Summary

斑马鱼<em>原位</em>脊髓制备用于脊髓感觉和运动神经元的电生理记录

Published: April 18, 2017
doi:

Summary

这手稿描述了从斑马鱼胚胎和幼虫的脊髓神经元的电生理记录的方法。制备维持神经元在原地 ,往往涉及到最小的清扫。这些方法允许对多种脊髓神经元的电生理学研究中,从通过早期幼体阶段的初始电兴奋采集。

Abstract

斑马鱼,首先介绍了作为一个发展模式,在许多领域都得到了普及。饲养迅速发展生物体的大数自如,与胚胎光学透明度结合,担任该模型的初始吸引力属性。在过去的二十年里,这种模式的成功得到了进一步的顺从大规模诱变筛选,并通过易于转基因的推动。最近,基因编辑方法扩展了模型的力量。

对于神经发育研究,斑马鱼胚胎和幼虫提供到可应用多种方法的模型。在这里,我们专注于让神经元,电兴奋性的本质属性的研究方法。我们对斑马鱼的脊髓神经元的电生理研究编制涉及使用兽医缝合的胶水,以确保准备录音室。记录替代方法从斑马鱼胚胎和幼虫涉及使用细钨针1,2,3,4,5的制备腔室的连接。钨销是最经常用于安装在横向取向的制备中,虽然它已被用于安装幼虫背的一面朝上4。缝合胶水已被用于安装在两个方向的胚胎和幼虫。使用胶水,一个最小的夹层可以执行,允许访问脊髓神经元而不使用酶处理的,从而避免了任何所得的损坏。然而,对于幼虫,有必要施加一个简短酶处理以除去脊髓周围的肌肉组织。这里所描述的方法已被用来在几个developmenta研究运动神经元的interneurons和感觉神经元的内在电性能升级6,7,8,9。

Introduction

乔治·斯特雷伊辛格率先使用斑马鱼 ,俗称斑马鱼作为脊椎动物发育10的遗传分析模型系统。该模型提供了几个优点,包括:(1)相对简单和便宜畜牧业; (2)外部受精,从而允许方便地从最早的发育阶段的胚胎;和(3)的透明胚胎,从而允许细胞,组织和器官的直接和重复观察,因为它们形成。

在随后的几十年里,进行了若干改进进一步增加了斑马鱼模型的力量。特别是,正向遗传学屏幕和全基因组测序工作发挥了重要作用的基因突变和基因关键的标识许多发育过程11,12,13,14,“> 15,16。Gateway克隆方法已经允许转基因的常规应用方法17,18。在基因组编辑的最新进展,通过转录激活状(TALEN的)中列举和群集规则间隔开的短回文重复序列(CRISPR)-Cas9核酸酶,允许有针对性地引入突变,以及敲除和敲入接近19,20,21,22,组合,这些方法使斑马鱼为底层的具体行为和一些人类疾病的遗传机制的研究提供强大的模型23,24,25,26,27。

今年工作重点放在发展心理调节和电活动的神经发育中的作用。重点是脊髓,为此,斑马鱼模型提供了几个优点。首先,它是相对容易的,在胚胎和幼体阶段访问斑马鱼;因此,可以在具有较少的神经元和较简单的电路28,29的发育阶段研究脊髓功能。此外,斑马鱼脊髓具有一组不同的神经元,类似于其他脊椎动物中,因子30,31,32,33,34,35所证明由转录的特性和特征图案。

旨在揭示背后脊髓电路的功能的机制的多数在斑马鱼的研究中,尤其是支持运动的,是可以理解集中于幼虫阶段36,37,38,39,40,41,42,43。然而,许多形成脊柱机车网络中的神经元的启动它们的分化在早期胚胎阶段,〜9-10小时后受精(HPF)44,45,46,47,48,49,50,51。鉴于此,了解如何脊髓神经元的形态学和电学性质发生和胚胎和幼虫阶段之间的变化是一个重要overa的运动电路形成和功能LL理解。

这里描述的解剖方法允许从脊髓神经元膜片钳记录,并在胚胎阶段(〜17-48 HPF)和幼体阶段(〜3-7天受精后[DPF])得到成功应用。该方法限制了解剖,以提供对感兴趣的神经元所需要的量。所述协议从多数的其它公开的方法的不同之从在兽医缝合胶斑马鱼脊髓神经元的记录使用的,而不是一个细钨针,胚胎或幼虫附着到记录室。的两种不同的方法的可用性( 即,缝合胶相对于钨针)用于安装所述斑马鱼的胚胎或幼虫用于电生理分析为研究人员提供替代方案,以实现其特定的实验目标。

首先,从弹出访问和记录过程初级感觉神经元的ulation,Rohon-胡子细胞,进行了描述。这些神经元的细胞体位于背侧脊髓内。 Rohon-胡子细胞存在于许多脊椎动物物种,在开发早期分化,和背后胚胎触摸响应6,44,47,48。

第二,用于访问和从脊髓运动神经元的记录程序的详细。期间神经发生的两个波斑马鱼脊髓运动神经元引起的。在出生较早初级运动神经元出现在原肠胚形成(〜9-16 HPF)的端部,只有3-4本每hemisegment 45,46,49初级运动神经元。相反,次级运动神经元的后出生人口是比较多,而且在长时间内产生,起始于14〜HPFEF“> 45,50。在中间躯干段二级运动神经元起源主要是由51 HPF 50完成。二级运动神经元都被认为是运动神经元的羊膜对方46。有趣的是,脊椎神经元,经由多巴胺,调节运动在胚胎的幼虫和次级运动神经元起源和年轻幼虫50,51。初级和次级的运动神经元的每一个包括若干不同的亚型。每个初级运动神经元亚型的项目的外围轴突其支配的特性的肌肉群,从而导致刻板,识别轴突轨迹。通常,次级运动神经元遵循先前由初级运动神经元建立的轴索途径。因此,相对于轴突轨迹,原发性和继发性运动神经元是相似的,不同之处在于厚度轴突胞体和尺寸的重新更大初级运动神经元45。

三,讨论从几个类型的interneurons的记录方法。然而,在这些情况下,需要去除其它脊髓细胞的有限量的,因此,脊髓比从Rohon-胡子细胞或运动神经元的录音少完好无损。

Protocol

(;实验动物资源处安舒茨科罗拉多大学医学校园IACUC)所有动物的程序是由机构动物护理和使用委员会的批准。 1.斑马鱼的饲养提高,并在28.5℃下在10小时黑暗/ 14小时光照循环,并用适当的水处理和交换52保持成年斑马鱼鱼(Danio rerio)。 提高斑马鱼胚胎/幼虫在28.5℃下在胚胎培养基,直到它们达到所要求的阶段( 例如,2 DPF)?…

Representative Results

( 图5A和5B),我们已经成功地从Rohon胡子神经元记录在17个HPF胚胎到7 DPF幼虫。当Rohon-胡子细胞记录,将制备物固定背的一面朝上。这种安装允许基于它们的表面背的位置和大胞体大小Rohon-胡子细胞的明确识别。识别另外由这些神经元的超极化的定型静息膜电位( 图5, 小图表 )6,54</su…

Discussion

这里描述的方法允许脊髓的最小解剖后斑马鱼胚胎的感觉和运动神经元的电和形态学特征。神经元至少1小时,强加给这些录音时间限制保持健康。神经元已经使用标准的全细胞构型记录,以及从核贴剂;后者的方法最小化,可以排除离子电流9的详细研究生物物理空间钳的问题。

一个重要的挑战是实现胚胎或幼虫室的坚定执着,以去除皮肤和执行提供访问感?…

Declarações

The authors have nothing to disclose.

Acknowledgements

这项工作是由来自美国国立卫生研究院(F32 NS059120到RLM和R01NS25217和P30NS048154到ABR)的资助。

Materials

Vacuum filter/Storage bottle, 0.22 mm pore Corning 431096
Syringe filter 0.2 mm Whatman 6780-2502
Tricaine Sigma A-5040 Ethyl 3-aminobenzoate methanesulfonate salt 
a-bugarotoxin Tocris 11032-79-4
Tetrodotoxin Tocris 4368-28-9
Alexa-549 hydrazine salt Molecular Probes A-10438 fluorescent dye
Spin-X centrifuge tube filter Corning 8161
Glass microscope slide Fisher 12-550C
Sylgard silicone elastomer kit Dow Corning 184 silicone elastomer
Petri dishes Falcon 351029
Borosilicate glass capillaries Harvard Apparatus 30-0038 inner and outer diameters of 0.78 and 1.0 mm (thin walled glass capillaries)
Borosilicate glass capillaries Drummond Scientific 1-000-1000-100 inner and outer diameters of 1.13 and 1.55 mm (thick walled glass capillaries)
Miniature barbed polypropylene fitting  Cole-Palmer 6365-90
Vetbond tissue adhesive  3M 1469SB
Collagenase XI Sigma C7657
Microelectrode puller Sutter Instruments  Model P-97 
Amplifier Molecular Devices Axopatch 200B
Head stage Molecular Devices CV203BU
Motorized micromanipulator   Sutter Instruments MP-285
Tygon tubing Fisher 14-169-1B ID 1/16 IN, OD 1/8 IN and WALL 1/32 IN (flexible laboratory tubing)
Electrode holder Molecular Devices 1-HC-U
Pharmaseal Three-Way Stopcocks  Baxter K75
Digitizer Axon Instruments Digidata 1440A
Inverted microscope  Zeiss Axioskop2 FS plus
40x/0.80w Achroplan objective Zeiss
Data acquisition and analysis software  Axon Instruments PClamp 10 – Clampex and Clampfit 
Micropipette puller Sutter Instruments Model P-97
Name Company Catalog Number Comments
Dissection and Recording Solutions(in mM)
All solutions, except the intracellular, are stable for ~ 2 – 3 months when filtered (0.22 mm filter cups) and stored at room temperature (RT).
The intracellular solution is filtered (0.2 mm syringe filters) and stored frozen (-20°C) in small aliquots that are individually thawed on the day of use. 
Dissection/Ringer’s solution 145 NaCl, 3 KCl, 1.8 CaCl2.2H2O, 10 HEPES; pH 7.4 (with NaOH)
Pipette (intracellular) recording solution 135 KCl, 10 EGTA-acid, 10 HEPES; pH 7.4 (with KOH).
Bath (extracellular) recording solution/voltage and current-clamp 125 NaCl, 2 KCl, 10 CaCl2.2H2O, 5 HEPES; pH 7.4 (with NaOH).
Alexa-594 hydrazine salt stock solution.  Prepare a 13.2 mM stock in ddH2O, aliquot (~ 100 µl) and store at -20°C. For use, dilute the stock solutiond 132 fold with pipette solution to a final concentration of 100 mM. After dilution, filter the Alexa-594 containing pipette solution  with a centrifuge tube filter.
Name Company Catalog Number Comments
Immobilizing agents
0.4 % ethyl 3-aminobenzoate methanesulfonate salt (Tricaine) Prepare a 0.4% stock solution in 0.2M Tris, pH9 (0.4 g Tricaine/100 ml 0.2 M Tris
Adjust pH to 7 with NaOH and store at -20°C.
For use, dilute the stock solution ~ 25 fold in embryo media
250 μM α-bungarotoxin Prepare a 250 μM stock in ddH2O (1 mg/500 μl), prepare 100 µl aliquots, and sotre at -20°C.
For use, dilute 2,500-fold with extracellular solution to a final concentration of 100 nM.
1 mM Tetrodotoxin Prepare a 1 mM stock in ddH2O (1 mg/3 ml), prepare 100 µl aliquots, and sotre at -20°C.
For use, dilute 2,000-fold with extracellular solution to a final concentration of 500 nM.

Referências

  1. Drapeau, P., Ali, D. W., Buss, R. R., Saint-Amant, L. In vivo recording from identifiable neurons of the locomotor network in the developing zebrafish. J Neurosci Methods. 88 (1), 1-13 (1999).
  2. Drapeau, P., Saint-Amant, L., Buss, R. R., Chong, M., McDearmid, J. R., Brustein, E. Development of the locomotor network in zebrafish. Prog Neurobiol. 68 (2), 85-111 (2002).
  3. Saint-Amant, L., Drapeau, P. Whole cell patch-clamp recordings from identified spinal neurons in the zebrafish embryo. Methods Cell Sci. 25, 59-64 (2003).
  4. Masino, M. A., Fetcho, J. R. Fictive Swimming Motor Patterns in Wild Type and Mutant Larval Zebrafish. J Neurophysiol. 93 (6), 3177-3188 (2005).
  5. Wen, H., Brehm, P. Paired motor neuron-muscle recordings in zebrafish test the receptor blockade model for shaping synaptic current. J Neurosci. 25 (35), 8104-8111 (2005).
  6. Ribera, A. B., Nüsslein-Volhard, C. Zebrafish Touch-Insensitive Mutants Reveal an Essential Role for the Developmental Regulation of Sodium Current. J Neurosci. 18, 9181-9191 (1998).
  7. Pineda, R. H., Heiser, R. A., Ribera, A. B. Developmental, molecular, and genetic dissection of INa in vivo in embryonic zebrafish sensory neurons. J Neurophysiol. 93, 3582-3593 (2005).
  8. Moreno, R. L., Ribera, A. B. Zebrafish motor neuron subtypes differ electrically prior to axonal outgrowth. J Neurophysiol. 102, 2477-2484 (2009).
  9. Moreno, R. L., Ribera, A. B. Spinal neurons require Islet1 for subtype-specific differentiation of electrical excitability. Neural Dev. 9 (1), 19 (2014).
  10. Streisinger, G., Walker, C., Dower, N., Knauber, D., Singer, F. Production of clones of homozygous diploid zebra fish (Brachydanio rerio). Nature. 291 (5813), 293-296 (1981).
  11. Driever, W., et al. A genetic screen for mutations affecting embryogenesis in zebrafish. Development. 123, 37-46 (1996).
  12. Haffter, P., et al. The identification of genes with unique and essential functions in the development of the zebrafish, Danio rerio. Development. 123, 1-36 (1996).
  13. Vogel, G. Genomics: Sanger will sequence zebrafish genome. Science. 290 (5497), 1671 (2000).
  14. Patton, E. E., Zon, L. I. The art and design of genetic screens: zebrafish. Nature Reviews Genetics. 2 (12), 956-966 (2001).
  15. Lawson, N. D., Wolfe, S. A. Forward and reverse genetic approaches for the analysis of vertebrate development in the zebrafish. Dev Cell. 21 (1), 48-64 (2011).
  16. Yates, A., et al. Ensembl. Nucleic Acids Res. 44 (D1), D710-D716 (2016).
  17. Kawakami, K. Transgenesis and gene trap methods in zebrafish by using the Tol2 transposable element. Methods Cell Biol. 77, 201-222 (2004).
  18. Kwan, K. M., et al. The Tol2kit: a multisite gateway-based construction kit for Tol2 transposon transgenesis constructs. Dev Dyn. 236 (11), 3088-3099 (2007).
  19. Huang, P., Xiao, A., Zhou, M., Zhu, Z., Lin, S., Zhang, B. Heritable gene targeting in zebrafish using customized TALENs. Nat Biotechnol. 29, 699-700 (2011).
  20. Sander, J. D., et al. Targeted gene disruption in somatic zebrafish cells using engineered TALENs. Nat Biotechnol. 29 (8), 697-698 (2011).
  21. Chang, N., et al. Genome editing with RNA-guided Cas9 nuclease in zebrafish embryos. Cell Res. 23 (4), 465-472 (2013).
  22. Hwang, W. Y., et al. Efficient genome editing in zebrafish using a CRISPR-Cas system. Nat Biotechnol. 31 (3), 227-229 (2013).
  23. Lieschke, G. J., Currie, P. D. Animal models of human disease: zebrafish swim into view. Nature Rev Genet. 8 (5), 353-367 (2007).
  24. Levin, E. D., Cerutti, D. T., Buccafusco, J. J. Behavioral neuroscience of zebrafish. Methods of behavior analysis in neuroscience. , (2009).
  25. Stewart, A., Gaikwad, S., Kyzar, E., Green, J., Roth, A., Kalueff, A. Modeling anxiety using adult zebrafish: A conceptual review. Neuropharmacology. 62, 135-143 (2012).
  26. Mushtaq, M. Y., Verpoorte, R., Kim, H. K. Zebrafish as a model for systems biology. Biotechnol Genet Eng Rev. 29 (2), 187-205 (2013).
  27. Phillips, J. B., Westerfield, M. Zebrafish models in translational research: tipping the scales toward advancements in human health. Dis Model Mech. 7 (7), 739-743 (2014).
  28. Weis, J. S. Analysis of the development of nervous system of the zebrafish, Brachydanio rerio. I. The normal morphology and development of the spinal cord and ganglia of the zebrafish. J Embryol Exp Morphol. 19 (2), 109-119 (1968).
  29. Bernhardt, R. R., Chitnis, A. B., Lindamer, L., Kuwada, J. Y. Identification of spinal neurons in the embryonic and larval zebrafish. J Comp Neurol. 302 (3), 603-616 (1990).
  30. Tsuchida, T., et al. Topographic organization of embryonic motor neurons defined by expression of LIM homeobox genes. Cell. 79 (6), 957-970 (1994).
  31. Guillemot, F. Spatial and temporal specification of neural fates by transcription factor codes. Development. 134, 3771-3780 (2007).
  32. Goulding, M. Circuits controlling vertebrate locomotion: Moving in a new direction. Nat Rev Neurosci. 10, 507-518 (2009).
  33. Fetcho, J. R., McLean, D. L. Some principles of organization of spinal neurons underlying locomotion in zebrafish and their implications. Ann N Y Acad Sci. 1198, 94-104 (2010).
  34. Del Barrio, M. G., et al. A transcription factor code defines nine sensory interneuron subtypes in the mechanosensory area of the spinal cord. PLoS One. 8 (11), (2013).
  35. Satou, C., Kimura, Y., Hirata, H., Suster, M. L., Kawakami, K., Higashijima, S. Transgenic tools to characterize neuronal properties of discrete populations of zebrafish neurons. Development. 140 (18), 3927-3931 (2013).
  36. McLean, D. L., Fan, J., Higashijima, S., Hale, M. E., Fetcho, J. R. A topographic map of recruitment in spinal cord. Nature. 446, 71-75 (2007).
  37. McLean, D. L., Masino, M. A., Koh, I. Y., Lindquist, W. B., Fetcho, J. R. Continuous shifts in the active set of spinal interneurons during changes in locomotor speed. Nat. Neurosci. 11, 1419-1429 (2008).
  38. McLean, D. L., Fetcho, J. R. Spinal interneurons differentiate sequentially from those driving the fastest swimming movements in larval zebrafish to those driving the slowest ones. J. Neurosci. 29, 13566-13577 (2009).
  39. Ampatzis, K., Song, J., Ausborn, J., El Manira, A. Separate microcircuit modules of distinct V2a interneurons and motoneurons control the speed of locomotion. Neuron. 83, 934-943 (2014).
  40. Ljunggren, E. E., Haupt, S., Ausborn, J., Ampatzis, K., El Manira, A. Optogenetic activation of excitatory premotor interneurons is sufficient to generate coordinated locomotor activity in larval zebrafish. J. Neurosci. 34, 134-139 (2014).
  41. Menelaou, E., VanDunk, C., McLean, D. L. Differences in the morphology of spinal V2a neurons reflect their recruitment order during swimming in larval zebrafish. J Comp Neurol. 522, 1232-1248 (2014).
  42. Hubbard, J. M., et al. Intraspinal Sensory Neurons Provide Powerful Inhibition to Motor Circuits Ensuring Postural Control during Locomotion. Curr Biol. 26 (21), 2841-2853 (2016).
  43. Song, J., Ampatzis, K., Björnfors, E. R., El Manira, A. Motor neurons control locomotor circuit function retrogradely via gap junctions. Nature. 529 (7586), 399-402 (2016).
  44. Lamborghini, J. E. Rohon-beard cells and other large neurons in Xenopus embryos originate during gastrulation. J. Comp. Neurol. 189, 323-333 (1980).
  45. Myers, P. Z., Eisen, J. S., Westerfield, M. Development and axonal outgrowth of identified motoneurons in the zebrafish. J Neurosci. 6, 2278-2289 (1986).
  46. Kimmel, C. B., Westerfield, M., Edelman, G. M., Gall, W. E., Cowan, W. M. Primary neurons of the zebrafish. Signals and Sense: Local and Global Order in Perceptual Maps. , 561-588 (1990).
  47. Metcalfe, W. K., Myers, P. Z., Trevarrow, B., Bass, M. B., Kimmel, C. B. Primary neurons that express the L2/HNK-1 carbohydrate during early development in the zebrafish. Development. 110 (2), 491-504 (1990).
  48. Rossi, C. C., Kaji, T., Artinger, K. B. Transcriptional control of Rohon-Beard sensory neuron development at the neural plate border. Dev Dyn. 238, 931-943 (2009).
  49. Lewis, K. E., Eisen, J. S. From cells to circuits: development of the zebrafish spinal cord. Progress in Neurobiology. 69 (6), 419-449 (2003).
  50. Reimer, M. M., et al. Dopamine from the brain promotes spinal motor neuron generation during development and adult regeneration. Dev Cell. 25 (5), 478-491 (2013).
  51. Lambert, A. M., Bonkowsky, J. L., Masino, M. A. The conserved dopaminergic diencephalospinal tract mediates vertebrate locomotor development in zebrafish larvae. J Neurosci. 32 (39), 13488-13500 (2012).
  52. Westerfield, M. . The zebrafish book. A guide for the laboratory use of zebrafish (Danio rerio). , (1995).
  53. Oesterle, A. . Pipette Cookbook 2015 P-97 & P-1000 Micropipette pullers. , (2015).
  54. Spitzer, N. C. The ionic basis of the resting potential and a slow depolarizing response in Rohon-Beard neurons of Xenopus tadpoles. J Physiol. 255 (1), 105-135 (1976).
  55. Higashijima, S., Hotta, Y., Okamoto, H. Visualization of cranial motor neurons in live transgenic zebrafish expressing green fluorescent protein under the control of the islet-1 promoter/enhancer. J Neurosci. 20, 206-218 (2000).
  56. Blader, P., Plessy, C., Strahle, U. Multiple regulatory elements with spatially and temporally distinct activities control neurogenin1 expression in primary neurons of the zebrafish embryo. Mech Dev. 120, 211-218 (2003).
  57. Palanca, A. M., et al. New transgenic reporters identify somatosensory neuron subtypes in larval zebrafish. Dev Neurobiol. 73, 152-167 (2013).
  58. Flanagan-Street, H., Fox, M. A., Meyer, D., Sanes, J. R. Neuromuscular synapses can form in vivo by incorporation of initially aneural postsynaptic specializations. Development. 132, 4471-4481 (2005).
  59. Arkhipova, V., Wendik, B., Devos, N., Ek, O., Peers, B., Meyer, D. Characterization and regulation of the hb9/mnx1 beta-cell progenitor specific enhancer in zebrafish. Dev Biol. 365, 290-302 (2012).
  60. Balciunas, D., Davidson, A. E., Sivasubbu, S., Hermanson, S. B., Welle, Z., Ekker, S. C. Enhancer trapping in zebrafish using the Sleeping Beauty transposon. BMC Genomics. 5 (1), 62 (2004).
  61. Meng, A., Tang, H., Ong, B. A., Farrell, M. J., Lin, S. Promoter analysis in living zebrafish embryos identifies a cis-acting motif required for neuronal expression of GATA-2. Proc. Natl. Acad. Sci. USA. 94, 6267-6272 (1997).
  62. Menelaou, E., McLean, D. L. A gradient in endogenous rhythmicity and oscillatory drive matches recruitment order in an axial motor pool. J Neurosci. 32, 10925-10939 (2012).
  63. Rohrbough, J., Pinto, S., Mihalek, R. M., Tully, T., Broadie, K. latheo, a Drosophila gene involved in learning, regulates functional synaptic plasticity. Neuron. 23 (1), 55-70 (1999).
  64. McKeown, K. A., Moreno, R., Hall, V. L., Ribera, A. B., Downes, G. B. Disruption of Eaat2b, a glutamate transporter, results in abnormal motor behaviors in developing zebrafish. Dev Biol. 362 (2), 162-171 (2012).
  65. Carmean, V., et al. pigk mutation underlies macho behavior and affects Rohon-Beard cell excitability. J Neurophysiol. 114 (2), 1146-1157 (2015).
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Moreno, R. L., Josey, M., Ribera, A. B. Zebrafish In Situ Spinal Cord Preparation for Electrophysiological Recordings from Spinal Sensory and Motor Neurons. J. Vis. Exp. (122), e55507, doi:10.3791/55507 (2017).

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