这项工作是由韦尔奇基金会（I - 1657年）和美国国立卫生研究院（RO1EY18884）的赠款支持。公屋是尤金麦克德莫特在生物医学研究的学者。

1。简介在这段视频中，我们将演示如何准备实时成像与感光细胞的焦点免疫组织化学 ，果蝇的大脑和视网膜。首先，我们将展示如何准备的大脑解剖。下一步，我们将演示进行免疫组化常用的两种基本解剖，现场准备的基础上。这两种制剂是成人的大脑和眼睛解剖。下一步，我们将展示用于实时成像强调实时成像感光器以及它们的使用成人和发育中的大脑的解剖。最后，我们将描述怎样的形象活组织，并采用谐振共聚焦显微镜实时成像显示一些例子。 2。夹层准备<ol><li>为了获得良好的解剖，你需要锋利的镊子。使用锐化块或非常细的砂纸（1500），每边轻轻地传递镊子来回，直到入不敷出在细点。我们使用一个标准的磨刀石。我们将不包括这里的锐化技术，但需要注意的是尖锐的钳现场解剖的基本。我们增强在每次解剖钳。 </li><li>对于成人的大脑解剖，苍蝇，以CO 2垫麻醉他们，并整理出所需的基因型。蛹的解剖，只需达到进入瓶中，并与您的镊子小心取出蛹，要小心，以避免破坏蛹的情况下。 </li><li>与水浸湿Kimwipe。您将使用在解剖，以消除碎片从您的镊子。一个立体舞台上位置的解剖盘和填充HL3解决方案 1 。在HL3将进行全部清扫，以确保该组织仍然活着，健康在解剖过程。此外，我们使用一个夹层的菜，有一个Sylgard底部覆盖，以保护在解剖钳。 </li><li>现在，准备修复解决方案，如果您计划执行免疫组织化学。广场180μLHL3成500μL的离心管。添加37％的甲醛20μL获得3.7％的甲醛溶液。 </li><li>最后，适当的手的位置是很重要的，良好的解剖。良好的手的位置，您的镊子将稳定和能够控制的，细微的动作。首先，将你的拇指侧的镊子。直降等，手指的尖端在于在上面，然后把食指。现在停留在中指一侧一侧的镊子，解剖盘和移动。你的拇指和中指，使菜的身体接触。休息时你的拇指和中指上的菜，厂房显微镜舞台上你的手腕。通过这种方式，你有三点种植坚决表面上在解剖和现在有可能使镊子非常微妙的操纵。这种技术可能会觉得有点尴尬，但通过练习，你很快就会掌握的大脑解剖。 </li></ol> 3。成人的大脑<ol><li>在CO 2垫，东方成人飞腹侧与远离你的手头部。抢头正下​​方的胸部。如果做得好，腿和喙将延长。一个立体下观察时，与您钳抢延长长鼻删除头部。丢弃的身体和淹没头部。重新聚焦到淹没头。整个剥离，将执行下的解决方案。到头部与钳在任何时候，这是非常重要。否则，头部将float和难以检索。如果头移动在夹层中的重点，只需移动到焦平面不调整显微镜。这看似简单的任务，可能很难在第一，但保持你的头在焦点将随着时间的推移变得更容易。 </li><li>第一撕裂的长鼻和眼睛之间的结缔组织开始清扫。同时，在任何时候都至少有一个钳坚决眼睛撕裂。一定镊子脚趾下方的视网膜或角质层，以避免损坏大脑的下方。左，右交替，使用您的镊子撕离视网膜，朝后脑勺工作。撕毁了视网膜会增加成功的机会，椎板剥离附着在整个视叶。继续围绕的后脑勺，流泪，一路上的角质层。撕裂通过另一只眼睛，开始拉远角质层。只要你抓住的角质层，你知道，你是不是破碎的大脑！考虑到这一点，继续拉角质层和视网膜，直到大脑透露。 </li><li>一旦大脑是可见的，你就可以开始删除气管连接到大脑（图1A）。继续拉在气管和角质层，直到大脑是孤立的。此时，你应该重新调整你的菜吨的底部他其余的步骤。除去剩余的气管因为你的大脑在一夜之间抗体清洗，否则可能会浮动。成像感光终端，它是可取的保留椎板，但您需要删除的视网膜，其中包含的胞体和强烈autofluorescent颜料。要做到这一点，小心地将在浓密的色素细胞仍然不破坏椎板。这是一个更精细的技能和需要练习。 </li><li>成人的大脑，可以保持几个小时甚至数天或使用条件下，我们将讨论在第6活着。进行免疫组化，成人的大脑现在已经准备好要固定甲醛（2）使用P20的pipetteman设置到5μL移动成人的大脑你已经准备到固定的解决方案。继续20分钟，成人的大脑解剖，产量4-10大脑。保留一个额外的40分钟的大脑修复，但此期间可能会有所不同最优的主要抗体染色。去除甲醛溶液和洗脑筋400μL含有PBS和0.4％TritonX - 100，以下简称为PBS，海卫一的解决方案，每5分钟的3倍。在彻底清洗修复方案后，你可以增加抗体的解决方案。 </li></ol> 4。成人眼<ol><li>为成人大脑中所述，开始剥离。淹没头和重新定位，在显微镜后，撕裂长鼻和眼睛之间的结缔组织开始。上面的长鼻，从眼睛分开的角质层。现在，单独从眼睛下方的角质层，走向的后脑勺。一个镊子，抓住中心的后脑勺。另一方面，抓住眼球拉。让眼睛定居的菜的底部（图1B）。椎板仍然可能会被贴在眼睛上，可在这个时候使用完全保存完好的感光细胞的终端的实时成像。要继续与实时成像感光细胞组织，去到4.5节。接下来的三节解释固定眼免疫组化。 </li><li>免疫组化，移动到固定的解决方案使用的是P20的pipetteman设置到5μL的眼睛。继续解剖共10分钟的成人眼睛。保留在一个额外的30分钟的修复的眼睛。删除的修复方案和进行清洗，如在成人大脑解剖中所述。 </li><li>如果希望最终图像的眼睛ommatidial结构的椎板现在必须拆除。返回固定的眼睛，您的解剖盘，并消除任何可能仍附在气管或脂肪细胞。下一步，而眼外的一个钳，删除与其他层。它应该在一块脱落，露出cellbodies下方（图1B）。小心只抢椎板！否则，你的风险分离从视网膜的光感受器。 </li><li>使用P20的pipetteman，移动眼睛400uL PBS，海卫一在500uL管。轻轻摇动在4℃过夜，以除去autofluorescent从红色颜料的光感受器。翌日，你可以丢弃的洗涤液，并增加抗体的解决方案。 </li><li>实时成像成人的眼睛，我们只使用pharate成蝇，即前不久后期蛹羽化。在这个阶段，机构在剥离椎板终端分开的感光细胞。 </li><li> Pharate成人蛹有良好定义的翅膀，眼睛，和角质层通过蛹的情况下可见。选择一个蛹和删除前蛹的情况下部分露出头部区域。小心取出，此外包围发展中国家飞薄的内膜。抓斗钳和拉头的基地。丢弃蛹的休息，同时保持头部淹没。 </li><li>按照免疫组织化学眼清扫证明。 pharate成年阶段，然而，感光细胞机构将分离更容易从椎板，让您的形象依偎在视网膜上的细胞机构。此外，椎板仍然视叶，让您远端椎板形象。如果你想形象近端椎板可以看出，其中的感光终端墨盒，然后执行椎板贴在眼睛上更强烈的成人蝇眼清扫。 </li></ol> 5。眼睛光盘脑复合物<ol><li>在20％以上的蛹发展（20％）的眼睛光盘实时成像在我们的实验室已成为最喜爱的，因为这比较大的发展细胞单层容易观察使用共聚焦显微镜。 </li><li>要选择一个20％的蛹，为马氏管，一个绿点出现蛹的情况下之下，并避免蛹，有一个“黑暗的身体”4 。选择一个P 20％的蛹，淹没它在您的解剖盘在HL3后，小心取出蛹麻袋的前部，小心不穿透薄薄的内膜。抓住内膜与您的镊子，并拉开。 </li><li>通过发布的内容仔细搜索，发现发育中的大脑和眼睛的光盘还相当突出（图1C）。眼睛光盘脑复杂变态在这个​​阶段是很容易分离。隔离的大脑解剖盘底部。仔细分离视神经叶的中央大脑。视神经叶/眼磁盘将用于实时成像。 </li></ol> 6。实时成像：在显微镜<ol><li>如果您希望只有一个或两个解决方案的时间在短期内的图像，你可以在玻片上的形象您的组织。在这种情况下，第一涂层表面，使用制造商的指示与Sylgard准备的幻灯片。放置在幻灯片中20μLHL3。您的解剖组织运输HL3额外20μL到幻灯片。组织绝不能暴露在空气中。此外，任何压力或应变移液器处理或溶液的表面张力必须避免。 </li><li>实时成像，组织必须以最小的处理和联系坐稳。我们安全的位置大脑使用水聚合手术胶GluShield通过一个电极中的应用，这种技术经常用于胚胎电5准备。获得像拉电生理记录的玻璃电极。断绝提示GluShield使用口产生负压，并填写结束。仅折断足够的电极，胶可能进入。现在，使用正压申请下降了HL3少量的胶水的Sylgard。现在，迅速成立组织上胶，保持实时成像所需的方向。现在可以使用一个浸水镜头成像。如果你想添加一个膜渗透性染料，你可以将它添加到洗澡，而成像。 </li><li>我们使用一个共振扫描共聚焦显微镜，可减少漂白6以上的时间或交换解决方案的长期完全或多次成像，我们用它连接到一个蠕动泵灌注室（哈佛IC30共焦成像室）快速成像慢慢perfuses活组织培养液（图2A）。恒定的液体交换和最小GluShield接触的运动，减少扁平化的组织已观察了较长时期的时间7。请注意，发育时间段的成像，眼脑光盘复杂，需要保持不变，并应尽量避免接触胶水。 </li><li>使用下降了Sylgard是剃光呈现薄和平面涂盖玻片。免除到盖玻片和胶水活组织的Sylgard中心20μLHL3。生成垫片，使气泡室通过不活组织暴露在空气中（图2B）。垫片应足够厚，以避免破碎的组织，但薄足以使组织共聚焦显微镜镜头，我们在我们的设置使用250微米。组装灌注室，注意保持完全淹没在任何时候都组织。最初开始灌注率接近每分钟使用蠕动泵100μL。这个速度是相当快垫片的厚度取决于定义室高度。成像上的小时刻度，您可能需要添加20 - 羟基和抗生素，但不抗真​​菌，并执行在低得多的速度大约每分钟加入10μl灌注。最后，我们到培养液中，供应灌注腔泡沫的氧气。 </li></ol> 7。代表性的成果： 在我们的视频协议结束时，我们使用在第5节的准备，在发展中国家感光细胞实时成像的例子。这种准备利用果蝇的转基因细胞特异性表达荧光记者。在所示的例子中，两个荧光记者表示下控制特定的感光- GMR - Gal4的驱动程序8：膜标签myrRFP和内体标记2xFYVE - GFP 9。双通道3D的70x70x17.5μm的边界框和一个137x137x700nm（512x512x26）的体素大小的数据集，每1分钟。影片中展示了在这个4D数据集，揭示了胞内体的贩运和囊泡融合事件的动态，随着时间的推移选定的平面。代表固定的组织准备工作的眼睛和椎板的图像如图所示​​。 3。

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T. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Ebony+protein+in+the+Drosophila+nervous+system:+optic+neuropile+expression+in+glial+cells.">Ebony protein in the Drosophila nervous system: optic neuropile expression in glial cells.</a> J Comp Neurol. 452 (1), 93-102 (2002).</li><li>Beronja, S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Essential+function+of+Drosophila+Sec6+in+apical+exocytosis+of+epithelial+photoreceptor+cells.">Essential function of Drosophila Sec6 in apical exocytosis of epithelial photoreceptor cells.</a> J Cell Biol. 169 (4), 635-646 (2005).</li><li>Mehta, S. Q. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Mutations+in+Drosophila+sec15+reveal+a+function+in+neuronal+targeting+for+a+subset+of+exocyst+components.">Mutations in Drosophila sec15 reveal a function in neuronal targeting for a subset of exocyst components.</a> Neuron. 46 (2), 219-232 (2005).</li></ol>

用于成像的荧光探针在这里，我们描述了果蝇视觉系统准备从三个类荧光探针的选择的长处和局限性：（1）探针被添加到一个活组织编制外生的;（2）现场和基因编码的荧光蛋白固定成像;（3）添加到固定的免疫组织化学组织的荧光染料。 1。外感探头： Lysotracker绿色免打扰- 26或红DND - 99（Invitrogen公司，公司），这些均为市售透水膜的荧光化合物积累在细胞中，最突出的溶酶体强烈酸化车厢，后期内涵体和自噬。在纳摩尔浓度（我们通常使用50纳米）Lysotracker完全渗透在不到一分钟的L3和P +20％眼光盘。由于Lysotracker不断积累，对碱性的影响，我们的形象，在不到5分钟。相反的眼睛光盘，Lysotracker不渗透不破坏组织大脑。然而，细心的外膜撕裂允许几个细胞直径在视神经叶的渗透。 Lysosensor DND - 189对比Lysotracker（Invitrogen公司，公司），Lysosensor不会积聚在酸化车厢，但在酸性的展品增加荧光。 Lysosensor已渗透的特点是非常相似Lysotracker的。我们使用浓度为1μmLysosensor。 2。基因编码的探头： 对于许多实时成像实验中，我们表达GFP，TagRFP 10或pH敏感pHluorin 11的二进制Gal4/UAS表达系统 12控制下的各种荧光分子标记的基因。 GAL4 -线，在发展中的光感受器表达包括GMR - Gal4的（感光器）和mDelta0.5 Gal4的（R4和R7的开发）13。几个亚型特异性Gal4的驱动线存在使用不同的视紫红质的推动者14。特别有用的荧光探针实时成像photoconvertible蛋白质。我们已经成功地使用Dendra2，单体绿色到红色photoconvertable蛋白质15。请注意，Dendra2使用488nm氩激光线的设计是photoconvertible。然而，我们无法实现转换，使用蓝色可见光，无论是传统的或共振的共焦扫描模式设置。相比之下，405纳米的紫外激光的转换效率。 3。免疫组化： 对于固定的成人的眼睛，我们往往会选择罗丹明鬼笔环肽（Invitrogen公司，公司）或抗Chaoptin（一个光感受器特异性抗体 16）以可视化rhabdomeric的结构（图1A ）。分析椎板墨盒结构的一个好方法是要结合反chaoptin 16，胶质标记反乌梅17，和反sec6 18（图1B）。如图所示。 3，这种结合的抗体区分的（chaoptin）主要突触前和突触后（sec6）进程以及上皮细胞的神经胶质细胞中的19层（乌木）。 组装灌注腔实时成像，我们采用了灌注腔慢交换的解决方案，允许在不扰民的组织准备。灌注室组装示范，包括视频和示意图如图2所示。大会灌注室如图所示。 2A。开始铺设垫片首先在试样，然后继续与组装，如图所示。垫片的目的是创建一个室的基础和盖子，在里面的组织是密封的空间，并通过该解决方案将流之间的空间。空间内的垫片必须包括入口（这里的解决方案进入），组织准备，并通过该解决方案离开会议厅的插座。垫片的两个特性是至关重要的：首先，垫圈必须足够厚的大脑还薄，足以让一个高分辨率的镜头成像。理想的垫片的厚度取决于设置特定的参数。其次，在垫片的缺口提供了一个隔间，让泡沫，通过无接触标本（图2B）。 显微镜的成像我们用一个63X（光圈1.3）的甘油浸没透镜灌注室或直接进入幻灯片上的培养基中一个63X的水浸泡镜头。甘油的镜头提供多油镜头的工作距离。我们使用一个共振扫描共聚焦显微镜扫描快得多的速度比传统的扫描仪（8000赫兹与1400赫兹），分别。共振扫描可随着时间的推移的快速三维录音ER帧速率。此外，更快的扫描速度显着降低激光光毒性6这是一个主要关注的活组织反复扫描。激光强度和PMT电压（增益）的平衡，必须实现最大化的信号，同时最大限度地降低了噪声和漂白。一个重要的进一步审议，激光影响某一区域上的金额是由分辨率和变焦决定。例如，选择的地区是在相同的分辨率在256x256的，变焦2X 1X变焦，512 × 512扫描，但最大可能的实时成像速度快4倍，在256 × 256，放大2倍。为了记录快速移动的细胞内的车厢的活动，我们已经成功地记录了以下设置：在一率5％的Z - Stack帧，每10秒的Z - Stack与8000赫兹和2X线平均扫描速度。对于长成像会议上的小时刻度，我们通常会降低帧速率以每几分钟，选择面积较大的地区，准备更可能转向。 果蝇视觉系统解剖学图3显示了解剖果蝇成人眼（图3A），成人的大脑（图3B），以及20％-25％的蛹眼盘/脑复杂（图3C）。图3A显示椎板和附着的脂肪细胞和无成人的眼睛。在成人眼清扫术，椎板，依偎在由感光细胞机构（左）组成的一个碗眼的中心，必须删除，而不会干扰下（右）的胞体。成人大脑解剖，应该能够分辨感光细胞组织，椎板，视叶，中央的大脑，和气管（图3B）。需要删除，同时使视神经叶椎板剥离的光感受器和气管。为了帮助确定在20-25％的蛹眼盘/大脑复杂，是一个实时影像图3C。 <img src="/files/ftp_upload/1936/1936fig1.jpg" alt="图1" /> 图1（A）成人的大脑。 （二）成人眼。 （三）20％-25％蛹eye-disc/brain复杂。 <img src="/files/ftp_upload/1936/1936fig2.jpg" alt="图2" /> 图2：（一）灌注室大会和（二）垫片定位相对标本。 <img src="/files/ftp_upload/1936/1936fig3.jpg" alt="图3" /> 图3：（一）成人rhabdomere染色用抗chaoptin 。 （二）使用反chaoptin（绿色，感光器），反乌梅（红色，神经胶质细胞），并sec6（蓝色，interneurons）成人椎板染色。

专用设备： <ol><li>夹层范围：徕卡数码相机Mz12.5 </li><li>徕卡TCS SP5的共焦串联扫描仪传统和共振扫描显微镜直播 成像显微镜，包括蓝（488nm），绿（561nm），橙色（594nm）和远红光（633纳米）激光器以及特殊的63X，1.4甘油和63X，1.3水浸泡镜片。 </li><li>哈佛仪器共焦成像商会RC - 30和蠕动泵型号720 </li><li>萨特移液器拉马的P - 97 </li><li> 5.2软件：阿米拉，Visage的影像</li></ol>

preparation of developing and adult drosophila brains and retinae for live imaging

“<em&gt;果蝇</em&gt;大脑和视觉系统广泛使用的模型系统来研究神经元的发育，功能和变性。在这里，我们展示三个准备的大脑和视觉系统，涵盖了从发展中国家眼睛光盘大脑复杂的发展中蛹个人从成蝇的眼睛和大脑清扫范围。所有协议优化生活文化的准备。但是，我们也存在固定组织的免疫组织化学法在适用的条件。最后，我们这些准备在灌注室使用传统和谐振4D现场共焦成像显示实时成像条件。总之，这些协议提供了在不同的时间尺度，从规模分钟多小时的规模发展或退化过程的功能的细胞内检测实时成像的基础上。

Watch this Scientific Journal Video about Preparation of Developing and Adult Drosophila Brains and Retinae for Live Imaging at JoVE.com

Preparation of Developing and Adult Drosophila Brains and Retinae for Live Imaging

本协议描述了三个<em&gt;果蝇</em&gt;准备工作：1）成人大脑清扫，2）成人视网膜剥离和3）发展中国家眼睛光盘大脑复合物清扫。重点是放在特殊的制备技术和条件，实时成像，可用于固定组织的免疫组织化学，但所有的准备工作。

发展和成人的制备果蝇大脑和视网膜实时成像

The Drosophila brain and visual system are widely utilized model systems to study neuronal development, function and degeneration. Here we show three ...

preparation-developing-adult-drosophila-brains-retinae-for-live

Research

JoVE Journal

Biology

Preparation of Developing and Adult Drosophila Brains and Retinae for Live Imaging

35.2K 次观看.  University of Texas Southwestern Medical Center. 本协议描述了三个果蝇准备工作：1）成人大脑清扫，2）成人视网膜剥离和3）发展中国家眼睛光盘大脑复合物清扫。重点是放在特殊的制备技术和条件，实时成像，可用于固定组织的免疫组织化学，但所有的准备工作。

视频: Preparation of Developing and Adult Drosophila Brains and Retinae for Live Imaging

Primary Neuronal Cultures from the Brains of Late Stage Drosophila Pupae

In this video, we demonstrate the preparation of primary neuronal cultures from the brains of late stage Drosophila pupae. The procedure begins with the removal of brains from animals at 70-78 hrs after puparium formation. The isolated brains are shown after brief incubation in papain followed by several washes in serum-free growth medium. The process of mechanical dissociation of each brain in a 5 ul drop of media on a coverslip is illustrated. The axons and dendrites of the post-mitotic neurons are sheered off near the soma during dissociation but the neurons begin to regenerate processes within a few hours of plating. Images show live cultures at 2 days. Neurons continue to elaborate processes during the first week in culture. Specific neuronal populations can be identified in culture using GAL4 lines to drive tissue specific expression of fluorescent markers such as  GFP or RFP. Whole cell recordings have demonstrated the cultured neurons form functional, spontaneously active cholinergic and GABAergic synapses. A short video segment illustrates calcium dynamics in the cultured neurons using Fura-2 as a calcium indicator dye to monitor spontaneous calcium transients and nicotine evoked calcium responses in a dish of cultured neurons. These pupal brain cultures are a useful model system in which genetic and pharmacological tools can be used to identify intrinsic and extrinsic factors that influence formation and function of central synapses.

This video demonstrates the preparation of primary neuronal cultures from the brains of late stage Drosophila pupae. Views of live cultures show neurite outgrowth and imaging of calcium levels using Fura-2.

从晚期果蝇蛹的大脑原发性神经文化

In this video, we demonstrate the preparation of primary neuronal cultures from the brains of late stage Drosophila pupae. The procedure begins with the ...

科研

生物学

The Preparation of Drosophila Embryos for Live-Imaging Using the Hanging Drop Protocol

Green fluorescent protein (GFP)-based timelapse live-imaging is a powerful technique for studying the genetic regulation of dynamic processes such as tissue morphogenesis, cell-cell adhesion, or cell death. Drosophila embryos expressing GFP are readily imaged using either stereoscopic or confocal microscopy. A goal of any live-imaging protocol is to minimize detrimental effects such as dehydration and hypoxia. Previous protocols for preparing Drosophila embryos for live-imaging analysis have involved placing dechorionated embryos in halocarbon oil and sandwiching them between a halocarbon gas-permeable membrane and a coverslip1-3. The introduction of compression through mounting embryos in this manner represents an undesirable complication for any biomechanical-based analysis of morphogenesis. Our method, which we call the hanging drop protocol, results in excellent viability of embryos during live imaging and does not require that embryos be compressed. Briefly, the hanging drop protocol involves the placement of embryos in a drop of halocarbon oil that is suspended from a coverslip, which is, in turn, fixed in position over a humid chamber. In addition to providing gas exchange and preventing dehydration, this arrangement takes advantage of the buoyancy of embryos in halocarbon oil to prevent them from drifting out of position during timelapse acquisition. This video describes in detail how to collect and prepare Drosophila embryos for live imaging using the hanging drop protocol. This protocol is suitable for imaging dechorionated embryos using stereomicroscopy or any upright compound fluorescence microscope.

The Preparation of Drosophila Embryos for Live-Imaging Using the Hanging Drop Protocol

A simple, inexpensive, and effective method of preparing Drosophila embryos for live-imaging analysis is presented. Our protocol provides humidity and gas exchange and does not compress the Drosophila embryo. This method is suitable for GFP-based live imaging of Drosophila embryos using a stereomicroscope or upright compound microscope.

编制果蝇实时成像的胚胎使用悬滴协议

Green fluorescent protein (GFP)-based timelapse live-imaging is a powerful technique for studying the genetic regulation of dynamic processes such as ...

Mesoscopic Fluorescence Tomography for In-vivo Imaging of Developing Drosophila

Visualizing developing organ formation as well as progession and treatment of disease often heavily relies on the ability to optically interrogate molecular and functional changes in intact living organisms. Most existing optical imaging methods are inadequate for imaging at dimensions that lie between the penetration limits of modern optical microscopy (0.5-1mm) and the diffusion-imposed limits of optical macroscopy (&gt;1cm) [1]. Thus, many important model organisms, e.g. insects, animal embryos or small animal extremities, remain inaccessible for in-vivo optical imaging. 
Although there is increasing interest towards the development of nanometer-resolution optical imaging methods, there have not been many successful efforts in improving the imaging penetration depth. The ability to perform in-vivo imaging beyond microscopy limits is in fact met with the difficulties associated with photon scattering present in tissues. Recent efforts to image entire embryos for example [2,3] require special chemical treatment of the specimen, to clear them from scattering, a procedure that makes them suitable only for post-mortem imaging. These methods however evidence the need for imaging larger specimens than the ones usually allowed by two-photon or confocal microscopy, especially in developmental biology and in drug discovery.
We have developed a new optical imaging technique named Mesoscopic Fluorescence Tomography [4], which appropriate for non-invasive in-vivo imaging at dimensions of 1mm-5mm. The method exchanges resolution for penetration depth, but offers unprecedented tomographic imaging performance and it has been developed to add time as a new dimension in developmental biology observations (and possibly other areas of biological research) by imparting the ability to image the evolution of fluorescence-tagged responses over time. As such it can accelerate studies of morphological or functional dependencies on gene mutations or external stimuli, and can importantly, capture the complete picture of development or tissue function by allowing longitudinal time-lapse visualization of the same, developing organism.
The technique utilizes a modified laboratory microscope and multi-projection illumination to collect data at 360-degree projections. It applies the Fermi simplification to Fokker-Plank solution of the photon transport equation, combined with geometrical optics principles in order to build a realistic inversion scheme suitable for mesoscopic range. This allows in-vivo whole-body visualization of non-transparent three-dimensional structures in samples up to several millimeters in size.
We have demonstrated the in-vivo performance of the technique by imaging three-dimensional structures of developing Drosophila tissues in-vivo and by following the morphogenesis of the wings in the opaque Drosophila pupae in real time over six consecutive hours.

Mesoscopic Fluorescence Tomography for In-vivo Imaging of Developing Drosophila

Mesoscopic fluorescence tomography operates beyond the penetration limits of tissue-sectioning fluorescence microscopy. The technique is based on multi-projection illumination and a photon transport description. We demonstrate in-vivo whole-body 3D visualization of the morphogenesis of GFP-expressing wing imaginal discs in Drosophila melanogaster.

为介观荧光体层摄影术在体内成像的发展果蝇</em

Visualizing  developing organ formation as well as progession and treatment of disease often  heavily relies on the ability to optically interrogate ...

Live Imaging Of Drosophila melanogaster Embryonic Hemocyte Migrations

Many studies address cell migration using in vitro methods, whereas the physiologically relevant environment is that of the organism itself. Here we present a protocol for the mounting of Drosophila melanogaster embryos and subsequent live imaging of fluorescently labeled hemocytes, the embryonic macrophages of this organism. Using the Gal4-uas system1 we drive the expression of a variety of genetically encoded, fluorescently tagged markers in hemocytes to follow their developmental dispersal throughout the embryo. Following collection of embryos at the desired stage of development, the outer chorion is removed and the embryos are then mounted in halocarbon oil between a hydrophobic, gas-permeable membrane and a glass coverslip for live imaging. In addition to gross migratory parameters such as speed and directionality, higher resolution imaging coupled with the use of fluorescent reporters of F-actin and microtubules can provide more detailed information concerning the dynamics of these cytoskeletal components.

Live Imaging Of Drosophila melanogaster Embryonic Hemocyte Migrations

Drosophila hemocytes disperse over the entirety of the developing embryo. This protocol demonstrates how to mount and image these migrations using embryos with fluorescently labelled hemocytes.

实时成像果蝇胚胎血细胞迁移

Many studies address cell migration using in vitro methods, whereas the physiologically relevant environment is that of the organism itself. Here we ...

Paraffin-Embedded and Frozen Sections of Drosophila Adult Muscles

The molecular characterization of muscular dystrophies and myopathies in humans has revealed the complexity of muscle disease and genetic analysis of muscle specification, formation and function in model systems has provided valuable insight into muscle physiology. Therefore, identifying and characterizing molecular mechanisms that underlie muscle damage is critical. The structure of adult Drosophila multi-fiber muscles resemble vertebrate striated muscles 1 and the genetic tractability of Drosophila has made it a great system to analyze dystrophic muscle morphology and characterize the processes affecting muscular function in ageing adult flies 2. Here we present the histological technique for preparing paraffin-embedded and frozen sections of Drosophila thoracic muscles. These preparations allow for the tissue to be stained with classical histological stains and labeled with protein detecting dyes, and specifically cryosections are ideal for immunohistochemical detection of proteins in intact muscles. This allows for analysis of muscle tissue structure, identification of morphological defects, and detection of the expression pattern for muscle/neuron-specific proteins in Drosophila adult muscles. These techniques can also be slightly modified for sectioning of other body parts.

Paraffin-Embedded and Frozen Sections of Drosophila Adult Muscles

Identification of mechanisms underlying muscle damage is crucial. Here we present the histological technique for preparing paraffin-embedded and frozen sections of Drosophila thoracic muscles. This allows analysis of muscle morphology and localization of protein and other muscle cell components.

石蜡包埋及冷冻的部分果蝇成人肌肉

The molecular characterization of muscular dystrophies and myopathies in humans has revealed the complexity of muscle disease and genetic analysis of ...

Live Imaging of Cell Extrusion from the Epidermis of Developing Zebrafish

Homeostatic maintenance of epithelial tissues requires the continual removal of damaged cells without disrupting barrier function. Our studies have found that dying cells send signals to their live neighbors to form and contract a ring of actin and myosin that ejects it out from the epithelial sheet while closing any gaps that might have resulted from its exit, a process termed cell extrusion1. The optical clarity of developing zebrafish provides an excellent system to visualize extrusion in living epithelia. Here we describe a method to induce and image extrusion in the larval zebrafish epidermis. To visualize extrusion, we inject a red fluorescent protein labeled probe for F-actin into one-cell stage transgenic zebrafish embryos expressing green fluorescent protein in the epidermis and induce apoptosis by addition of G418 to larvae. We then use time-lapse imaging on a spinning disc confocal microscope to observe actin dynamics and epithelial cell behaviors during the process of apoptotic cell extrusion. This approach allows us to investigate the extrusion process in live epithelia and will provide an avenue to study disease states caused by the failure to eliminate apoptotic cells.

Dying cells are extruded from epithelial tissues by concerted contraction of neighboring cells without disrupting barrier function. The optical clarity of developing zebrafish provides an excellent system to visualize extrusion in living epithelia. Here we describe methods to induce and image extrusion in the larval zebrafish epidermis at cellular resolution.

发展中国家斑马鱼的表皮细胞挤压实时成像

Homeostatic maintenance of epithelial tissues requires the continual removal of damaged cells without disrupting barrier function.  Our studies have found ...

Preparation of Adult Drosophila Eyes for Thin Sectioning and Microscopic Analysis

Drosophila has long been used as model system to study development, mainly due to the ease with which it is genetically tractable. Over the years, a plethora of mutant strains and technical tricks have been developed to allow sophisticated questions to be asked and answered in a reasonable amount of time. Fundamental insight into the interplay of components of all known major signaling pathways has been obtained in forward and reverse genetic Drosophila studies. The fly eye has proven to be exceptionally well suited for mutational analysis, since, under laboratory conditions, flies can survive without functional eyes. Furthermore, the surface of the insect eye is composed of some 800 individual unit eyes (facets or ommatidia) that form a regular, smooth surface when looked at under a dissecting microscope. Thus, it is easy to see whether a mutation might affect eye development or growth by externally looking for the loss of the smooth surface ('rough eye' phenotype; Fig. 1) or overall eye size, respectively (for examples of screens based on external eye morphology see e.g.1). Subsequent detailed analyses of eye phenotypes require fixation, plastic embedding and thin-sectioning of adult eyes.
The Drosophila eye develops from the so-called eye imaginal disc, a bag of epithelial cells that proliferate and differentiate during larval and pupal stages (for review see e.g. 2). Each ommatidium consists of 20 cells, including eight photoreceptors (PR or R-cells; Fig. 2), four lens-secreting cone cells, pigment cells ('hexagon' around R-cell cluster) and a bristle. The photoreceptors of each ommatidium, most easily identified by their light sensitive organelles, the rhabdomeres, are organized in a trapezoid made up of the six "outer" (R1-6) and two "inner" photoreceptors (R7/8; R8 [Fig. 2] is underneath R7 and thus only seen in sections from deeper areas of the eye). The trapezoid of each facet is precisely aligned with those of its neighbors and the overall anteroposterior and dorsoventral axes of the eye (Fig. 3A). In particular, the ommatidia of the dorsal and ventral (black and red arrows, respectively) halves of the eye are mirror images of each other and correspond to two chiral forms established during planar cell polarity signaling (for review see e.g. 3).
The method to generate semi-thin eye sections (such as those presented in Fig. 3) described here is slightly modified from the one originally described by Tomlinson and Ready4. It allows the morphological analysis of all cells except for the transparent cone cells. In addition, the pigment of R-cells (blue arrowheads in Fig. 2 and 3) can be used as a cell-autonomous marker for the genotype of a R-cell, thus genetic requirements of genes in a subset of R-cells can readily be determined5,6.

Preparation of Adult Drosophila Eyes for Thin Sectioning and Microscopic Analysis

A standard approach to prepare adult Drosophila eyes for semi-thin sectioning and light microscopic analysis is presented here. The protocol can be used for gross morphological analysis of eye defects, or with the indicated adjustments can be used to determine genetic requirements of genes in specific cell types of the eye (e.g. clonal analysis of photoreceptors) or for electron microscopic analysis.

成人的制备果蝇</em薄切片和显微镜分析>的眼睛

Drosophila has long been used as model system to study development, mainly due to the ease with which it is genetically tractable. Over the years, 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, ...

Live Imaging of Apoptotic Cell Clearance during Drosophila Embryogenesis

The proper elimination of unwanted or aberrant cells through apoptosis and subsequent phagocytosis (apoptotic cell clearance) is crucial for normal development in all metazoan organisms. Apoptotic cell clearance is a highly dynamic process intimately associated with cell death; unengulfed apoptotic cells are barely seen in vivo under normal conditions. In order to understand the different steps of apoptotic cell clearance and to compare 'professional' phagocytes - macrophages and dendritic cells to 'non-professional' - tissue-resident neighboring cells, in vivo live imaging of the process is extremely valuable. Here we describe a protocol for studying apoptotic cell clearance in live Drosophila embryos. To follow the dynamics of different steps in phagocytosis we use specific markers for apoptotic cells and phagocytes. In addition, we can monitor two phagocyte systems in parallel: 'professional' macrophages and 'semi-professional' glia in the developing central nervous system (CNS). The method described here employs the Drosophila embryo as an excellent model for real time studies of apoptotic cell clearance.

Live Imaging of Apoptotic Cell Clearance during Drosophila Embryogenesis

Here we describe an effective method for studying dynamics of apoptotic cell clearance in vivo. This method employs live Drosophila embryos as a powerful model for monitoring phagocytosis of apoptotic cells using specific labeling of apoptotic cells and phagocytes.

凋亡细胞清除过程中实时成像<em&gt;果蝇</em&gt;胚胎发育

The  proper elimination of unwanted or aberrant cells through apoptosis and  subsequent phagocytosis (apoptotic cell clearance) is crucial for normal  ...

The Tomato/GFP-FLP/FRT Method for Live Imaging of Mosaic Adult Drosophila Photoreceptor Cells

The Drosophila eye is widely used as a model for studies of development and neuronal degeneration. With the powerful mitotic recombination technique, elegant genetic screens based on clonal analysis have led to the identification of signaling pathways involved in eye development and photoreceptor (PR) differentiation at larval stages. We describe here the Tomato/GFP-FLP/FRT method, which can be used for rapid clonal analysis in the eye of living adult Drosophila. Fluorescent photoreceptor cells are imaged with the cornea neutralization technique, on retinas with mosaic clones generated by flipase-mediated recombination. This method has several major advantages over classical histological sectioning of the retina: it can be used for high-throughput screening and has proved an effective method for identifying the factors regulating PR survival and function. It can be used for kinetic analyses of PR degeneration in the same living animal over several weeks, to demonstrate the requirement for specific genes for PR survival or function in the adult fly. This method is also useful for addressing cell autonomy issues in developmental mutants, such as those in which the establishment of planar cell polarity is affected.

The Tomato/GFP-FLP/FRT Method for Live Imaging of Mosaic Adult Drosophila Photoreceptor Cells

The Tomato/GFP-FLP/FRT method involves visualizing mosaic photoreceptor cells in living Drosophila. It can be used to follow individual photoreceptor cell fates in the retina for days or weeks. This method is ideal for studies of retinal degeneration and neurodegenerative diseases or photoreceptor cell development.

番茄/ GFP-FLP / FRT方法马赛克成人的实时成像<em&gt;果蝇</em&gt;感光细胞

The Drosophila eye is widely used as a model  for studies of development and neuronal degeneration. With the powerful mitotic  recombination technique, ...