Name: Time-lapse Microscopy of Early Embryogenesis in Caenorhabditis elegans (Video) | JoVE Journal
Uploaded: 2011-08-25T12:03:00.000+00:00
Duration: 7 min 58 s
Description: Learn time-lapse microscopy for visualizing early embryogenesis in Caenorhabditis elegans using fluorescent labeling and confocal imaging. This protocol enables researchers to study subcellular dynamics and organelle movements in live embryos.

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5 minutes

We would like to acknowledge funding from the National Institutes of Health (R15 GM065444-03 to L.B.) and the Intramural Research Program of the National Institutes of Health (NIH) and the National Institute of Diabetes and Digestive and Kidney Diseases (K. O.). An NSF award (DBI-0923402) funded the acquisition of the LSM700 confocal microscope. We would also like to acknowledge general assistance and helpful comments on the manuscript from Dr. Andy Golden. Credit goes to the entire C. elegans community for sharing resources, protocols, and ideas.

1. Nematode cultures
<ol>
	<li>Obtain the appropriate C. elegans strain from the Caenorhabditis Genetics Stock Center (CGC) or from a colleague.</li>
	<li>Grow nematodes on NGM agar plates seeded with an OP50 bacterial lawn (3). For analysis of GFP strains growth at 25°C is recommended.</li>
	<li>The day before your microscopy experiment, pick at least 40 L4 larvae onto seeded plates and place the plates at 25°C overnight. These worms will be young adults for the experiment.</li>
</ol>
2. Injections
If it is desirable to view structures such as lysosomes or mitochondria, adults can be injected with fluorescent dye prior to visualization.
<ol>
	<li>Prepare dried agarose injection pads. (this can be done one to many days beforehand)
		<ol>
			<li>Prepare molten 2% agarose in water. With a Pasteur pipette put 2 drops onto a 22X54 mm coverglass. Place another coverglass cross-wise on top of the drop. In order to achieve the proper thickness, press on the top coverglass until the diameter of the pad is around 1.5 cm. Let sit for 5-10 minutes.</li>
			<li>Remove top coverglass. Dehydrate the agarose pad by placing it in an 80°C oven for 1 hour or allow to sit on benchtop overnight.</li>
		</ol>
	</li>
	<li>Prepare Mitotracker or Lysotracker solution.
		<ol>
			<li>Dilute fluorescent agent in Egg Buffer. We typically use a 1:10 dilution of Mitotracker or Lysotracker.</li>
			<li>The injection needle is made from a 1.2 OD glass capillary. Pull the injection needle using a needle puller.</li>
			<li>Use a pulled Pastuer pipet to back fill the needle with the diluted dye.</li>
			<li>Place needle in a light-free, humidified chamber until use.</li>
		</ol>
	</li>
	<li>Pre-Injection: Setting up the microscope and needle.
		<ol>
			<li>Mount injection needle onto injection apparatus. Our apparatus is a Narishige direct drive micromanipulator mounted onto the stage of a Nikon TE200 inverted microscope equipped with DIC imaging capability.</li>
			<li>Connect the needle to a 1.2 mm ID tube that is connected to the pressure regulator. Either compressed air or nitrogen can be used as an external pressure source. Set the regulator to 20-25 psi.</li>
			<li>Place 2 drops of heavy mineral oil (Parafin oil) onto the injection pad.</li>
			<li>Place the injection pad onto the microscope and lower the needle into the oil. Check to make sure fluid flows freely from the needle when pressure is applied. Apply injection pressure and observe whether fluid flows out of the needle. If not, you will need to gently break the end of the needle. The needle can be broken by gently driving the tip into a small bit of broken coverglass placed on the injection pad. After the needle is flowing, move on to the next step.</li>
		</ol>
	</li>
	<li>Injection.
		<ol>
			<li>Approximately 1 hour prior to viewing embryos, transfer young adult worms into the oil drop on the injection pad. Perform this transfer using the dissecting microscope. Use a platinum wire worm pick with a pointed tip for transferring worms.</li>
			<li>Arrange 3 – 10 worms so that they are lying directly on the pad. For injection, it is easiest if the worms are lined up parallel to one another and a little less than one worm length apart. Once the worms are on the injection pad it is important to work quickly and complete the injection process before the worms perish from desiccation.</li>
			<li>Place the coverglass with worms onto the microscope stage of the injection microscope. Focus into the central part (rachis) of the distal gonad tube. Then use the micromanipulator to move the tip of the needle into the same focal plane. Move the stage horizontally so that the worm is punctured by the needle. Once the tip of the needle is inside the rachis, apply pressure and allow the gonad to fill with the dye mixture. Inject both gonad arms of all worms on the injection pad.</li>
			<li>After injection, use a pulled Pasteur pipet to deliver about 0.5 ml of Egg Buffer to the worms. This will keep them from dying of dehydration and allow them to recover from the injection procedure.</li>
			<li>Allow the worms to recover for 1-2 hours at room temperature in a light-free, humidified chamber.</li>
		</ol>
	</li>
</ol>
3. Agarose pad for viewing embryos
<ol>
	<li>Prepare molten 2% agarose in Egg Buffer.</li>
	<li>With a Pasteur pipet, put 3 drops of molten agarose onto a clean microscope slide.</li>
	<li>Place slide between two other slides that have one layer of labeling tape covering them. These serve as spacers that will ensure that your agarose pad is the correct thickness.</li>
	<li>Place a second clean microscope slide perpendicularly down onto the agarose drops. Press down until top slide rests on tape from guide slides.</li>
	<li>Just before you are ready to mount embryos for observation, remove the top slide.</li>
</ol>
4. Mounting embryos for viewing
<ol>
	<li>If worms have not been injected, follow the next two steps. Otherwise skip them.
		<ol>
			<li>Choose young adult worms from the plate that was prepared the day before. You should be able to see embryos inside the young adults.</li>
			<li>Pick 5-20 worms onto an unseeded NGM plate. Allow the worms to crawl around on the unseeded plate for a few minutes so that the worms will lose most of the bacteria that is stuck to them.</li>
		</ol>
	</li>
	<li>Obtain young embryos from adult worms
		<ol>
			<li>Place a 5-20 μl drop of Egg Buffer in the middle of a 18 mm2 glass coverglass (a coverglass thickness of 1.5 is preferable).</li>
			<li>Move worms from the unseeded plate or injection pad into the drop of Egg Buffer.</li>
			<li>Cut open worms with a 26 gauge hypodermic needle. Cut open the worm near the vulva. You should see embryos spill out of the worm.</li>
		</ol>
	</li>
	<li>Mount onto microscope slide
		<ol>
			<li>Invert the microscope slide that was prepared with the agarose pad and lower it onto the coverglass with embryos.</li>
			<li>If extended time lapse is required, use petroleum jelly to seal the edges of the coverglass to prevent dessication.</li>
		</ol>
	</li>
</ol>
5. Microscopy
<ol>
	<li>Finding the appropriate embryos.
		<ol>
			<li>Place slide onto microscope stage. Scan with 10X lens to find embryos.</li>
			<li>To find early stage embryos, search for embryos that are in the process of completing maternal meiosis. Meiotic embryos can be distinguished from later embryos because they have not undergone shortening. (After embryos have completed meiosis, you can see a significant gap between the cell membrane and the egg shell at the anterior end.)</li>
			<li>Once you have identified an appropriately staged embryo, move to a higher magnification lens for recording of embryogenesis.</li>
		</ol>
	</li>
	<li>Imaging
		<ol>
			<li>In this example, we are imagining embryos that express the mCherry::H2B and GFP::Ub tagged proteins. We are using the Zeiss LSM700 conventional laser scanning microscope.</li>
			<li>Images are collected using the 63X 1.4NA lens. The 488 and 555 lasers are used with minimal laser power and maximum gain. A Z stack is collected every 10-15 seconds. Depending on the extent of photobleaching, 30-60 time points are collected. Z stack images are collapsed into a maximum projection image for the final time-lapse video.</li>
		</ol>
	</li>
</ol>
6. Representative Results:
<img src="/files/ftp_upload/2852/2852fig1.jpg" alt="Meiosis process diagram; phases with mCherry H2B, GFP Ubiquitin fluorescence microscopy results." data-altupdatedat="1788235205000" data-alt="Figure 1"> 
Figure 1. Fertilization to First Mitosis in C. elegans.
<img src="/files/ftp_upload/2852/2852fig2.jpg" alt="Gonad injection process, microscope images: surface view, rachis, needle insertion, post-injection result." data-altupdatedat="1788235205000" data-alt="Figure 2"> 
Figure 2. Microinjection procedure.
Supplemental Video 1. Time-lapse video of GFP:ubiquitin plus Lysotracker Blue in a meiotic embryo. <a href="https://www.jove.com/files/ftp_upload/2852/2852fig3.avi">Click here to view the video.</a>

<ol><li>Berkowitz, L. A., Knight, A. L., Caldwell, G. A., Caldwell, K. A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Generation+of+stable+transgenic+C.+elegans+using+microinjection%5BTitle%5D)+AND+%22J+Vis+Exp%22%5BJournal%5D)">Generation of stable transgenic C. elegans using microinjection.</a> J Vis Exp. (18), e833-e833 (2008).</li>
<li>Hochbaum, D., Ferguson, A. A., Fisher, A. L. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Generation+of+Transgenic+C.+elegans+by+Biolistic+Transformation%5BTitle%5D)+AND+%22J+Vis+Exp%22%5BJournal%5D)">Generation of Transgenic C. elegans by Biolistic Transformation.</a> J Vis Exp. (42), e2090-e2090 (2010).</li>
<li>Brenner, S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((The+Genetics+of+Caenrohabditis+elegans%5BTitle%5D)+AND+%22Genetics%22%5BJournal%5D)">The Genetics of Caenrohabditis elegans.</a> Genetics. 77, 71-94 (1974).</li>
<li>Cell division WormBook. Oegema, K., Hyman, A. A. , WormBook. (2006).</li>
<li>McNally, K. L., McNally, F. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Fertilization+initiates+the+transition+from+anaphase+I+to+metaphase+II+during+female+meiosis+in+C.+elegans%5BTitle%5D)+AND+%22Dev+Biol%22%5BJournal%5D)">Fertilization initiates the transition from anaphase I to metaphase II during female meiosis in C. elegans.</a> Dev Biol. 282, 218-230 (2005).</li>
<li>Sato, K., Sato, M., Audhya, A., Oegema, K., Schweinsberg, P., Grant, B. D. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Dynamic+regulation+of+caveolin-1+trafficking+in+the+germ+line+and+embryo+of+Caenorhabditis+elegans%5BTitle%5D)+AND+%22Mol+Biol+Cell%22%5BJournal%5D)">Dynamic regulation of caveolin-1 trafficking in the germ line and embryo of Caenorhabditis elegans.</a> Mol Biol Cell. 17, 3085-3094 (2006).</li>
<li>Stitzel, M. L., Pellettieri, J., Seydoux, G. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((The+C.+elegans+DYRK+Kinase+MBK-2+Marks+Oocyte+Proteins+for+Degradation+in+Response+to+Meiotic+Maturation%5BTitle%5D)+AND+%22Curr+Biol%22%5BJournal%5D)">The C. elegans DYRK Kinase MBK-2 Marks Oocyte Proteins for Degradation in Response to Meiotic Maturation.</a> Curr Biol. 16, 56-62 (2006).</li>
<li>Schetter, A., Askjaer, P., Piano, F., Mattaj, I., Kemphues, K. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Nucleoporins+NPP-1%2C+NPP-3%2C+NPP-4%2C+NPP-11+and+NPP-13+are+required+for+proper+spindle+orientation+in+C.+elegans%5BTitle%5D)+AND+%22Dev.+Biol%22%5BJournal%5D)">Nucleoporins NPP-1, NPP-3, NPP-4, NPP-11 and NPP-13 are required for proper spindle orientation in C. elegans.</a> Dev. Biol. 289, 360-371 (2006).</li>
<li>Samuel, A. D., Murthy, V. N., Hengartner, M. O. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Calcium+dynamics+during+fertilization+in+C.+elegans%5BTitle%5D)+AND+%22BMC+Dev+Biol%22%5BJournal%5D)">Calcium dynamics during fertilization in C. elegans.</a> BMC Dev Biol. 1, (2001).</li>
<li>Badrinath, A. S., White, J. S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Contrasting+patterns+of+mitochondrial+redistribution+in+the+early+lineages+of+Caenorhabditis+elegans+and+Acrobeloides+sp.+PS1146%5BTitle%5D)+AND+%22Dev.+Biol%22%5BJournal%5D)">Contrasting patterns of mitochondrial redistribution in the early lineages of Caenorhabditis elegans and Acrobeloides sp. PS1146.</a> Dev. Biol. 258, 270-275 (2003).</li>
<li>Johnston, W. L., Krizus, A., Dennis, J. W. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((The+eggshell+is+required+for+meiotic+fidelity%2C+polar-body+extrusion+and+polarization+of+the+C.+elegans+embryo%5BTitle%5D)+AND+%22BMC+Biol%22%5BJournal%5D)">The eggshell is required for meiotic fidelity, polar-body extrusion and polarization of the C. elegans embryo.</a> BMC Biol. 4, 35-35 (2006).</li>
<li>Parry, J. M., Velarde, N. V., Lefkovith, A. J., Zegarek, M. H., Hang, J. S., Ohm, J., Klancer, R., Maruyama, R., Druzhinina, M. K., Grant, B. D., Piano, F., Singson, A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((EGG-4+and+EGG-5+Link+Events+of+the+Oocyte-to-Embryo+Transition+with+Meiotic+Progression+in+C.+elegans%5BTitle%5D)+AND+%22Curr+Biol%22%5BJournal%5D)">EGG-4 and EGG-5 Link Events of the Oocyte-to-Embryo Transition with Meiotic Progression in C. elegans.</a> Curr Biol. 19, 1752-1757 (2009).</li>
<li>Centonze, V. E., White, J. G. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Multiphoton+excitation+provides+optical+sections+from+deeper+within+scattering+specimens+than+confocal+imaging%5BTitle%5D)+AND+%22Biophys+J%22%5BJournal%5D)">Multiphoton excitation provides optical sections from deeper within scattering specimens than confocal imaging.</a> Biophys J. 75, 2015-2024 (1998).</li>
<li>Zinselmeyer, B. H., Dempster, J., Wokosin, D. L., Cannon, J. J., Pless, R., Parker, I., Miller, M. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Two-photon+microscopy+and+multidimensional+analysis+of+cell+dynamics%5BTitle%5D)+AND+%22Methods+Enzymol%22%5BJournal%5D)">Two-photon microscopy and multidimensional analysis of cell dynamics.</a> Methods Enzymol. 461, 349-378 (2009).</li>
</ol>

No conflicts of interest declared.

After fusion with the sperm, the zygote undergoes a series of dynamic events. These events transform the oocyte from a relatively 
static cell into the rapidly developing embryo. In many organisms, cytoplasmic rearrangements are critical for establishing embryonic polarity and for 
egg activation. The C. elegans embryo provides an excellent opportunity for observing these early events of egg activation and embryogenesis. The 
embryo is transparent and early developmental events occur relatively rapidly after fertilization. C. elegans researchers have generated many useful 
transgenic strains that express fluorescently labeled proteins involved in early development. Using these strains along with RNAi or mutants provides 
a powerful system for dissecting the molecular pathways that underlie development of a multicellular organism (5, 6, 7, 8). This video article presents 
a practical approach to using these tools and techniques to record events during early embryogenesis in C. elegans. 
The mature C. elegans oocyte is arrested in prophase of meiosis I and is fertilized in the spermatheca of the adult 
hermaphrodite. After fertilization, the oocyte proceeds through the remainder of meiosis I and II. These stages can be observed using fluorescently 
labeled histone H2B (5). During completion of maternal meiosis, the sperm DNA remains condensed (see Figure 1). After the completion of meiosis, the 
maternal and paternal DNA becomes decondensed and the two pronuclei form. The maternal pronucleus migrates towards the paternal pronucleus and they 
join together near the center of the embryo. Mitosis quickly ensues after pronuclear meeting. All of these events occur in less than an hour. Thus, 
time-lapse microscopy of this sequence of events can be readily accomplished. Performing this technique requires certain facility in nematode culturing 
as well as basic microscopy skills in addition to access to a confocal microscope.
The example presented here utilizes a transgenic C. elegans strain that expresses two fluorescent proteins, GFP::ubiquitin and 
mCherry::H2B. A confocal laser scanning microscope is used to observe the dynamic localization of these two proteins. In addition, we show that injection of 
fluorescently labeled Lysotracker can be used to follow the fate of lysosomes in the embryo after fertilization. Injection of a labeled tracker can also be 
performed for visualization of mitochondria or local calcium concentrations (9). In theory, the injection protocol could be used to view any type of 
fluorescently labeled molecule in the embryo. This might include labeled small molecules, proteins, lipids, etc. In some cases, it may not be necessary 
to actually inject labeled trackers as the worms will readily uptake some dyes such as mitotracker (10). However, we have tried both soaking and injection 
of lysotracker and have found much better results with injection for visualization in embryos.
Technical Considerations:
The technical challenges presented with this procedure include the microinjection technique as well as the difficulty in imaging very early embryos. Regarding microinjections, the thickness of the agarose injection pad is critical for successful injections. If the pad is too thick, the worms will dessicate and die quickly. If the pad is too thin, the worms will not stick to the pad during the injection process.
The early embryos can be sensitive to environmental conditions such as temperature and buffer conditions. Wild type embryos should initiate decondensation of the sperm DNA and pronuclear migration within a minute or so of completing meiosis II (4). Within 20 minutes, the first cytokinesis should begin. If embryos are exposed to very high laser intensity or are overheated or crushed during mounting, this process may be disrupted. Such embryos will arrest development. If this occurs, the experiment should be repeated with decreased laser power or a thicker agarose pad.
C. elegans embryos do not secrete their egg shell until shortly after fertilization. During the short time between 
fertilization and the secretion of the egg shell, embryos are not viable outside of the mother (11). Some labs have avoided this challenge by imaging 
embryos in utero (5, 12). In utero imaging offers certain advantages such as the ability to capture events that occur during or just after fertilization. 
However, this technique requires the use of a microscopy system that allows for deep tissue imaging. Systems such as two-photon or multi-photon are especially 
suited for this (13, 14). When using a conventional laser-scanning or spinning disk confocal microscope, the best images are obtained from embryos that have 
been cut out of the adult worm as described here.
Imaging System Considerations:
The type of confocal imaging system used depends upon the users' needs. In general, laser scanning systems are recommended if one wishes to acquire images with the highest resolution possible. On the other hand, spinning disk systems are better suited for imaging highly dynamic processes as image acquisition is relatively rapid and photobleaching/phototoxicity is reduced.
The specific imaging parameters will vary depending upon the experimental design and the confocal imaging system employed. In multidimensional imaging, one needs to consider the number of channels, focal planes, and time-points being collected. Due to the limiting effects of photobleaching and phototoxicity, a finite number of images can be acquired in any given experiment. Thus, if you choose to increase the number of channels to be acquired, you will likely need to reduce the number of focal planes and/or time points and vice versa. The total number of images that can be acquired will be influenced by the intensity of the fluorophore being imaged as well as the sensitivity of the confocal microscope being used. More intense fluorophores and more sensitive instruments will require shorter exposure times and thus produce less photobleaching and phototoxicity.
A large number or imaging systems are available. We will give details on the systems that have been used in our own laboratories. For imaging worm embryos by spinning disk confocal microscopy, we use a Nikon TE2000U inverted microscope equipped with a 60X/1.4 N.A. Plan Apo objective. The microscope is connected to a Yokogawa CSU10 spinning disk unit, a Hamamatsu C9100-13 EM CCD camera, and a Spectral Applied Research LMM5 laser merge module containing four solid state lasers with output at 405, 491, 561 and 655 nm. Using this system we can capture one or two channels, six to twelve focal planes, and 25 to 50 timepoints (at 30 to 60 second intervals). A typical exposure time is 100 to 200 milliseconds.
For imaging embryos with laser scanning confocal microscopy, we use a Zeiss LSM700 equipped with 2 channels and four solid state lasers (405, 488, 555, and 635 nm). This system is attached to a Zeiss Axio Observer inverted microscope with a Plan-Apo 63X/1.4NA objective used for embryo imaging. The system uses the Zeiss ZEN software for image acquisition which also provides limited image processing and analysis. When imaging with two fluorochromes, we typically gather 1-5 focal planes (~0.5 &mu;m apart) and collect a Z stack every 15 seconds. Collections are continued until significant photobleaching has occurred. To generate the final time-lapse video, Z stack images are consolidated using the maximum projection tool in the ZEN software. Videos are exported from the software as avi files.

<table><tbody><tr><td>Egg Buffer </td><td></td><td></td><td>5mM Hepes pH 7.2, 110mM NaCl, 4mM KCl, 5mM Mg Acetate, 5mM CaCl&lt;sub&gt;2&lt;/sub&gt;</td></tr><tr><td>Mitotracker </td><td>Invitrogen</td><td>M-7512</td><td></td></tr><tr><td>Lysotracker</td><td>Invitrogen</td><td>L-7525</td><td></td></tr><tr><td>Injection Capillary Pipet</td><td>Harvard Apparatus</td><td>GC120F-10</td><td></td></tr><tr><td>Needle Puller</td><td>Kopf Instruments</td><td>Model 700C</td><td></td></tr><tr><td>Microinjector Apparatus</td><td>Tritech Research, Inc.</td><td>MINJ-1 microinjection regulator</td><td></td></tr></tbody></table>

time-lapse microscopy of early embryogenesis in caenorhabditis elegans

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.

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 ...

The overall goal of the following experiment is to record subcellular events during early embryogenesis Insino Ratu elegance. This is achieved by using transgenic strains with multiple fluorescently labeled proteins that allow the visualization of DNA as well as subsets of vesicles In the early embryo, specific organelles are labeled by a microinjection of fluorescent dyes, such as Lyo tracker or MIT tracker into the gonad of the adult worm. Next embryos are cut out of adult worms and observed with a confocal microscope to obtain time-lapse recordings of the movements of the labeled proteins in organelles.Ultimately, the protocol reveals interesting movement of lysosomes and GFP tagged vesicles during early development in CL allergans. My name is Connie Hedger, and today I'll be showing you how we perform time-lapse microscopy in our lab, the Boyd lab at the University of Alabama in Huntsville. For this experiment, gro nematodes on NGM agri plates, seated with an OP 50 bacterial lawn at 25 degrees Celsius the day before the experiment, pick at least 40 L four larvae onto seated plates and incubate them overnight at 25 degrees Celsius.These worms will be young adults. For the experiment. Make an injection pad by sandwiching, two drops of 2%aros in water from a past pipette between two cover glass slips, and applying pressure to shape the agros into a 1.5 centimeter diameter circle.Remove the top cover slip and dehydrate the pad by placing it in an 80 degree Celsius oven for one hour, or allow it to sit on the benchtop overnight. On the day of the injection, prepare a diluted MIT tracker or lyo tracker dye solution and pull injection needles from a 1.2 od glass capillary. Backfill the needles with a diluted dye solution and store them in a dark humidified chamber mount injection needle onto the micro manipulator and connect the needle to a pressure regulator with a 1.2 millimeter internal diameter tube.Now place two drops of heavy mineral oil onto the injection pad. Place the injection pad onto the microscope and lower the needle into the oil. Apply injection pressure and observe whether fluid flows out of the needle.If not, you will need to gently break the end of the needle by gently driving the tip into a small bit of broken cover glass placed on the injection pad. After the needle is flowing, proceed with the injection a pro one hour prior to viewing embryos. Use a pic to transfer young adult worms into the oil drop on the injection pad.Arrange three to 10 worms in parallel and a little less than one worm length apart. It is now important to inject the worms quickly before the worms desiccate. Move the pad with worms onto the microscope stage.Focus onto the central part of the distal gonad. Then use the micro manipulator to move the tip of the needle into the same focal plane. Puncture the worm by moving the stage horizontally.When the needle tip is inside the cus apply pressure to fill the gonad with the dye mixture. The gonad will expand with the volume of the injection. Proceed with injecting.Both go net arms of each worm after injection. Cover the worms with 0.5 milliliters of egg buffer applied. Using a pulled pasture pipette, then allow the worms to recover for one to two hours at room temperature in a light free humidified chamber.Begin by preparing a new agros pad for viewing the injected embryos with a past pipette. Put three drops of molten agros onto a clean microscope slide and place that slide between two other slides wrapped in one layer of labeling. Tape the tape controls the thickness of the pad.Next, press the clean microscope slide down onto the agros drops so it rests on the tape slides. Do not remove the top slide until the embryos are ready to mount. Now drop 20 microliters of egg buffer onto the center of a cover glass and transfer the injected worms into the buffer Under a dissecting microscope, cut the worms open near the vulva.Using a 26 gauge hypodermic needle, the embryos will spill out of the worm. Now remove the top slide of the imaging slide and lower the aros pad upside down onto the cover glass with the embryos. If the embryos will be imaged over an extended period of time, seal the edges of the cover glass with petroleum jelly to prevent desiccation.Now, begin imaging by locating embryos under the 10 x objective to find early stage embryos, search for embryos that are in the process of completing maternal meiosis. Myotic embryos can be distinguished from later embryos because they have not undergone shortening. After completing meiosis, there's a significant gap between the cell membrane and the eggshell at the anterior end of the embryo.Once an appropriately staged embryo is identified, move to a higher magnification lens for recording of embryogenesis. This embryo expresses both M cherry histone two B and GFP ubiquitin tagged proteins. The maternal and paternal DNA is visible via the M cherry H two B, and thus shows the stages of early development.This embryo was injected with OC Tracker blue and expresses GFP Ubiquitin. The images were collected using a 63 x 1.4 NA lens with 4 88 and 5 55 lasers set at minimal power and maximum gain. Typically, Zacks are collected every 10 to 15 seconds for up to 60 time points before photobleaching occurs.For this TimeLapse video, the Z stacks were collapsed into maximum projection images While performing time-lapse. Microscopy embryos should begin pronuclear migration within a few minutes of completing myosis two, and within 20 minutes. The first cytokinesis should begin.If you observe arrested development of your embryos, try decreasing the laser intensity or use a thicker augurous pad pad.

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Research

JoVE Journal

Biology

The C. elegans embryo is a powerful system for studying cell biology and development.  We present a protocol for live imaging of C. elegans embryos utilizing DIC optics or fluorescence using readily available epifluorescent microscopes and open-source software.

Imaging C. elegans Embryos using an Epifluorescent Microscope and Open Source Software

The easiness of maintaining and propagating the nematode C. elegans make it a nice model organism to work with. The possibility of synchronizing worms allows the work with a significant amount of subjects at the same developmental stage, what facilitates the study of one particular process in many animals.

Basic Caenorhabditis elegans Methods: Synchronization and Observation

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.

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

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

Here we detail protocols specifically designed to monitor morphogenic defects that occur during early and late phases of embryonic elongation of the nematode Caenorhabditis elegans. Ultimately, these protocols are designed to identify genes that regulate these phases and to characterize their differential requirements along the antero-posterior axis of the embryo.

Novel Metrics to Characterize Embryonic Elongation of the Nematode Caenorhabditis elegans

In this article, we demonstrate live imaging of individual worms employing a custom microfluidic device. In the device, multiple worms are individually confined to separate chambers, allowing multiplexed longitudinal surveillance of various biological processes.

A Microfluidic Platform for Longitudinal Imaging in Caenorhabditis elegans

Here, we present a combinatorial approach using high-resolution microscopy, computational tools, and single-cell labeling in living C. elegans embryos to understand single cell dynamics during neurodevelopment.

Isotropic Light-Sheet Microscopy and Automated Cell Lineage Analyses to Catalogue Caenorhabditis elegans Embryogenesis with Subcellular Resolution

This work describes a semi-high-throughput protocol that allows simultaneous 3D time-lapse imaging of embryogenesis in 80–100 C. elegans embryos in a single overnight run. Additionally, image processing and visualization tools are included to streamline data analysis. The combination of these methods with custom reporter strains enables detailed monitoring of embryogenesis.

A Semi-high-throughput Imaging Method and Data Visualization Toolkit to Analyze C. elegans Embryonic Development

Here, we present a protocol for preparing and mounting Caenorhabditis elegans embryos, recording development under a 4D microscope and tracing cell lineage.

4D Microscopy: Unraveling Caenorhabditis elegans Embryonic Development Using Nomarski Microscopy

Here, we present protocols and tools for visualizing calcium during fertilization and early embryogenesis using a genetically encoded calcium reporter expressed in the germline of the model nematode C. elegans.

In Vivo Visualization of Calcium Transients during Fertilization and Early Development in C. elegans

This protocol describes microfluidic-based C. elegans time-lapse imaging across the entire post-embryonic development.

High-Resolution C. elegans Imaging Across All Larval Stages

Learn time-lapse microscopy for visualizing early embryogenesis in Caenorhabditis elegans using fluorescent labeling and confocal imaging. This protocol enables researchers to study subcellular dynamics and organelle movements in live embryos.

Time-lapse Microscopy of Early Embryogenesis in Caenorhabditis elegans

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