesion of the sacral spinal cord
    1. Prepare a clean and disinfected area dedicated to rodent surgery with only the equipment related to surgery (Figure 1A). Place the surgical instruments in an area which can be easily accessed during the procedure (Figure 1B).
    2. Prepare a glass pipette with a diameter of 50-100 μm from a glass capillary (Figure 1C). The glass capillary is pulled, and the tip is then manually adapted to the spinal cord. Connect the glass pipette to a vacuum pump through a series of silicon tubes of increasing diameter. The pulled glass pipette will be used in the surgery for suction-transection of the spinal cord.
    3. Weigh the mouse before surgery. Monitor the weight of the animal for the next 15 days to evaluate for the potential loss of weight after surgery.
    4. Place the mouse in a sealed induction chamber with 5% isoflurane/95% oxygen until it is deeply anesthetized (Figure 1A.2).
    5. Move the mouse from the induction chamber to the area dedicated to the surgery and prepare it for the surgery (Figure 1D):
      1. Position the mouse ventral side down on a heating pad to maintain body temperature (37 °C) constant during the entire procedure (Figure 1D.1).
      2. Use 2% isofluorane for the entire period of the surgery. Deliver isofluorane to the mouse through a facial mask (Figure 1D.2).
      3. Check reflexes of the animal to verify an appropriate state of anesthesia. There should be no pinch-evoked reflexes. We assessed the pedal withdrawal reflex by pinching the tail and the metacarpal region of the hind foot.
      4. Secure the animal to make sure it will not move during the surgery (as might be caused by touching the dorsal roots). Secure the animal with strips of tape attached to the limbs. Avoid excessive stretching of the limbs, which may damage joints as well as impair the animal’s breathing (Figure 1D.3).
      5. Shave the back of the mouse along the rostrocaudal axis of the spinal column. Apply sodium iodine to the shaved area and leave it for 5 min (Figure 1D.4). Apply eye ointment to protect the eyes during surgery.
      6. Remember to avoid resting your hands or instruments on the mouse thorax. The external pressure may interfere with respiration and/or blood circulation.
      7. Apply a surgical cover to the body of the mouse, leaving a window at the point of incision (Figure 1D.5). 
    6. Localization of the second sacral segment and transection:
      1. Use two fingers to localize the T12 vertebral body. The T12 vertebra has the longest spinous process of all vertebrae, and if the spine of the mouse is put into flexion, the T12 vertebra protrudes outward in the spinal column. With a scalpel, make a longitudinal incision of the skin from approximately the T12 to the L4 vertebral bodies (Figure 1E).
      2. The second sacral segment (S2) of the spinal cord lies beneath the rostral part of L2 vertebral body, on the boundaries between the L1 and L2 vertebral body. The spinous process of L2 points rostrally, and should be used as a landmark for making a deep vertical incision with a small eye scissor. If the cut is performed vertically, it will reveal the ligamentum flavum between the L1 and L2 vertebral bodies (Figure 1F). For a better understanding of the anatomical landmarks, and especially the relationship between lumbar vertebral bodies and the spinal cord in adult mice, refer to Harrison et al., 2013.
      3. Cut the ligamentum flavum with the eye scissor. The spinal cord will appear with a dorsal artery lying in the midline (Figure 1G).
      4. Apply Xilocaine (1%) on the top of the cord to prevent movements elicited by touching the spinal cord or the dorsal roots. Wait for the drug to take effect (about half a minute) and then use fine forceps to position the dorsal roots as lateral as possible. In the case of other structural damage (e.g., bones, arteries, tendons), the surgeon should consider discluding the animal from subsequent analysis.
      5. If the cut to the ligamentum flavum is performed correctly, no other damage will be caused to the surrounding tissue (muscles, ligament or skin), and no blood should be visible.
      6. Starting on one side of the cord, use the glass pipette attached to vacuum suction to remove the S2 spinal tissue. Keep aspirating tissue until a complete discontinuity is observed between the rostral and the caudal ends of the cord, in total corresponding to one segment.
    7. Suture the surgical wound and let the animal recover:
      1. Suture the muscle around the spinal column at the injury site to protect the cord, and use veterinary glue close the skin.
      2. Give post-surgery treatment of Buprenorphine (0.1 mg/kg), Carprofen (5 mg/kg) and, if necessary, 0.3 ml of sterile physiologic solution subcutaneously for 2 to 5 days post-surgery.
      3. Turn the anesthesia off and place the mouse back in the cage with a heating pad to keep the animal warm for a period of 1-2 h. The animal may be housed alone for the first week, and thereafter, if the recovery is complete, it may be housed with another animal. Special cage bedding is not necessary.
      4. Monitor the animal daily for signs of distress, including weight loss (a > 10% drop in body weight should be avoided), dehydration, or infection. In any of these cases, the surgeon should consult with a veterinarian for suggestions and solutions.
      5. The injury should only cause paralysis of the tail muscles, and should not affect the bladder or hindlimbs. However, during daily postoperative care it is important to monitor for bladder dysfunction, which can sometimes occur if the injury site is too rostral. 
    Note: The limited visibility caused by a small working area during the surgery makes it difficult to reliably evaluate the completeness of the lesion. Therefore, all lesions should be evaluated visually after the end the experiment after dissection of the cord (Figures 1I-1L). 

    Figure 1. Lesion of the sacral spinal cord in adult mice. A. Area prepared for the surgery with easy access to the necessary equipment: 1) Dissection microscope; 2) Induction chamber for anesthesia; 3) Facial mask for anesthesia; 4) Isofluorane vaporizer; 5) Heating pad; 6) Glass pipette for aspirating the spinal cord connected to the vacuum pump; 7) Vacuum cleaner; 8) Device for surgical instrument sterilization. B. Surgical instruments for lesioning the sacral spinal cord with (from left to right): scissors, forceps, eye ointment, veterinary glue, and suture. C. Glass pipette and vacuum pump for aspirating the spinal cord. D. A mouse prepped for the lesioning procedure. 1) Heating pad; 2) Facial mask to deliver anesthesia; 3) Strips of tape for preventing sudden movements; 4) Area of interest shaved and prepped with sodium iodine; 5) Green cover for the mouse body with a work window in the area of interest. E. Surgical incision of the skin in the area of interest. F. Schematic of the surgery area with the vertebral body L1 and L2 with the second sacral segment of the spinal cord. G. Incision at the level of the L2 vertebral body after cutting the ligamentum flavum. H. Aspiration of the spinal cord using the glass pipette. I-L. Examples of dissected, lesioned sacral cords two months after SCI with either an incomplete (I) or complete (L) lesion.

  2. Dissection of the sacral spinal cord of adult chronic spinalized mice
    1. Prepare a clean disinfected area dedicated to rodent surgery with easy access to the equipment necessary for isolation of the spinal cord (similar to that of Figure 1A).
    2. Place the animal in a chamber for induction of anesthesia (5% isofluorane/95% oxygen), and move the animal from the induction chamber to the dissection table when deeply anesthetized. Check reflexes as in Step A4c.
    3. Place the mouse on a bench liner paper (Scienceware) and apply isofluorane (2%) through the facial mask. Check the reflexes of an appropriate state of anesthesia as in Step A5c. Apply strips of tape to secure the limbs, shave the back, and clean the area with alcohol (Figure 2A).
      Note: In this step, the animal does not have to be positioned on a heating pad as a lower temperature will decrease the metabolism, improving the dissection of the cord; eye ointment is not necessary since the procedure will last few minutes.
    4. Laminectomy along the site of interest:
      1. Identify T12 as described above.
      2. Cut the skin from about the T12 vertebral body to L5 vertebral body. Keep in mind that the second sacral segment of the spinal cord is beneath the second vertebral body of the lumbar spinal cord (L2).
      3. Expose the spinal column by cutting the muscles and tendons around it (Figure 2B).
      4. Localize the spinous process of the T13 vertebral body (the T13 vertebra has the last pair of ribs attached), and start a dorsal laminectomy (a surgical procedure removing the dorsal portion of the vertebrae) in the rostro-caudal direction.
      5. Begin to perfuse the spinal column with cold mACSF (20 ml/min) to slow down metabolism and reduce blood flow in the site of interest.
      6. Proceed with the laminectomy by cutting the left and right sides of the vertebral body with the scissor.
      7. Avoid damaging the spinal cord with the scissor, which can sometimes occur when moving the tip of the scissor from the left to right sides of the vertebrae (or vice versa). Damage may result in contusion or bruises of the spinal cord.
      8. Use a continuous flow (20 ml/min) of cold (~4 °C), oxygenated mACSF on the spinal cord.
    5. Isolation of the sacral spinal cord:
      1. When the spinal column is exposed form the caudal lumbar segments to the cauda equina, the laminectomy is complete (Figure 2C).
      2. Give pure oxygen to the mouse through the facial mask for about 5 min to increase blood oxygenation levels.
      3. Cut the skin at the level of the abdominal muscles and save the abdominal artery. This cut will cause a decrease in blood pressure, preventing an overflow of blood at the level of the cord during isolation.
      4. Cut the cord at the level of the caudal lumbar segments and proceed cutting the ventral roots on the right and left sides of the cord to isolate it from the spinal column.
      5. Pay special attention when you reach the site of the lesion. At the level of the injury, the dura mater is often attached to the vertebral body, requiring careful detachment of the spinal cord and dura from the rest of the spinal column.
      6. Once the cord is completely detached, move it in a dissection chamber with a continuous flow (20 ml/min) of cold oxygenated-mACSF.
      7. Carefully and completely remove the dura mater from the spinal cord to allow greater diffusion of oxygenation into the tissue. Cut the spinal cord, the dorsal root and the ventral roots to decrease the length and allow an easier recognition of the different segments and roots during the experiment (Figure 2C).
      8. Once all roots of the cord are cut and the isolation of the cord is complete, the spinal cord can be moved to a perfusion chamber covered with Sylgard (Figure 2D). Provide a continuous flow of mACSF at room temperature (3-7 ml/min). 
    6. In vitro preparation of the sacral spinal cord for simultaneous calcium imaging of spinal interneurons and ventral root recording:
      1. Move the spinal cord to the recording chamber. The recording chamber has a Sylgard bottom and a Sylgard ‘bridge’ attached which allows the cord to be placed in an L-shaped position (Figure 2E), such that the coronal plane of the spinal cord can be imaged with an objective lens.
      2. Keep a continuous flow of oxygenated Ringer solution for the duration of the experiment.
      3. The cord is pinned down ventral side up from the most caudal end to ensure mechanical stability. The rostral end of the cord leans onto the bridge and a bended minute pin is used as a hook to keep the cord in place (Figure 2E).
      4. Suction electrodes are used for recording motor activity (S4-Co1) and stimulating dorsal roots.
      5. Calcium imaging is performed by lowering the objective over the region of interest (Figure 2F) once the glass suction electrodes are connected to the roots.
    1. The procedure from the Step B5 to the Step B6 should not take more than a minute otherwise the preparation’s viability may be compromised.
    2. The dissection of the sacral spinal cord in lesioned mice (Procedure B) should be performed > 2 months after lesion for studying chronic spinal cord injury. 

      Figure 2. Dissection of the sacral spinal cord from adult mouse. A. Adult lesioned mouse under anesthesia and prepped for dissection of the sacral spinal cord. B. Incision of the skin and isolation of the spinal column from muscles and tendons. C. Isolated sacral spinal cord after dissection. D. Recording chamber for simultaneous calcium imaging of spinal interneurons and recording of motor activity. E. Sacral spinal cord positioned in the recording chamber with the transverse cut facing the microscope. F. Example of spinal neurons from a spinal cord of a Vglut2Cre (Borgius et al., 2010):: Rosa26-LSL-GCaMP3 (Ai38) mouse during calcium imaging. Scale bar = 20 µm.

Data analysis

The effect of the injury, its reproducibility, and quantification of the electrophysiological and calcium imaging data are conceptualized and quantified in our study ‘Spatiotemporal correlation of spinal network dynamics underlying spasms in chronic spinalized mice’ (Bellardita et al., 2017).


  1. The quality of the lesion is largely dependent on the manual skills of the surgeon. These techniques require experience with identification of anatomical structures, and it can be common for newly trained surgeons to disturb the lumbar dorsal/ventral root–with negative consequence to the sensory/motor function of the hindlimbs.
  2. During aspiration of the cord, the main dorsal artery should be left intact. Severing the dorsal artery during the procedure can result in degeneration of the spinal cord below the injury site. This effect may present behaviorally as flaccidity of the tail, and is typically referred to as dead tail syndrome.
  3. For obtaining high quality recordings in the in vitro preparation, special care should be used in maintaining the temperature of the solution during the laminectomy close to 4 °C to reduce cellular death.


  1. Ringer’s solution
    111 mM NaCl
    3 mM KCl
    11 mM glucose
    25 mM NaHCO3
    1.25 mM MgSO4
    1.1 mM KH2PO4
    2.5 mM CaCl2
    Oxygenated in 95% O2 and 5% CO2 to obtain a pH of 7.4 and maintained at 22-24 °C
  2. Oxygenated modified artificial cerebrospinal fluid
    125 mM Choline-Cl
    1.9 mM KCl
    1 mM CaCl2
    7 mM MgCl2
    1.2 mM KH2PO4
    10 mM HEPES
    25 mM glucose

Note: Storing of the solutions is important. The solutions can be saved in refrigeration < 10 °C and must be transparent. In case of any concern about the possibility of contamination or bacterial growth, the solution should be replaced.


This work was supported by the European Research Council (LocomotorIntegration), NINDS, Novo Nordisk Foundation, Laureate Program. The authors declare that they have no conflicts or competing interests.


  1. Bellardita, C., Caggiano, V., Leiras, R., Caldeira, V., Fuchs, A., Bouvier, J., Low, P. and Kiehn, O. (2017). Spatiotemporal correlation of spinal network dynamics underlying spasms in chronic spinalized mice. Elife 6:e23011.
  2. Bennett, D. J., Gorassini, M., Fouad, K., Sanelli, L., Han, Y. and Cheng, J. (1999). Spasticity in rats with sacral spinal cord injury. J Neurotrauma 16: 69-84.
  3. Bennett, D. J., Li, Y. and Siu, M. (2001). Plateau potentials in sacrocaudal motoneurons of chronic spinal rats, recorded in vitro. J Neurophysiol 86: 1955-1971.
  4. Borgius, L., Restrepo, C. E., Leao, R. N., Saleh, N. and Kiehn, O. (2010). A transgenic mouse line for molecular genetic analysis of excitatory glutamatergic neurons. Mol Cell Neurosci 45: 245-257.
  5. Harrison, M., O'Brien, A., Adams, L., Cowin, G., Ruitenberg, M. J., Sengul, G. and Watson, C. (2013). Vertebral landmarks for the identification of spinal cord segments in the mouse. Neuroimage 68: 22-29.
  6. Jiang, M. C. and Heckman, C. J. (2006). In vitro sacral cord preparation and motoneuron recording from adult mice. J Neurosci Methods 156: 31-36.
  7. Kiehn, O. (2016). Decoding the organization of spinal circuits that control locomotion. Nat Rev Neurosci 17(4): 224-38.
  8. Ritz, L. A., Friedman, R. M., Rhoton, E. L., Sparkes, M. L. and Vierck, C. J., Jr. (1992). Lesions of cat sacrocaudal spinal cord: a minimally disruptive model of injury. J Neurotrauma 9: 219-230.
  9. Sharif-Alhoseini, M., Khormali, M., Rezaei, M., Safdarian, M., Hajighadery, A., Khalatbari, M. M., Safdarian, M., Meknatkhah, S., Rezvan, M., Chalangari, M., Derakhshan, P. and Rahimi-Movaghar, V. (2017). Animal models of spinal cord injury: a systematic review. Spinal Cord 55(8): 714-721.


脊髓损伤(SCI)的特点是由不同的潜在神经机制引起的多种感觉/运动损伤。 将特定的感觉/运动障碍与神经机制联系起来受限于缺乏对这些神经回路的直接实验访问。 在这里,我们描述了一个解决这个缺点的实验模型。 我们产生了一种慢性脊髓损伤的小鼠模型,其可靠地再现SCI后观察到的痉挛状态,同时允许研究运动损伤体内和体外制剂 的脊髓。 该模型允许将体外和体内条件下的小鼠遗传与高级成像,行为分析和详细的电生理学技术相结合,这些技术在常规SCI中不易应用 楷模。

【背景】脊髓损伤导致了毁灭性的感官 - 运动障碍。动物模型中的大量工作已经研究了SCI后脊髓电路的病理生理状态。大多数脊髓损伤研究是在动物模型中进行的,这些动物模型与脊髓损伤后的运动损伤和恢复的临床评估相关(Sharif-Alhoseini等人,2017)。这些模型的主要限制之一是难以将感觉运动功能障碍的临床特征与特定的细胞机制联系起来。在过去的十年中,对脊髓损伤后运动障碍可能的细胞机制的新认识来自脊髓损伤的横断模型。在这里,手术横切骶骨脊髓,仅导致尾部肌肉瘫痪。该模型首先在猫(Ritz等人,1992)和后来的大鼠(Bennett等,1999)中引入,并且提供了对细胞机制的见解(Bennett等人,2001)。尽管如此,用于操纵猫和大鼠神经元活动的遗传工具非常有限。相反,小鼠允许将电生理学与遗传学相结合以鉴定和操纵脊髓中特定神经元的活性(Jiang和Heckman,2006; Kiehn,2016)。在最近的努力中,这些优势使得光学方法(例如,光遗传学,限定的神经元亚型的钙成像)能够用于解剖SCI后肌肉痉挛的神经机制(Bellardita等人。,2017)。在该协议中,我们描述了成年小鼠骶脊髓横断的技术方面,以及随后使用体外骶脊髓制备物直接检查导致慢性脊髓痉挛的神经机制伤害。

关键字:脊髓损伤, 完全横断, 体外制备, 骶髓, 痉挛状态


  1. 玻璃微量移液器(Harvard Apparatus,产品目录号:GC150F-10)
  2. 木棒棉签(Medline Scientific,目录号:300230S)
  3. 未消毒的纱布拭子(Kruuse,目录号:160120)

  4. 手术覆盖60 x 90厘米(Kruuse,目录号:141770)
  5. 吸收性拭子(Kettenbach,目录号:001911)
  6. 缝合,直切割针,不可吸收(eSutures,Ethicon,目录号:K889H)
  7. 皮下注射针头26 G棕色16毫米(BD,Microlance,目录号:304300)
  8. 23号针头(PrecisionGlide IM; BD,目录号:305145)
  9. Vetbond组织粘合剂(3M,目录号:1469C)
  10. 手术胶(3M,Vetbond,目录号:372146)
  11. 面部纸巾(VWR,目录号:115-0600)
  12. 工作台衬纸(ScienceWare,VWR,目录号:470145-292)
  13. 8周龄小鼠(JAX小鼠品系; THE JACKSON LABORATORY,目录号:000664)
  14. 100%氧气(O 2)
  15. 异氟醚(Baxter,目录号:1001936060)
  16. 利多卡因盐酸盐(Sigma-Aldrich,目录号:BP214)
  17. Betadine®外科磨砂(聚维酮碘,7.5%)
  18. 西利卡因(1%)
  19. 丁丙诺啡盐酸盐(Reckitt Benckiser Healthcare,0.3mg / ml)
  20. Carprofen(Canidryl,目录号:122964)
  21. 生理溶液(0.9%氯化钠)(Grovet,Braun Ecotainer,目录号:99512)
  22. Viscotears用于干眼症的液体凝胶(Novartis,目录号:2082642)
  23. 70%乙醇
  24. Sylgard(Sigma-Aldrich,目录号:761028-5EA)
  25. 氯化钠(NaCl,Sigma-Aldrich,目录号:433209)
  26. 氯化钾(KCl,Sigma-Aldrich,目录号:P9541)
  27. 葡萄糖(Sigma-Aldrich,目录号:G8270)
  28. 碳酸氢钠(NaHCO 3,Sigma-Aldrich,目录号:S5761)
  29. 硫酸镁七水合物(MgSO 4·7H 2 O,Sigma-Aldrich,目录号:63138)
  30. 磷酸二氢钾(KH 2 PO 4,Sigma-Aldrich,目录号:92214)
  31. 氯化钙(CaCl 2,Sigma-Aldrich,目录号:449709)
  32. 氯化镁(MgCl 2,Sigma-Aldrich,目录号:V000149)
  33. HEPES(Sigma-Aldrich,目录号:H3375)
  34. 林格的解决方案(见食谱)
  35. 含氧改性人造脑脊液(mACSF,见食谱)


  1. 镊子(精细科学工具,目录号:11251-10)
  2. 齿镊(Fine Science Tools,目录号:11154-10)
  3. Vannas弹簧剪刀 - 微锯齿(Fine Science Tools,目录号:15007-08)
  4. 杜蒙第2号椎板切除钳(Fine Science Tools,目录号:11223-20)
  5. 精细剪刀 - 碳化钨(Fine Science Tools,目录号:14568-09)
  6. 精细剪刀 - 碳化钨&amp; ToughCut(精细科学工具,目录号:14558-11)
  7. 充电式动物剪(Wahl Arco)
  8. 手术刀(精细科学工具,目录号:10020-00)
  9. 温控可变热垫(K&amp; H Manufacturing,型号:1009)
  10. 膜片式真空泵 - 润滑 - 单级(环保快递,目录号:EE0753280)
  11. 异氟醚蒸发器(Soarmed,型号:MSS-3)


  1. 完整的骶脊髓病变
    1. 准备一个专门用于啮齿动物手术的清洁和消毒区域,仅使用与手术有关的设备(图1A)。
    2. 从玻璃毛细管中准备直径为50-100μm的玻璃移液管(图1C)。拉动玻璃毛细管,然后手动调整尖端至脊髓。通过一系列直径不断增加的硅管将玻璃吸管连接到真空泵。被拉的玻璃吸管将被用于脊髓横断的手术。
    3. 手术前称重小鼠。在接下来的15天内监测动物的体重,以评估手术后潜在的体重减轻。
    4. 将鼠标置于带有5%异氟烷/ 95%氧气的密封诱导室中,直至深度麻醉(图1A.2)。
    5. 将小鼠从诱导室移动到专门用于手术的区域并准备进行手术(图1D):
      1. 将整个过程中鼠标腹侧放在加热板上以保持体温(37°C)不变(图1D.1)。
      2. 在整个手术期间使用2%异氟烷。
      3. 检查动物的反射来验证麻醉的适当状态。不应该出现夹捏诱发的反应。我们通过捏尾部和后足的掌骨区域评估了踏板收回反射。
      4. 确保动物在手术期间不会移动(可能是由于触摸背根造成的)。用附着在四肢上的条带固定动物。避免四肢过度伸展,这可能会损伤关节并损害动物的呼吸(图1D.3)。
      5. 沿着脊柱的rostrocaudal轴剃去老鼠的背部。将钠碘应用于剃毛区域并保持5分钟(图1D.4)。
      6. 切记避免将手或器械放在鼠标胸部。外部压力可能会干扰呼吸和/或血液循环。

      7. 在鼠标本体上涂上外科手术套,在切口处留下一个窗口(图1D.5)。&nbsp;
    6. 第二骶节段和横断的定位:
      1. 用两根手指定位T12椎体。 T12椎骨具有所有椎骨中最长的棘突,并且如果小鼠的脊柱屈曲,则T12椎骨向外突出到脊柱中。使用手术刀,从大约T12椎体到L4椎体纵向切开皮肤(图1E)。
      2. 脊髓的第二骶节段(S2)位于L2椎体的头侧部分之下,L1和L2椎体之间的边界处。 L2的棘突指向直立,并且应该被用作用小眼睛剪刀进行深度垂直切口的标志。如果切割垂直进行,则会显示L1和L2椎体之间的黄韧带(图1F)。为了更好地了解成年小鼠的解剖标志,特别是腰椎体和脊髓之间的关系,请参阅Harrison等人,2013年。
      3. 用眼睛剪刀切开黄韧带。
      4. 在绳索顶部涂上1%利多卡因,以防止接触脊髓或背根引起的运动。等待药物生效(大约半分钟),然后用精细的镊子将背根定位为尽可能侧向。在其他结构性损伤(例如,骨骼,动脉,肌腱)的情况下,外科医生应考虑将动物排除在后续分析之外。
      5. 如果对黄韧带的切割正确执行,则不会对周围组织(肌肉,韧带或皮肤)造成其他伤害,也不会看到血迹。
      6. 从软线的一侧开始,使用连接到真空吸力的玻璃吸管去除S2脊椎组织。保持抽吸组织,直至在绳索的尾端和尾端之间观察到完全不连续,总共对应于一个部分。
    7. 缝合手术伤口并让动物恢复:

      1. 在伤口部位缝合脊柱周围的肌肉以保护绳索,并使用兽医胶水密封皮肤。

      2. 术后给予丁丙诺啡(0.1 mg / kg),卡洛芬(5 mg / kg),必要时皮下注射0.3 ml无菌生理溶液2至5天。
      3. 关闭麻醉,将鼠标放回笼中,加热垫使动物保持1-2小时的温度。该动物可以在第一周内单独饲养,之后如果恢复完成,可以将它与另一只动物一起饲养。
      4. 每天监测动物的窘迫迹象,包括体重减轻(应该避免体重下降> 10%),脱水或感染。在任何这些情况下,外科医生都应该咨询兽医的建议和解决方案。
      5. 伤害只应引起尾部肌肉瘫痪,不应影响膀胱或后肢。然而,在日常的术后护理中,监测膀胱功能障碍是很重要的,如果损伤部位过于安全,有时会发生膀胱功能障碍。&nbsp;

    图1.成年小鼠的骶脊髓损伤。 :一种。为手术准备的区域容易获得必要的设备:1)解剖显微镜; 2)麻醉诱导室; 3)麻醉面膜; 4)异氟醚蒸发器; 5)加热垫; 6)用于抽吸连接到真空泵的脊髓的玻璃吸管; 7)吸尘器; 8)手术器械消毒装置。 B.用(从左到右)损伤骶脊髓的手术器械:剪刀,镊子,眼膏,兽用胶和缝合线。 C.吸取脊髓的玻璃吸管和真空泵。 D.准备用于损伤程序的小鼠。 1)加热垫; 2)提供麻醉的面罩; 3)防止突然移动的胶带条; 4)剃光并用碘化钠预处理的感兴趣区域; 5)鼠标身体的绿色覆盖物,在感兴趣的区域有一个工作窗口。 E.感兴趣区域的皮肤切口。 F.手术区域的示意图,其中椎体L1和L2与脊髓的第二骶节段。 G.切开黄韧带后切开L2椎体。 H.使用玻璃吸管抽吸脊髓。我-L。在脊髓损伤后2个月时切开的,损伤的骶带的例子有不完整的(I)或完成(L)病变。

  2. 解剖成年慢性脊髓小鼠的骶脊髓
    1. 准备一个专门用于啮齿动物手术的清洁消毒区域,方便使用隔离脊髓所需的设备(类似于图1A)。
    2. 将动物置于麻醉诱导室(5%异氟烷/ 95%氧气)中,当深度麻醉时将动物从诱导室移至解剖台。按照步骤A4c检查反射。
    3. 将鼠标放在工作台衬纸(Scienceware)上并通过面膜敷用异氟醚(2%)。如步骤A5c中检查适当麻醉状态的反射。用胶带固定四肢,剃刮背部,并用酒精清洁该区域(图2A)。
    4. 沿着感兴趣的部位进行椎板切除术:
      1. 如上所述识别T12。
      2. 将皮肤从T12椎体切到L5椎体。请记住,脊髓的第二骶节段位于腰脊髓第二椎体(L2)的下方。

      3. 通过切割周围的肌肉和腱来暴露脊柱(图2B)。
      4. 定位T13椎体的棘突(T13椎骨上有最后一对肋骨),并开始背侧椎板切除术(去除椎骨背侧部分的外科手术)。
      5. 开始用冷mACSF(20毫升/分钟)灌注脊柱,以减缓新陈代谢并减少目标部位的血流量。

      6. 用剪刀剪下椎体的左右两侧进行椎板切除术。
      7. 避免使用剪刀损伤脊髓,当剪刀尖从左侧移动到右侧时(或反之亦然),有时会发生剪刀。损伤可能导致脊髓挫伤或挫伤。
      8. 冷(〜4°C),含氧的mACSF在脊髓上连续流动(20毫升/分钟)。
    5. 骶髓的分离:
      1. 当脊柱从尾部腰段暴露于马尾时,椎板切除完成(图2C)。

      2. 通过面膜给小鼠提供纯氧,持续约5分钟以增加血液氧合水平。
      3. 切开腹部肌肉的水平并保存腹部动脉。这种切割会导致血压下降,防止在隔离期间血液在绳索处溢出。
      4. 切断尾骶腰段水平线,并切断脊髓右侧和左侧的腹侧根部,将其与脊柱隔离。
      5. 当你到达病变部位时要特别注意。在损伤的水平上,硬膜常与椎体相连,需要从脊柱其余部位仔细分离脊髓和硬脑膜。
      6. 一旦电线完全脱离,将其移入具有连续流动(20 ml / min)的冷氧合-mACSF的解剖室。
      7. 小心并彻底地从脊髓中取出硬脑膜,使氧合作用更多地扩散到组织中。切割脊髓,背根和腹侧根部以减少长度,并在实验过程中更容易识别不同部分和根部(图2C)。
      8. 一旦切断了所有脊髓根部并完成脊髓的分离,脊髓就可以移动到Sylgard覆盖的灌注室(图2D)。
    6. 用于脊髓中间神经元和腹侧根记录的同时钙成像的骶脊髓的体外制备
      1. 将脊髓移至录音室。记录室具有Sylgard底部和附接的Sylgard'桥',其允许线被置于L形位置(图2E),使得脊柱的冠状平面可以用物镜成像。 br />

      2. 在实验过程中保持连续流通的含氧林格溶液。
      3. 将绳索从最尾端向上固定,以确保机械稳定性。绳索的头端靠在桥上,弯曲的分针用作钩子以保持绳索就位(图2E)。
      4. 吸气电极用于记录运动活动(S4-Co1)并刺激背根。
      5. 一旦玻璃吸入电极连接到根部,钙成像通过降低感兴趣区域上的物镜(图2F)进行。
    1. 从步骤B5到步骤B6的程序不应超过一分钟,否则准备的可行性可能受损。
    2. 在病变小鼠中解剖骶脊髓(程序B)应当>研究慢性脊髓损伤病变后2个月。&nbsp;

      图2.解剖成年小鼠骶髓。 :一种。在麻醉下成人损伤的小鼠并准备解剖骶脊髓。 B.切开皮肤并从肌肉和肌腱中分离脊柱。 C.解剖后的孤立的骶脊髓。 D.记录室用于脊髓中间神经元的同时钙成像和记录运动活动。 E.骶骨脊髓位于记录室内,横切面向显微镜。 F.来自Vglut2 Cre的脊髓的脊髓神经元的实例(Borgius等人,2010) ::在钙成像期间的Rosa26-LSL-GCaMP3(Ai38)小鼠。比例尺= 20微米。


我们的研究“慢性脊髓性小鼠脊髓网络动力学基础痉挛的时空相关性”( Bellardita ,2017 )。


  1. 病变的质量在很大程度上取决于外科医生的手工技能。这些技术需要解剖结构的识别经验,新接受训练的外科医生通常会扰乱腰背/腹根,这对后肢的感觉/运动功能具有负面影响。
  2. 在抽出绳索的过程中,主背动脉应保持完整。手术过程中切断背侧动脉可导致损伤部位下方的脊髓变性。这种效应可能在行为上表现为尾部松弛,通常称为死尾症候群。
  3. 为了在体外制备中获得高质量的记录,应特别注意在椎板切除术期间保持溶液温度接近4°C以减少细胞死亡。


  1. 林格的解决方案
    111 mM NaCl
    3 mM KCl
    11 mM葡萄糖
    25mM NaHCO 3 3/2 1.25mM MgSO 4
    1.1mM KH 2 PO 4 4/2 2.5 mM CaCl 2 2/2 在95%O 2和5%CO 2中进行氧化以获得7.4的pH并保持在22-24℃。
  2. 含氧改性人造脑脊液
    1.9 mM KCl
    1 mM CaCl 2 2 /
    7mM MgCl 2·/ 2 1.2mM KH 2 PO 4 4/2 10 mM HEPES
注意:存储解决方案非常重要。解决方案可以保存在制冷系统中&nbsp;&lt; 10°C,并且必须是透明的。如果担心污染或细菌生长的可能性,应该更换解决方案。




  1. Bellardita,C.,Caggiano,V.,Leiras,R.,Caldeira,V.,Fuchs,A.,Bouvier,J.,Low,P。和Kiehn,O。(2017)。
  2. Bennett,D.J。,Gorassini,M.,Fouad,K.,Sanelli,L.,Han,Y。和Cheng,J。(1999)。 骶髓损伤大鼠的痉挛状态 J Neurotrauma 16:69-84。
  3. Bennett,D.J.,Li,Y。和Siu,M。(2001)。 体外记录的慢性脊髓大鼠的骶尾运动神经元的高原潜能。
  4. Borgius,L.,Restrepo,C.E.,Leao,R.N.,Saleh,N.and Kiehn,O。(2010)。 一种用于兴奋性谷氨酸能神经元分子遗传分析的转基因小鼠系。 Cell Neurosci 45:245-257。
  5. Harrison,M.,O'Brien,A.,Adams,L.,Cowin,G.,Ruitenberg,M.J。,Sengul,G。和Watson,C。(2013)。 脊椎标志用于鉴定小鼠脊髓节段。 Neuroimage 68:22-29。
  6. Jiang,M.C。和Heckman,C.J。(2006)。 成年小鼠体外骶髓制备和运动神经元记录 a> J Neurosci Methods 156:31-36。
  7. Kiehn,O.(2016)。 解码控制运动的脊髓回路的组织。 Nat Rev Neurosci 17(4):224-38。
  8. Ritz,L.A.,Friedman,R.M.,Rhoton,E.L。,Sparkes,M.L。和Vierck,C.J.,Jr。(1992)。 猫骶尾脊髓损伤:最小破坏性损伤模型 J Neurotrauma 9:219-230。
  9. Sharif-Alhoseini,M.,Khormali,M.,Rezaei,M.,Safdarian,M.,Hajighadery,A.,Khalatbari,MM,Safdarian,M.,Meknatkhah,S.,Rezvan,M.,Chalangari,M. ,Derakhshan,P.和Rahimi-Movaghar,V.(2017)。 脊髓损伤的动物模型:系统综述 脊髓55(8):714-721。
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引用: Readers should cite both the Bio-protocol article and the original research article where this protocol was used:
  1. Bellardita, C., Marcantoni, M., Löw, P. and Kiehn, O. (2018). Sacral Spinal Cord Transection and Isolated Sacral Cord Preparation to Study Chronic Spinal Cord Injury in Adult Mice. Bio-protocol 8(7): e2784. DOI: 10.21769/BioProtoc.2784.
  2. Bellardita, C., Caggiano, V., Leiras, R., Caldeira, V., Fuchs, A., Bouvier, J., Low, P. and Kiehn, O. (2017). Spatiotemporal correlation of spinal network dynamics underlying spasms in chronic spinalized mice. Elife 6:e23011.