We thank Mark Terasaki for initial advice on using a two-photon microscope to create local tissue damage, Kathy Joubin for advice on mounting for time-lapse imaging, and the Sagasti and Portera-Cailliau labs for discussions. Initial experiments were performed by AS as a Grass Foundation Fellow at the Marine Biological Labs in Woods Hole, MA. Work in the Sagasti lab was supported by grants from the Whitehall Foundation, the Klingenstein Foundation, and a Burroughs Wellcome Fund Career Award in the Biological Sciences.

Part 1: Mounting zebrafish embryos for long-term imaging
<ol>
<li>Prepare 1% low melt agarose solution for embedding. Dissolve agarose in DI water by heating in a microwave, aliquot into small tubes and store in a heating block at 42 degrees Celsius.</li>
<li>Select embryos for imaging and remove their chorions by gently pulling the chorion apart with forceps. Embryos can be placed into a 5% PTU Ringers solution at 22-24 hours post fertilization (hpf) to inhibit the formation of pigment. This improves the clarity of imaging and is essential for performing two-photon axotomies of sensory axons. If the natural pigment is allowed to form, autofluorescence obscures axons on the two-photon microscope. Ringers solution for zebrafish contains 116 mM NaCl, 2.9 mM KCl, 1.8 mM CaCl2, and 5mM HEPES, pH 7.2.</li>
<li>Anesthetize embryos by adding ~0.02% tricaine to the PTU Ringers. Make sure animals are not responsive to touch before proceeding.</li>
<li>Prepare the long-term imaging chamber by applying vacuum grease to one side of a glass or Teflon ring and fixing it onto a glass cover slip.</li>
<li>Transfer one embryo into the 42 degree agarose solution with a glass pipet, taking care not to transfer much of the PTU Ringers, then transfer the embryo with one drop of agarose onto a prepared cover slip.</li>
<li>Position the embryo as desired as the agarose hardens. Keep in mind that the imaging chamber will be flipped when you are done so that the cover slip will be the top surface.</li>
<li>If you have a motorized stage and can image multiple locations at once, repeat steps 1.4-1.6 for each embryo.</li>
<li>Allow the agarose to completely solidify, then fill the ring with 0.02% tricaine PTU Ringers.</li>
<li>Apply vacuum grease to the other side of the ring(s), then cover the ring(s) with a glass slide. Flip the sealed chamber(s) over. These chambers can be used for imaging on upright or inverted microscopes.</li>
</ol>
Part 2: Two-photon axotomy using a custom built two-photon microscope with a Chameleon Ti-Sapphire laser
<ol>
<li>Prepare for imaging. Place the mounted embryo(s) on a slide holder under the microscope. Focus on one embryo with a 40X (0.8 NA) water objective. Turn on the laser. We have been able to reproducibly visualize and damage GFP expressing neurons using the following parameters. To visualize axons at a non-damaging power, set laser to a wavelength of 910 nanometers (nm) and a power of 30 milliwatts (mW listed are the amount of power at the sample). Open the imaging software. We use ScanImage Software developed in Karel Svoboda's laboratory 2, 3.</li>
<li>Press focus to scan the embryo with the two-photon laser, locate the axon you wish to axotomize, and capture an image of this axon. Mark the first and last Z positions, acquire the image, and make a maximum projection of the Z stacks.</li>
<li>Zoom in 70X on the branch of the axon you wish to axotomize and stop scanning. Increase the mW at the sample to the damaging power of 180 mW, and perform a single scan with the laser. We do this by setting the number of Z slices to 1 and pressing &ldquo;Grab&rdquo;. This should be sufficient to sever the axon. See discussion for methods to optimize this procedure for your microscope and experimental goal.</li>
<li>Zoom out, reduce the power to 30 mW and take an image.</li>
</ol>
Part 3: Two-photon axotomy on Zeiss 510 confocal/two-photon microscope
<ol>
<li>Place mounted embryo onto the stage and bring it into focus using a 25X water objective or other suitable objective.</li>
<li>Turn on the two-photon (910 nm) and Argon (488 nm) lasers in a multi-track setting so that it is possible to switch from one to the other. Although both lasers are used to detect GFP, the two-photon emission is visualized with red, and the argon laser emission with green to differentiate the two.</li>
<li>Use the Argon laser to identify an axon to injure. Under Z settings mark the first and last optical sections. Take a confocal image and create a maximum projection of the Z stack.</li>
<li>Choose the region of the axon to be injured and bring this region into focus. Turn off the Argon laser and turn on the two-photon laser. Scan the specimen with the two-photon at a laser intensity of ~9% transmission to make sure that the axon is still in focus.</li>
<li>Click the &ldquo;stop&rdquo; button so that the crop tool will be available. Use crop to zoom in on the area of interest. Usually we zoom to ~70X (zoom can be checked under the &ldquo;mode&rdquo; tab). Under the &ldquo;channels&rdquo; tab change the intensity of the two-photon from ~9% transmission to 15-20% transmission.</li>
<li>To sever an axon activate the &ldquo;fast XY&rdquo; button for about 1 second and then press the stop button to avoid excess damage. The axon should be seen as scattered debris if the procedure worked.</li>
<li>To ensure that the axon was damaged switch back to the 488 nm Argon laser, take another confocal image, and create a maximum projection.</li>
</ol>
Part 4: Confocal time-lapse imaging on Zeiss LSM 510
<ol>
<li>Prepare for imaging. If you are using a heated stage, be sure to turn it on at least 30 minutes before setting up your time-lapse movie. Place the mounted embryo(s) on a slide holder under the microscope. Focus on one embryo with desired air objective. We use a 20X, 0.5 NA objective. Open Zeiss LSM 510 imaging software. Turn on the appropriate lasers and set up the desired configuration. We will now describe a detailed protocol for long term imaging with Zeiss LSM Multi Time software. These methods can be adapted for use on your own microscope with your imaging software.</li>
<li>Define the position and configuration of your first embryo in the Multi Time window. Open the scan, stage, and Multi Time windows. Activate fast scan on the scan window. Move the stage to the desired XY position. Mark the first and last Z positions in the scan window. Press stop, then Mid, also in the scan window. Wait for the scan of the middle Z slice to finish, then press mark position in the stage window. If you are only imaging one embryo, click on the &ldquo;Single Location&rdquo; tab in the Multi Time window. Click on &ldquo;Replace XYZ&rdquo; and then on &ldquo;Save Configuration&rdquo;. Agree when asked to overwrite the previous configuration. Proceed to step 4.4. If you have a mechanized stage, you can set up multiple locations to image multiple embryos. In this case, you should select the &ldquo;Multiple Locations&rdquo; tab before defining the XYZ and configuration for your first location. Also, be sure that the pull-down menu just below the &ldquo;Multiple Locations&rdquo; tab is on location 1, and that the pull-down menu in the configuration section says multi loc 1, before you click on save configuration.</li>
<li>Define the position and configuration of your remaining embryos in the Multi Time window. Locate your next embryo, then press fast scan, move the stage into the desired XY position, and mark your first and last Z positions in the scan window. Press stop, then Mid, also in the scan window. Wait for the scan of the middle Z slice to finish, then press mark position in the stage window. Click on &ldquo;Replace XYZ&rdquo;. Check that the pull-down menu just below the &ldquo;Multiple Locations&rdquo; tab is on location 2, and that the pull-down menu in the configuration section says multi loc 2, then click on &ldquo;Save Configuration&rdquo;. Agree when asked to overwrite the previous configuration. Repeat this process for the remaining embryos.</li>
<li>Set up the parameters for image acquisition in the Multi Loc window. Choose the GR-G-L option in the &ldquo;List of Blocks&rdquo; section, then enter the desired number of group repetitions. This is the number of times the confocal will acquire an image at each location. Enter the desired wait interval in the Parameters section. This is the amount of time from the start of one imaging repetition to the start of the next repetition. Select &ldquo;ZStackXY&rdquo; in the configuration section. Enter you base file name in the bottom section. Click on &ldquo;Select Image DB&rdquo; and choose your mdb. Click on the &ldquo;Options&rdquo; button. Select your mdb folder to save temp files. Check &ldquo;keep final image open&rdquo;, &ldquo;save final image&rdquo;, and &ldquo;middle of the z stack&rdquo;. Select wait interval. Click ok to close the options window. Click on &ldquo;start time&rdquo;. Leave Multi Time window open for duration of imaging.</li>
<li>Analyze your data. When the time-lapse imaging has finished, the Multi Time software will compile all of your temporary files into a summary file for each location. If you want to stop the movie before it is complete, be sure to press &ldquo;Finish&rdquo; instead of &ldquo;Stop&rdquo;, so that a summary file will be created. The temp files and summary files should be automatically saved to the designated mdb folder. From the LSM software menu, click on &ldquo;3D View&rdquo;, then &ldquo;Projection&rdquo;. With your summary file selected in the pull-down menu, click on &ldquo;Apply&rdquo;. This will generate maximum projections of the Z stack for each confocal image. On the LSM software menu, click on &ldquo;File&rdquo;, then &ldquo;Export&rdquo;. Save the maximum projections as a series of tif files. You may then create a movie out of the series of tif files using ImageJ or QuickTime Pro. Axons in the LSM images can be traced in three dimensions using image analysis software (e.g. NeuroLucida from MicroBrightfield) to generate detailed quantitative information about axon morphology. </li>
</ol>
Part 5: Representative results
A successful experiment will result in an accurate representation of the cellular dynamics of axon recovery from injury. Your embryos will be healthy after imaging, with no visible degeneration and a strong heartbeat. The axotomy should result in precise damage, only severing the defined branch of the axon. There should be no injury to surrounding axons and minimal cell death. We believe we see the death of a single epidermal cell directly over the site of axotomy in ~50% of experiments. If you observe more damage, you should optimize your two-photon protocol as described in the discussion.

<ol>
	<li>Sagasti, A., Guido, M. R., Raible, D. W., Schier, A. F. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Repulsive+interactions+shape+the+morphologies+and+functional+arrangement+of+zebrafish+peripheral+sensory+arbors.">Repulsive interactions shape the morphologies and functional arrangement of zebrafish peripheral sensory arbors.</a> Curr Biol. 15, 804-814 (2005).</li><li>Pologruto, T. A., Sabatini, B. L., Svoboda, K. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=ScanImage:+flexible+software+for+operating+laser+scanning+microscopes.">ScanImage: flexible software for operating laser scanning microscopes.</a> Biomed Eng Online. 2 (13), (2003).</li><li>Svoboda, K., Yasuda, R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Principles+of+two-photon+excitation+microscopy+and+its+applications+to+neuroscience.">Principles of two-photon excitation microscopy and its applications to neuroscience.</a> Neuron. 50, 823-839 (2006).</li><li>Sacconi, L. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=In+vivo+multiphoton+nanosurgery+on+cortical+neurons.">In vivo multiphoton nanosurgery on cortical neurons.</a> J Biomed Opt. 12, 050502-050502 (2007).</li><li>Galbraith, J. A., Terasaki, M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Controlled+damage+in+thick+specimens+by+multiphoton+excitation.">Controlled damage in thick specimens by multiphoton excitation.</a> Mol Biol Cell. 14, 1808-1817 (2003).</li><li>Yanik, M. F. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Neurosurgery:+functional+regeneration+after+laser+axotomy.">Neurosurgery: functional regeneration after laser axotomy.</a> Nature. 432, 822-822 (2004).</li><li>Quinto-Su, P. A., Venugopalan, V. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Mechanisms+of+laser+cellular+microsurgery.">Mechanisms of laser cellular microsurgery.</a> Methods Cell Biol. 82, 113-151 (2007).</li></ol>

We have used the methods described to precisely axotomize peripheral sensory axons and to directly observe regeneration in the living zebrafish embryo. Long-term time-lapse confocal imaging in zebrafish can be used to observe many developmental processes in vivo. The two-photon axotomy procedure described can be modified for many different experimental goals. We have used the same general procedure to ablate entire trigeminal sensory neuron cell bodies, by zooming in on the cell body rather than on a branch o...

two-photon axotomy and time-lapse confocal imaging in live zebrafish embryos

Zebrafish have long been utilized to study the cellular and molecular mechanisms of development by time-lapse imaging of the living transparent embryo. Here we describe a method to mount zebrafish embryos for long-term imaging and demonstrate how to automate the capture of time-lapse images using a confocal microscope. We also describe a method to create controlled, precise damage to individual branches of peripheral sensory axons in zebrafish using the focused power of a femtosecond laser mounted on a two-photon microscope. The parameters for successful two-photon axotomy must be optimized for each microscope. We will demonstrate two-photon axotomy on both a custom built two-photon microscope and a Zeiss 510 confocal/two-photon to provide two examples.

Zebrafish trigeminal sensory neurons can be visualized in a transgenic line expressing GFP driven by a sensory neuron specific promoter 1. We have adapted this zebrafish trigeminal model to directly observe sensory axon regeneration in living zebrafish embryos. Embryos are anesthetized with tricaine and positioned within a drop of agarose as it solidifies. Immobilized embryos are sealed within an imaging chamber filled with phenylthiourea (PTU) Ringers. We have found that embryos can be continuously imaged in these chambers for 12-48 hours. A single confocal image is then captured to determine the desired site of axotomy. The region of interest is located on the two-photon microscope by imaging the sensory axons under low, non-damaging power. After zooming in on the desired site of axotomy, the power is increased and a single scan of that defined region is sufficient to sever the axon. Multiple location time-lapse imaging is then set up on a confocal microscope to directly observe axonal recovery from injury.

Watch this Scientific Journal Video about Two-photon axotomy and time-lapse confocal imaging in live zebrafish embryos at JoVE.com

Two-photon axotomy and time-lapse confocal imaging in live zebrafish embryos

Here we describe a method for mounting zebrafish embryos for long-term imaging, two-photon imaging and tissue-damage techniques, and time-lapse confocal imaging.

Zebrafish have long been utilized to study the cellular and molecular mechanisms of development by time-lapse imaging of the living transparent embryo.  ...

two-photon-axotomy-time-lapse-confocal-imaging-live-zebrafish

Research

JoVE Journal

Biology

14.1K Görüntüleme.  University of California, Los Angeles. Burada, uzun vadeli görüntüleme, iki foton görüntüleme ve doku hasarı teknikleri ve zaman atlamalı konfokal görüntüleme zebrafish embriyolar montaj için bir yöntem açıklanmaktadır.

Video: Two-photon axotomy and time-lapse confocal imaging in live zebrafish embryos

Live Imaging of the Zebrafish Embryonic Brain by Confocal Microscopy

In this video, we demonstrate the method our lab has developed to analyze the cell shape changes and rearrangements required to bend and fold the developing zebrafish brain (Gutzman et al, 2008). Such analysis affords a new understanding of the underlying cell biology required for development of the 3D structure of the vertebrate brain, and significantly increases our ability to study neural tube morphogenesis. The embryonic zebrafish brain is shaped beginning at 18 hours post fertilization (hpf) as the ventricles within the neuroepithelium inflate. By 24 hpf, the initial steps of neural tube morphogenesis are complete. Using the method described here, embryos at the one cell stage are injected with mRNA encoding membrane-targeted green fluorescent protein (memGFP). After injection and incubation, the embryo, now between 18 and 24 hpf, is mounted, inverted, in agarose and imaged by confocal microscopy. Notably, the zebrafish embryo is transparent making it an ideal system for fluorescent imaging. While our analyses have focused on the midbrain-hindbrain boundary and the hindbrain, this method could be extended for analysis of any region in the zebrafish to a depth of 
80-100 &mu;m.

In this video, we demonstrate a method by which to analyze the developing vertebrate brain in live zebrafish embryos at single cell resolution by confocal microscopy. This includes the method by which we inject the single-cell zebrafish embryo and subsequently mount and image the developing brain.

In this video, we demonstrate the method our lab has developed to analyze the cell shape changes and rearrangements required to bend and fold the ...

Live Imaging of Cell Motility and Actin Cytoskeleton of Individual Neurons and Neural Crest Cells in Zebrafish Embryos

The zebrafish is an ideal model for imaging cell behaviors during development in vivo. Zebrafish embryos are externally fertilized and thus easily accessible at all stages of development. Moreover, their optical clarity allows high resolution imaging of cell and molecular dynamics in the natural environment of the intact embryo. We are using a live imaging approach to analyze cell behaviors during neural crest cell migration and the outgrowth and guidance of neuronal axons.
Live imaging is particularly useful for understanding mechanisms that regulate cell motility processes. To visualize details of cell motility, such as protrusive activity and molecular dynamics, it is advantageous to label individual cells. In zebrafish, plasmid DNA injection yields a transient mosaic expression pattern and offers distinct benefits over other cell labeling methods. For example, transgenic lines often label entire cell populations and thus may obscure visualization of the fine protrusions (or changes in molecular distribution) in a single cell. In addition, injection of DNA at the one-cell stage is less invasive and more precise than dye injections at later stages.
Here we describe a method for labeling individual developing neurons or neural crest cells and imaging their behavior in vivo. We inject plasmid DNA into 1-cell stage embryos, which results in mosaic transgene expression. The vectors contain cell-specific promoters that drive expression of a gene of interest in a subset of sensory neurons or neural crest cells. We provide examples of cells labeled with membrane targeted GFP or with a biosensor probe that allows visualization of F-actin in living cells1.
Erica Andersen, Namrata Asuri, and Matthew Clay contributed equally to this work.

This protocol describes imaging of individual neurons or neural crest cells in living zebrafish embryos. This method is used to examine cellular behaviors and actin localization using fluorescence confocal time-lapse microscopy.

The zebrafish is an ideal model for imaging cell behaviors during development in vivo. Zebrafish embryos are externally fertilized and thus easily ...

Time-lapse Imaging of Mitosis After siRNA Transfection

Changes in cellular organization and chromosome dynamics that occur during mitosis are tightly coordinated to ensure accurate inheritance of genomic and cellular content. Hallmark events of mitosis, such as chromosome movement, can be readily tracked on an individual cell basis using time-lapse fluorescence microscopy of mammalian cell lines expressing specific GFP-tagged proteins. In combination with RNAi-based depletion, this can be a powerful method for pinpointing the stage(s) of mitosis where defects occur after levels of a particular protein have been lowered. In this protocol, we present a basic method for assessing the effect of depleting a potential mitotic regulatory protein on the timing of mitosis. Cells are transfected with siRNA, placed in a stage-top incubation chamber, and imaged using an automated fluorescence microscope. We describe how to use software to set up a time-lapse experiment, how to process the image sequences to make either still-image montages or movies, and how to quantify and analyze the timing of mitotic stages using a cell-line expressing mCherry-tagged histone H2B. Finally, we discuss important considerations for designing a time-lapse experiment. This strategy is complementary to other approaches and offers the advantages of 1) sensitivity to changes in kinetics that might not be observed when looking at cells as a population and 2) analysis of mitosis without the need to synchronize the cell cycle using drug treatments. The visual information from such imaging experiments not only allows the sub-stages of mitosis to be assessed, but can also provide unexpected insight that would not be apparent from cell cycle analysis by FACS.

Here we describe a basic protocol to image and quantify the mitotic timing of live mammalian tissue culture cells after siRNA transfection.

Changes in cellular organization and chromosome dynamics that occur during mitosis are tightly coordinated to ensure accurate inheritance of genomic and ...

Two-Photon-Based Photoactivation in Live Zebrafish Embryos

Photoactivation of target compounds in a living organism has proven a valuable approach to investigate various biological processes such as embryonic development, cellular signaling and adult physiology. In this respect, the use of multi-photon microscopy enables quantitative photoactivation of a given light responsive agent in deep tissues at a single cell resolution. As zebrafish embryos are optically transparent, their development can be monitored in vivo. These traits make the zebrafish a perfect model organism for controlling the activity of a variety of chemical agents and proteins by focused light. Here we describe the use of two-photon microscopy to induce the activation of chemically caged fluorescein, which in turn allows us to follow cell's destiny in live zebrafish embryos. We use embryos expressing a live genetic landmark (GFP) to locate and precisely target any cells of interest. This procedure can be similarly used for precise light induced activation of proteins, hormones, small molecules and other caged compounds.

Multiphoton microscopy allows control of low energy photons with deep optical penetration and reduced phototoxicity. We describe the use of this technology for live cell labeling in zebrafish embryos. This protocol can be readily adapted for photo-induction of various light-responsive molecules.

Photoactivation of target compounds in a living organism has proven a valuable approach to investigate various biological processes such as embryonic ...

Transplantation of GFP-expressing Blastomeres for Live Imaging of Retinal and Brain Development in Chimeric Zebrafish Embryos

Cells change extensively in their locations and property during embryogenesis. These changes are regulated by the interactions between the cells and their environment. Chimeric embryos, which are composed of cells of different genetic background, are great tools to study the cell-cell interactions mediated by genes of interest. The embryonic transparency of zebrafish at early developmental stages permits direct visualization of the morphogenesis of tissues and organs at the cellular level. Here, we demonstrate a protocol to generate chimeric retinas and brains in zebrafish embryos and to perform live imaging of the donor cells. The protocol covers the preparation of transplantation needles, the transplantation of GFP-expressing donor blastomeres to GFP-negative hosts, and the examination of donor cell behavior under live confocal microscopy. With slight modifications, this protocol can also be used to study the embryonic development of other tissues and organs in zebrafish. The advantages of using GFP to label donor cells are also discussed. 

We demonstrate a protocol to generate chimeric zebrafish embryos for live imaging cellular behavior during embryogenesis.

Cells change extensively in their locations and property during embryogenesis. These changes are regulated by the interactions between the cells and their ...

Multicolor Time-lapse Imaging of Transgenic Zebrafish: Visualizing Retinal Stem Cells Activated by Targeted Neuronal Cell Ablation

High-resolution time-lapse imaging of living zebrafish larvae can be utilized to visualize how biological processes unfold (for review see 1). Compound transgenic fish which express different fluorescent reporters in neighboring cell types provide a means of following cellular interactions 2 and/or tissue-level responses to experimental manipulations over time. In this video, we demonstrate methods that can be used for imaging multiple transgenically labeled cell types serially in individual fish over time courses that can span from minutes to several days. The techniques described are applicable to any study seeking to correlate the "behavior" of neighboring cells types over time, including: 1) serial 'catch and release' methods for imaging a large number of fish over successive days, 2) simplified approaches for separating fluorophores with overlapping excitation/emission profiles (e.g., GFP and YFP), 3) use of hypopigmented mutant lines to extend the time window available for high-resolution imaging into late larval stages of development, 4) use of membrane targeted fluorescent reporters to reveal fine morphological detail of individual cells as well as cellular details in larger populations of cells, and 5) a previously described method for chemically-induced ablation of transgenically targeted cell types; i.e., nitroreductase (NTR) mediated conversion of prodrug substrates, such as metronidazole (MTZ), to cytotoxic derivatives 3,5.
As an example of these approaches, we will visualize the ablation and regeneration of a subtype of retinal bipolar neuron within individual fish over several days. Simultaneously we will monitor several other retinal cell types, including neighboring non-targeted bipolar cells and potential degeneration-stimulated retinal stem cells (i.e., M&#971;ller glia). This strategy is being applied in our lab to characterize cell- and tissue-level (e.g., stem cell niche) responses to the selective loss and regeneration of targeted neuronal cell types.

In this video, techniques for multicolor confocal time-lapse imaging and targeted cell ablation are provided. Time-lapse imaging is used to monitor the behavior of multiple cell types of interest in vivo. Targeted cell ablation facilitates the study neural circuit function and cell-specific neuronal regeneration paradigms.

High-resolution time-lapse imaging of living zebrafish larvae can be utilized to visualize how biological processes unfold (for review see 1). Compound ...

Time-lapse Microscopy of Early Embryogenesis in Caenorhabditis elegans

Caenorhabditis elegans has often been used as a model system in studies of early developmental processes. The transparency of the 
embryos, the genetic resources, and the relative ease of transformation are qualities that make C. elegans an excellent model for early embryogenesis. Laser-based 
confocal microscopy and fluorescently labeled tags allow researchers to follow specific cellular structures and proteins in the developing embryo. For example, 
one can follow specific organelles, such as lysosomes or mitochondria, using fluorescently labeled dyes. These dyes can be delivered to the early embryo by means 
of microinjection into the adult gonad. Also, the localization of specific proteins can be followed using fluorescent protein tags. Examples are presented here 
demonstrating the use of a fluorescent lysosomal dye as well as fluorescently tagged histone and ubiquitin proteins. The labeled histone is used to visualize 
the DNA and thus identify the stage of the cell cycle. GFP-tagged ubiquitin reveals the dynamics of ubiquitinated vesicles in the early embryo. Observations 
of labeled lysosomes and GFP:: ubiquitin can be used to determine if there is colocalization between ubiquitinated vesicles and lysosomes. A technique for 
the microinjection of the lysosomal dye is presented. Techniques for generating transgenenic strains are presented elsewhere (1, 2). For imaging, embryos 
are cut out of adult hermaphrodite nematodes and mounted onto 2% agarose pads followed by time-lapse microscopy on a standard laser scanning confocal 
microscope or a spinning disk confocal microscope. This methodology provides for the high resolution visualization of early embryogenesis.

Time-lapse Microscopy of Early Embryogenesis in Caenorhabditis elegans

This article describes a technique for the visualization of the early events of embryogenesis in the nematode Caenorhabditis elegans.

Caenorhabditis elegans has often been used as a model system in studies of early developmental processes.  The transparency of the 
embryos, the genetic ...

Fluorescence-based Measurement of Store-operated Calcium Entry in Live Cells: from Cultured Cancer Cell to Skeletal Muscle Fiber

Store operated Ca2+ entry (SOCE), earlier termed capacitative Ca2+ entry, is a tightly regulated mechanism for influx of extracellular Ca2+ into cells to replenish depleted endoplasmic reticulum (ER) or sarcoplasmic reticulum (SR) Ca2+ stores1,2. Since Ca2+ is a ubiquitous second messenger, it is not surprising to see that SOCE plays important roles in a variety of cellular processes, including proliferation, apoptosis, gene transcription and motility. Due to its wide occurrence in nearly all cell types, including epithelial cells and skeletal muscles, this pathway has received great interest3,4. However, the heterogeneity of SOCE characteristics in different cell types and the physiological function are still not clear5-7.
The functional channel properties of SOCE can be revealed by patch-clamp studies, whereas a large body of knowledge about this pathway has been gained by fluorescence-based intracellular Ca2+ measurements because of its convenience and feasibility for high-throughput screening. The objective of this report is to summarize a few fluorescence-based methods to measure the activation of SOCE in monolayer cells, suspended cells and muscle fibers5,8-10. The most commonly used of these fluorescence methods is to directly monitor the dynamics of intracellular Ca2+ using the ratio of F340nm and F380nm (510 nm for emission wavelength) of the ratiometric Ca2+ indicator Fura-2. To isolate the activity of unidirectional SOCE from intracellular Ca2+ release and Ca2+ extrusion, a Mn2+ quenching assay is frequently used. Mn2+ is known to be able to permeate into cells via SOCE while it is impervious to the surface membrane extrusion processes or to ER uptake by Ca2+ pumps due to its very high affinity with Fura-2. As a result, the quenching of Fura-2 fluorescence induced by the entry of extracellular Mn2+ into the cells represents a measurement of activity of SOCE9. Ratiometric measurement and the Mn+2 quenching assays can be performed on a cuvette-based spectrofluorometer in a cell population mode or in a microscope-based system to visualize single cells. The advantage of single cell measurements is that individual cells subjected to gene manipulations can be selected using GFP or RFP reporters, allowing studies in genetically modified or mutated cells. The spatiotemporal characteristics of SOCE in structurally specialized skeletal muscle can be achieved in skinned muscle fibers by simultaneously monitoring the fluorescence of two low affinity Ca2+ indicators targeted to specific compartments of the muscle fiber, such as Fluo-5N in the SR and Rhod-5N in the transverse tubules9,11,12.

The extent of store-operated Ca2+ entry (SOCE) can be monitored using fluorescent Ca2+ indicators. Mn2+ quenching of such indicators assays SOCE in cultured cells and skeletal muscle fibers. A technique allowing spatial and temporal resolution of SOCE by confocal imaging of mechanically skinned muscle fibers is also described.

Store operated Ca2+ entry (SOCE), earlier termed capacitative Ca2+ entry, is a tightly regulated mechanism for influx of extracellular Ca2+ into cells to ...

Use of Time Lapse Microscopy to Visualize Anoxia-induced Suspended Animation in C. elegans Embryos

Caenorhabdits elegans has been used extensively in the study of stress resistance, which is facilitated by the transparency of the adult and embryo stages as well as by the availability of genetic mutants and transgenic strains expressing a myriad of fusion proteins1-4. In addition, dynamic processes such as cell division can be viewed using fluorescently labeled reporter proteins. The study of mitosis can be facilitated through the use of time-lapse experiments in various systems including intact organisms; thus the early C. elegans embryo is well suited for this study. Presented here is a technique by which in vivo imaging of sub-cellular structures in response to anoxic (99.999% N2; &lt;2 ppm O2) stress is possible using a simple gas flow through setup on a high-powered microscope. A microincubation chamber is used in conjunction with nitrogen gas flow through and a spinning disc confocal microscope to create a controlled environment in which animals can be imaged in vivo. Using GFP-tagged gamma tubulin and histone, the dynamics and arrest of cell division can be monitored before, during and after exposure to an oxygen-deprived environment. The results of this technique are high resolution, detailed videos and images of cellular structures within blastomeres of embryos exposed to oxygen deprivation.

Use of Time Lapse Microscopy to Visualize Anoxia-induced Suspended Animation in C. elegans Embryos

Described here is an in vivo technique to image sub-cellular structures in animals exposed to anoxia using a gas flow through microincubation chamber in conjunction with a spinning disc confocal microscope. This method is straightforward and flexible enough to suit a variety of experimental parameters and model systems.

Caenorhabdits elegans has been used extensively in the study of stress resistance, which is facilitated by the transparency of the adult and embryo stages ...

Long-term, High-resolution Confocal Time Lapse Imaging of Arabidopsis Cotyledon Epidermis during Germination

Imaging in vivo dynamics of cellular behavior throughout a developmental sequence can be a powerful technique for understanding the mechanics of tissue patterning. During animal development, key cell proliferation and patterning events occur very quickly. For instance, in Caenorhabditis elegans all cell divisions required for the larval body plan are completed within six hours after fertilization, with seven mitotic cycles1; the sixteen or more mitoses of Drosophila embryogenesis occur in less than 24 hr2. In contrast, cell divisions during plant development are slow, typically on the order of a day 3,4,5 . This imposes a unique challenge and a need for long-term live imaging for documenting dynamic behaviors of cell division and differentiation events during plant organogenesis. Arabidopsis epidermis is an excellent model system for investigating signaling, cell fate, and development in plants. In the cotyledon, this tissue consists of air- and water-resistant pavement cells interspersed with evenly distributed stomata, valves that open and close to control gas exchange and water loss. Proper spacing of these stomata is critical to their function, and their development follows a sequence of asymmetric division and cell differentiation steps to produce the organized epidermis (Fig. 1).
This protocol allows observation of cells and proteins in the epidermis over several days of development. This time frame enables precise documentation of stem-cell divisions and differentiation of epidermal cells, including stomata and epidermal pavement cells. Fluorescent proteins can be fused to proteins of interest to assess their dynamics during cell division and differentiation processes. This technique allows us to understand the localization of a novel protein, POLAR6, during the proliferation stage of stomatal-lineage cells in the Arabidopsis cotyledon epidermis, where it is expressed in cells preceding asymmetric division events and moves to a characteristic area of the cell cortex shortly before division occurs. Images can be registered and streamlined video easily produced using public domain software to visualize dynamic protein localization and cell types as they change over time.

Long-term, High-resolution Confocal Time Lapse Imaging of Arabidopsis Cotyledon Epidermis during Germination

We describe a protocol using chamber slides and media to immobilize plant cotyledons for confocal imaging of the epidermis over several days of development, documenting stomatal differentiation. Fluorophore-tagged proteins can be tracked dynamically by expression and subcellular localization, increasing understanding of their possible roles during cell division and cell-type differentiation.

Imaging in vivo dynamics of cellular behavior throughout a developmental sequence can be a powerful technique for understanding the mechanics of tissue ...

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Two-photon axotomy and time-lapse confocal imaging in live zebrafish embryos