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Obtaining Acute Brain Slices

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May 2017



Obtaining acute brain slices for electrophysiology or amperometric recordings has become a routine procedure in most labs in the field of neuroscience. Yet, protocols describing the step by step process are scarce, in particular for routine acute preparations such as from the mouse hippocampus. Here we provide a detailed protocol for the dissection, extraction and acute slicing of the mouse brain, including tips and list of material required.

Keywords: Acute brain slices (急性脑切片), Hippocampus (海马), Dissection (解剖), Brain extraction (脑提取), Electrophysiology (电生理学), Mouse (小鼠)


With the democratization of in vitro electrophysiology and amperometry recording techniques, obtaining acute slices from rodent brains has become a classic and pivotal procedure in neuroscience research. Yet, the know-how required to achieving this procedure is typically passed on verbally, and most labs have developed home-made recipes and adapted the most important steps to their own needs or region of interest, such that there is a lack of protocols describing how to obtain high-quality acute brain slices in a step by step manner. While some protocols can be found describing particularly challenging preparations, such as acute slicing of adult mouse spinal cord (Garre et al., Bioprotocol 2017:, a description of the basic procedure for more routine preparations (e.g., hippocampal slices) is particularly lacking. Aside from the practical aspect, this is a major problem because extracellular field recordings from hippocampal slice have become widely employed, due to their relative simplicity and little equipment-requirement, including by labs without electrophysiology and acute brain slices preparation expertise. Given that the slices’ quality is the limiting factor to obtaining reliable electrophysiological recordings, this poses a major challenge for the reproducibility of results within and across labs. In light of these needs and caveats, we here provide a detailed protocol for the dissection, brain extraction and acute slicing of the mouse hippocampus, including tips and list of material required. It will allow beginner and non-experts to obtain acute hippocampal brain slices of the required quality for follow-up studies such as field recordings, patch-clamp recordings or amperometric recordings (Papouin et al., 2017).

Materials and Reagents


  1. For the ‘nest beaker’
    1. Nylon tights
    2. Instant superglue (such as Scotch Super Glue, 3M, catalog number: AD124 )
    3. 15 ml tubes (such as VWR, catalog number: 89039-670 US, 525-0450 Europe)
    4. Disposable 6 cm diameter plastic Petri dish (such as Thermo Fisher Scientific, Thermo ScientificTM, catalog number: 123TS1 )

  2. For dissection
    1. Large kitchen scissors or guillotine
    2. Straight fine scissors (such as Fine Science Tools, catalog number: 14060-11 )
    3. Curved spatula (such as Fine Science Tools, catalog number: 10092-12 )
    4. Scalpel (such as Fine Science Tools, catalog number: 91003-12 )
    5. Glass disposable Pasteur pipet (such as Fisher Scientific, FisherBrand, catalog number: 13-678-6A )
    6. Dropper bulb (such as Fisher Scientific, FisherBrand, catalog number: 03-448-25 )
    7. Plastic container, about 2.5 cm high and 150 ml, such as the lid of a pipet tip box or a large glass Petri dish (Cole-Parmer Instrument, catalog number: EW-34551-06 )
    8. Whatman paper (GE Healthcare, Whatman, catalog number: 1001-090 )
    9. Disposable Razor blade (such as Personna Double Edge Razor Blades [Amazon, PERSONNA, catalog number: BP9020 ])


  1. Glucose (Sigma-Aldrich, catalog number: G7021 )
  2. Sodium chloride (NaCl) (Sigma-Aldrich, catalog number: S7653 )
  3. Sodium phosphate monobasic anhydrous (VWR, catalog number: 470302-666 )
    Manufacturer: ALDON, catalog number: SS0756-500GR .
  4. Sodium bicarbonate (NaHCO3) (Sigma-Aldrich, catalog number: S5761 )
  5. Potassium chloride (KCl) (Sigma-Aldrich, catalog number: P9333 )
  6. Magnesium chloride solution (1 M) (Sigma-Aldrich, catalog number: 63069 )
  7. Calcium chloride solution (1 M) (Sigma-Aldrich, catalog number: 21115 )
  8. Stock artificial cerebrospinal fluid (ACSF) solution (see Recipes)
  9. Ice-cold Slicing ACSF solution (see Recipes)
  10. Recovery ACSF solution (see Recipes)
  11. Experimental ACSF (see Recipes)


  1. 250 ml Pyrex beaker (such as VWR, catalog number: 10754-952 )
  2. Straight spring scissors (such as Fine Science Tools, catalog number: 15018-10 )
  3. Curved fine forceps (such as Fine Science Tools, catalog number: 11152-10 )
  4. 600 ml Pyrex beaker (such as VWR, catalog number: 10754-956 )
  5. 95% O2/5% CO2 tank (such as AirGas, catalog number: Z02OX9522000043 )
  6. Vibratome (such as Leica, model: Leica VT 1200 S , catalog number: 14048142066)
  7. Bath heater (such as Thermo Fisher Scientific, Thermo Scientific, model: Precision 180 , catalog number: 51221073)


Before starting:

  1. Assemble the ‘nest beaker’ (Figure 1) and a modified Pasteur pipet dropper (Figure 2).
    1. ‘Nest beaker’ preparation (Figure 1)
      1. Using nylon tights, a 6 cm plastic Petri dish, a 15 ml tube, superglue and a 250 ml beaker, prepare a ‘nest beaker’ in which slices will be incubated during and after recovery. Using kitchen scissors (or a ‘Dremel’ if you have one) cut out or simply open the base of the Petri dish, preferably without breaking the wall.
      2. Stretch the nylon around the open Petri dish to form a firm mesh base, and secure it by tying it or by using elastic bands. Glue the nylon on the outside wall of the Petri dish. Do not use excessive amounts of glue as this can be toxic to slices and would prevent proper drying. Let dry for 24 h.
      3. Using a scalpel or fine scissors, cut out the excess nylon. Rinse abundantly and soak in clear water overnight (we had instances where ‘fresh’ glue revealed toxic to slices). Cut and discard the conical end of the 15 ml tube, and cut out a rectangular window near the bottom end.
      4. Assemble all three elements as shown. Plastic tubing from the 95% CO2/5% O2 tank will be lowered into the 15 ml tube to maintain appropriate pH and oxygenation.

        Figure 1. Nest beaker. Using nylon tights, a 6 cm plastic Petri dish, a 15 ml tube, superglue and a 250 ml beaker, prepare a ‘nest beaker’ in which slices will be incubated during and after recovery.

    2. Modified Pasteur pipette dropper (Figure 2)

      Figure 2. Modifier Pasteur pipette dropper. Break the thinnest end of a Glass disposable Pasteur pipet, and insert that end in a Dropper bulb.

  2. Prepare 1 L of ACSF the day prior and store overnight at 4 °C.

Setting up:

  1. On the day of the experiment, prepare 300 ml of ice-cold slicing ACSF (see Recipes) in a 600 ml Pyrex beaker (by adding 2 mM of MgCl2 and 1 mM of CaCl2 to 300 ml of stock ACSF) and place in a -20 °C freezer for about 20-30 min or until a thin layer of ice on the walls of the beaker and at the surface forms. Agitate vigorously to break the ice into a homogeneous icy solution. Avoid over-freezing as this will drastically change the osmolarity of the solution and reduce the quality of the slices. However, the amount of ice should be enough that the solution remains at 0-1 °C throughout the entire slicing procedure. Therefore it is recommended to adjust the iciness of the solution to your need/speed. We recommend against placing the beaker of slicing ACSF in a -80 °C freezer.
  2. While the ice-cold slicing ACSF is in the freezer, prepare 150 ml of recovery ACSF in the ‘nest beaker’ (by adding 1.5 mM of MgCl2 and 2 mM of CaCl2 to 150 ml of stock ACSF). Warm up in the heated bath at 33 °C while oxygenating with 95% O2/5% CO2 for at least 25 min before to start (Figure 3B).
  3. Prepare the vibratome by placing a mix of ice and water in the tray surrounding the slicing chamber. Cut a razor blade in half with the kitchen scissors (or use two blades): place one half in the blade holder of the vibratome and keep the other half for the dissection Procedure B.
  4. Familiarize yourself with the Procedures A to C below and prepare your tools accordingly, to optimize the process. Typically, this consists in placing the tools in the following order (right to left but adjust depending on your dominant hand): Large kitchen scissors, scalpel, fine scissors, fine forceps, plastic container or large glass Petri dish with a piece of Whatman paper at the bottom (this will help increasing visual contrast and providing greater surface grip) and the curved spatula nearby, ready-to-grab second half of the razor blade, vibratome cutting plate and tube of glue, modified Pasteur pipet dropper and spring scissors (Figure 3A).

    Figure 3. Set-up. A. The tools on the dissection and slicing bench are set up from right to left according to the sequence of steps described in this protocol. B. Close up view of the nest beaker filled with recovery ACSF bubbling with 95%O2/5%CO2 and incubating at 33 °C.
  1. Quickly extract the brain
    1. Pour about half (150 ml) of the ice-cold slicing ACSF into the plastic container or large glass Petri dish and oxygenate.
    2. Anesthetize the mouse using isoflurane, check for the absence of reflex upon tail or paw pinching and quickly decapitate the mouse using a small guillotine or large kitchen scissors (Figures 4A and 4B). Expose the skull with a large incision through the skin down the midline (Figure 4C) and cut the auditory conducts on each side (Figures 4D and 4E). Pull the skin toward the nose of the animal to fully expose the skull (Figure 4F). This will also provide a better grip of the head.
    3. Place the head in ice-cold oxygenated slicing ACSF. Leave it submerged for 10 sec to chill.
    4. While making sure the head remains submerged at all times, with fine scissors, open the back of the skull by making a cut immediately caudal to the cerebellum (Figures 4G and 4H), and then cut the skull open along the midline from the caudal end working your way up to the olfactory bulbs (Figures 4I-4L). Avoid putting pressure on the skull and make sure no damage is made to the brain underneath with the lower scissors tip. We also recommend making a lateral cut at the base of the skull through the jaw bones, this will help to extract the brain (Figure 4M).
    5. Using fine forceps grab the open edge of the skull on one side of the midline, hold firmly and open to the side while steadily holding down the head with the other hand (Figures 4N and 4O, ideally, your index and thumb should be on each side of the mouse ‘face’, roughly on the eyes. Having the skin tight under your fingers usually helps).
    6. Then proceed to the other side (Figures 4P and 4Q). Using a curved spatula, and being extremely gentle reach under the brain (let the floor of the skull guide you) and gently scoop out the brain, without pulling (Figures 4R-4T). The optic nerve, on the ventral part, and the cranial nerves, caudally, might need to be cut with the fine scissors or directly with the spatula to completely free the brain. Leave the extracted brain in the ice-cold slicing ACSF (Figure 4U). Make sure you keep the brain submerged in the ice-cold ACSF throughout this entire procedure.

      Figure 4. Extracting the brain from the skull. A. Decapitate the anesthetized animal; B-F. Using the scalpel, incise the skin, cut the auditory conducts and pull the skin to expose the skull. G-H. Open the back of the skull by making a cut immediately caudal to the cerebellum. I-L. Using fine scissors, carefully cut through the skull along the midline. Angle the scissors to minimize potential damage by the tip to the brain underneath. M. If animals used are adults, we recommend making a lateral cut at the base of the skull through the jaw bones. N-Q. Using fine tweezers, grab the open edge of the skull on one side, hold firmly and open to the side while firmly holding down the head by the ‘nose’ with the other hand. Then proceed to the other side. R-U. Using the curved spatula, carefully reach under the brain (let the floor of the skull guide you) and gently scoop out the brain. Cut through the optic chiasma with fine scissors or directly with the spatula (T). For clarity purpose, there is no ice and no bubbling in the solution in Panels G-U.

  2. Isolate the region of interest
    1. With the razor blade, remove the unwanted parts of the brain, rostral and caudal to the region of interest. In the case of the hippocampus: place the brain ventral side down, locate the superior colliculi, make a transverse cut and discard the caudal part (cerebellum, Figure 5A).
      Note: Make sure the cut is perpendicular to the rostro-caudal axis as this face will be glued on the cutting-plate of the vibratome.
    2. Then flip the brain ventral side up, locate the optic chiasma and make a transverse cut (Figures 5B and 5D). This should expose the fimbria of the fornix, which is immediately rostral to the hippocampus (i.e., the hippocampus lies under it). Spread just enough glue on the cutting plate (make sure the plate is dry beforehand, Figure 5E).
    3. Using the curved spatula and your index and thumb as an abutment (do not grab the brain with your fingers!), pick up the brain rostral side up and ventral side facing you (Figures 5F and 5G). Gently place the bottom of the spatula on a paper towel, to drain the excess of ACSF by capillarity (do not touch the brain with the paper towel). Place the spatula immediately above the glue (without touching it) and gently transfer the brain on the glue in a single motion by pushing it off the spatula with your finger (Figure 5H).
      Note: Being slow or hesitant will ‘stretch’ the brain and reduce the quality of the slices. Do not press down on the brain, tap the plate or wait and let the brain dry off, or this will dramatically reduce the quality of the slices as well.

      Figure 5. Isolate the region of interest. A. While the brain is dorsal side up, locate the inferior colliculi and, using a razor blade, make a cut immediately above them perpendicular to the rostro-caudal axis. Discard the part containing the cerebellum. B-C. Flip the brain ventral side up, locate the optic chiasma and using the razor blade, make a cut perpendicular to the rostro-caudal axis. Discard the frontal part. This should free the fimbria of the fornix, which lies immediately above the hippocampus. E-H. Apply just the required amount of glue on the vibratome plate. F. Using the curved spatula and your index and thumb as an abutment (do not grab the brain with your fingers!) pick up the brain. Use your finger to push the brain off the spatula immediately above the glue and delicately ‘drop’ in on the glue in a single motion. For clarity purpose, there is no ice and no bubbling in the solution in Panels A-D and F.

  3. Obtain brain slices
    1. Immediately transfer the plate into the slicing chamber (Figure 6A), with the ventral part of the brain facing you and the dorsal part (i.e., the surface of cortex) facing the back of the vibratome. Gently pour the rest of the ice-cold oxygenated slicing ACSF (Figure 6B).
    2. Lower the blade in the solution and, using the vibratome control panel, set up a fairly narrow yet safe ‘slicing window’ (~2 mm on each side). Lower the blade to the surface of the brain (do not press the blade on the brain) and start the slicing to obtain 300-350 μm hippocampal coronal slice (thickness should be pre-set).
      1. Avoid pouring the ACSF directly onto the brain and be careful not to drop pieces of ice onto the brain. Oxygenate while ensuring that the agitation caused by the bubbling is not excessive as this may be a problem once slices come unattached.
      2. To optimize the quality of the slices and minimize the total duration of the procedure, we recommend adjusting the speed of the vibratome to medium (0.12-0.16 mm/sec on Leica VT1200s) when the blade is not in any region of interest, and to low when the blade is in the hippocampus or region of interest (0.08-0.1 mm/sec on Leica VT1200s).
      3. Once the blade reaches the last ventral micrometers, the optic chiasma or meninges can resist and prevent full detachment of the slice. This can also distort or ‘pull’ the slice before it is entirely freed. In most cases, gently holding the slice onto the blade with the spring scissors without applying any pressure (Figure 6C) will suffice to help the blade cut through the chiasma or meninges. In extreme cases, we recommend quickly but very carefully sniping the meninges or the remaining part if the slice with spring scissors. In any case, be extremely careful not to push on the vibrating blade, on the brain underneath, or to pull the slice while still attached. Any sort of mechanical pressure (‘pulling’) will damage the slice. This could also cause the brain to come unglued.
    3. Once the first slice is freed (Figure 6D), with spring scissors, separate both hemispheres (Figure 6E) and, using the Pasteur pipet dropper, transfer them into the nest beaker containing the recovery ACSF (see Recipes), incubating at 33 °C in the bath heater (Figure 6F).
    4. Repeat until all slices are obtained and all hemi-slices are transferred in the nest beaker containing the recovery ACSF. Incubate at 33 °C for an additional 30 min.
    5. Carefully remove the nest beaker from the heating bath and let recover at room temperature for 45 min.
    6. Slices are now ready to be used for electrophysiology or other procedures such as Bio-protocol ‘D-serine measurement in brain slices or other tissue explants’ (Papouin and Haydon, 2018).

      Figure 6. Obtain brain slices. A. Transfer the plate with the glued brain in the slicing chamber of the vibratome and lower the blade holder. B. Poor the remaining of the ice-cold Slicing aCSF. C. If meninges or the optic chiasma are an issue during slicing, gently hold the slice onto the blade with the spring scissors (without applying any pressure) to help cut through. D. Once the slice is freed, separate the two hemispheres with the spring scissors (you can use the fine forceps to hold the bottom of the slice) and transfer to the nest beaker with Recovery ACSF incubating and bubbling at 33 °C. For clarity purpose, there is no ice and no bubbling in the solution in Panels B-E.

Data analysis

The quality of slices (notably hippocampal slices) can be very easily assessed visually with the 5x objective of any given microscope (Figure 7). Typically, healthy slices show stark contrasts and differential coloring across layers and regions. The stratum oriens and radiatum will have a bright orange color. The stratum lacunasorum molecular generally appears much darker (deep brown to deep grey). The pyramidal layer, while clear in comparison, will appear thin or ‘compact’ and, depending on the angle of the slicing, can be delineated from the s. oriens and radiatum by thin dark lines. Unhealthy slices take greyish and uniform tints. The pyramidal layer of an unhealthy slice appears exceedingly white or transparent and usually ‘swollen’.

Figure 7. A healthy hippocampal slice. Healthy, brain/hippocampal slices typically show orange coloring with obvious differences in tints and contrasts across layers. Layers are also evident in the cortex.
Note: The two sets of horizontal ‘strings’ are from the ‘harp’ system that holds the slice down under the microscope (not described in this protocol).


  1. From our experience, Steps A2 (once the mouse is decapitated) to C4 (when the last slice is extracted) should be achieved within 10 min for optimal brain slices quality. In particular, the complete extraction of the brain should be achieved in less than 2 min, i.e., no more than ~2-3 min should elapse between the decapitation of the animal and the transfer of the brain into the vibratome chamber.
  2. Please note that contrary to most of labs or protocols we strongly advice against using a sucrose-based slicing solution. We found that reducing calcium concentration while increasing that of magnesium and ensuring that the procedure is performed rapidly (see above) and in ice-cold ACSF throughout is the most efficient way to reduce excitotoxicity and the best guarantee of good slice quality.
  3. Please note that contrary to most of labs or protocols we also strongly advice against using a paint brush to manipulate slices. While they feel soft to the touch of one’s finger, at the scale of a 350 µm slice, paint brushes are the equivalent of many small knives bundle together and result in multiple stab wounds to slices.


  1. Stock artificial cerebrospinal fluid (ACSF) solution (1 L, store at 4 °C)
    Glucose 10 mM (1.8 g for 1 L)
    Potassium chloride 3.2 mM (0.23 g for 1 L)
    Sodium chloride 120 mM (7 g for 1 L)
    Sodium phosphate monobasic anhydrous 1 mM (0.119 g for 1 L)
    Sodium bicarbonate 26 mM (2.18 g for 1 L)
    Make up to 1 L with ddH2O
    Verify and adjust pH to 7.3 and osmolarity to 290-300 mOsm L-1
  2. Ice-cold slicing ACSF (~300 ml)
    Stock ACSF
    2 mM magnesium chloride
    1 mM calcium chloride
  3. Recovery ACSF (~150 ml)
    Stock ACSF
    1.5 mM magnesium chloride
    2 mM calcium chloride
  4. Experimental ACSF (~550 ml) for follow-up electrophysiological or amperometric recordings
    Stock ACSF
    1.3 mM magnesium chloride
    2 mM calcium chloride


This work was supported by two Philippe Foundation grants and a Human Frontier Science Program long-term fellowship (LT000010/2013) awarded to T.P., and two NIH/NINDS R01 grants (NS037585 and AA020183) awarded to P.G.H. who is also the founder of GliaCure. Authors declare no conflict of interest. We thank Jaclyn M. Dunphy for her careful proofreading of this protocol.


  1. Garré, J. M., Yang, G., Bukauskas, F. F. and Bennett, M. V. (2017). An acute mouse spinal cord slice preparation for studying glial activation ex vivo. Bio-protocol 7(2): e2102.
  2. Papouin, T., Dunphy, J. M., Tolman, M., Dineley, K. T. and Haydon, P. G. (2017). Septal cholinergic neuromodulation tunes the astrocyte-dependent gating of hippocampal NMDA receptors to wakefulness. Neuron 94(4): 840-854 e847.
  3. Papouin, T and Haydon, P. G. (2018). D-serine measurements in brain slices or other tissue explants. Bio-protocol 8(2): e2698.


在神经科学领域的大多数实验室中,获得用于电生理学或电流记录的急性脑切片已经成为常规手术。 然而,描述逐步过程的方案是稀缺的,特别是对于例如来自小鼠海马的常规急性制剂。 在这里,我们提供了一个详细的协议,解剖,提取和急性切片的小鼠大脑,包括技巧和所需材料清单。

【背景】随着体外电生理和电流记录技术的民主化,从啮齿动物大脑获取急性切片已经成为神经科学研究中经典和关键的步骤。然而,实现这一程序所需的技术诀窍通常是口头传递的,大多数实验室已经开发了自制的食谱,并根据自己的需要或感兴趣的区域调整了最重要的步骤,从而缺乏描述如何逐步获得高质量的急性脑切片。虽然可以发现一些方案描述特别具有挑战性的准备,例如成年小鼠脊髓的急性切片(Garre等人,Bioprotocol 2017: ),描述了更多日常准备的基本程序( ,海马片)特别缺乏。除了实际情况之外,这是一个主要的问题,因为海马切片的细胞外场记录由于其相对简单和设备要求少,包括没有电生理学的实验室和急性脑切片制备专业知识而被广泛使用。鉴于切片的质量是获得可靠的电生理记录的限制因素,这对于实验室内和实验室之间的结果的重复性提出了重大挑战。鉴于这些需求和注意事项,我们在这里提供了解剖,脑部提取和急性切片的小鼠海马的详细协议,包括提示和所需材料清单。这将允许初学者和非专家获得具有所需质量的急性海马脑切片,用于后续研究,例如野外记录,膜片钳记录或电流记录(Papouin等人,2017) 。

关键字:急性脑切片, 海马, 解剖, 脑提取, 电生理学, 小鼠



  1. 对于“窝烧杯”
    1. 尼龙紧身衣
    2. 即时超强胶(如苏格兰超级胶,3M,目录号:AD124)
    3. 15毫升管(如VWR,目录号:89039-670美国,525-0450欧洲)
    4. 一次性6厘米直径的塑料培养皿(如Thermo Fisher Scientific,Thermo Scientific TM,产品目录号:123TS1)

  2. 解剖
    1. 大厨房剪刀或断头台
    2. 直的罚款剪刀(如精细科学工具,目录号:14060-11)
    3. 弯曲的铲子(如Fine Science Tools,目录号:10092-12)
    4. 手术刀(如Fine Science Tools,目录号:91003-12)
    5. 玻璃一次性巴斯德吸管(如Fisher Scientific,FisherBrand,目录号:13-678-6A)
    6. 滴管灯泡(如Fisher Scientific,FisherBrand,目录编号:03-448-25)
    7. 塑料容器,高约2.5厘米,150毫升,如吸管尖端盒或大玻璃培养皿的盖子(Cole-Parmer Instrument,目录号:EW-34551-06)
    8. Whatman纸(GE Healthcare,Whatman,目录号:1001-090)
    9. 一次性剃刀刀片(如Personna双刃剃刀刀片[亚马逊,PERSONNA,目录号:BP9020])


  1. 葡萄糖(Sigma-Aldrich,目录号:G7021)
  2. 氯化钠(NaCl)(Sigma-Aldrich,目录号:S7653)
  3. 无水磷酸二氢钠(VWR,目录号:470302-666)
  4. 碳酸氢钠(NaHCO 3)(Sigma-Aldrich,目录号:S5761)
  5. 氯化钾(KCl)(Sigma-Aldrich,目录号:P9333)
  6. 氯化镁溶液(1M)(Sigma-Aldrich,目录号:63069)
  7. 氯化钙溶液(1M)(Sigma-Aldrich,目录号:21115)
  8. 股票人造脑脊液(ACSF)的解决方案(见食谱)
  9. 冰冷切片ACSF解决方案(见食谱)
  10. 恢复ACSF解决方案(见食谱)
  11. 实验ACSF(见食谱)


  1. 250毫升派热克斯烧杯(如VWR,目录号:10754-952)
  2. 直弹簧剪刀(如Fine Science Tools,目录编号:15018-10)
  3. 弯曲细镊子(如Fine Science Tools,目录号:11152-10)
  4. 600毫升派热克斯烧杯(如VWR,目录号:10754-956)
  5. 95%O 2/5%CO 2罐(例如AirGas,目录号:Z 0 OXX9522000043)。
  6. 颤音(如Leica,型号:Leica VT 1200 S,目录号:14048142066)
  7. 浴加热器(如Thermo Fisher Scientific,Thermo Scientific,型号:Precision 180,目录号:51221073)



  1. 组装'巢烧杯'(图1)和改进的巴斯德吸管滴管(图2)。
    1. “巢烧杯”的准备(图1)
      1. 使用尼龙紧身衣,一个6厘米的塑料培养皿,一个15毫升管,强力胶和一个250毫升的烧杯,准备一个'巢烧杯',其中切片将在恢复期间和之后孵化。使用厨房剪刀(如果有的话)或者简单地打开培养皿的底部,最好不要打破墙壁。
      2. 在开放的培养皿周围拉伸尼龙以形成牢固的网状基底,并通过捆绑或使用弹性带将其固定。在培养皿的外壁上粘上尼龙。不要使用过量的胶水,因为这可能对切片有毒,并会妨碍适当的干燥。让干24小时。
      3. 用手术刀或精细剪刀剪掉多余的尼龙。大量冲洗,并在清水中浸泡过夜(我们有“新鲜”胶水显示对切片有毒的情况)。切割并丢弃15毫升管的圆锥形末端,并在底端附近切出一个矩形窗口。
      4. 如图所示组装所有三个元素。将95%CO 2/5%O 2储罐中的塑料管放入15ml试管中以保持合适的pH值和充氧量。


    2. 改良的巴斯德吸管滴管(图2)

      图2.修饰符巴斯德吸管滴管。 打破玻璃一次性巴斯德吸管的最薄端,并将其插入滴管灯泡。

  2. 前一天准备1L ACSF,并在4°C过夜。


  1. 在实验当天,在600ml派热克斯(Pyrex)烧杯中(通过加入2mM MgCl 2和1mM CaCl 2)制备300ml冰冷切片ACSF(参见食谱)至300ml的ACSF储备液),并置于-20℃冰箱中约20-30分钟,或直到烧杯壁上和表面形成一薄层冰。大力搅拌,将冰块打成均匀的冰块。避免过度冻结,因为这将彻底改变溶液的渗透压,并降低切片的质量。但是,在整个切片过程中,冰量应足以使溶液保持在0-1℃。因此,建议根据您的需求/速度调整解决方案的冰冷度。我们建议不要将切片ACSF的烧杯放在-80°C的冷冻箱中。
  2. 在冷冻切片ACSF在冰箱中时,在“巢式烧杯”中制备150ml的回收ACSF(通过加入1.5mM的MgCl 2和2mM的CaCl 2 2) 150毫升ACSF)。在33℃的加热浴中预热至少25分钟,同时用95%O 2/5%CO 2氧化至少25分钟(图3B) br />
  3. 通过在切片室周围的托盘中放入冰和水的混合物来准备vibratome。用厨房剪刀(或使用两个刀片)将刀片切成两半:将一半放在vibratome的刀架中,另一半放在夹层程序B中。
  4. 熟悉下面的程序A到C,并相应地准备工具,以优化过程。通常情况下,这是按照以下顺序放置工具(从右到左,但根据你的优势手调整):大厨房剪刀,手术刀,细剪刀,细镊子,塑料容器或大玻璃培养皿和一张沃特曼纸在底部(这将有助于增加视觉对比度和提供更大的表面抓地力)和附近的弯曲的刮刀,准备抢下半年的刀片,vibratome切割板和胶管,改进的巴斯德吸管滴管和弹簧剪刀(图3A)。

    图3.设置。 :一种。解剖和切片台上的工具按照本协议所述步骤的顺序从右到左设置。 B.填充有95%O 2/5%CO 2的回收ACSF鼓泡并在33℃孵育的巢式烧杯的近视图。
  1. 快速提取大脑
    1. 将大约一半(150毫升)的冰冷切片ACSF倒入塑料容器或大玻璃培养皿和含氧化合物中。
    2. 使用异氟醚麻醉小鼠,检查尾巴或爪子捏反射的情况下,使用小型断头台或大型厨房剪刀快速断开鼠标(图4A和4B)。通过皮肤沿中线(图4C)的大切口暴露颅骨,切开每侧的听觉传导(图4D和4E)。将皮肤拉向动物的鼻子以完全暴露头骨(图4F)。这也将提供更好的头部抓地力。
    3. 将头放入冰冷的含氧切片ACSF中。让它淹没10秒冷却。
    4. 在确保头部始终处于水下的情况下,用精细的剪刀,通过立刻切开小脑的尾部(图4G和4H),打开颅骨的后部,然后沿着中线从尾部切开颅骨按照你的方式去嗅球(图4I-4L)。避免对头骨施加压力,并确保下面的剪刀尖不会对下面的大脑造成伤害。我们还建议通过颚骨在颅骨的底部做一个侧面切口,这将有助于提取大脑(图4M)。
    5. 用细镊子抓住中线一侧颅骨的开口边缘,牢固地握住并向侧面开放,同时用另一只手稳稳地压住头部(图4N和4O,理想的是,你的指标和拇指应该在每个老鼠脸上的一面,大致在眼睛上,手指紧贴皮肤通常是有帮助的)。
    6. 然后转到另一边(图4P和4Q)。使用弯曲的铲子,并且非常温和地到达大脑下面(让头骨的地板引导你),轻轻地舀出大脑,而不用拉动(图4R-4T)。视神经,腹部和颅神经,可能需要用精细的剪刀或直接用刮刀切割,以使脑完全释放。将提取的大脑留在冰冷切片ACSF中(图4U)。

      图4.从颅骨中提取大脑:一种。捣毁被麻醉的动物; B-F。使用手术刀,切开皮肤,切开听觉导管,拉动皮肤暴露头骨。 G-小时。打开后面的头骨通过切割立即尾小脑。我-L。用精细的剪刀沿着中线小心地切开头骨。将剪刀倾斜以最小化尖端对下方大脑的潜在损害。 M.如果使用的动物是成年人,我们建议在颅骨的底部通过颚骨进行侧切。 ñ-Q。用镊子抓住一侧头骨的开放边缘,用一只手牢固地握住头部,一边用鼻子牢牢按住头部。然后转到另一边。 R-Ú。使用弯曲的小铲,小心地到达大脑下面(让头骨的地板引导你),然后轻轻舀出大脑。用细剪刀或直接用抹刀(T)切开视交叉。为了清晰起见,在G-U面板的解决方案中没有冰块和起泡。

  2. 隔离感兴趣的区域
    1. 用剃刀刀片,去除感兴趣区域的大脑,嘴唇和尾部不需要的部分。在海马的情况下:将大脑腹侧向下放置,找到上丘,做一个横切和尾部(小脑,图5A)。
    2. 然后翻转大脑腹侧,找到视交叉,并做横切(图5B和5D)。这应该暴露穹窿的海马立即在海马(即,海马在其下面)的穹窿。在切割板上涂上足够的粘合剂(确保板预先干燥,图5E)。
    3. 使用弯曲的铲子和你的指标和拇指作为基台(不要用手指抓住大脑!),将大脑嘴侧朝上,腹侧朝向你(图5F和5G)。轻轻地将抹刀的底部放在纸巾上,通过毛细作用排出多余的ACSF(不要用纸巾擦大脑)。将刮刀放在胶水的正上方(不要接触它),用手指将其从刮刀上推下(图5H),轻轻地将胶囊上的大脑一次性移动到胶水上。

      图5.隔离感兴趣的区域。 :一种。当大脑背侧朝上时,找到下丘,并用剃刀刀片在垂直于尾骨轴的正上方进行切割。丢弃包含小脑的部分。公元前。翻转大脑腹侧,找到视交叉,并使用剃刀刀片,垂直于rostro-caudal轴进行切割。丢弃正面部分。这应该释放位于海马正上方的穹窿的伞状。 E-H。在vibratome板上涂上所需量的胶水。 F.使用弯曲的铲子和您的指标和拇指作为基台(不要用手指抓住大脑!)拿起大脑。用手指将大脑从胶水上方的刮刀上推下来,然后轻轻地将胶水“一滴一滴地”滴在胶水上。为了清楚起见,在A-D和F组的溶液中没有冰和不起泡。

  3. 获取大脑切片
    1. 立即将平板转移到切片室(图6A),使大脑的腹侧部分面向您,并将背部(即,皮层表面)朝向振动切片的背部。轻轻倒入其余的冰冷含氧切片ACSF(图6B)。
    2. 降低解决方案的刀片,并使用vibratome控制面板,建立一个相当狭窄但安全的“切片窗口”(每边约2毫米)。将刀片下降到大脑表面(不要按压大脑的刀片),开始切片,获得300-350微米的海马冠状切片(厚度应该是预先设定的)。
      1. 避免将ACSF直接倒在大脑上,注意不要把冰块掉到大脑上。在确保由起泡引起的搅动不会过度的同时进行充氧,因为一旦切片未连接,这可能是一个问题。
      2. 为了优化切片的质量并尽量减少整个手术的持续时间,我们建议当刀片不在任何感兴趣的区域时,将vibratome的速度调整到中等(在Leica VT1200上为0.12-0.16 mm / sec) ,当刀片处于海马或感兴趣的区域(Leica VT1200s为0.08-0.1毫米/秒)时为低。
      3. 一旦叶片到达最后的腹径微米,视交叉或脑膜可抵抗并防止切片完全分离。这也可以在切片被完全释放之前扭曲或“拉”切片。在大多数情况下,不用施加任何压力(图6C)用弹簧剪刀轻轻地将切片保持在刀片上就足以帮助刀片切开交叉或脑膜。在极端情况下,我们建议快速但非常小心地狙击脑膜或其余部分,如果用弹簧剪刀片。无论如何,要特别小心,不要推动振动刀片,在下面的大脑上,或者在连接时拉动切片。任何形式的机械压力(“拉动”)都会损坏切片。这也可能导致大脑脱胶。
    3. 一旦释放第一个切片(图6D),用弹簧剪刀分离两个半球(图6E),并使用巴斯德吸管滴管,将它们转移到含有回收ACSF的巢烧杯中(参见食谱),在33℃在浴缸加热器(图6F)。
    4. 重复,直到获得所有切片,并且所有半切片在包含恢复ACSF的巢式烧杯中转移。在33°C孵育另外30分钟。
    5. 小心地从加热浴中取出巢烧杯,并在室温下恢复45分钟。
    6. 切片现在准备用于电生理学或其他程序,如生物协议'脑切片或其他组织外植体中的D-丝氨酸测量'(Papouin and Haydon,2018)。

      图6.获取大脑切片。 :一种。将粘有大脑的板转移到vibratome的切片室中,然后放下刀架。 B.把剩下的冰块切成片状。 C.如果在切片过程中脑膜或视交叉是一个问题,用弹簧剪刀(不施加任何压力)将切片轻轻地夹在刀片上以帮助切断。 D.一旦切片被释放,用弹簧剪刀将两个半球分开(您可以用细钳子夹住切片的底部),并在33℃下用恢复ACSF孵育和鼓泡转移到巢烧杯中。为了清晰起见,在B-E面板的解决方案中没有冰块,也没有鼓泡。


用任何给定的显微镜的5倍物镜可以非常容易地评估切片(特别是海马切片)的质量(图7)。通常情况下,健康的切片显示形成鲜明的对比,跨层和区域的差异着色。 地层和 radiatum 将具有明亮的橙色。 lamusasorum分子层通常看起来更暗(深棕色到深灰色)。相比之下,金字塔层将显得薄或“紧凑”,并且根据切片的角度,可以从该图案划定。 oriens 和 radiatum 。不健康的切片采取灰色和统一的色彩。不健康的切片金字塔层显得非常白或透明,通常“肿”。



  1. 根据我们的经验,步骤A2(一旦鼠标被斩首)到C4(当最后一个切片被提取时)应该在10分钟内达到最佳脑切片质量。特别是,在2分钟内完成大脑的提取应该在动物断头和大脑转移之间不超过2-3分钟的时间内完成vibratome室。
  2. 请注意,与大多数实验室或协议相反,我们强烈建议不要使用基于蔗糖的切片溶液。我们发现,减少钙浓度,同时增加镁的含量,确保程序迅速执行(见上文)和冰冷ACSF是减少兴奋性毒性的最有效方法,也是切片质量最好的保证。 >
  3. 请注意,与大多数实验室或协议相反,我们强烈建议不要使用画笔来操作切片。虽然他们的手指摸起来很软,在350微米的切片尺度上,油漆刷相当于许多小刀捆在一起,导致多次刺伤切片。


  1. 股票人造脑脊液(ACSF)的解决方案(1升,存储在4°C)
    调节pH值至7.3,渗透压浓度为290-300 mOsm L-1
    用ddH <2:O>补足1L
  2. 冰冷切片ACSF(约300毫升)
    2 mM氯化镁
    1 mM氯化钙
  3. 恢复ACSF(〜150毫升)
    1.5 mM氯化镁
    2 mM氯化钙
  4. 用于后续电生理或电流记录的实验性ACSF(约550 ml)
    1.3 mM氯化镁
    2 mM氯化钙


这项工作得到了两项菲利普基金会的资助以及授予T.P.的人类前沿科学计划长期奖学金(LT000010 / 2013)和两项授予P.G.H.的NIH / NINDS R01奖学金(NS037585和AA020183)的支持。谁也是GliaCure的创始人。作者声明不存在利益冲突。我们感谢Jaclyn M. Dunphy对本协议的仔细校对。


  1. Garré,J.M.,Yang,G.,Bukauskas,F.F。和Bennett,M.V。(2017)。 用于研究神经胶质激活的急性小鼠脊髓切片准备 ex vivo 。 Bio-protocol 7(2):e2102。
  2. Papouin,T.,Dunphy,J.M.,Tolman,M.,Dineley,K.T。和Haydon,P.G.(2017)。 中隔胆碱能神经调节调节海马NMDA受体的星形胶质细胞依赖性门控以调整清醒状态 < Neuron 94(4):840-854 e847。
  3. Papouin,T和Haydon,P.G。(2018)。 脑切片或其他组织外植体的D-丝氨酸测量 Bio-protocol 8(2):e2698。
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Copyright: © 2018 The Authors; exclusive licensee Bio-protocol LLC.
引用:Papouin, T. and Haydon, P. G. (2018). Obtaining Acute Brain Slices. Bio-protocol 8(2): e2699. DOI: 10.21769/BioProtoc.2699.