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第1部分：设备和用品<ol><li>一个良好的解剖镜下胚胎解剖;建议40X放大倍率25X眼睛件，最大限度地增加放大。 </li><li>胚胎及胚胎devitellinization手动选择需要精细镊子（5号）。 </li><li>制定和修改细玻璃针的设备是必需的。拉玻璃针都需要剥离。我们喜欢实心玻璃夹层（持续更长的时间），但工程，以及标准的厚壁玻璃管（外径1-1.5毫米）。有些人用细的钨针，电解所需的清晰度在1M的NaOH使用自耦变压器电池10 + AMP浴。锐利的钨清扫针，他们是比较有弹性和长期的利益。中空玻璃针（形成类似膜片钳电极）连接到塑料管中，并在清扫吸力和生理盐水驱逐，以及胶水的控制下交付使用。各种玻璃车夫可用于制造玻璃针。电脑编程车夫可预置拉各种形状和大小。布朗阻燃模型（萨特仪器有限公司）的广泛青睐。一个复合式显微镜（20 - 40X的目标）应可用来检查和修改电极在使用前。 </li><li>两种类型的解决方案通常用于沐浴在剥离过程中胚胎组织：1）（月和1月，1976a）“标准”或“修订标准”（Broadie 2000年）salines，根据记录在其他无脊椎动物系统常用的解决方案，2）“血淋巴样”（HL）的salines（Stewart等，1994），标准生理盐水和在果蝇血淋巴测量离子浓度之间的妥协。应该指出，这些salines都没有准确地模仿在体内的血淋巴中的化学成分（Broadie 2000年）。标准生理盐水（毫米）：135氯化钠，氯化钾，4 2 1.8氯化镁2，氯化钙，72蔗糖，5工商业污水附加费，pH值7.2。实验者也可能要考虑商业化准备的昆虫生理盐水（如施耐德公司的昆虫媒体），这是不合适的电生理记录，但最有可能的，以保护暴露组织的健康。 </li></ol>第2部分：胚胎分期和解剖<ol><li>最佳鸡蛋收集，保持年轻（&lt;7天），健康雄性和雌性果蝇的新鲜奠定盆2-3天（20-40）。铺设盆通常由100毫升塑料烧杯（三倒）穿孔，以提供足够的空气流通，涵盖了60毫米琼脂平板。琼脂平板上包含苹果或葡萄汁与琼脂硬化。多个配方存在，但其中一个例子如下：700毫升的水，琼脂25-30克，300毫升的浓缩果汁（葡萄或苹果），0.5克甲基苯甲酸酯（P - hydroxymethylbenzoate），和30克糖。高压灭菌的水和琼脂，并分别熬糖和果汁的P - hydroxymethylbenzoate。混合的琼脂和果汁解决方案，并迅速倒入60毫米的钢板，消除泡沫。 </li><li>鸡蛋收集之前，苍蝇喂酵母膏（9毫升的水储存于4 ° C，面包酵母7克）新鲜的板块，每天至少两次，至少两天改变。第3天，好锅应每小时100-200鸡蛋。更好的产蛋观察划伤板琼脂在其一侧保持了锅。许多胚胎将放置在旁边的划痕。 </li><li>为了收集大量的胚胎，轻轻地从琼脂平板上转移到一个篮子用画笔胚胎。一篮子可与15或50 mL离心管。切断底部留出足够的表面积陷阱之间的盖子和管顶部的屏幕（70微米网状）。与卫生署2 0冲洗胚胎放入一个培养皿新鲜的50％至100％的漂白粉去除外层的绒毛膜。此外，鸡蛋可以收集单独使用罚款钳，并对其进行手动放入漂白水下降dechorionated。 chorionation可以从30秒到2分钟（新鲜漂白快得多），并应解剖显微镜下监测。去除绒毛膜公开透明的光泽，卵黄膜。 Dechorionated鸡蛋应简单地冲洗，或置于生理盐水中，作为漂白剂迅速破坏devitellinized组织。 </li><li>仔细到所需的发育年龄阶段的胚胎，这一点至关重要。定义的果蝇胚胎阶段（1-17;坎波斯奥尔特加和Hartenstein，1985年）是没有用的，所有功能的神经肌肉发展发生在后期的16或阶段17的。因此，胚胎发育小时上演25 °在培养箱中小时后受精或，更普遍的，产蛋后（AEL）。在这些条件下，胚胎持续21 + / - 1小时25 ° C。 </li><li>鸡蛋从一个定时蛋打下（1-2小时S），是适当的年龄在旁边观看的形态学标准。 DH 2 O的广泛的洗涤后，dechorionated鸡蛋都放在一个塑料培养皿（卫生署辖下 2）查看。可以确认正确的临时使用反射光的形态与解剖显微镜（坎波斯奥尔特加和Hartenstein，1985年）的标准。成熟后期胚胎孵化的边缘（22 - 24小时机场快线）是普遍认可的角质层和虚增气管分割。胚胎也可以使用GFP平衡器和一个适当的过滤器的荧光解剖镜下基因型。 </li><li>清扫，dechorionated和发育上演的胚胎附着在盖玻片下的生理盐水下降（见第1.5步）（背侧）。胚胎从卵黄膜与玻璃微吸管或钨针刺破膜附近的前或后结束，然后轻轻地从膜释放的胚胎。在准备解剖，胚胎应定位于背水面达（背侧了清扫将公开腹侧神经系统和实验neuromusculature盖玻片，但是，胚胎显然是定位在其他方面，以方便访问其他组织。） </li><li>早期胚胎（&lt;16小时机场快线）直接连接到干净的玻璃或镀膜玻璃与多聚赖氨酸（聚- L -赖氨酸氢溴酸; Broadie和巴特，1993，B）。胚胎（机场快线&gt; 16小时）后，角质层的形成，必须粘，最好Sylgard涂层（道康宁）盖玻片。使用一个聚合水的氰基丙烯酸酯胶外科或兽医（Broadie和巴特，1993c，D）。胶是通过一个小玻璃电极吸管口压力控制的胶水流连接橡胶管（10-20微米内径TIP）交付。必须小心胶交付期间，胶水联系生理盐水后迅速聚合。请注意，也不能执行，粘合没有或低的二价离子浓度的盐水，胶水，需要二价离子聚合。首次使用少量的胶水牢固地附着在头部和尾部。 </li><li>一旦胚胎的两端粘盖玻片，切口是沿背中线与玻璃电极或削尖的钨针。这是轻轻射孔切口线，晋级决赛之前变得更加容易。切口的两侧，然后连接到盖玻片更多的胶水，轻轻蔓延胚胎单位。内部器官，包括肠，脂肪体，可选，气管，然后由使用橡胶管（Broadie和贝特1993 - C）连接到一个玻璃吸管吸删除。现在应该附加胚胎持平盖玻片，表皮腹中枢神经系统，周围神经和躯体肌肉暴露实验。 </li></ol>

胚胎的精确分期是由于功能特性的只是几个小时的时间当然的迅速成熟的关键。几个问题复杂化分期。首先，大多数研究人员使用定时鸡蛋奠定阶段的胚胎，但产蛋时间可以千差万别，在不同的条件（Broadie等，1992）在动物。特别是，在有限的食物的女性倾向于保留奠定前长时间的受精卵。因此，至关重要的丰富的酵母饮食喂养前至少2天收集蛋（Broadie等，1992）“明确的”女性。此外，老年女性?...

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		<div>
		<table border="1">
		<thead>
		<tr>
		<th>Material Name</th>
		<th>Type</th>
		<th>Company</th>
		<th>Catalogue Number</th>
		<th>Comment</th>
		</tr>
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<tr>
<td>Embryo Collection Cages</td>
<td>&nbsp;</td>
<td>Genesee Scientific (www.flystuff.com)</td>
<td>59-100 (for 60 mm dish; other sizes available)</td>
<td>Cages can also be home made using punctured tri-pour beakers, as shown in video</td>
</tr>
<tr>
<td>Sylgard 184</td>
<td>&nbsp;</td>
<td>Dow Corning</td>
<td>Available from various companies</td>
<td>Surgical glue adheres better to sylgard-coated coverslips</td>
</tr>
<tr>
<td>Fine glass tubing, outer diameter 1.0-1.5 mm</td>
<td>&nbsp;</td>
<td>various</td>
<td>&nbsp;</td>
<td>For pulling into fine glass needles for dissection and tubes for glue delivery</td>
</tr>
<tr>
<td>Plastic tubing, to attach to glass pipette for mouth suction and glue application</td>
<td>&nbsp;</td>
<td>Tygon</td>
<td>&nbsp;</td>
<td>Tubing inner diameter needs to match glass outer diameter.</td>
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harvesting and preparing drosophila embryos for electrophysiological recording and other procedures

果蝇胚胎发育和功能的神经科学的研究是首屈一指的遗传模型。传统上，这些领域都相当彼此孤立的，很大程度上是独立的历史和科学界。然而，这些通常是不同的领域之间的接口是发展计划的基本功能的电子信号的属性和功能的化学突触在神经回路形成的最后阶段分化的收购。该接口是一个极为重要的领域进行调查。在果蝇中，这些功能开发阶段发生在胚胎发育的最后三分之一的&lt;8小时期间（在25 ° C） 。后期发展时期，长期被视为棘手的调查，由于一个艰难的，不透水的表皮角质层的沉积。一个突破性的进展是，可以在本地应用的角质层，使后期胚胎的控制清扫手术，聚合水胶中的应用。胚胎背侧纵切口，可以平放，露出腹侧神经线和体壁肌肉的实验研究。该系统已大量用于分离和鉴定基因突变体，损害胚胎的突触的形成，从而揭示了突触连接和功能的突触信号特性的规范和分化的分子机制。

Watch this Scientific Journal Video about Harvesting and Preparing Drosophila Embryos for Electrophysiological Recording and Other Procedures at JoVE.com

Harvesting and Preparing Drosophila Embryos for Electrophysiological Recording and Other Procedures

这种技术公开的果蝇胚胎neuromusculature免疫组织化学或电生理记录。它是有用的，为研究神经肌肉发展的早期事件，或执行中的突变体，不能孵化电。

收获和准备果蝇胚胎的电生理记录和其他程序

Drosophila is a premier genetic model for the study of both embryonic development and functional neuroscience. Traditionally, these fields are quite ...

harvesting-preparing-drosophila-embryos-for-electrophysiological

Research

JoVE Journal

Biology

Harvesting and Preparing Drosophila Embryos for Electrophysiological Recording and Other Procedures

13.3K 次观看.  University of Illinois. 这种技术公开的果蝇胚胎neuromusculature免疫组织化学或电生理记录。它是有用的，为研究神经肌肉发展的早期事件，或执行中的突变体，不能孵化电。

视频: Harvesting and Preparing Drosophila Embryos for Electrophysiological Recording and Other Procedures

Preparation of Neuronal Cultures from Midgastrula Stage Drosophila Embryos

This video illustrates the procedure for making primary neuronal cultures from midgastrula stage Drosophila embryos. The methods for collecting embryos and their dechorionation using bleach are demonstrated. Using a glass pipet attached to a mouth suction tube, we illustrate the removal of all cells from single embryos. The method for dispersing cells from each embyro into a small (5 l) drop of medium on an uncoated glass coverslip is demonstrated. A view through the microscope at 1 hour after plating illustrates the preferred cell density. Most of the cells that survive when grown in defined medium are neuroblasts that divide one or more times in culture before extending neuritic processes by 12-24 hours. A view through the microscope illustrates the level of neurite outgrowth and branching expected in a healthy culture at 2 days in vitro. The cultures are grown in a simple bicarbonate based defined medium, in a 5% CO2 incubator at 22-24&deg;C. Neuritic processes continue to elaborate over the first week in culture and when they make contact with neurites from neighboring cells they often form functional synaptic connections. Neurons in these cultures express voltage-gated sodium, calcium, and potassium channels and are electrically excitable. This culture system is useful for studying molecular genetic and environmental factors that regulate neuronal differentiation, excitability, and synapse formation/function.

This video demonstrates the preparation of primary neuronal cultures from midgastrula stage Drosophila embryos. Views of live cultures show cells 1 hour after plating and differentiated neurons after 2 days of growth in a bicarbonate-based defined medium. The neurons are electrically excitable and form synaptic connections.

从Midgastrula舞台果蝇胚胎神经文化的制备

This video illustrates the procedure for making primary neuronal cultures from midgastrula stage Drosophila embryos. The methods for collecting embryos ...

科研

生物学

Electrophysiological Methods for Recording Synaptic Potentials from the NMJ of Drosophila Larvae

In this video, we describe the electrophysiological methods for recording synaptic transmission at the neuromuscular junction (NMJ) of Drosophila larva. The larval neuromuscular system is a model synapse for the study of synaptic physiology and neurotransmission, and is a valuable research tool that has defined genetics and is accessible to experimental manipulation. Larvae can be dissected to expose the body wall musculature, central nervous system, and peripheral nerves. The muscles of Drosophila and their innervation pattern are well characterized and muscles are easy to access for intracellular recording. Individual muscles can be identified by their location and orientation within the 8 abdominal segments, each with 30 muscles arranged in a pattern that is repeated in segments A2 - A7. Dissected drosophila larvae are thin and individual muscles and bundles of motor neuron axons can be visualized by transillumination1. Transgenic constructs can be used to label target cells for visual identification or for manipulating gene products in specific tissues. In larvae, excitatory junction potentials (EJP&#8217;s) are generated in response to vesicular release of glutamate from the motoneurons at the synapse. In dissected larvae, the EJP can be recorded in the muscle with an intracellular electrode. Action potentials can be artificially evoked in motor neurons that have been cut posterior to the ventral ganglion, drawn into a glass pipette by gentle suction and stimulated with an electrode. These motor neurons have distinct firing thresholds when stimulated, and when they fire simultaneously, they generate a response in the muscle. Signals transmitted across the NMJ synapse can be recorded in the muscles that the motor neurons innervate. The EJP&#8217;s and minature excitatory junction potentials (mEJP&#8217;s) are seen as changes in membrane potential. Electrophysiological responses are recorded at room temperature in modified minimal hemolymph-like solution2 (HL3) that contains 5 mM Mg2+ and 1.5 mM Ca2+. Changes in the amplitude of evoked EJP&#8217;s can indicate differences in synaptic function and structure. Digitized recordings are analyzed for EJP amplitude, mEJP frequency and amplitude, and quantal content.

Here we describe electrophysiological methods for measuring synaptic transmission at the neuromuscular junction of Drosophila larva. Evoked release is initiated artificially by stimulating the motor neuron axons, and transmission through the NMJ can be measured by the postsynaptic response evoked in the muscle.

电生理方法记录从果蝇幼虫NMJ突触潜力

In this video, we describe the electrophysiological methods for recording synaptic transmission at the neuromuscular junction (NMJ) of Drosophila larva. ...

Electrophysiological Recording in the Drosophila Embryo

Drosophila is a premier genetic model for the study of both embryonic development and functional neuroscience. Traditionally, these fields are quite isolated from each other, with largely independent histories and scientific communities. However, the interface between these usually disparate fields is the developmental programs underlying acquisition of functional electrical signaling properties and differentiation of functional chemical synapses during the final phases of neural circuit formation. This interface is a critically important area for investigation. In Drosophila, these phases of functional development occur during a period of &lt;8 hours (at 25&deg;C) during the last third of embryogenesis. This late developmental period was long considered intractable to investigation owing to the deposition of a tough, impermeable epidermal cuticle. A breakthrough advance was the application of water-polymerizing surgical glue that can be locally applied to the cuticle to enable controlled dissection of late-stage embryos. With a dorsal longitudinal incision, the embryo can be laid flat, exposing the ventral nerve cord and body wall musculature to experimental investigation. Whole-cell patch-clamp techniques can then be employed to record from individually-identifiable neurons and somatic muscles. These recording configurations have been used to track the appearance and maturation of ionic currents and action potential propagation in both neurons and muscles. Genetic mutants affecting these electrical properties have been characterized to reveal the molecular composition of ion channels and associated signaling complexes, and to begin exploration of the molecular mechanisms of functional differentiation. A particular focus has been the assembly of synaptic connections, both in the central nervous system and periphery. The glutamatergic neuromuscular junction (NMJ) is most accessible to a combination of optical imaging and electrophysiological recording. A glass suction electrode is used to stimulate the peripheral nerve, with excitatory junction current (EJC) recordings made in the voltage-clamped muscle. This recording configuration has been used to chart the functional differentiation of the synapse, and track the appearance and maturation of presynaptic glutamate release properties. In addition, postsynaptic properties can be assayed independently via iontophoretic or pressure application of glutamate directly to the muscle surface, to measure the appearance and maturation of the glutamate receptor fields. Thus, both pre- and postsynaptic elements can be monitored separately or in combination during embryonic synaptogenesis. This system has been heavily used to isolate and characterize genetic mutants that impair embryonic synapse formation, and thus reveal the molecular mechanisms governing the specification and differentiation of synapse connections and functional synaptic signaling properties.

Electrophysiological Recording in the Drosophila Embryo

Electrophysiological recordings from Drosophila embryos allow analyses of developing muscle and neuron electrical properties, as well as characterization of functional synaptogenesis at the glutamatergic neuromuscular junction and central cholinergic and GABAergic synapses.

在电生理记录果蝇胚胎

Preparation of embryos for Electron Microscopy of the Drosophila embryonic heart tube

The morphogenesis of the Drosophila embryonic heart tube has emerged as a valuable model system for studying cell migration, cell-cell adhesion and cell shape changes during embryonic development. One of the challenges faced in studying this structure is that the lumen of the heart tube, as well as the membrane features that are crucial to heart tube formation, are difficult to visualize in whole mount embryos, due to the small size of the heart tube and intra-lumenal space relative to the embryo. The use of transmission electron microscopy allows for higher magnification of these structures and gives the advantage of examining the embryos in cross section, which easily reveals the size and shape of the lumen. In this video, we detail the process for reliable fixation, embedding, and sectioning of late stage Drosophila embryos in order to visualize the heart tube lumen as well as important cellular structures including cell-cell junctions and the basement membrane.

Preparation of embryos for Electron Microscopy of the Drosophila embryonic heart tube

We describe a process for fixation, embedding, sectioning, and imaging of late stage Drosophila embryos for Trasmission Electron Microscopy of the embryonic heart tube. This technique allows for the visualization of the heart tube lumen as well as the basement membrane, which lines the lumen of the heart.

电子显微镜对胚胎的制备果蝇胚胎心管

The morphogenesis of the Drosophila  embryonic heart tube has emerged as a valuable model system for studying cell migration, cell-cell adhesion and cell ...

In-vivo Centrifugation of Drosophila Embryos

A major strategy for purifying and isolating different types of intracellular organelles is to separate them from each other based on differences in buoyant density. However, when cells are disrupted prior to centrifugation, proteins and organelles in this non-native environment often inappropriately stick to each other. Here we describe a method to separate organelles by density in intact, living Drosophila embryos. Early embryos before cellularization are harvested from population cages, and their outer egg shells are removed by treatment with 50% bleach. Embryos are then transferred to a small agar plate and inserted, posterior end first, into small vertical holes in the agar. The plates containing embedded embryos are centrifuged for 30 min at 3000g. The agar supports the embryos and keeps them in a defined orientation. Afterwards, the embryos are dug out of the agar with a blunt needle. Centrifugation separates major organelles into distinct layers, a stratification easily visible by bright-field microscopy. A number of fluorescent markers are available to confirm successful stratification in living embryos. Proteins associated with certain organelles will be enriched in a particular layer, demonstrating colocalization. Individual layers can be recovered for biochemical analysis or transplantation into donor eggs. This technique is applicable for organelle separation in other large cells, including the eggs and oocytes of diverse species.

In-vivo Centrifugation of Drosophila Embryos

We describe a method to separate organelles by density in living Drosophila embryos. Embryos are embedded in agar and centrifuged. This technique yields reproducible separation of major organelles along the anterior-posterior embryo axis. This method facilitates colocalization experiments and yields organelle fractions for biochemical analysis and transplantation experiments.

果蝇胚胎的体内离心

A major strategy for purifying and isolating different types of intracellular organelles is to separate them from each other based on differences in ...

Upright Imaging of Drosophila Embryos

Several well-known morphogenetic gradients and cellular movements occur along the dorsal/ventral axis of the Drosophila embryo. However, the current techniques used to view such processes are somewhat limited. The following protocol describes a new technique for mounting fixed and labeled Drosophila embryos for coronal viewing with confocal imaging. This method consists of embedding embryos between two layers of glycerin jelly mounting media, and imaging jelly strips positioned upright. The first step for sandwiching the embryos is to make a thin bedding of glycerin jelly on a slide. Next, embryos are carefully aligned on this surface and covered with a second layer of jelly. After the second layer is solidified, strips of jelly are cut and flipped upright for imaging. Alternatives are described for visualizing the embryos depending upon the type of microscope stand to be used. Since all cells along the dorsal-ventral axis are imaged within a single confocal Z-plane, our method allows precise measurement and comparison of fluorescent signals without photobleaching or light scattering common to 3D reconstructions of longitudinally mounted embryos.

Upright Imaging of Drosophila Embryos

Here we present a mounting protocol for stained Drosophila embryos in an upright position that allows imaging of cross-sections using Confocal microscopy.

直立成像果蝇胚胎

Several well-known morphogenetic gradients and cellular movements occur along the dorsal/ventral axis of the Drosophila embryo. However, the current ...

Optimized Transfection Strategy for Expression and Electrophysiological Recording of Recombinant Voltage-Gated Ion Channels in HEK-293T Cells

The in vitro expression and electrophysiological recording of recombinant voltage-gated ion channels in cultured human embryonic kidney cells (HEK-293T) is a ubiquitous research strategy. HEK-293T cells must be plated onto glass coverslips at low enough density so that they are not in contact with each other in order to allow for electrophysiological recording without confounding effects due to contact with adjacent cells. Transfected channels must also express with high efficiency at the plasma membrane for whole-cell patch clamp recording of detectable currents above noise levels. Heterologous ion channels often require long incubation periods at 28&deg;C after transfection in order to achieve adequate membrane expression, but there are increasing losses of cell-coverslip adhesion and membrane stability at this temperature. To circumvent this problem, we developed an optimized strategy to transfect and plate HEK-293T cells. This method requires that cells be transfected at a relatively high confluency, and incubated at 28&deg;C for varying incubation periods post-transfection to allow for adequate ion channel protein expression. Transfected cells are then plated onto glass coverslips and incubated at 37&deg;C for several hours, which allows for rigid cell attachment to the coverslips and membrane restabilization. Cells can be recorded shortly after plating, or can be transferred to 28&deg;C for further incubation. We find that the initial incubation at 28&deg;C, after transfection but before plating, is key for the efficient expression of heterologous ion channels that normally do not express well at the plasma membrane. Positively transfected, cultured cells are identified by co-expressed eGFP or eGFP expressed from a bicistronic vector (e.g. pIRES2-EGFP) containing the recombinant ion channel cDNA just upstream of an internal ribosome entry site and an eGFP coding sequence. Whole-cell patch clamp recording requires specialized equipment, plus the crafting of polished recording electrodes and L-shaped ground electrodes from borosilicate glass. Drug delivery to study the pharmacology of ion channels can be achieved by directly micropipetting drugs into the recording dish, or by using microperfusion or gravity flow systems that produce uninterrupted streams of drug solution over recorded cells.

Reliable method for highly efficient in vitro expression and subsequent electrophysiological recording of recombinant voltage-gated ion channels in cultured human embryonic kidney cells (HEK-293T).

优化转染的HEK - 293T细胞中的表达和重组的电压门控离子通道的电生理记录策略

The in vitro expression and electrophysiological recording of recombinant voltage-gated ion channels in cultured human embryonic kidney cells (HEK-293T) ...

Isolation and Purification of Kinesin from Drosophila Embryos

Motor proteins move cargos along microtubules, and transport them to specific sub-cellular locations. Because altered transport is suggested to underlie a variety of neurodegenerative diseases, understanding microtubule based motor transport and its regulation will likely ultimately lead to improved therapeutic approaches. Kinesin-1 is a eukaryotic motor protein which moves in an anterograde (plus-end) direction along microtubules (MTs), powered by ATP hydrolysis. Here we report a detailed purification protocol to isolate active full length kinesin from Drosophila embryos, thus allowing the combination of Drosophila genetics with single-molecule biophysical studies. Starting with approximately 50 laying cups, with approximately 1000 females per cup, we carried out overnight collections. This provided approximately 10 ml of packed embryos. The embryos were bleach dechorionated (yielding approximately 9 grams of embryos), and then homogenized. After disruption, the homogenate was clarified using a low speed spin followed by a high speed centrifugation. The clarified supernatant was treated with GTP and taxol to polymerize MTs. Kinesin was immobilized on polymerized MTs by adding the ATP analog, 5'-adenylyl imidodiphosphate at room temperature. After kinesin binding, microtubules were sedimented via high speed centrifugation through a sucrose cushion. The microtubule pellet was then re-suspended, and this process was repeated. Finally, ATP was added to release the kinesin from the MTs. High speed centrifugation then spun down the MTs, leaving the kinesin in the supernatant. This kinesin was subjected to a centrifugal filtration using a 100 KD cut off filter for further purification, aliquoted, snap frozen in liquid nitrogen, and stored at -80 &deg;C. SDS gel electrophoresis and western blotting was performed using the purified sample. The motor activity of purified samples before and after the final centrifugal filtration step was evaluated using an in vitro single molecule microtubule assay. The kinesin fractions before and after the centrifugal filtration showed processivity as previously reported in literature. Further experiments are underway to evaluate the interaction between kinesin and other transport related proteins.

Isolation and Purification of Kinesin from Drosophila Embryos

This is a protocol to isolate active full length Kinesin from Drosophila embryos for single-molecule biophysical studies. We show how to collect embryos, make the embryo lysate, and then polymerize microtubules (MTs). Kinesin is purified by immobilizing it on the MTs, spinning down the Kinesin-MT complexes, and then releasing the kinesin from the MTs via ATP addition.

kinesin的分离和纯化果蝇胚胎

Motor proteins move cargos along microtubules, and transport them to specific sub-cellular locations. Because altered transport is suggested to underlie a ...

Preparing Individual Drosophila Egg Chambers for Live Imaging

Live cell imaging is an important technique applied to a number of Drosophila tissues used as models to investigate topics such as axis specification, cell differentiation and organogenesis 1. Correct preparation of the experimental samples is a crucial, often neglected, step. The goal of preparation is to ensure physiological relevance and to establish optimal imaging conditions. To maintain tissue viability, it is critical to avoid dehydration, hypoxia, overheating or medium deterioration 2. 
 The Drosophila egg chamber is a well established system for examining questions relating, but not limited, to body patterning, mRNA localization and cytoskeletal organization 3,4. For early- and mid-stage egg chambers, mounting in halocarbon oil is good for survival in that it allows free diffusion of oxygen, prevents dehydration and hypoxia and has superb optical properties for microscopy. Imaging of fluorescent proteins is possible through the introduction of transgenes into the egg chamber or physical injection of labeled RNA, protein or antibodies 5-7. For example, addition of MS2 constructs to the genome of animals enables real time observation of mRNAs in the oocyte 8. These constructs allow for in vivo labeling of mRNA through utilization of the MS2 bacteriophage RNA stem loop interaction with its coat protein 9. 
 Here, we present a protocol for the extraction of ovaries as well as isolating individual ovarioles and egg chambers from the female Drosophila. For a detailed description of Drosophila oogenesis see Allan C. Spradling (1993, reprinted 2009) 10.

Preparing Individual Drosophila Egg Chambers for Live Imaging

The Drosophila egg chamber is an excellent model for studying the mechanisms of mRNA localization. In order to capture the dynamic events that underpin the processes of localization, rapid high resolution imaging of live tissue is required. Here, we present a protocol for dissection and imaging of live samples with minimal disruption.

准备个人果蝇蛋钱伯斯实时成像

Live cell imaging is an important technique applied to a number of Drosophila tissues used as models to investigate topics such as axis specification, ...

Studying Mitotic Checkpoint by Illustrating Dynamic Kinetochore Protein Behavior and Chromosome Motion in Living Drosophila Syncytial Embryos

The spindle assembly checkpoint (SAC) mechanism is an active signal, which monitors the interaction between chromosome kinetochores and spindle microtubules to prevent anaphase onset until the chromosomes are properly connected. Cells use this mechanism to prevent aneuploidy or genomic instability, and hence cancers and other human diseases like birth defects and Alzheimer's1. A number of the SAC components such as Mad1, Mad2, Bub1, BubR1, Bub3, Mps1, Zw10, Rod and Aurora B kinase have been identified and they are all kinetochore dynamic proteins2. Evidence suggests that the kinetochore is where the SAC signal is initiated. The SAC prime regulatory target is Cdc20. Cdc20 is one of the essential APC/C (Anaphase Promoting Complex or Cyclosome) activators3 and is also a kinetochore dynamic protein4-6. When activated, the SAC inhibits the activity of the APC/C to prevent the destruction of two key substrates, cyclin B and securin, thereby preventing the metaphase to anaphase transition7,8. Exactly how the SAC signal is initiated and assembled on the kinetochores and relayed onto the APC/C to inhibit its function still remains elusive.
Drosophila is an extremely tractable experimental system; a much simpler and better-understood organism compared to the human but one that shares fundamental processes in common. It is, perhaps, one of the best organisms to use for bio-imaging studies in living cells, especially for visualization of the mitotic events in space and time, as the early embryo goes through 13 rapid nuclear division cycles synchronously (8-10 minutes for each cycle at 25 &deg;C) and gradually organizes the nuclei in a single monolayer just underneath the cortex9.
Here, I present a bio-imaging method using transgenic Drosophila expressing GFP (Green Fluorescent Protein) or its variant-targeted proteins of interest and a Leica TCS SP2 confocal laser scanning microscope system to study the SAC function in flies, by showing images of GFP fusion proteins of some of the SAC components, Cdc20 and Mad2, as the example.

Studying Mitotic Checkpoint by Illustrating Dynamic Kinetochore Protein Behavior and Chromosome Motion in Living Drosophila Syncytial Embryos

The kinetochore is where the SAC initiates its signal monitoring the mitotic segregation of the sister chromatids. A method is described to visualize the recruitment and turnover of one of the kinetochore proteins and its coordination with the chromosome motion in Drosophila embryos using a Leica laser scanning confocal system.