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A Controlled Cortical Impact Mouse Model for Mild Traumatic Brain Injury
轻度脑外伤模型建立:小鼠脑皮层可控性撞击模型   

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Abstract

Traumatic brain injury (TBI) affects millions of people worldwide; however, the immediate impact of TBI and the secondary injury mechanisms are still not fully understood. TBI can cause devastating neuromotor deficits in both acute and chronic stages. Time course studies utilizing animal models of focal TBI have provided essential insight into TBI neuropathology. Here, we describe a surgical technique for creating a mouse model of focal, mild TBI (Dixon et al., 1991; Smith et al., 1995; Bolkvadze and Pitkanen, 2012). Furthermore, we provide protocols for validating TBI models using behavioral tests that examine post-traumatic neuromotor deficits resulting from TBI neuropathology (Fujimoto et al., 2004; Febinger et al., 2015; Smith et al., 1995; Bolkvadze and Pitkanen, 2012).

Keywords: TBI(TBI), Concussion(脑震荡), Head trauma(头部外伤)

Materials and Reagents

  1. Sterile cotton tipped applicators
  2. Gauze pads (ATC Medical, CurityTM, catalog number: 686132 )
  3. Nolvasan surgical scrub (Zoetis Inc., catalog number: 8NOL413 )
  4. Adult C57BL6/J laboratory mice (6 weeks old)
  5. 70% ethanol
  6. Isoflurane
  7. Opthalmic ointment
  8. 3.0% hydrogen peroxide solution
  9. Sterile saline
  10. VetBond glue (3M, catalog number: 1469SB )
  11. 70% isopropanol

Equipment

  1. Surgical Tools
    1. Scalpel handle (Fine Science Tools, catalog number: 10003-12 )
    2. Sterile scalpel blades (Fine Science Tools, catalog number: 10010-00 )
    3. Dumont forceps (Fine Science Tools, catalog number: 11251-10 )
    4. 5.0-mm diameter trephine (Figure 1) (Fine Science Tools, catalog number: 18004-50 )
    5. Pin vise (Figure 1) (ProEdge, catalog number: PRO58111 )


    Figure 1. Trephine in pin vise

  2. Hot bead sterilizer (Fine Science Tools, catalog number: 18000-45 )
  3. Warm water recirculator (Kent Scientific, catalog number: TP-700 )
  4. Polystyrene weighing boat
  5. Controlled cortical impact (CCI) device (Impact One Stereotaxic Impactor for CCI) (Leica Biosystems, catalog number: 39463920 )
    1. 3.0-mm diameter metal piston (Leica Biosystems , catalog number: 39463920)
  6. Small animal stereotaxic instrument with mouse adapters (Kopf Instruments, model: 900 and 923-B )
  7. Scale
  8. Leather hole punch, sharp (Anytime Tools)
  9. Incandescent lamp (50-75 watt)
  10. Inclined plane device, custom made (Figure 2) (The angle board device is 50 x 60 cm in size and is covered with a vertically grooved rubber mat. The platform was attached to a hinge, allowing the researcher to manually adjust the angle of the board.)


    Figure 2. Inclined plane (custom made)

Procedure

  1. Animal preparation
    1. Autoclave surgical tools, cotton tipped applicators, and gauze pads. Disinfect the polystyrene weighing boat with 70% ethanol.
    2. Prepare the surgical area. Turn on the warm water recirculator, and the hot bead sterilizer so that they can warm up. It is recommended to bead sterilize each surgical tool before use. Be sure to cool the tool before using it on the mouse.
      1. Impactor device should be placed near surgical area. Screw in 3.0-mm diameter metal piston and set the impact velocity to 5.0 m/s and dwell time of 100 ms from the dura for a mild injury (Figure 3).


        Figure 3. Stereotaxic setup

    3. Weigh the animal and record its mass in grams.
      Note: Recording the mouse’s body weight prior to performing surgery is important for monitoring post-operative behavior.
    4. Anesthesia (Figure 4): Place the mouse in the induction box and turn on the oxygen tank so that it is flowing at approximately 1.0 L per minute. Turn on the isoflurane vaporizer to 4.0%. Be sure to monitor the mouse as it is becoming unconscious. After approximately 2-3 min, the animal should exhibit slow, even, quiet breathing. To determine if the mouse is fully anesthetized, pinch the mouse’s toe. If the mouse draws its paw towards its body, it is not appropriately anesthetized. If there is no reaction, turn down the isoflurane to 2.0%-2.5%, and route the isoflurane to the stereotaxic device.


      Figure 4. Isoflurane vaporizer and induction box

    5. Stereotaxic device: Place the mouse’s mouth on the bite bar with the teeth in the hole, and move the anesthesia cone over the animal’s nose for a snug fit. Guide the two ear bars into the auditory canals and secure. Use the scales to ensure both ear bars are equally placed. The head should be completely secure and level so that it will not move during the craniotomy. At this time, place ophthalmic ointment generously onto the eyes. To maintain the mouse’s body temperature, a heat source needs to be provided. Placing the mouse on a pad that is temperature controlled with a warm water recirculator is recommended. 
    6. Continue to monitor the depth of anesthesia, and adjust the vaporizer as needed. 1.25%-1.75% should be the proper range of maintenance anesthesia for most mice. 
    7. Remove fur from the mouse’s head using clippers to expose the skin. 
    8. Prepare the surgical site by cleaning the head area with Nolvasan surgical scrub using sterile cotton tipped applicators.

  2. Surgical procedure
    1. Using a scalpel blade, make a midline skin incision.
    2. Using the Dumont forceps, remove as much fascia as possible. Then, apply 3.0% hydrogen peroxide to the skull to clean and dry the skull area.
      Note: Be sure to flush the hydrogen peroxide with sterile saline when done. 
    3. Once the skull has been dried, create a 5-mm diameter craniotomy over the left parietal cortex (between lambda and bregma) using the 5-mm diameter trephine. Apply gentle, uniform pressure when using the trephine. Once the area feels loose, carefully remove the fragment using Dumont forceps.
    4. At this point, flush the area with saline to remove any bone fragments that may have accumulated. Be sure to keep the brain moist throughout the surgery. Check the integrity of the dura (Figure 5). If the dura is damaged, it will appear as a detached, light layer. Damaging the dura can confound studies and potentially affect behavior test outcomes.


      Figure 5. Craniotomy

    5. Attach the controlled cortical impact (CCI) device to the stereotax. Attach the contact sensor to one of the ear bars. Lower the piston until it just touches the dura (the machine will beep when contact is made). Retract the piston and then move down the CCI device 0.5 mm for a mild injury. Then, hit the impact switch on the CCI device (Figures 6 and 7).


      Figure 6. CCI impact


      Figure 7. CCI impact

    6. After the impact, flush the area again with sterile saline, and stop any bleeding that may have occurred by applying light pressure with a sterile cotton tipped applicator.
    7. Then, cover the exposed brain with a 5-mm disc created from a polystyrene weighing boat using the leather hold punch. Apply a small amount of VetBond glue along the edges of the disc. The trick is to keep the brain area moist, but to dry the skull area as much as possible. This will fix the plastic disc to the skull but not the brain (Figure 8). Make sure to remove any VetBond that has adhered to the skin.


      Figure 8. Covering the site of impact

    8. Once the plastic disc is fixed over the injury site, close the incision area with sutures.
    9. Turn off the isoflurane, and administer analgesics subcutaneously. Remove the mouse from the stereotactic frame, and place the animal in a clean cage. Position an incandescent lamp (50-75 watt) approximately 12 inches away from the animal. Be sure to place the lamp so that only half of the cage is illuminated and the rodent can escape the light if desired. Be sure to monitor the rodent until it has fully woken up and is ambulatory.

  3. Composite neuroscore behavioral tasks
    The composite neuroscore test (CN) was implemented to evaluate posttraumatic neurological deficits using a 28-point system (Fujimoto et al., 2004). For each task, animals are scored on a scale of 0-4, with 0 valued at severely impaired and 4 at no impairment (Fujimoto et al., 2004). Mice undergo a battery of tests: right and left forelimb and hindlimb flexion, right and left lateral pulsion, and inclined plane test. The scores for these tests are added together to calculate the CN.
    1. Hindlimb and forelimb flexion
      1. Grasp the mouse at the base of the tail while the forepaws remain on a flat surface. 
      2. Observe the hindlimb position for 10 sec. An uninjured animal will characteristically extend its hindlimbs back and out with full extension and spread toes, whereas an injured animal will retract its hindlimbs towards the abdomen and curl its toes (Figures 9 and 10) (Fujimoto et al., 2004). Scoring should be as follows for the left and right limb:
        1. 4 (no impairment)-Hindlimbs splayed outward, away from the animal's body and toes are spread.
        2. 3-If the hindlimb briefly retracts towards the abdomen less than 25% of the time, and toes are spread. 
        3. 2-If the hindlimb retracts towards the abdomen more than 25% of the time, but less than 50% of the time, and toes are curled. 
        4. 1-If the hindlimb retracts towards the abdomen more than 50% of the time, and toes are curled. 
        5. 0 (severely impaired)-If the hindlimb is entirely retracted while touching the abdomen and toes are curled for the entire time while lifted. 


        Figure 9. Healthy hindlimb position


        Figure 10. Impaired hindlimb flexion

      3. Forelimb flexion is similar to hindlimb flexion test (Fujimoto et al., 2004). Grasp the mouse by the base of the tail and lift the mouse. Gently and slowly lower the mouse head first towards a flat surface. Uninjured animals will immediately extend their forelimbs towards the mat (Figure 11) (Fujimoto et al., 2004). Use the following parameters to score the left and right forelimbs: 
        1. 4 (no impairment)-Forelimb is extended forward, away from the animal’s body and toes are spread. 
        2. 3-If the forelimb briefly retracts towards the abdomen less than 25% of the time, and toes are spread. 
        3. 2-If the forelimb retracts towards the abdomen more than 25% of the time, but less than 50% of the time, and toes are curled. 
        4. 1-If the forelimb retracts towards the abdomen more than 50% of the time, and toes are curled. 
        5. 0 (severely impaired)-If the forelimb is entirely retracted while touching the abdomen and toes are curled for the entire time while lifted.


        Figure 11. Forelimb flexion

    2. Lateral pulsion task-The lateral pulsion task tests the animal's ability to coordinate the movements of all four limbs when pressure is applied laterally.
      1. Place the mouse facing away from the observer. Wipe a pen down with 70% isopropanol and apply gentle, lateral pressure to the mouse's midsection. Uninjured animals will show complete resistance to falling, while injured animals will show little resistance (Figure 12) (Fujimoto et al., 2004). Score the behavior using the following parameters: 
        1. 4 (no impairment)-Complete resistance to falling and no steps taken away from stimulus. 
        2. 3-Moderate resistance, and one step taken with forelimb away from the stimulus. 
        3. 2-Moderate resistance, and more than one step taken away from stimulus. 
        4. 1-Slight resistance, but mouse eventually falls laterally. 
        5. 0 (severely impaired)-No resistance, and mouse falls laterally. 


        Figure 12. Lateral pulsion

    3. Inclined plane test-The difference between the highest angles an animal can stay on an inclined plane without falling at baseline and after injury typically correlates with the severity of the brain injury (Fujimoto et al., 2004). The angle board device is 50 x 60 cm in size and is covered with a vertically grooved rubber mat (Figure 2). Baseline values should be collected the day before CCI surgery. These values are compared to post-TBI scores.
      1. Baseline scoring: Set the angle board at 40° and make sure it is secure. Clean any hair on the board using the vacuum. 
        Note: Do not clean the board with alcohol or any other liquid. Note the room temperature.
      2. Place the mouse in the vertical direction first, then the left, and then the right. A successful completion is determined when the animal is able to stand still in that direction for 5 sec without being held by the tail. Each direction is allotted three attempts (Figure 13).


        Figure 13. Inclined plane test

      3. If the mouse succeeds in standing on the board, increase the angle by 2.5° and repeat step C2 for each direction. Continue increasing the angle until the mouse can no longer stand on the board in any of the directions. Note the maximum angles.
      4. Post-TBI scoring: Post-injured animals start testing at 10° below their lowest maximum baseline value. For example, if a mouse fell in the right direction at 47.5°, the left direction at 50°, and the vertical direction at 55°, the starting angle for post-injury testing would be 37.5° (10° less than the right direction maximum angle). 
      5. Scores are assigned for each direction by comparing the post-injury values to the baseline maximum angles. The three scores are then averaged for a final score. Use the following parameters to score the performances: 
        1. 4 (no impairment)-There is no difference or the angle was higher than the baseline maximum angle. 
        2. 3-There is a 2.5° decrease from the baseline maximum angle. 
        3. 2-There is a 5° decrease from the baseline maximum angle. 
        4. 1-There is a 7.5° decrease from the baseline maximum angle. 
        5. 0 (severely impaired)-There is a 10° or more decrease from the baseline maximum angle. 

The composite neuroscore is generated by combining the scores of all seven tests.

Acknowledgments

This study was supported by the Department of Anesthesiology and Pain Medicine of the University of Washington and by NIH SC1GM095426. This protocol was adapted from what was described in Dixon et al., 1991.

References

  1. Bolkvadze, T. and Pitkanen, A. (2012). Development of post-traumatic epilepsy after controlled cortical impact and lateral fluid-percussion-induced brain injury in the mouse. J Neurotrauma 29(5): 789-812.
  2. Dixon, C. E., Clifton, G. L., Lighthall, J. W., Yaghmai, A. A. and Hayes, R. L. (1991). A controlled cortical impact model of traumatic brain injury in the rat. J Neurosci Methods 39(3): 253-262. 
  3. Febinger, H. Y., Thomasy, H. E., Pavlova, M. N., Ringgold, K. M., Barf, P. R., George, A. M., Grillo, J. N., Bachstetter, A. D., Garcia, J. A., Cardona, A. E., Opp, M. R. and Gemma, C. (2015). Time-dependent effects of CX3CR1 in a mouse model of mild traumatic brain injury. J Neuroinflammation 12: 154. 
  4. Fujimoto, S. T., Longhi, L., Saatman, K. E., Conte, V., Stocchetti, N. and McIntosh, T. K. (2004). Motor and cognitive function evaluation following experimental traumatic brain injury. Neurosci Biobehav Rev 28(4): 365-378. 
  5. Smith, D. H., Soares, H. D., Pierce, J. S., Perlman, K. G., Saatman, K. E., Meaney, D. F., Dixon, C. E. and McIntosh, T. K. (1995). A model of parasagittal controlled cortical impact in the mouse: cognitive and histopathologic effects. J Neurotrauma 12(2): 169-178.

简介

创伤性脑损伤(TBI)影响全世界数百万人; 然而,TBI和继发性损伤机制的直接影响仍然没有完全理解。 TBI可以在急性和慢性阶段引起毁灭性的神经性运动缺陷。 使用局灶性TBI动物模型的时程研究提供了对TBI神经病理学的重要见解。 在这里,我们描述了用于产生局灶性,轻度TBI的小鼠模型的外科技术(Dixon等人,1991; Smith等人,1995; Bolkvadze和Pitkanen, 2012)。 此外,我们提供用于验证TBI模型的方案,其使用行为测试,其检查由TBI神经病理学产生的创伤后神经运动缺陷(Fujimoto等人,2004; Febinger等人, 2015; Smith 等人,1995; Bolkvadze和Pitkanen,2012)。

关键字:TBI, 脑震荡, 头部外伤

材料和试剂

  1. 无菌棉签
  2. 纱布垫(ATC Medical,Curity TM ,目录号:686132)
  3. Nolvasan手术擦洗(Zoetis Inc.,目录号:8NOL413)
  4. 成年C57BL6/J实验室小鼠(6周龄)
  5. 70%乙醇
  6. 异氟烷
  7. 眼用软膏
  8. 3.0%过氧化氢溶液
  9. 无菌盐水
  10. VetBond胶(3M,目录号:1469SB)
  11. 70%异丙醇

设备

  1. 外科工具
    1. 手术刀柄(Fine Science Tools,目录号:10003-12)
    2. 无菌手术刀刀片(Fine Science Tools,目录号:10010-00)
    3. Dumont镊子(Fine Science Tools,目录号:11251-10)
    4. 5.0-mm直径环钻(图1)(Fine Science Tools,目录号:18004-50)
    5. 固定钳(图1)(ProEdge,目录号:PRO58111)


    图1.钉头钳钻孔

  2. 热珠灭菌器(Fine Science Tools,目录号:18000-45)
  3. 温水再循环器(Kent Scientific,目录号:TP-700)
  4. 聚苯乙烯称船
  5. 受控皮质影响(CCI)装置(用于CCI的Impact One Stereotaxic Impactor)(Leica Biosystems,目录号:39463920)
    1. 3.0-mm直径金属活塞(Leica Biosystems,目录号:39463920)
  6. 使用小鼠适配器的小动物立体定位仪(Kopf Instruments,型号:900和923-B)
  7. 规模
  8. 皮革打孔,锋利(随时工具)
  9. 白炽灯(50-75瓦)
  10. 倾斜平面装置,定做(图2)(角板装置的尺寸为50×60厘米,并覆盖有垂直沟槽橡胶垫。平台连接到铰链,允许研究人员手动调整角度 板。)


    图2.倾斜平面(自定义)

程序

  1. 动物准备
    1. 高压灭菌外科工具,棉签和纱布垫。 用70%乙醇消毒聚苯乙烯称量船。
    2. 准备手术区。 打开温水再循环器和热珠灭菌器,以便他们可以预热。 建议在使用前对每个手术工具进行珠消毒。 在将鼠标用于鼠标之前,请务必冷却该工具。
      1. 撞击装置应放置在手术区附近。 拧入3.0毫米直径的金属活塞,将冲击速度设置为5.0米/秒,从硬脑膜的轻度损伤的停留时间为100毫秒(图3)。


        图3.立体定位设置

    3. 称重动物并记录其质量(克)。
      注意:在手术前记录鼠标的体重对于监测术后行为是很重要的
    4. 麻醉(图4):将小鼠放在感应箱中,打开氧气罐,使其以每分钟大约1.0升的速度流动。打开异氟烷蒸发器至4.0%。确保监视鼠标,因为它变得无意识。大约2-3分钟后,动物应该表现出缓慢,平静,安静的呼吸。要确定鼠标是否完全麻醉,捏住鼠标的脚趾。如果小鼠将其爪朝向其身体,则不能适当地麻醉。如果没有反应,将异氟烷调低至2.0%-2.5%,然后将异氟烷送至立体定位装置。


      图4.异氟烷蒸发器和感应箱

    5. 立体定位装置:将小鼠的嘴放在咬合棒上,牙齿在孔中,并将麻醉锥移动到动物的鼻子上以适合紧贴。引导两个耳杆到听觉运河和安全。使用秤来确保两个耳杆均匀放置。头部应该是完全安全和水平的,以便它在开颅手术期间不会移动。此时,将眼膏大量地置于眼睛上。为了保持鼠标的体温,需要提供热源。建议将鼠标放在用温水再循环器控制温度的垫子上。
    6. 继续监测麻醉深度,并根据需要调节蒸发器。 1.25%-1.75%应该是大多数小鼠的适当维持麻醉范围。
    7. 使用剪刀从鼠标的头部去除毛皮,露出皮肤。
    8. 准备手术部位,通过清洁头部区域与Nolvasan手术擦洗使用无菌棉头胶枪。

  2. 外科手术
    1. 使用解剖刀刀片,做一个中线皮肤切口。
    2. 使用杜蒙镊子,尽可能多的筋膜。 然后,应用3.0%的过氧化氢到头骨清洁和干颅骨区域。
      注意:完成后,请务必用无菌盐水冲洗过氧化氢。
    3. 一旦头骨已经干燥,使用直径5毫米的环钻在左顶叶皮层(λ和前囟)之间创建一个5mm直径的开颅手术。 使用时使用温和均匀的压力 环钻。一旦区域感觉松动,使用杜蒙镊子小心地删除片段。
    4. 此时,用盐水冲洗该区域以清除可能积聚的任何骨碎片。一定要保持大脑在整个手术潮湿。检查硬脑膜的完整性(图5)。如果硬膜损伤,它将显示为一个分离的,轻的层。损伤硬脑膜会混淆研究结果,并可能影响行为测试结果

      图5.撕裂术

    5. 将受控皮层撞击(CCI)设备连接到立体声。将接触传感器连接到其中一个耳杆。降下活塞,直到它刚好接触硬脑膜(接触时机器会发出哔声)。收回活塞,然后向下移动CCI装置0.5 mm,轻度受伤。然后,点击CCI设备上的冲击开关(图6和7)。


      图6. CCI影响


      图7. CCI影响

    6. 冲击后,用无菌盐水再次冲洗该区域,并停止任何可能通过用无菌棉签施加器施加轻压而发生的出血。
    7. 然后,使用皮革夹持冲头,用由聚苯乙烯称量船创建的5-mm圆盘覆盖暴露的脑。沿光盘边缘涂少量VetBond胶水。诀窍是保持大脑区域潮湿,但尽可能干燥颅骨区域。这将修复塑料光盘到头骨,但不是大脑(图8)。确保清除粘附在皮肤上的任何VetBond。


      图8.覆盖影响网站

    8. 一旦塑料盘固定在损伤部位上,用缝线闭合切口区域。
    9. 关闭异氟醚,并皮下给予镇痛药。从立体定位框架上删除鼠标,并将动物放在一个干净的笼子里。放置距离动物约12英寸的白炽灯(50-75瓦特)。一定要放置灯泡,使笼子只有一半被照亮,如果需要啮齿动物可以逃避光线。确保监测啮齿动物,直到它完全唤醒,并且 走动。

  3. 复合神经系统行为任务
    使用28点系统进行复合神经评分试验(CN)以评价创伤后神经功能缺陷(Fujimoto等人,2004)。 对于每个任务,以0-4的等级对动物进行评分,0表示严重受损,4表示无损害(Fujimoto等人,2004)。 小鼠经历一系列测试:左右前肢和后肢屈曲,左右横向脉动和倾斜平面测试。 将这些测试的得分加在一起以计算CN。
    1. 后肢和前肢屈曲
      1. 抓住鼠标在尾巴的根部,而前爪保持在平坦的表面上。
      2. 观察后肢位置10秒。未受伤的动物将特征性地将其后肢以完全伸展和伸展的脚趾向后延伸,而受伤的动物将使其后肢向腹部收回并卷曲其脚趾(图9和10)(Fujimoto等人 。,2004)。左肢和右肢的评分应如下:
        1. 4(无损伤) - 后肢向外张开,远离动物的身体和脚趾传播。
        2. 3 - 如果后肢短暂地向腹部收缩小于25%的时间,脚趾会蔓延。
        3. 2 - 如果后肢超过25%的时间缩回腹部,但小于50%的时间,脚趾会卷曲。
        4. 1 - 如果后肢向腹部收缩超过50%的时间,脚趾会卷曲。
        5. 0(严重受损) - 如果后肢在接触腹部时完全收缩,并且脚趾在提起时卷曲整个时间。 


        图9.健康后肢位置


        图10.受损的后肢屈曲

      3. 前肢屈曲类似于后肢屈曲试验(Fujimoto等人,2004)。抓住鼠标的尾部,抬起鼠标。轻轻地,慢慢地将鼠标头降低到平坦的表面。未受伤的动物将立即将其前肢向垫子延伸(图11)(Fujimoto等人,2004)。使用以下参数对左右前肢进行评分:
        1. 4(无损伤) - 前肢向前伸展,远离动物的身体,脚趾伸展。
        2. 3 - 如果前肢短暂地向腹部缩回不到25%的时间,脚趾会蔓延。
        3. 2 - 如果前肢向腹部收缩超过25%的时间,但小于50%的时间,脚趾会卷曲。
        4. 1 - 如果前肢向腹部收缩超过50%的时间,脚趾会卷曲。
        5. 0(严重受损) - 如果前肢在接触腹部时完全缩回,并且脚趾在提起时卷曲整个时间。


        图11.前肢屈曲

    2. 横向脉动任务 - 横向脉动任务测试动物在横向施加压力时协调所有四肢运动的能力。
      1. 将鼠标置于远离观察者的位置。 用70%异丙醇擦拭笔,对小鼠的中部施加温和的侧向压力。 未受伤的动物将表现出完全的抗下落,而受伤的动物将显示出很小的抗性(图12)(Fujimoto等人,2004)。 使用以下参数对行为进行评分:
        1. 4(无损害) - 完全抗性下降,没有步骤从刺激。
        2. 3 - 中等的阻力,和前肢远离刺激采取一步。
        3. 2 - 中等的阻力,以及一步以上的刺激。
        4. 1 - 轻微的阻力,但鼠标最终横向下降。
        5. 0(严重受损) - 无抵抗力,且小鼠横向跌落。


        图12.侧向脉动

    3. 倾斜平面测试 - 动物可以停留在倾斜平面上的最高角度之间的差异,在基线和损伤后不下降通常与脑损伤的严重程度相关(Fujimoto等人,2004)。 角板装置尺寸为50×60厘米,并覆盖有垂直沟槽橡胶垫(图2)。 基线值应在CCI手术前一天收集。 将这些值与TBI后得分进行比较。
      1. 基线评分:将角板设置在40°,并确保其牢固。使用真空清洁板上的任何头发。
        注意:不要用酒精或任何其他液体清洁电路板。请注意室温 。
      2. 首先将鼠标放在垂直方向,然后是左边,然后是右边。当动物能够在该方向上静止5秒而不被尾巴握持时,确定成功完成。每个方向分配三次尝试(图13)。


        图13.倾斜平面测试

      3. 如果鼠标成功站在板上,将角度增加2.5°,并对每个方向重复步骤C2。继续增加角度,直到鼠标不再在任何方向站在板上。注意最大角度。
      4. 后TBI评分:受伤动物在低于最低最大基线值10°时开始测试。例如,如果鼠标在47.5°处向右方向,在50°处向左方向,在55°处垂直方向,则用于损伤后测试的起始角度将是37.5°(比右侧方向小10°)最大角度)。
      5. 通过将损伤后的值与基线最大角度进行比较,为每个方向分配得分。 然后将三个分数平均以得到最终分数。 使用以下参数对效果进行评分:
        1. 4(无损伤) - 没有差异或角度高于基线最大角度。
        2. 3 - 从基线最大角度减少2.5°。
        3. 2 - 从基线最大角度减少5°。
        4. 1 - 从基线最大角度减去7.5°。
        5. 0(严重受损) - 从基线最大角度减少10°以上。 

通过组合所有七个测试的分数产生复合神经分数。

致谢

这项研究由华盛顿大学麻醉和疼痛医学部和NIH SC1GM095426支持。 该方案改编自Dixon等人1991年描述的内容。

参考文献

  1. Bolkvadze,T。和Pitkanen,A。(2012)。 在受控皮层冲击和侧向流体冲击后发展创伤后癫痫,撞击诱导的小鼠的脑损伤。 J Neurotrauma 29(5):789-812。
  2. Dixon,C.E.,Clifton,G.L.,Lighthall,J.W.,Yaghmai,A.A。和Hayes,R.L。(1991)。 大鼠创伤性脑损伤的受控皮质影响模型。/a> J Neurosci Methods 39(3):253-262。
  3. Fujimoto,S.T.,Longhi,L.,Saatman,K.E.,Conte,V.,Stocchetti,N.and McIntosh,T.K。(2004)。 实验性创伤性脑损伤后的运动和认知功能评估。 Neurosci Biobehav Rev 28(4):365-378。 
  4. 这些研究结果表明,这些研究结果表明,该方法可以有效地降低患者的生存率,提高患者的生存质量。 )。 CX3CR1在轻度创伤性大脑的小鼠模型中的时间依赖性作用 J Neuroinflammation 12:154。 
  5. Smith,DH,Soares,HD,Pierce,JS,Perlman,KG,Saatman,KE,Meaney,DF,Dixon,CE和McIntosh, TK(1995)。 小鼠的旁瓣对照皮质影响模型:认知和组织病理学 J Neurotrauma 12(2):169-178。
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引用:Febinger, H. Y., Thomasy, H. E. and Gemma, C. (2016). A Controlled Cortical Impact Mouse Model for Mild Traumatic Brain Injury. Bio-protocol 6(16): e1901. DOI: 10.21769/BioProtoc.1901.
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