Name: imaging c. elegans embryos using an epifluorescent microscope and open source software
Uploaded: 2011-03-24T13:00:00
Duration: 8 min 32 s
Description: Watch this Scientific Journal Video about Imaging C. elegans Embryos using an Epifluorescent Microscope and Open Source Software at JoVE.com

Internal institutional funds and the Biological Sciences Scholars Program at the University of Michigan supported this work.

Worm Preparation
<ol>
	<li>Transfer L4 larval worms to appropriate plates 18-24hrs prior to imaging22. This will ensure you have egg-bearing adults for an ample supply of early embryos.</li>
	<li>Prepare tubes of Vaseline and Agarose. We prepare several 15mL falcon tubes by melting Vaseline in a beaker in the microwave and dispensing into 4-5 mL aliquot per tube. Similarly melt 2-3% (w/v) Agarose in Buffer (M9 or Egg Buffer) and aliquot 4-5 mL into falcon tubes. Be careful not to boil to excess in order to minimize evaporation. On day of imaging, melt 1 tube of Vaseline in a 65-70&deg;C heat block and melt 1 tube of agarose in the microwave, again be careful to minimize boiling over. Store the tubes in the heat block to keep the Vaseline and the Agarose in the molten state.</li>
	<li>Make Agarose pad. Position a new microscopy slide between 2 guide slides, which are slides covered by a single layer of common laboratory tape adhered to the top and bottom of each guide slide. Use a Pasteur pipette with end broken off to place 2-3 drops of melted Agarose on the new slide. Cover with a 2nd slide perpendicular to the original slide, so that it covers and helps to spread the Agarose to create a thin square pad. The 2nd slide should rest above the tape on the guide slides. The Agarose concentration may be increased to 5% for thicker pads.</li>
	<li>Using a dissecting scope and a platinum wire "pick", transfer 2 adult worms to a 9-12&mu;L drop of M9 or Egg Buffer on an 18 mm x 18 mm coverslip.</li>
	<li>Cut worms open with a needle (we use 27G1/2 needles) or scalpel to release embryos. Try to cut in the middle of the worm.</li>
	<li>Lower Agarose pad (with the agar side down) onto the 18x18 mm coverslip containing the embryos. Trim agar extending from the edge of the cover slip with a razor blade.</li>
	<li>Seal Coverslip by using a paintbrush or thin wooden stick to apply the melted Vaseline to all four edges of the coverslip.</li>
</ol>
Alternative to agar mounts are hanging drops23 if embryos are susceptible to pressure (i.e. if the egg shell does not form properly). Imaging can also be performed in utero to follow fertilization or early events such as meiosis. Worms should be anesthetized with Levamisole24. In order to increase the number of embryos from a particular stage of development, several worms may be dissected at one time and the embryos positioned together gently using a fine-tip pipette or an eyelash brush or by mouth pipetting.
4D Nomarski DIC (Differential Interference Contrast) Movies
<ol start="8">
	<li>Turn on the Olympus BX61 microscope. Because the camera is sensitive to EM waves emitted by other devices it is turned on last, the mercury bulb (if necessary for GFP or fluorescence) first. Start Micro-Manager once everything is on. Choose the appropriate configuration file for DIC or fluorescent imaging. The major difference is that the DIC file does not include control of fluorescent shutter. Each file also selects the appropriate filter cube and camera gain setting.</li>
	<li>Find embryos on the microscope using 10x objective. Look near worm bodies for groups of young embryos in order to get multiple embryos within the imaging field. Avoid embryos inside the worm for best images.</li>
	<li>Using the software switch to a higher magnification (40x to 100x) and resolution (N.A. 0.9 to 1.4) objective for imaging.</li>
	<li>Adjust Nomarski DIC Optics (The following web sites contain helpful explanation of DIC optics and the terminology; <a href="http://www.olympusmicro.com/primer/techniques/dic/dichome.html">http://www.olympusmicro.com/primer/techniques/dic/dichome.html</a> 
	http://microscopy.berkeley.edu/Resources/instruction/DIC.html)
	<ol class="lalpha">
		<li>On our Olympus BX61 the important components are:
		<ol>
			<li>Polarizer - slider below the condenser</li>
			<li>Condenser Wollaston Prism - turret on the condenser, just below the stage.</li>
			<li>Objective Wollaston Prism - slider below the filter cubes</li>
			<li>2nd Polarizer (Analyzer) - filter cube in the DIC position</li>
		</ol>
		</li>
		<li>Switch Filter Cube to DIC to position the Analyzer (2nd polarizer) in the light path (software should do this on start up).</li>
		<li>Make sure that the two polarizers are properly aligned (crossed). When both Wollaston prisms are out of the light path the field of view should be dark. If necessary, rotate the polarizer below the condenser until this is correct.</li>
		<li>Insert the objective (upper) Wollaston Prism into the light path.</li>
		<li>Rotate the turret on the condenser to select the Wollaston prism that matches the objective you are using.</li>
		<li>Adjust slider below this turret to match NA of lens.</li>
		<li>Focus the condenser for optimal Koehler illumination.</li>
		<li>Adjust the upper Wollaston Prism to get optimal contrast by rotating the knob on the slider.</li>
		<li>Other settings controlled by software startup include camera gain (set to 0) and switch to saving files as 8-bit not 16-bit tiff files.</li>
	</ol>
	</li>
	<li>Adjust light level to give nice image. (usually use 4.7 Volts) (controlled via software). We prefer to illuminate the sample so that the brightest pixels are just below saturation levels. 
	Good DIC optics results in an image with a 3-dimensional look where the yolk granules in the embryo stand out clearly. Alternatively this can be described as if the light illuminating the sample appears to shine from an angle so that one side of the sample (or features in the sample) is lit brightly and the opposite side is in shadows and appears darker.</li>
	<li>Select Region Of Interest (ROI) to include the embryos you want to image, click set ROI button to reduce window.</li>
	<li>Open the multi-dimensional acquisition window. Select Z-stacks (slices) and Time Points.</li>
	<li>Use microscope focus knob to find the bottom of the embryo (last focal plane with some yolk granules in focus). Click on Select bottom. Repeat to identify and set the top of the embryo. We usually use 1 micron step size and end up with ~20 focal planes. We generally collect multiple focal planes at each time point because the embryos and cells are relatively thick. In addition especially when cells begin to divide in different orientations they can displace other cells around in the embryo moving them in or out of the original focal plane. If your microscope does not have a focus motor you can manually adjust the focus to follow the cell or process you are interested.</li>
	<li>Input number of time points and time interval needed. We usually use 15 seconds interval and set a max of 500 time points. We start with more time points than we'll need and stop the acquisition when desired. If you are imaging multiple focal planes, make sure there is enough time to acquire all of the focal planes in the interval between time points.</li>
	<li>Select or create a location to save the data. Make sure that software is set to display last image only. Give the file a name and click Acquire. Monitor the automated image acquisition for the first few time points to ensure it is operating properly.</li>
	<li>Click stop when you wish to stop the acquisition, switch to 10x objective and prepare next slide.</li>
	<li>When done imaging remove DIC components from the light path. Clean oil off objective if necessary. Turn everything off starting with the camera.</li>
</ol>
GFP Movies
<ol start="20">
	<li>Start microscope and find embryos as above. Use configuration file that includes software control of fluorescent shutter.</li>
	<li>Make sure that the upper DIC prism (slider) is not in the light path.</li>
	<li>Switch Filter Cube to FITC/GFP (automatically set on startup)</li>
	<li>Change Camera gain to 255 and image file to 16-bit Tiff (automatically set on startup with this configuration file).</li>
	<li>Turn off white light.</li>
	<li>Click Live Imaging to preview image. Work quickly to minimize light exposure. Alternatively use Snap Image to collect single image.</li>
	<li>Adjust Mercury bulb power/neutral density filters, exposure length to get a good image. In general, you should use minimum light reaching the sample to acquire the data you want to reduce photodamage to the cell and photobleaching of the fluorescence signals.</li>
	<li>Set up time interval, number of time points and number of focal planes. We often use 5-10 second intervals and 1-3 focal plane.</li>
	<li>Select ROI and set parameters for image acquisition as described earlier in Steps 13-19.</li>
</ol>
Analyzing Movies
<ol start="29">
	<li>Data will be saved to folder with the name you entered and contain a series of tiff files for each image of the 4D stack, a text file and an xml file containing metadata such as exposure levels and time intervals.</li>
	<li>The tiff images can be opened by ImageJ via two methods. The first (method a) is the simplest and will also read the metadata of each file. The second (method b) is able to use virtual memory to open files that are larger than the amount of memory allocated to ImageJ, but it does not include the metadata.
	<ol class="lalpha">
		<li>Using Micro-Manager Plugin
		<ol>
			<li>Micro-Manager is built on and uses ImageJ (it installs a new version of ImageJ which will be different from any other version that you may have previously installed) and includes a plugin to open data. Alternatively you can copy the content of the Micro-Manager-1.3/plugins folder to the plugins folder for another version of ImageJ.</li>
			<li>Go to Plugins menu, select Micro-Manager and then Open Micro-Manager File.</li>
			<li>Select the folder containing the data you wish to view and click ok.</li>
			<li>Play through the Z and T dimensions.</li>
		</ol>
		</li>
		<li>Using LOCI plugin to open movies with BioFormats Importer.
		<ol>
			<li>Transfer the xml and txt files from the folder containing the tiff files, and rename the two files to match the name of the data set.</li>
			<li>Within ImageJ, go to the Plugins pull down menu, select LOCI and then either Data Browser or Bio-Formats Importer.</li>
			<li>Find the folder with your tiff stacks. Click on the first tiff file, then open.</li>
			<li>View stack with Hyperstack. Under Dataset organization: Select Group Files with Similar names, Swap dimensions and Open all series. Under Memory management: Select: Use virtual stack (especially for large data sets). Click ok</li>
			<li>The plugin will determine the dimensions of the data set (# of time points and focal planes) and display the range between a set of &lt;&gt; brackets. Click ok or alter the number of time points and/or focal planes to open.</li>
			<li>Confirm which dimension corresponds to each series.</li>
			<li>Play through the Z and T dimensions.</li>
		</ol>
		</li>
	</ol>
	</li>
</ol>
Representative Results
<img src="/files/ftp_upload/2625/2625fig1.jpg" alt="Figure 1" /> 
Figure 1. Stills from a Nomarski DIC movie illustrating normal cellular events during early C. elegans embryo development: pronuclear migration (0-5:45), asymmetric first division (6:45-14:45) and asynchronous second division (14:45-27:00). Data was collected with a 60x 1.35 NA lens. 20 focal planes at 1 micron intervals were collected every 15 seconds with 40 &mu;s exposure. Images were rotated so that posterior is to the right and ventral is down. Scale Bar represent 10 microns.
<img src="/files/ftp_upload/2625/2625fig2.jpg" alt="Figure 2" /> 
Figure 2. Stills from a fluorescent movie illustrating the dynamic movements of a GFP-labeled subunit of the chromosome passenger complex (AIR-2) present on chromosomes from metaphase to early anaphase (A to C) before appearing on the spindle midzone in anaphase until cytokinesis completes (C to F). Data was collected with a 60x 1.35 NA lens. A single focal plane was collected every 10 seconds with 50 &mu;sec exposure. Images were rotated so that posterior is to the right. Scale Bar represent 10 microns.
Movie 1 Nomarski Movie of Wild-Type Embryo. 
Movie corresponding to Figure 1. Posterior is to the bottom right and ventral is to the lower left. Shows a single focal plane of original data set. Image was collected every 15 seconds and play back is set at 14 frames per second. <a href="https://www.jove.com/files/ftp_upload/2625/Movie1.mp4">Click here to watch Video</a>
Movie 2 Fluorescent Movie of Wild-Type embryo expressing GFP::AIR-2. 
Movie corresponding to Figure 2. Posterior is to the upper right. Movie was collected as a single focal plane. Image was collected every 10 seconds and play back is set at 14 frames per second. <a href="https://www.jove.com/files/ftp_upload/2625/Movie2.mp4">Click here to watch Video</a>

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No conflicts of interest declared.

A major consideration for live time-lapse imaging is to preserve the integrity and viability of the cell and the organism. DIC microscopy provides the benefit that the sample is not exposed to ultraviolet light and excessive heating from the lighting source. DIC microscopy is well suited to detect cell migration and cell shape changes such as cytokinesis and to identify some subcellular structures, such as the mitotic spindle and nuclear membrane. Fluorescence imaging of reporter proteins complements DIC microscopy by identifying additional subcellular compartments and allowing visualization of specific proteins. To mitigate the hazards of phototoxicity and damage to the embryo during fluorescence imaging, we usually increase the digital gain of the camera to offset the reduced UV light intensity and duration of illumination. For faint fluorescent transgenic reporters, deconvolution algorithms to reassign out-of-focus light or deblurring algorithms to reduce background can improve the quality of the captured images. While advanced microscopes such as confocal or multiphoton systems are sometimes necessary, we have found that many fluorescent reporters can be imaged using this epifluorescent microscope setup, yielding high quality images.
Micro-Manager (http://www.micro-manager.org), in addition to being free, saves files directly to tiff format instead of proprietary file formats of many commercial software packages. This significantly simplifies the analyses of our data by eliminating the file conversion step needed to read the image files in ImageJ, Photoshop and other image editing software.

Olympus BX61 microscope with Hamamatsu Orca-ER camera. 10x Plan Fluorite (NA 0.3) and 60x PlanAchromat (NA 1.35) objective lenses
Micro-Manager 1.3.46(http://www.micro-manager.org)
Image J 1.43 (http://rsbweb.nih.gov/ij)
loci_tools.jar (http://www.loci.wisc.edu)
Agarose - molecular biology/gel electrophoresis grade
Fisher Superfrost/Plus Microscope Slides (Catalog # 12-550-15)
Laboratory tape (Fisher Catalog # 15-901 Series)
18 mm x 18 mm #1.5 Coverslips
M9 (242 mM KH2PO4, 40 mM Na2HPO4, 9 mM g NaCl, 19 mM NH4Cl MgSO4)
(http://130.15.90.245/wormlab_recipe_book.htm).
Egg Buffer (118 mM NaCl, 48 mM KCl, 2 mM CaCl2, 2 mM MgCl2, 25 mM HEPES, pH 7.3)25.
Vaseline
Brush/Stick
Razor blades
Heat block
Dissecting microscope
Worms22
Platinum wire pick22
27G1/2 Precision Glide Needle (Beckton Dickinson) or a scalpel with a small blade

imaging c. elegans embryos using an epifluorescent microscope and open source software

Cellular processes, such as chromosome assembly, segregation and cytokinesis,are inherently dynamic. Time-lapse imaging of living cells, using fluorescent-labeled reporter proteins or differential interference contrast (DIC) microscopy, allows for the examination of the temporal progression of these dynamic events which is otherwise inferred from analysis of fixed samples1,2. Moreover, the study of the developmental regulations of cellular processes necessitates conducting time-lapse experiments on an intact organism during development. The Caenorhabiditis elegans embryo is light-transparent and has a rapid, invariant developmental program with a known cell lineage3, thus providing an ideal experiment model for studying questions in cell biology4,5and development6-9. C. elegans is amendable to genetic manipulation by forward genetics (based on random mutagenesis10,11) and reverse genetics to target specific genes (based on RNAi-mediated interference and targeted mutagenesis12-15). In addition, transgenic animals can be readily created to express fluorescently tagged proteins or reporters16,17. These traits combine to make it easy to identify the genetic pathways regulating fundamental cellular and developmental processes in vivo18-21. In this protocol we present methods for live imaging of C. elegans embryos using DIC optics or GFP fluorescence on a compound epifluorescent microscope. We demonstrate the ease with which readily available microscopes, typically used for fixed sample imaging, can also be applied for time-lapse analysis using open-source software to automate the imaging process.

Watch this Scientific Journal Video about Imaging C. elegans Embryos using an Epifluorescent Microscope and Open Source Software at JoVE.com

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

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

Cellular processes, such as chromosome assembly, segregation and cytokinesis,are inherently dynamic. Time-lapse imaging of living cells, using ...

imaging-c-elegans-embryos-using-an-epifluorescent-microscope-open

Research

JoVE Journal

Biology

Generation of Stable Transgenic C. elegans Using Microinjection

Transgenic Caenorhabditis elegans can be readily created via microinjection of a DNA plasmid solution into the gonad 1. The plasmid DNA rearranges to form extrachromosomal concatamers that are stably inherited, though not with the same efficiency as actual chromosomes 2. A gene of interest is co-injected with an obvious phenotypic marker, such as rol-6 or GFP, to allow selection of transgenic animals under a dissecting microscope. The exogenous gene may be expressed from its native promoter for cellular localization studies. Alternatively, the transgene can be driven by a different tissue-specific promoter to assess the role of the gene product in that particular cell or tissue. This technique efficiently drives gene expression in all tissues of C. elegans except for the germline or early embryo 3. Creation of transgenic animals is widely utilized for a range of experimental paradigms. This video demonstrates the microinjection procedure to generate transgenic worms. Furthermore, selection and maintenance of stable transgenic C. elegans lines is described.

This video demonstrates the technique of microinjection into the gonad of C. elegans to create transgenic animals.

Transgenic Caenorhabditis elegans can be readily created via microinjection of a DNA plasmid solution into the gonad 1.  The plasmid DNA rearranges to ...

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

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

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

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

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

Assembly, Tuning and Use of an Apertureless Near Field Infrared Microscope for Protein Imaging

This paper aims to instruct the reader in the assembly and operation of an infrared near-field microscope for imaging beyond the diffraction limit.  The apertureless near-field microscope is a light scattering-type instrument that provides infrared spectra at circa 20 nm resolution. A complete list of components and a step-by-step protocol for use is provided.  Common errors in assembly and instrument tuning are discussed.  A representative data set that shows the secondary structure of an amyloid fibril is presented.  

The assembly of a nearfield infrared microscope for imaging protein aggregates is described.

This paper aims to instruct the reader in the assembly and operation of an infrared near-field microscope for imaging beyond the diffraction limit.  The ...

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 ...

C. elegans Chemotaxis Assay

Many organisms use chemotaxis to seek out food sources, avoid noxious substances, and find mates. Caenorhabditis elegans has impressive chemotaxis behavior.
 The premise behind testing the response of the worms to an odorant is to place them in an area and observe the movement evoked in response to an odorant. Even with the many available assays, optimizing worm starting location relative to both the control and test areas, while minimizing the interaction of worms with each other, while maintaining a significant sample size remains a work in progress 1-10. The method described here aims to address these issues by modifying the assay developed by Bargmann et al.1. A Petri dish is divided into four quadrants, two opposite quadrants marked "Test" and two are designated "Control". Anesthetic is placed in all test and control sites. The worms are placed in the center of the plate with a circle marked around the origin to ensure that non-motile worms will be ignored. Utilizing a four-quadrant system rather than one 2 or two 1 eliminates bias in the movement of the worms, as they are equidistant from test and control samples, regardless of which side of the origin they began. This circumvents the problem of worms being forced to travel through a cluster of other worms to respond to an odorant, which can delay worms or force them to take a more circuitous route, yielding an incorrect interpretation of their intended path. This method also shows practical advantages by having a larger sample size and allowing the researcher to run the assay unattended and score the worms once the allotted time has expired.

C. elegans Chemotaxis Assay

A method of quantitatively evaluating the chemotactic response of Caenorhabditis elegans is described. A chemotactic index (CI) was employed as a way to precisely evaluate the response of worms to certain targets, and serve as a platform of comparison between strains and compounds of interest.

Many organisms use chemotaxis to seek out food sources, avoid noxious substances, and find mates.  Caenorhabditis elegans has impressive chemotaxis ...

Antibody Staining in C. Elegans Using &quot;Freeze-Cracking&quot;

To stain C. elegans with antibodies, the relatively impermeable cuticle must be bypassed by chemical or mechanical methods. "Freeze-cracking" is one method used to physically pull the cuticle from nematodes by compressing nematodes between two adherent slides, freezing them, and pulling the slides apart. Freeze-cracking provides a simple and rapid way to gain access to the tissues without chemical treatment and can be used with a variety of fixatives. However, it leads to the loss of many of the specimens and the required compression mechanically distorts the sample. Practice is required to maximize recovery of samples with good morphology. Freeze-cracking can be optimized for specific fixation conditions, recovery of samples, or low non-specific staining, but not for all parameters at once. For antibodies that require very hard fixation conditions and tolerate the chemical treatments needed to chemically permeabilize the cuticle, treatment of intact nematodes in solution may be preferred. If the antibody requires a lighter fix or if the optimum fixation conditions are unknown, freeze-cracking provides a very useful way to rapidly assay the antibody and can yield specific subcellular and cellular localization information for the antigen of interest.

Antibody Staining in C. Elegans Using &quot;Freeze-Cracking&quot;

"Freeze-cracking," a method for exposing the inner tissues of the nematode C. elegans to antibodies for protein localization, is demonstrated.

To stain C.  elegans with antibodies, the relatively impermeable cuticle must be  bypassed by chemical or mechanical methods.  "Freeze-cracking" is one ...

Measuring the Effects of Bacteria on C. Elegans Behavior Using an Egg Retention Assay

C. elegans egg-laying behavior is affected by environmental cues such as osmolarity1 and vibration2. In the total absence of food C. elegans also cease egg-laying and retain fertilized eggs in their uterus3. However, the effect of different sources of food, especially pathogenic bacteria and particularly Enterococcus faecalis, on egg-laying
	behavior is not well characterized. The egg-in-worm (EIW) assay is a useful tool to quantify the effects of different types of bacteria, in this case E. faecalis, on egg- laying behavior.
	
	EIW assays involve counting the number of eggs retained in the uterus of C. elegans4. The EIW assay involves bleaching staged, gravid adult C. elegans to remove the cuticle and separate the retained eggs from the animal. Prior to bleaching, worms are exposed to bacteria (or any type of environmental cue) for a fixed period of time. After bleaching, one is very easily able to count the number of eggs retained inside the uterus of the worms. In this assay, a quantifiable increase in egg retention after E. faecalis exposure can be easily measured. The EIW assay is a behavioral assay that may be used to screen for potentially pathogenic bacteria or the presence of environmental toxins. In addition, the EIW assay may be a tool to screen for drugs that affect neurotransmitter signaling since egg-laying behavior is modulated by neurotransmitters such as serotonin and acetylcholine5-9.

Measuring the Effects of Bacteria on C. Elegans Behavior Using an Egg Retention Assay

An egg-in-worm (EIW) assay is a useful method to quantify egg-laying behavior. Alterations in egg laying can be a behavioral response of the model organism Caenorhabditis elegans to potentially harmful environmental substances such as those produced by pathogenic bacteria.

C. elegans egg-laying behavior is  affected by environmental cues such as osmolarity1 and  vibration2. In the total absence of food C. elegans also cease ...

An Anoxia-starvation Model for Ischemia/Reperfusion in C. elegans

Protocols for anoxia/starvation in the genetic model organism C. elegans simulate ischemia/reperfusion. Worms are separated from bacterial food and placed under anoxia for 20 hr&#160;(simulated ischemia), and subsequently moved to a normal atmosphere with food (simulated reperfusion). This experimental paradigm results in increased death and neuronal damage, and techniques are presented to assess organism viability, alterations to the morphology of touch neuron processes, as well as touch sensitivity, which represents the behavioral output of neuronal function. Finally, a method for constructing hypoxic incubators using common kitchen storage containers is described. The addition of a mass flow control unit allows for alterations to be made to the gas mixture in the custom incubators, and a circulating water bath allows for both temperature control and makes it easy to identify leaks. This method provides a low cost alternative to commercially available units.

An Anoxia-starvation Model for Ischemia/Reperfusion in C. elegans

A protocol is described that uses anoxia/starvation in C. elegans to model ischemia/reperfusion. Functional outcomes include increased mortality, visible abnormalities in GFP-labeled neuronal processes, and impaired behavioral responses that require neuronal function.

Protocols for anoxia/starvation in the genetic model organism C. elegans simulate ischemia/reperfusion. Worms are separated from bacterial food and placed ...

Setting Up a Simple Light Sheet Microscope for In Toto Imaging of C. elegans Development

Fast and low phototoxic imaging techniques are pre-requisite to study the development of organisms in toto. Light sheet based microscopy reduces photo-bleaching and phototoxic effects compared to confocal microscopy, while providing 3D images with subcellular resolution. Here we present the setup of a light sheet based microscope, which is composed of an upright microscope and a small set of opto-mechanical elements for the generation of the light sheet. The protocol describes how to build, align the microscope and characterize the light sheet. In addition, it details how to implement the method for in toto imaging of C. elegans embryos using a simple observation chamber. The method allows the capture of 3D two-colors time-lapse movies over few hours of development. This should ease the tracking of cell shape, cell divisions and tagged proteins over long periods of time.

Setting Up a Simple Light Sheet Microscope for In Toto Imaging of C. elegans Development

This protocol describes the setup of a light sheet microscope and its implementation for in vivo imaging of C. elegans embryos.

Fast and low phototoxic imaging techniques are pre-requisite to study the development of organisms in toto. Light sheet based microscopy reduces ...

Workflow for High-content, Individual Cell Quantification of Fluorescent Markers from Universal Microscope Data, Supported by Open Source Software

Advances in understanding the control mechanisms governing the behavior of cells in adherent mammalian tissue culture models are becoming increasingly dependent on modes of single-cell analysis. Methods which deliver composite data reflecting the mean values of biomarkers from cell populations risk losing subpopulation dynamics that reflect the heterogeneity of the studied biological system. In keeping with this, traditional approaches are being replaced by, or supported with, more sophisticated forms of cellular assay developed to allow assessment by high-content microscopy. These assays potentially generate large numbers of images of fluorescent biomarkers, which enabled by accompanying proprietary software packages, allows for multi-parametric measurements per cell. However, the relatively high capital costs and overspecialization of many of these devices have prevented their accessibility to many investigators.
Described here is a universally applicable workflow for the quantification of multiple fluorescent marker intensities from specific subcellular regions of individual cells suitable for use with images from most fluorescent microscopes. Key to this workflow is the implementation of the freely available Cell Profiler software1&#160;to distinguish individual cells in these images, segment them into defined subcellular regions and deliver fluorescence marker intensity values specific to these regions. The extraction of individual cell intensity values from image data is the central purpose of this workflow and will be illustrated with the analysis of control data from a siRNA screen for G1 checkpoint regulators in adherent human cells. However, the workflow presented here can be applied to analysis of data from other means of cell perturbation (e.g., compound screens) and other forms of fluorescence based cellular markers and thus should be useful for a wide range of laboratories.

Presented is a flexible informatics workflow enabling multiplexed image-based analysis of fluorescently labeled cells. The workflow quantifies nuclear and cytoplasmic markers and computes marker translocation between these compartments. Procedures are provided for perturbation of cells using siRNA and reliable methodology for marker detection by indirect immunofluorescence in 96-well formats.

Advances in understanding the control mechanisms governing the behavior of cells in adherent mammalian tissue culture models are becoming increasingly ...