铜银染色.doc

铜银染色操作步骤1. 试剂：水合氯醛，多聚甲醛，二甲砷酸，蔗糖，氯化钠，右旋葡萄糖，无水氯化钙，明胶，丙酮，浓盐酸，硝酸银，硝酸铜，尿囊素，硼酸，四硼酸钠，吡啶，无水乙醇，氢氧化钠，氢氧化铵，37%甲醛，柠檬酸，铁氰化钾，硫代硫酸钠，去离子水，多聚赖氨酸，中性树胶，二甲苯2. 切片：20 um3. 溶液配制2.1 1%硝酸铜溶液硝酸铜 1 g去离子水 100 mL充分混均2.2 0.1%尿囊素尿囊素 0.1 g去离子水 100 mL 充分混均2.3 0.4%氢氧化钠氢氧化钠 0.4 g去离子水 100 mL 充分混均2.5 硼酸缓冲液（光敏性）试剂A：0.2M硼酸硼酸 1.236 g去离子水 100 mL充分混均试剂B：0.05M硼酸钠硼酸钠 1.908 g去离子水 100 mL往试剂A用加试剂B直到pH至8.52.6 铜银溶液： 去离子水 120 mL硝酸银 1.05 g1%硝酸铜 1 mL0.1%尿囊素 10 mL吡啶 6 mL无水乙醇 12 mL硼酸缓冲液 6 mL每次使用前于通风橱下新鲜配制，在光线较暗处进行配制。2.7 银氨溶液去离子水 60 mL硝酸银 12 g0.4%氢氧化钠 30 ml氢氧化铵 15 mL去离子水 60 ml每次使用于通风橱下新鲜配制，较暗出进行。2.8 还原剂溶液去离子水 135 mL无水乙醇 15 mL或95%乙醇 16 mL37%甲醛 180 uL无水柠檬酸(精确称量) 10.5 mg或柠檬酸(精确称量) 11.5 mg2.9 铁氰化钾漂白溶液：铁氰化钾 450 mg去离子水 150 mL充分混均。新鲜配制使用2.10 硫代硫酸钠溶液：硫代硫酸钠 150 mg 去离子水 150 mL充分混均。新鲜配制使用4. 染色步骤4.1 用镊子从多聚甲醛溶液中取一张含脑组织或脊髓组织切片的载玻片，置于含去离子水的染色缸内，浸泡1-5 min。每隔30 s摇晃一次。然后用镊子取出载玻片，擦干载玻片上多余的液体。连续5次。4.2 用镊子将载玻片置于铜银溶液中，在40条件下，于摇床中慢摇（54转/min）45-50 min。用镊子取出载玻片后，擦干载玻片上多余的液体。4.3 于丙酮溶液中浸泡1min。用镊子立即取出载玻片。4.4 置于氨银溶液中，在室温下，于摇床上慢摇（54转/min）28-32 min，用镊子取出载玻片后，迅速擦干载玻片上多余的液体。4.5 快速置于还原液中，于室温下，浸泡45 s（注意观察切片颜色，如果颜色较浅，可适当延长时间），取出后擦干载玻片上多余的液体。4.6 用镊子将载玻片置于漂白液中，于摇床上，慢摇3-5 min。取出后擦干载玻片上多余的液体。4.7 用镊子将载玻片置于去离子水中，浸泡1 min。取出后擦干载玻片上多余的液体。4.8 用镊子将载玻片置于去离子水中，浸泡1 min。取出后擦干载玻片上多余的液体。4.9 用镊子将载玻片置于去离子水中，浸泡1 min。取出后擦干载玻片上多余的液体。4.10 用镊子将载玻片置于硫代硫酸钠溶液中，浸泡1 min。取出后擦干载玻片上多余的液体。4.11 用镊子将载玻片置于去离子水中，浸泡1 min。取出后擦干载玻片上多余的液体。4.12 用镊子将载玻片置于去离子水中，浸泡1 min。取出后擦干载玻片上多余的液体。4.13 用镊子将载玻片置于去离子水中，浸泡1 min。取出后擦干载玻片上多余的液体。4.14 用镊子取出载玻片，于50%乙醇中浸泡5 min。4.15 用镊子取出载玻片，于70%乙醇中浸泡5 min。4.16 用镊子取出载玻片，于95%乙醇中浸泡5 min。4.17 用镊子取出载玻片，于95%乙醇中浸泡5 min。4.18 用镊子取出载玻片，于100%乙醇中浸泡5 min。4.19 用镊子取出载玻片，于100%乙醇中浸泡5 min。4.20 用镊子取出载玻片，于二甲苯溶液中浸泡5 min。4.21 用镊子取出载玻片，于二甲苯溶液中浸泡5 min。4.22 用镊子取出载玻片后，于切片左侧滴1-2滴中性树胶，然后盖玻片沿45方向慢慢的将切片封盖。4.23 用记号笔标记载玻片，于显微镜下观察病理变化。结果：溃变神经元及其轴突：黑色，正常组织：金黄色备注：所有溶剂都应该在干净的烧杯中，用搅拌棒搅拌，然后在干净的玻璃瓶中储存。采用50%的硝酸对所有玻璃器皿进行清洗（在通风橱下进行），然后迅速用蒸馏水冲洗，最后用去离子水冲洗。对每一步操作进行精确测量。储存液可以在室温下于化学贮藏柜中保存至一年。Amino-Cupric-Silver TechniqueReagents: cupric nitrate, Allantoin, Sodium Hydroxide, Ammonium hydroxide, boric acid, sodium tetraborate, Sodium thiosulfate, potassium Ferricyanide, Anhydrous citric acid, Formaldehyde, Pyridine, AgNO3.Section：20umSolution：1. 1%CuNO3 (light sensitive).cupric nitrate 1g d H20 100mL Mixture well2. 0.1% Allantoin.Allantoin 0.1g d H20 100 mlMixture well3. 0.4% Sodium Hydroxide （NaOH）.Sodium Hydroxide 0.4gd H2O 100 ml Mixture well4Borate buffer (light sensitive):Solution A: 0.2 m boric acidboric acid 1.236 g d H20 100 mL Mixture wellSolution B: 0.05M sodium tetraboratesodium tetraborate 1.908g dH20 100mL add sol. B to sol. A until ph = 8.5 5. Cupric-silver solution: Deionized water 120mLAgNO3 1.05g1% CuNO3 1mL0.1% Allantoin 10mLPyridine 6mLAbsolute ethanol 12mLBorate buffer 6mLMake fresh in the hood for each use, with the hood lights off, as it is light sensitive.6. Silver diamine incubation solution:Deionized water 60mLAgNO3 12 g0.4% NaOH 30mLNH4OH 15mLDeionized water (total 120 mL) 60mLMake fresh for each use, in the hood with the hood lights off.7. Reducing agent solution:Deionized water 135mL15mL Absolute ethanol (see Note 3)or 95% ethanol 16mL37% w/w Formaldehyde 180uLAnhydrous citric acid 10.5 mg(or citric acid if it is monohydrous) 11.5 mgMixture well. Make fresh for each use.8. 0.3% K3Fe（CN）6:K3Fe（CN）6 450 mgdistilled water 150mLMixture well.Make fresh for each use.9. Thiosulfate solution:Sodium thiosulfate 150 mgdistilled water 150mLMixture well. Make fresh for each use.Staining1. Take the slides with brain or spinal cord sections from 4%paraformaldehyde solution.2. Wash the sections five times with deionized water for 15 min each, by transfer through five successive dishes, shaking occasionally. Blot the tray well on paper towels (but be careful not to dry it) before the next step.3. Place the slide in cupric-silver solution, at 40for 1 h in oven, then remove it from the oven and leave it in the hood at room temperature overnight.4. The next day, blot the slide well on paper towels (but again, be careful not to dry it).5. Dip the slide in a 100% acetone for 45 s 6. Quickly transfer the slide into silver diamine incubation solution and incubate for 35 min. 7. Remove the slide from the silver diamine incubation solution and blot it very well with paper towels (again, be careful not to dry it).8. Transfer the staining tray to a dish of reducing agent solution for 14 min 9. Transfer the slide to ferricyanide bleaching solution and incubate for approximately 1 min with continuous gentle agitation. Watch this step very carefully. The tissue should end up a dark straw color. If the tissue remains too dark, it can be left in this solution for up to 30 min. Blot well.10. Transfer the slide through three distilled water washes for three times, spending 1 min in each. Blot well between each transfer.11. Transfer to a stabilizing solution composed of 0.1% sodium thiosulfate for 1 min. Blot well.12. Again Transfer the slide through three distilled water washes for three times, spending 1 min in each. Blot well between each transfer.13. Dehydrate through 50% alcohol for 5 minutes each.14. Dehydrate through 70% alcohol for 5 minutes each.15. 2 changes of absolute alcohol for 5 minutes each.16. Clear in 2 changes of xylene for 5 minutes each.17. Mark and examine slides under bright-field microscopyResult : degenerating neron:black. Back ground：gold-yellowNotes1. Many of the solutions are toxic, volatile, corrosive, and/or carcinogenic. Use appropriate precautions for safety, storage, and disposal at all stages. Do not remove the preincubation cupric-silver solution from the hood; it contains pyridine (a carcinogen). Store the silver waste solutions and potassium ferricyanide solution in disposable containers. The glassware, stir bars, thermometer, and hemostat should be cleaned by soaking in 50% nitric acid for at least 1hr, followed by rinsing with distilled then dionized water.2. All the measurements should be very precise. If this solution does not clear completely, it may mean that the ammonium hydroxide is not fresh (the most common reason) or that the water or dishes are not clean.3. Use 15mL alcohol only if the bottle has been opened for the first time, otherwise use 16 mL.4. From this point in the procedure on, deionized water does not need to be used; ordinary distilled water is adequate.5. The best fixative for de Olmos staining is 4% paraformaldehyde in 0.1M cacodylate buffer, pH 7.4; this fixative is also perfectly fine for light microscopy and immunocytochemistry. Alternatively, 4% paraformaldehyde in 0.1M Tris buffer, pH 7.4, is also effective and less toxic than cacodylate buffer. Other tested buffers resulted in unacceptably high backgrounds.6. If the sections are overcrowded, the quality of the staining is not goodespecially, the bleaching step is perturbed.7. The sections can be distributed horizontally or vertically, so every row or every column corresponds to sections from one brain. Placement of the sections in rows allows processing of 12 sections from six brains at the same time.8. You should watch the color of the sections. The color should be dark brown velvet. Sometimes, the color does not correspond to the time.9. This is the most critical step referring to the background of the staining. Watch the color of the sections and continuously agitate the tray. The bleaching of young tissue generally proceeds much faster (for neonatal mouse brains, 2 min).10. Sections from experimental brains should be compared with age-matched control brains that are processed in parallel.Note：All solutions should