Name: 基因功能分析和可视化的纤毛产生的流体流动的在枯否囊泡中 (视频) | JoVE Journal
Uploaded: 2013-03-31T14:00:00.000+00:00
Duration: 8 min 11 s
Description: 学习在斑马鱼胚胎枯否囊泡中调控基因功能并可视化纤毛产生的流体流动的方法。该技术利用荧光微珠和显微成像定量分析不对称流体动力学，为研究左右不对称模式形成提供重要工具。

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We thank Fiona Foley for excellent lab support and zebrafish care. This work was supported an AHA predoctoral fellowship to G.W. (11PRE5730027) and NHLBI grants to H.J.Y. (R01HL66292) and J.D.A. (R01HL095690).

Overview of Stage-Specific Zebrafish Embryo Injections
Antisense morpholino oligonucleotides (MO), which bind to a targeted mRNA and disrupt protein expression from that transcript, are widely used in gene knockdown (loss-of-function) studies in zebrafish13,14. Gene Tools, LLC offers MOs that are tagged with either carboxyfluorescein (emits green fluorescence) or lissamine (emits red fluorescence) to detect MO in injected embryos using fluorescent microscopy. By injecting MO into the yolk cell at different stages of zebrafish development, it is possible to deliver the MO to specific compartments of the embryo (Figures 1A-F). MO injected into the yolk between the 1-4 cell stages (0-1 hpf) enters all embryonic cells (Figures 1A, D, D') via connections with the yolk that persist until the 32-cell stage15 to facilitate global knockdown. MO injected into the yolk during midblastula stages (2.5-3 hpf) can enter the progenitors of the DFCs (Figures 1B, E, E') likely through cytoplasmic bridges8 and knockdown gene function specifically in the DFC/KV cell lineage12, without entering most other embryonic cell lineages. As an important control to test whether gene function is required in DFC/KV or also in yolk12,16, it is also possible to restrict MO to the yolk cell by injecting between the dome-30% epiboly stages (~4.5 hpf) after all cytoplasmic bridges have closed (Figures 1C, F, F'). These injections have been used in combination to analyze gene function in DFC/KV cells17-24. To assess fluid flow in KV, fluorescent microbeads are injected into the KV lumen between the 6-10 somite stages (12-14 hpf) and then immediately imaged using videomicroscopy (Figures 1G-J). Microbeads are available that emit red or green fluorescence (or both), so it is possible to use different channels to image fluorescent microbeads and MO.
1. Stage-specific Injection of Morpholinos (MO)
Injection of MO into zebrafish embryos has been demonstrated previously25,26. Here, we briefly describe stage-specific MO injections. Following all injections, embryos are transferred to a Petri dish and incubated at 28.5 &deg;C.
<ol>
	<li>Global MO injection: Collect embryos immediately after fertilization and load them into an injection plate. Load fluorescent MO into a capillary needle and mount the needle onto a microinjector (e.g. Harvard Apparatus PLI-90 Pico-injector). Break the needle tip and adjust injection settings (pressure and/or time) to create an injection drop that has a volume of 1 nl as described25,26.Using a dissecting stereomicroscope, inject 1 nl of MO into the yolk (Figure 1A) of &ge;50 embryos per experiment. Work quickly to complete injections by the 4-cell stage.</li>
	<li>DFC-targeted MO injection: Collect embryos immediately after fertilization and incubate at 28.5 &deg;C. When the embryos have reached the 256-cell stage (~2.5 hpf), quickly load them into an injection plate and then inject 1 nl of fluorescent MO into the yolk of &ge;100 embryos between the 256-cell and 1,000-cell stages (Figure 1B).</li>
	<li>Yolk-targeted MO injection: Collect embryos immediately after fertilization and incubate at 28.5 &deg;C. At the dome stage (~4 hpf), load the embryos into an injection plate and then inject 1 nl of fluorescent MO in the yolk of &ge;100 embryos between the dome and 30% epiboly stages (Figure 1C).</li>
</ol>
2. Selecting Injected Embryos for Analysis
<ol>
	<li>When MO injected embryos reach the 75%-epiboly stage (8 hpf), remove unfertilized and dead embryos and then screen living embryos under a fluorescent dissecting microscope for distribution of the fluorescent MO. Then allow selected embryos to develop at 28.5 &deg;C.
	<ol>
		<li>For global MO injections, select embryos that have fluorescence evenly distributed in throughout all embryonic cells (Figure 1D; Figure 2A).</li>
		<li>For DFC-targeted MO injections, select embryos in which fluorescent MO has diffused throughout the yolk and appears concentrated at the dorsal blastoderm margin (DFCs) (Figure 1E; Figure 2B). At this stage, it can be difficult to visualize fluorescent DFCs due to bright fluorescence in the underlying yolk. Take care to exclude embryos in which the fluorescent MO has incorporated into embryonic cells other than DFCs or remains aggregated at the injection site (Figure 2D).</li>
		<li>For yolk-targeted MO injections, select embryos in which MO fluorescence is evenly distributed throughout the yolk and not observed in any embryonic cells (Figure 1F;Figure 2C). Again, remove embryos in which the fluorescent MO remains aggregated at the injection site.</li>
	</ol>
	</li>
	<li>Between the 2-4-somite stages (~11 hpf), screen embryos a second time under a fluorescent microscope using higher magnification. Then allow selected embryos to develop at 28.5 &deg;C.
	<ol>
		<li>For global MO injections, ensure selected embryos have MO fluorescence evenly distributed in KV and all embryonic cells (Figure 1D'; Figure 2E).</li>
		<li>For DFC-targeted MO injections, carefully select embryos that have MO fluorescence only in KV cells and the yolk (Figure 1E'). Transgenic embryos that express GFP in KV cells, such as Tg(Dusp6:d2EGFP)27, can aid in the identification of embryos in which MO has been successfully delivered to KV cells (Figure 2F). Discard embryos with MO fluorescence in embryonic cells other that KV cells.</li>
		<li>For yolk-targeted MO injections, select embryos that have MO fluorescence exclusively in the yolk (Figure 1F': Figure 2G).</li>
	</ol>
	</li>
</ol>
3. Mounting Embryos to Analyze Fluid Flow in KV
<ol>
	<li>To analyze asymmetric fluid flow in KV of global MO, DFC-targeted MO or yolk-targeted MO injected embryos, carefully dechorionate ~20 embryos between 4-6 somite stages (~11-12 hpf) with sharp forceps.</li>
	<li>Prepare 50 ml of 1% low-melting point agarose and aliquot into 5 ml tubes to be maintained as a liquid in a dry bath at 42 &deg;C.</li>
	<li>Transfer one embryo to a glass depression slide (United Scientific Supplies, Inc.) and remove as much of the water as possible. Immobilize the embryo by adding enough warm 1% low-melting point agarose to just cover the embryo (too much agarose can interfere with subsequent imaging). As the agarose solidifies, use forceps to position the embryo such that KV is facing up (dorsal view) (Figure 1H). Mount 10 embryos with one embryo per slide. Work quickly to ensure all injections in the next step are completed by the 10 somite stage (14 hpf).</li>
</ol>
4. Injection of Fluorescent Microbeads into KV
<ol>
	<li>Dilute Fluoresbrite Multifuorescent 0.5 micron diameter Microspheres (Polysciences, Inc.) to 1:50 in sterile water in a 1.5 ml Eppendorf tube.</li>
	<li>Mix the 1:50 dilution of microbeads thoroughly by tapping the tube and load 3 &mu;l into a capillary needle. Using the same microinjector used for MO injections (e.g. Harvard Apparatus PLI-90 Pico-injector), break the needle tip with forceps and adjust the injection settings to generate the smallest possible (&lt;0.5 nl) injection drop.</li>
	<li>Under a dissecting microscope, align the needle with the KV lumen of the first mounted embryo. Insert the needle into the lumen and inject a small volume (&lt; 0.5 nl) of beads (Figure 1G). A small needle tip and small injection volume are critical to avoid damaging KV.</li>
	<li>Inject all mounted embryos. Once an embryo has been injected, a drop of sterile water can be added on top of the agarose to prevent it from drying out. During the course of the injections, the beads will often obstruct the needle opening. This requires re-breaking the needle tip and adjusting the injection volume by modulating the pressure or time setting of the microinjection apparatus. Replace the needle if it becomes blunt enough to cause significant damage during injection.</li>
	<li>Screen the injected embryos for successful delivery of beads under an upright fluorescent compound microscope (e.g. Zeiss AxioImager M1) using a 20X objective. First, determine whether KV structure is intact using brightfield illumination (Figure 1J), and then use fluorescence to observe the beads. Select embryos in which beads are present and moving inside the KV lumen. Discard embryos with KV damage or no floating beads in KV. It is easy to miss KV and deliver beads to nearby tissue or the underlying yolk. If unsuccessful, mount another 10 embryos and repeat the injection procedure.</li>
</ol>
5. Visualization and Analysis of KV Fluid Flow
<ol>
	<li>For each selected embryo with beads inside KV, add a drop of sterile water to cover the agarose and then observe KV under the upright fluorescent compound microscope using a 63X water dipping objective (Figure 1I). Alternatively, a coverslip can be applied for use with non-water dipping objectives and/or inverted microscopes.</li>
	<li>Use a high-speed camera mounted on the microscope (e.g. Zeiss AxioCamHSm) to record a 10 sec movie. Record both the beads using the fluorescent channel (approximately 70 frames/sec) and the KV lumen using a brightfield or differential interference contrast (DIC) channel.</li>
	<li>To visualize all bead movements over time, import the movie of fluorescent beads into ImageJ software (free download at <a href="http://rsb.info.nih.gov/ij/" target="_blank">http://rsb.info.nih.gov/ij/</a>) and create a maximum projection of all fluorescent signals in the movie. To perform the maximum projection in ImageJ, click "Image&rarr;Stacks&rarr; Z project." In the "Z project" window, project all slices with projection type of "Max intensity". This image can be superimposed on a DIC image of the KV in which the beads were imaged (Figures 3A-B).</li>
	<li>The movements of individual beads can be tracked using ImageJ. A plugin ("Manual Tracking") can be downloaded at <a href="http://rsbweb.nih.gov/ij/plugins/track/track.html" target="_blank">http://rsbweb.nih.gov/ij/plugins/track/track.html</a>. Open the fluorescent beads movie in ImageJ and run the Manual Tracking function. In the window of Manual Tracking, check "show parameters" and input parameters for"time intervals" and "x/y calibration". Manually select beads that remain in the focal plane of the movie for &ge;50 frames and then track &ge;5 beads per embryo. The path and velocity of each bead is generated by the software (Figures 3 C-D).</li>
</ol>

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</ol>

No conflicts of interest declared.

Using stage-specific injections to target MO to the DFC/KV cell lineage is a useful approach to study cell-autonomy of gene function and avoid pleiotropic phenotypes caused by global gene knockdown. However, these injections can be technically challenging. Injection of MO between the 256-cell and 1,000-cell stages can result in three possible outcomes: 1) the MO remains aggregated at the injection site, 2) the MO diffuses throughout the yolk and enters DFC/KV cells or 3) the MO diffuses throughout the yolk and enters DFC...

<table><tbody><tr><td>Standard Control oligo-Lissamine tagged </td><td>Gene Tools, LLC</td><td></td><td></td></tr><tr><td>Custom Rock2b morpholino oligo</td><td>Gene Tools, LLC</td><td></td><td></td></tr><tr><td>Fluoresbrite Multifluorescent 0.5 micron Microspheres</td><td>Polysciences, Inc.</td><td>24054</td><td></td></tr></tbody></table>

analysis of gene function and visualization of cilia-generated fluid flow in kupffer's vesicle

Analysis of Gene Function and Visualization of Cilia-Generated Fluid Flow in Kupffer's Vesicle

Internal organs such as the heart, brain, and gut develop left-right (LR) asymmetries that are critical for their normal functions1. Motile cilia are involved in establishing LR asymmetry in vertebrate embryos, including mouse, frog, and zebrafish2-6. These 'LR cilia' generate asymmetric fluid flow that is necessary to trigger a conserved asymmetric Nodal (TGF-&beta; superfamily) signaling cascade in the left lateral plate mesoderm, which is thought to provide LR patterning information for developing organs7. Thus, to understand mechanisms underlying LR patterning, it is essential to identify genes that regulate the organization of LR ciliated cells, the motility and length of LR cilia and their ability to generate robust asymmetric flow.
In the zebrafish embryo, LR cilia are located in Kupffer's vesicle (KV)2,4,5. KV is comprised of a single layer of monociliated epithelial cells that enclose a fluid-filled lumen. Fate mapping has shown that KV is derived from a group of ~20-30 cells known as dorsal forerunner cells (DFCs) that migrate at the dorsal blastoderm margin during epiboly stages8,9. During early somite stages, DFCs cluster and differentiate into ciliated epithelial cells to form KV in the tailbud of the embryo10,11. The ability to identify and track DFCs&mdash;in combination with optical transparency and rapid development of the zebrafish embryo&mdash;make zebrafish KV an excellent model system to study LR ciliated cells.
Interestingly, progenitors of the DFC/KV cell lineage retain cytoplasmic bridges between the yolk cell up to 4 hr post-fertilization (hpf), whereas cytoplasmic bridges between the yolk cell and other embryonic cells close after 2 hpf8. Taking advantage of these cytoplasmic bridges, we developed a stage-specific injection strategy to deliver morpholino oligonucleotides (MO) exclusively to DFCs and knockdown the function of a targeted gene in these cells12. This technique creates chimeric embryos in which gene function is knocked down in the DFC/KV lineage developing in the context of a wild-type embryo. To analyze asymmetric fluid flow in KV, we inject fluorescent microbeads into the KV lumen and record bead movement using videomicroscopy2. Fluid flow is easily visualized and can be quantified by tracking bead displacement over time.
Here, using the stage-specific DFC-targeted gene knockdown technique and injection of fluorescent microbeads into KV to visualize flow, we present a protocol that provides an effective approach to characterize the role of a particular gene during KV development and function.

Stage-specific MO injections provide a useful approach to analyze gene function in specific compartments of the embryo. Figure 1 presents a flow chart of the injection strategies used to test gene function in DFC/KV cells and how to introduce fluorescent beads to visualize fluid flow in KV. The distribution of fluorescent MO in successful stage-specific injected embryos is shown schematically in Figures 1D-F and in live embryos in Figure 2. An unsuccessful MO injection, ...

Cilia-generated fluid flow in Kupffer&#x2019;s Vesicle (KV) controls left-right patterning of the zebrafish embryo. Here, we describe a technique to modulate gene function specifically in KV cells. In addition, we show how to deliver fluorescent beads into KV to visualize fluid flow.

Internal organs such as the  heart, brain, and gut develop left-right (LR) asymmetries that are critical for  their normal functions1. Motile cilia are ...

The aim of this procedure is to modulate the function genes in Kavas vesicle or the KV in zebrafish embryo that is responsible for left right patterning and analyze asymmetric fluid flow generated by motil clia by delivering fluorescent beads into the kv. This is accomplished by first injecting fluorescent morph oligonucleotides that repress expression of a specific gene of interest in the yolk cell of a mid blash Dilla stage embryo, so that they load into dorsal forerunner cells that give rise to the kv. Next injected embryos that have fluorescent morphos only in the yolk cell and the kv are selected for further analysis.Then the embryos are mounted in depression slides and fluorescent beads are injected into the fluid-filled vesicle lumen of the kv. Finally, video microscopy is used to record beads flowing inside the kv. Ultimately, results can be obtained that determine whether tissue specific depletion of a candidate gene in the KV alters asymmetric fluid flow through the quantitative analysis of bead movements.Viral demonstration of this semester is critical as stage specific molino injections and fluorescence bead injections are difficult to learn because both depend on propo

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Research

JoVE Journal

Biology

We present a method to visualize cuticle in live C. elegans using the red fluorescent lipophilic dye DiI (1,1'-dioctadecyl-3,3,3',3'-tetramethylindocarbocyanine perchlorate), which is commonly used in C. elegans to visualize environmentally exposed neurons. With this optimized protocol, alae and annular cuticular structures are stained by DiI and observed using compound microscopy.

Visualization of Caenorhabditis elegans Cuticular Structures Using the Lipophilic Vital Dye DiI

A simple microfluidic device has been developed to perform anesthetic free in vivo imaging of C. elegans, intact Drosophila larvae and zebrafish larvae. The device utilizes a deformable PDMS membrane to immobilize these model organisms in order to perform time lapse imaging of numerous processes such as heart beat, cell division and sub-cellular neuronal transport.  We demonstrate the use of this device and show examples of different types of data collected from different model systems.

Simple Microfluidic Devices for in vivo Imaging of C. elegans, Drosophila and Zebrafish

An easy and reliable technique for visualizing and quantifying airway cilia motility and cilia generated flow using mouse trachea is described.  This technique can be modified to determine how a wide range of factors influence cilia motility, including pharmacological agents, genetic factors, environmental exposures, and/or mechanical factors such as mucus load.

Ex vivo Method for High Resolution Imaging of Cilia Motility in Rodent Airway Epithelia

Sea lamprey lose the gall bladder and bile ducts during metamorphosis, a process similar to human biliary atresia. A new fixation and clarification method (CLARITY) was modified to visualize the entire biliary tree using laser scanning confocal microscopy. This method provides a powerful tool to study biliary degeneration.

A New Clarification Method to Visualize Biliary Degeneration During Liver Metamorphosis in Sea Lamprey (Petromyzon marinus)

Adaptive immunity is controlled by dynamic 'immunological synapses' formed between T cells and antigen presenting cells. This protocol describes methods for investigating endothelial cells both as understudied physiologic APCs and as a novel type of 'planar cellular APC model'.

An Endothelial Planar Cell Model for Imaging Immunological Synapse Dynamics

Here it is shown how to track and quantify developmental processes in C. elegans. The methods presented are based on open-source tools that can be easily implemented. It is demonstrated how to reconstruct 3D cell-shape models, how to manually track subcellular structures, and how to analyze cortical contractile flow.

Tracking and Quantifying Developmental Processes in C. elegans Using Open-source Tools

This protocol demonstrates a fluorescence-based method to visualize the vasculature and to quantify its complexity in Xenopus tropicalis. Blood vessels can be imaged minutes after the injection of a fluorescent dye into the beating heart of an embryo after genetic and/or pharmacological manipulations to study cardiovascular development in vivo.

Visualization and Quantitative Analysis of Embryonic Angiogenesis in Xenopus tropicalis

Cilia development is vital to proper organogenesis. This protocol describes an optimized method to label and visualize ciliated cells of the zebrafish.

Visualizing Multiciliated Cells in the Zebrafish Through a Combined Protocol of Whole Mount Fluorescent In Situ Hybridization and Immunofluorescence

Imaging flow cytometry provides an ideal approach to detect the morphological and functional alteration of cells at individual and populational levels. Disrupted endocytic function for lipid antigen presentation in pollutant-exposed human dendritic cells was demonstrated with a combined transcriptomic profiling of gene expression and morphological demonstration of protein trafficking.

Integrate Imaging Flow Cytometry and Transcriptomic Profiling to Evaluate Altered Endocytic CD1d Trafficking

In the early Drosophila embryo, many organelles are motile. In principle, they can be imaged live via specific fluorescent probes, but the eggshell prevents direct application to the embryo. This protocol describes how to introduce such probes via microinjection, and then analyze bulk organelle motion via particle image velocimetry.

Visualizing Cytoskeleton-Dependent Trafficking of Lipid-Containing Organelles in Drosophila Embryos

学习在斑马鱼胚胎枯否囊泡中调控基因功能并可视化纤毛产生的流体流动的方法。该技术利用荧光微珠和显微成像定量分析不对称流体动力学，为研究左右不对称模式形成提供重要工具。

Analysis of Gene Function and Visualization of Cilia-Generated Fluid Flow in Kupffer's Vesicle

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