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&deg;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&deg;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&deg;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 &#x2013; 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 &mu;l 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="Figure 1" /> 
Figure 1. Fertilization to First Mitosis in C. elegans.
<img src="/files/ftp_upload/2852/2852fig2.jpg" 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>

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	<li>Berkowitz, L. A., Knight, A. L., Caldwell, G. A., Caldwell, K. A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Generation+of+stable+transgenic+C.+elegans+using+microinjection.">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/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Generation+of+Transgenic+C.+elegans+by+Biolistic+Transformation.">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/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+Genetics+of+Caenrohabditis+elegans.">The Genetics of Caenrohabditis elegans.</a> Genetics. 77, 71-94 (1974).</li><li>Oegema, K., Hyman, A. A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Cell+division+WormBook.">Cell division WormBook.</a> , (2006).</li><li>McNally, K. L., McNally, F. 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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 border="1">
	<tbody>
		<tr>
			<th>Name of the reagent</th>
			<th>Company</th>
			<th>Catalogue number</th>
			<th>Composition</th>
		</tr>
		<tr>
			<td>Egg Buffer </td>
			<td>&nbsp;</td>
			<td>&nbsp;</td>
			<td>5mM Hepes pH 7.2, 110mM NaCl, 4mM KCl, 5mM Mg Acetate, 5mM CaCl2</td>
		</tr>
		<tr>
			<td>Mitotracker </td>
			<td>Invitrogen</td>
			<td>M-7512</td>
			<td>&nbsp;</td>
		</tr>
		<tr>
			<td>Lysotracker</td>
			<td>Invitrogen</td>
			<td>L-7525</td>
			<td>&nbsp;</td>
		</tr>
		<tr>
			<td>Injection Capillary Pipet</td>
			<td>Harvard Apparatus, Kent, UK</td>
			<td>GC120F-10</td>
			<td>&nbsp;</td>
		</tr>
		<tr>
			<td>Needle Puller</td>
			<td>Kopf</td>
			<td>Model 700C</td>
			<td>&nbsp;</td>
		</tr>
		<tr>
			<td>Microinjector Apparatus</td>
			<td>Tritech Research</td>
			<td>MINJ-1 microinjection regulator</td>
			<td>&nbsp;</td>
		</tr>
	</tbody>
</table>

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.

Watch this Scientific Journal Video about Time-lapse Microscopy of Early Embryogenesis in Caenorhabditis elegans at JoVE.com

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vertical immobilization method for time-lapse microscopy analysis in filamentous cyanobacteria

Vertical Immobilization Method for Time-Lapse Microscopy Analysis in Filamentous Cyanobacteria

We present a simple and accessible method for filamentous cyanobacterial visualization in the XY plane. A low-melting point agarose matrix was used, allowing the acquisition of images of proteins involved in the division, in a vertical orientation. Therefore, this methodology can be applied to any filamentous organism and different kinds of proteins.

manipulation of ploidy in caenorhabditis elegans

Manipulation of Ploidy in Caenorhabditis elegans

This method allows for the generation of tetraploid and triploid Caenorhabditis nematodes from any diploid strain. Polyploid strains generated by this method have been used to study chromosome interactions in meiotic prophase, and this method is useful for examining important basic questions in cell, developmental, evolutionary, and cancer...

Manipulation of Ploidy in Caenorhabditis elegans

a layered mounting method for extended time-lapse confocal microscopy of whole zebrafish embryos

A Layered Mounting Method for Extended Time-Lapse Confocal Microscopy of Whole Zebrafish Embryos

This article describes a method to mount fragile zebrafish embryos for extended time-lapse confocal microscopy. This low-cost method is easy to perform using regular glass-bottom microscopy dishes for imaging on any inverted microscope. The mounting is performed in layers of agarose at different...

specific labeling of mitochondrial nucleoids for time-lapse structured illumination microscopy

Specific Labeling of Mitochondrial Nucleoids for Time-lapse Structured Illumination Microscopy

The protocol describes specific labeling of mitochondrial nucleoids with a commercially available DNA gel stain, acquisition of time lapse series of live labeled cells by super-resolution structured illumination microscopy (SR-SIM), and automatic tracking of nucleoid...

automating aggregate quantification in caenorhabditis elegans

Automating Aggregate Quantification in Caenorhabditis elegans

The following protocol describes the development and optimization of a high-throughput workflow for worm culturing, fluorescence imaging, and automated image processing to quantify polyglutamine aggregates as an assessment of changes in proteostasis.

Automating Aggregate Quantification in Caenorhabditis elegans

label-free imaging of lipid storage dynamics in caenorhabditis elegans using stimulated raman scattering microscopy

Label-Free Imaging of Lipid Storage Dynamics in Caenorhabditis elegans using Stimulated Raman Scattering Microscopy

Stimulated Raman scattering (SRS) microscopy allows selective, label-free imaging of specific chemical moieties and it has been effectively employed to image lipid molecules in vivo. Here, we provide a brief introduction to the principle of SRS microscopy and describe methods for its use in imaging lipid storage in Caenorhabditis...

Label-Free Imaging of Lipid Storage Dynamics in Caenorhabditis elegans using Stimulated Raman Scattering Microscopy

measuring embryonic viability and brood size in caenorhabditis elegans

Measuring Embryonic Viability and Brood Size in Caenorhabditis elegans

Here, we present a general method to determine the embryonic viability and total number of embryos produced (brood) using the model organism C.

Measuring Embryonic Viability and Brood Size in Caenorhabditis elegans

ratiometric calcium imaging of individual neurons in behaving caenorhabditis elegans

Ratiometric Calcium Imaging of Individual Neurons in Behaving Caenorhabditis Elegans

This protocol describes the use of genetically encoded Ca2+ reporters to record changes in neural activity in behaving Caenorhabditis elegans worms.

Ratiometric Calcium Imaging of Individual Neurons in Behaving Caenorhabditis Elegans

measurement of oxygen consumption rates in intact caenorhabditis elegans

Measurement of Oxygen Consumption Rates in Intact Caenorhabditis elegans

Mitochondrial respiration is critical for organismal survival; therefore, oxygen consumption rate is an excellent indicator of mitochondrial health. In this protocol, we describe the use of a commercially available respirometer to measure basal and maximal oxygen consumption rates in live, intact, and freely-motile Caenorhabditis...

Measurement of Oxygen Consumption Rates in Intact Caenorhabditis elegans

cultivation of caenorhabditis elegans in three dimensions in the laboratory

Cultivation of Caenorhabditis elegans in Three Dimensions in the Laboratory

We present a simple method to construct 3D nematode cultivation systems called NGT-3D and NGB-3D. These can be used to study nematode fitness and behaviors in habitats that are more similar to natural Caenorhabditis elegans habitats than the standard 2D laboratory C. elegans culture...

Cultivation of Caenorhabditis elegans in Three Dimensions in the Laboratory

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 ...

Isolation and In vitro Activation of Caenorhabditis elegans Sperm

Males and hermaphrodites are the two naturally found sexual forms in the nematode C. elegans. The amoeboid sperm are produced by both males and hermaphrodites. In the earlier phase of gametogenesis, the germ cells of hermaphrodites differentiate into limited number of sperm - around 300 - and are stored in a small 'bag' called the spermatheca. Later on, hermaphrodites continually produce oocytes1. In contrast, males produce exclusively sperm throughout their adulthood. The males produce so much sperm that it accounts for &gt;50% of the total cells in a typical adult worm2. Therefore, isolating sperm from males is easier than from that of hermaphrodites.
Only a small proportion of males are naturally generated due to spontaneous non-disjunction of X chromosome3. Crossing hermaphrodites with males or more conveniently, the introduction of mutations to give rise to Him (High Incidence of Males) phenotype are some of strategies through which one can enrich the male population3.
Males can be easily distinguished from hermaphrodites by observing the tail morphology4. Hermaphrodite's tail is pointed, whereas male tail is rounded with mating structures.
Cutting the tail releases vast number of spermatids stored inside the male reproductive tract. Dissection is performed under a stereo microscope using 27 gauge needles. Since spermatids are not physically connected with any other cells, hydraulic pressure expels internal contents of male body, including spermatids2.
Males are directly dissected on a small drop of 'Sperm Medium'. Spermatids are sensitive to alteration in the pH. Hence, HEPES, a compound with good buffering capacity is used in sperm media. Glucose and other salts present in sperm media help maintain osmotic pressure to maintain the integrity of sperm.
Post-meiotic differentiation of spermatids into spermatozoa is termed spermiogenesis or sperm activation. Shakes5, and Nelson6 previously showed that round spermatids can be induced to differentiate into spermatozoa by adding various activating compounds including Pronase E. Here we demonstrate in vitro spermiogenesis of C. elegans spermatids using Pronase E.
Successful spermiogenesis is pre-requisite for fertility and hence the mutants defective in spermiogenesis are sterile. Hitherto several mutants have been shown to be defective specifically in spermiogenesis process7. Abnormality found during in vitro activation of novel Spe (Spermatogenesis defective) mutants would help us discover additional players participating in this event.

Isolation and In vitro Activation of Caenorhabditis elegans Sperm

A protocol for isolating and activating spermatids from male C. elegans is described here. Cutting the posterior end of male releases spermatids. The spermatids can be activated by addition of protease.

Males and hermaphrodites are the two naturally found sexual forms in the nematode C. elegans. The amoeboid sperm are produced by both males and ...

Quantitative Analysis of Random Migration of Cells Using Time-lapse Video Microscopy

Cell migration is a dynamic process, which is important for embryonic development, tissue repair, immune system function, and tumor invasion 1, 2. During directional migration, cells move rapidly in response to an extracellular chemotactic signal, or in response to intrinsic cues 3 provided by the basic motility machinery. Random migration occurs when a cell possesses low intrinsic directionality, allowing the cells to explore their local environment.
Cell migration is a complex process, in the initial response cell undergoes polarization and extends protrusions in the direction of migration 2. Traditional methods to measure migration such as the Boyden chamber migration assay is an easy method to measure chemotaxis in vitro, which allows measuring migration as an end point result. However, this approach neither allows measurement of individual migration parameters, nor does it allow to visualization of morphological changes that cell undergoes during migration.
Here, we present a method that allows us to monitor migrating cells in real time using video - time lapse microscopy. Since cell migration and invasion are hallmarks of cancer, this method will be applicable in studying cancer cell migration and invasion in vitro. Random migration of platelets has been considered as one of the parameters of platelet function 4, hence this method could also be helpful in studying platelet functions. This assay has the advantage of being rapid, reliable, reproducible, and does not require optimization of cell numbers. In order to maintain physiologically suitable conditions for cells, the microscope is equipped with CO2 supply and temperature thermostat. Cell movement is monitored by taking pictures using a camera fitted to the microscope at regular intervals. Cell migration can be calculated by measuring average speed and average displacement, which is calculated by Slidebook software.

This method allows monitoring of cells in real time and quantitative measurements of different cell migration parameters such as speed, displacement, and velocity. Unlike the traditional methods, this real time approach is not based on endpoint quantitative migration measurements; instead it allows monitoring and calculating different parameters continuously.

Cell migration is a dynamic process, which is important for embryonic development, tissue repair, immune system function, and tumor invasion 1, 2. During ...

Basic Caenorhabditis elegans Methods: Synchronization and Observation

Research into the molecular and developmental biology of the nematode Caenorhabditis elegans was begun in the early seventies by Sydney Brenner and it has since been used extensively as a model organism 1. C. elegans possesses key attributes such as simplicity, transparency and short life cycle that have made it a suitable experimental system for fundamental biological studies for many years 2. Discoveries in this nematode have broad implications because many cellular and molecular processes that control animal development are evolutionary conserved 3.
C. elegans life cycle goes through an embryonic stage and four larval stages before animals reach adulthood. Development can take 2 to 4 days depending on the temperature. In each of the stages several characteristic traits can be observed. The knowledge of its complete cell lineage 4,5 together with the deep annotation of its genome turn this nematode into a great model in fields as diverse as the neurobiology 6, aging 7,8, stem cell biology 9 and germ line biology 10. 
An additional feature that makes C. elegans an attractive model to work with is the possibility of obtaining populations of worms synchronized at a specific stage through a relatively easy protocol. The ease of maintaining and propagating this nematode added to the possibility of synchronization provide a powerful tool to obtain large amounts of worms, which can be used for a wide variety of small or high-throughput experiments such as RNAi screens, microarrays, massive sequencing, immunoblot or in situ hybridization, among others. 
Because of its transparency, C. elegans structures can be distinguished under the microscope using Differential Interference Contrast microscopy, also known as Nomarski microscopy. The use of a fluorescent DNA binder, DAPI (4',6-diamidino-2-phenylindole), for instance, can lead to the specific identification and localization of individual cells, as well as subcellular structures/defects associated to them.

Basic Caenorhabditis elegans Methods: Synchronization and Observation

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.

Research into the molecular and developmental biology of the nematode Caenorhabditis elegans was begun in the early seventies by Sydney Brenner and it has ...

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.

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 ...

Prostaglandin Extraction and Analysis in Caenorhabditis elegans

Caenorhabditis elegans is emerging as a powerful animal model to study the biology of lipids1-9. Prostaglandins are an important class of eicosanoids, which are lipid signals derived from polyunsaturated fatty acids (PUFAs)10-14. These signalling molecules are difficult to study because of their low abundance and reactive nature. The characteristic feature of prostaglandins is a cyclopentane ring structure located within the fatty acid backbone. In mammals, prostaglandins can be formed through cyclooxygenase enzyme-dependent and -independent pathways10,15. C. elegans synthesizes a wide array of prostaglandins independent of cyclooxygenases6,16,17. A large class of F-series prostaglandins has been identified, but the study of eicosanoids is at an early stage with ample room for new discoveries. Here we describe a procedure for extracting and analyzing prostaglandins and other eicosanoids. Charged lipids are extracted from mass worm cultures using a liquid-liquid extraction technique and analyzed by liquid chromatography coupled to electrospray ionization tandem mass spectrometry (LC-ESI-MS/MS). The inclusion of deuterated analogs of prostaglandins, such as PGF2 &alpha;-d4 as an internal standard is recommended for quantitative analysis. Multiple reaction monitoring or MRM can be used to quantify and compare specific prostaglandin types between wild-type and mutant animals. Collision-induced decomposition or MS/MS can be used to obtain information on important structural features. Liquid chromatography mass spectrometry (LC-MS) survey scans of a selected mass range, such as m/z 315-360 can be used to evaluate global changes in prostaglandin levels. We provide examples of all three analyses. These methods will provide researchers with a toolset for discovering novel eicosanoids and delineating their metabolic pathways.

Prostaglandin Extraction and Analysis in Caenorhabditis elegans

In this paper, we describe an optimized procedure for extracting and analyzing prostaglandins and other eicosanoids from C. elegans using LC-MS/MS.

Caenorhabditis elegans is emerging as a powerful animal model to study the biology of lipids1-9. Prostaglandins are an important class of eicosanoids, ...

The use of genetic model organisms such as Caenorhabditis elegans has led to seminal discoveries in biology over the last five decades. Most of what we know about C. elegans is limited to laboratory cultivation of the nematodes that may not necessarily reflect the environments they normally inhabit in nature. Cultivation of C. elegans in a 3D habitat that is more similar to the 3D matrix that worms encounter in rotten fruits and vegetative compost in nature could reveal novel phenotypes and behaviors not observed in 2D. In addition, experiments in 3D can address how phenotypes we observe in 2D are relevant for the worm in nature. Here, a new method in which C. elegans grows and reproduces normally in three dimensions is presented. Cultivation of C. elegans in Nematode Growth Tube-3D (NGT-3D) can allow us to measure the reproductive fitness of C. elegans strains or different conditions in a 3D environment. We also present a novel method, termed Nematode Growth Bottle-3D (NGB-3D), to cultivate C. elegans in 3D for microscopic analysis. These methods allow scientists to study C. elegans biology in conditions that are more reflective of the environments they encounter in nature. These can help us to understand the overlying evolutionary relevance of the physiology and behavior of C. elegans we observe in the laboratory.

We present a simple method to construct 3D nematode cultivation systems called NGT-3D and NGB-3D. These can be used to study nematode fitness and behaviors in habitats that are more similar to natural Caenorhabditis elegans habitats than the standard 2D laboratory C. elegans culture plates.

The use of genetic model organisms such as Caenorhabditis elegans has led to seminal discoveries in biology over the last five decades. Most of what we ...

Optimal mitochondrial function is critical for healthy cellular activity, particularly in cells that have high energy demands like those in the nervous system and muscle. Consistent with this, mitochondrial dysfunction has been associated with a myriad of neurodegenerative diseases and aging in general. Caenorhabditis elegans have been a powerful model system for elucidating the many intricacies of mitochondrial function. Mitochondrial respiration is a strong indicator of mitochondrial function and recently developed respirometers offer a state-of-the-art platform to measure respiration in cells. In this protocol, we provide a technique to analyze live, intact C. elegans. This protocol spans a period of ~7 days and includes steps for (1) growing and synchronization of C. elegans, (2) preparation&#160;of compounds to be injected and hydration of probes, (3) drug loading and cartridge equilibration, (4) preparation of worm assay plate and assay run, and (5) post-experiment data analysis.

Mitochondrial respiration is critical for organismal survival; therefore, oxygen consumption rate is an excellent indicator of mitochondrial health. In this protocol, we describe the use of a commercially available respirometer to measure basal and maximal oxygen consumption rates in live, intact, and freely-motile Caenorhabditis elegans.

Optimal mitochondrial function is critical for healthy cellular activity, particularly in cells that have high energy demands like those in the nervous ...

Mitochondrial nucleoids are compact particles formed by mitochondrial DNA molecules coated with proteins. Mitochondrial DNA encodes tRNAs, rRNAs, and several essential mitochondrial polypeptides. Mitochondrial nucleoids divide and distribute within the dynamic mitochondrial network that undergoes fission/fusion and other morphological changes. High resolution live fluorescence microscopy is a straightforward technique to characterize a nucleoid&#39;s position and motion. For this technique, nucleoids are commonly labeled through fluorescent tags of their protein components, namely transcription factor a (TFAM). However, this strategy needs overexpression of a fluorescent protein-tagged construct, which may cause artifacts (reported for TFAM), and is not feasible in many cases. Organic DNA-binding dyes do not have these disadvantages. However, they always show staining of both nuclear and mitochondrial DNAs, thus lacking specificity to mitochondrial nucleoids. By taking into account the physico-chemical properties of such dyes, we selected a nucleic acid gel stain (SYBR Gold) and achieved preferential labeling of mitochondrial nucleoids in live cells. Properties of the dye, particularly its high brightness upon binding to DNA, permit subsequent quantification of mitochondrial nucleoid motion using time series of super-resolution structured illumination images.

The protocol describes specific labeling of mitochondrial nucleoids with a commercially available DNA gel stain, acquisition of time lapse series of live labeled cells by super-resolution structured illumination microscopy (SR-SIM), and automatic tracking of nucleoid motion.

Mitochondrial nucleoids are compact particles formed by mitochondrial DNA molecules coated with proteins. Mitochondrial DNA encodes tRNAs, rRNAs, and ...