We wish to acknowledge the cooperative spirit of all Caldwell Lab members. Movement disorders research in the lab has been supported by the Bachmann-Strauss Dystonia &amp; Parkinson Foundation, United Parkinson Foundation, American Parkinson Disease Association, Parkinson's Disease Association of Alabama, the Michael J. Fox Foundation for Parkinson's Research, and an Undergraduate Research.

Expression Plasmid Construction
Two plasmids are required: one for tissue-specific expression of the gene of interest and a second as selectable transformation marker.
Experimental Plasmid
<ol>
<li>Select a promoter that is expressed in the tissue/cell type of interest, for example, a nerve or muscle-specific promoter. If one is to express multiple genes within the same tissue, a promoter cassette plasmid in the Gateway system (Invitrogen) can be used. This approach allows the insertion of any gene of interest downstream of the promoter by recombinational cloning 4. Generally, 2-5 kb of upstream sequences is sufficient to have correct and accurate expression. </li>
<li>The cDNA of the gene of interest can be cloned in two ways: <ol>
<li>Conventional cloning using restriction enzymes.</li>
<li>For Gateway cloning, it is PCR amplified using primers that contain the Gateway att B recombinational sequence. The amplified cDNA is first recombined into the donor vector pDONR201 to create the entry vector and then transformed into E. coli strain DH5a. Following selection of successful recombinants and mini-prep isolation of DNA, the entry vector is recombined into the promoter-containing destination vector to create the expression plasmid. Details on recombinational cloning are available from either the Invitrogen Gateway manual or in Caldwell et al. 5.</li>
</ol></li>
</ol>
Selectable Marker Plasmid
Examples of transgenic marker plasmids include rol-6 or use of the body wall muscle promoter unc-54 to drive fluorescent protein expression. These marker plasmids are usually available from within the research community upon request.
Microinjection of C. elegans
Preparation
<ol>
<li>Perform DNA isolation of the two plasmids that are to be injected. Quantitate them and mix together in a 1:1 ratio of 50 ng/&micro;l each. </li>
<li>Prepare agar pads: <ol>
<li>Make a 2% agarose solution in water; heat or microwave until dissolved.</li>
<li>Set out several 22 x 50 mm cover glasses on a benchtop.</li>
<li>Using a Pasteur pipette, place a drop of the melted agarose on one cover glass and immediately place a second cover glass over the drop at a 90 angle to the first. Repeat for several other cover glasses. The diameter of the flattened disc of agarose should be between 15-20 mm.</li>
<li>After the agarose has solidified (this only takes moments), remove the cover glass and allow the pad to air dry completely (several hours to overnight).</li>
<li>Store the pads in a cover glass box at room temperature. </li>
</ol></li>
<li>Make needle loaders for DNA solution: Needle loaders are created from 100 &micro;l capillary tubes that are heated in the middle over an open flame and rapidly pulled to approximately twice their length. The two halves are snapped apart once the capillary tube cools. Stockpile 10-20 needle loaders for injection. Store upright in a microcentrifuge tube rack. Use caution as to not impale oneself on the points. </li>
<li>Make microinjection needles: The needle is made from a special capillary tube that contains an internal glass filament; this filament serves as a wick for the DNA solution. These are pulled on a needle-puller machine. We use a Narishige PP-830 puller with a heat setting of 24.8. Several needles are pulled at a time and are stored in a small plastic box. Multiple needles can be readily "mounted" on a strip of modeling clay for long-term storage. </li>
<li>Needle tip "breakers": The tip of the microinjection needle is pulled so fine that it is actually closed at it end and should be broken open using small shards of cover glass. These are prepared by wrapping an 18 x 18 mm cover glass in a paper towel and gently applying pressure until it breaks into several pieces. Shards of a useful size are approximately 3 x 4 mm. </li>
<li>Grow wild type (N2) worms to the early adult stage for injection using standard procedures 6. Prepare about 100 properly staged worms/construct injected.</li>
</ol>
Procedure
<ol>
<li>Open the main valve on a helium tank that is connected to a microinjection needle arm and holder. Close the regulator valve to allow the gas to enter the line, bringing the pressure to 35 psi. Notice that the gas can be released using a foot pedal. </li>
<li>Insert a needle loader into a standard mouth pipetter tubing (found in containers of glass capillary tubes) and draw up a small amount of the plasmid mixture (~1 &micro;l). </li>
<li>The needle loader is carefully placed into the back end of an injection needle and gently inserted all the way through the length of the needle until resistance is felt. Expel the DNA solution by gently blowing, then remove the loader. </li>
<li>Insert the needle into the microinjection arm, being careful to place it within the small internal rubber gasket. </li>
<li>Place a needle-breaking shard of cover glass on one edge of an agar pad. Cover the shard, as well as the whole surface of the agar pad, with halocarbon oil. Place the agar pad on the stage of an inverted microscope. </li>
<li>Position the needle in the center of the viewing field using the 4X objective and bright field illumination, but do not yet lower it into the oil. Also, position the inner edge of the shard of cover glass in the center, then focus on it (still using 4X objective). Lower the needle into the oil, bringing it into the same focal plane as the shard of cover glass. Raise the magnification by switching to the 40X objective. Refocus on the edge of the shard and reposition the tip of the needle, if necessary. </li>
<li>To open up the tip of the injection needle, GENTLY nudge the needle against the edge of the cover glass shard, while at the same time applying a short pulse of gas via the foot pedal. The needle will be sufficiently broken open when small droplets of liquid escape the end of the needle. Excess liquid flow due to too large an opening is not desirable, as it will kill the animals. </li>
<li>Remove the agar pad from the stage by raising the needle up, but do not change the X- or Y-axis position. Slide the stage out from underneath the needle, remove the agar pad and place it onto a dissecting microscope. Transfer a worm onto the pad, below the surface of the oil. If the worm wriggles, GENTLY stroke it until it contacts the agar pad. The moist worm should adhere to the dry agarose. </li>
<li>Injection <ol>
<li>Quickly place the agar pad back onto the stage, centering the worm in the field of view.</li>
<li>Lower the needle into the same plane of focus as the worm.</li>
<li>Switch the filters to Hoffman Illumination (a type of contrast filter). This allows morphological detail of the worm to become visible. If Nomarski/DIC optics are available on the inverted scope that will work as well. Using the 40X objective, focus on the "grainy" syncytial center of the worm gonad,&nbsp; below the "honeycomb" pattern of the germ nuclei (choose whichever gonad is positioned on the side of the worm facing the needle).</li>
<li>Fine tune the position of the needle until the tip is also in focus (the same plane as the gonad "graininess").</li>
<li>Gently insert the needle into the center of the gonad and apply a brief pulse of gas pressure to expel a droplet or two of DNA into the gonad arm. You should observe a wave of liquid spread across the distal gonad. Do not assume more liquid is better, since too much liquid can flow into the proximal bend of the gonad arm, which can shut down oocyte production. If possible, depending on the position of the worm, inject the other gonad arm in the same manner.</li>
<li>It is important to work quickly while injecting a worm, as it can easily desiccate. The decision to inject the second gonad arm is dependent upon how quickly you can re-position the needle and worm. The entire process should ideally take less than 1-2 minutes. </li>
</ol></li>
<li>Remove the cover glass from the stage and place it on a dissecting scope. Apply 1&micro;l of M9 buffer (22mM KH2PO4, 42mM Na2HPO4, 85mM NaCl, 1mM MgSO4) directly onto the injected worm (this might entail submerging the pipette tip below the surface of the halocarbon oil). The buffer should rehydrate the worm; it will then float off the agarose pad surface. Transfer the worm to a regular plate using a worm pick. The agarose pad can be reused several more times until it has become too saturated with buffer and the worms no longer adhere.</li>
</ol>
Transgenic selection
<ol>
<li>Injected worms, the P0 generation, are transferred to individual fresh, bacterially seeded plates (60 mm) and allowed to reproduce. Typically, injected animals are incubated at 20&deg;C for 2-3 days until the offspring appear. </li>
<li>The F1 offspring are observed for evidence of the transgenic marker (such as fluorescence) using a fluorescent stereomicroscope. Each fluorescent, carrier F1, animal is separately cloned onto a new plate (since offspring from each F1 are considered a distinct line). The F1 hermaphrodites are allowed to reproduce. Again, this is typically performed at at 20&deg;C for 2-3 days until the offspring appear. </li>
<li>The F2 generation is observed for transgenic animals as well. F2 animals that have inherited and express the transgenic array are considered a "stable" line. NOTE: At the F1 stage there may be many transgenic animals, but they are not stable. The majority of F1 transgenic animals will not be propagated into F2 stable lines. NOTE: To control for the variability in the gene copy number, generate a minimum of three independent stable lines per construct injected. </li>
<li>Each independent stable line should be maintained as a separate line of worms. Each line can be propagated by transferring several transgenic animals to a new plate at each generation; this must be performed using a fluorescent stereomicroscope if the selectable marker is fluorescent.</li>
</ol>

<ol>
	<li>Stinchcomb, D. T., Shaw, J. E., Carr, S. H., Hirsh, D. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Extrachromosomal+DNA+transformation+of+Caenorhabditis+elegans.">Extrachromosomal DNA transformation of Caenorhabditis elegans.</a> Mol. Cell. Biol. 5, 3484-3496 (1985).</li><li>Mello, C. C., Kramer, J. M., Stinchcomb, D., Ambros, V. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Efficient+gene+transfer+in+C.elegans:+extrachromosomal+maintenance+and+integration+of+transforming+sequences.">Efficient gene transfer in C.elegans: extrachromosomal maintenance and integration of transforming sequences.</a> EMBO J. 10, 3959-3970 (1991).</li><li>Kelly, W. G., Xu, S., Montgomery, M. K., Fire, A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Distinct+requirements+for+somatic+and+germline+expression+of+a+generally+expressed+Caenorhabditis+elegans+gene.">Distinct requirements for somatic and germline expression of a generally expressed Caenorhabditis elegans gene.</a> Genetics. 146, 227-238 (1997).</li><li>Cao, S., Gelwix, C. C., Caldwell, K. A., Caldwell, G. A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Torsin-mediated+neuroprotection+from+cellular+stresses+to+dopaminergic+neurons+of+C.+elegans.">Torsin-mediated neuroprotection from cellular stresses to dopaminergic neurons of C. elegans.</a> J Neurosci. 25, 3801-3812 (2005).</li><li>Caldwell, G. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=.">.</a> Integrated Genomics: A Discovery-Based Laboratory Course. , (2006).</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+Caenorhabditis+elegans.">The genetics of Caenorhabditis elegans.</a> Genetics. 77, 71-94 (1974).</li></ol>

When doing this procedure, it is important to remember to: - inject directly into the center of the gonad - not inject too much liquid - work quickly to prevent dessication.
If you run into problems with the needle, such as it is broken too large, it is best to remake a fresh needle, rather than trying to inject with a less than ideal needle.
The generation of transgenic worms has many applications such as: - identification of mutant genes, in ...

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		<div>
		<table border="1">
		<thead>
		<tr>
		<th>Material Name</th>
		<th>Type</th>
		<th>Company</th>
		<th>Catalogue Number</th>
		<th>Comment</th>
		</tr>
		</thead>
		<tbody>
		
<tr>
<td>Agarose</td>
<td>Ultrapure</td>
<td>Invitrogen</td>
<td>15510-027</td>
<td>&nbsp;</td>
<tr>
<td>Halocarbon Oil, Voltalef</td>
<td>Hulle 10S</td>
<td>elfatochem, France</td>
<td>&nbsp;</td>
<td>&nbsp;</td>
<tr>
<td>Coverglass 18x18 mm</td>
<td>&nbsp;</td>
<td>Fisher</td>
<td>12-548-A</td>
<td>&nbsp;</td>
<tr>
<td>Coverglass 20x30 mm</td>
<td>&nbsp;</td>
<td>Fisher</td>
<td>12-548-5A</td>
<td>&nbsp;</td>
<tr>
<td>Coverglass 22x50 mm</td>
<td>&nbsp;</td>
<td>Fisher</td>
<td>12-545-E</td>
<td>&nbsp;</td>
<tr>
<td>100 ul Capillary</td>
<td>&nbsp;</td>
<td>VWR</td>
<td>53432-921</td>
<td>&nbsp;</td>
<tr>
<td>Glass Capillary for Needles</td>
<td>"Kwik-Fil"</td>
<td>World Precision Instruments</td>
<td>1B100F-4</td>
<td>&nbsp;</td>
<tr>
<td>Needle Puller</td>
<td>Tool</td>
<td>Narishige</td>
<td>Model PP-830</td>
<td>&nbsp;</td>
<tr>
<td>microINJECTOR System</td>
<td>&nbsp;</td>
<td>Tritech Research</td>
<td>MINJ-1000</td>
<td>Scope, Stage, Manipulator</td>
</tr>
<tr>
<td>Dissecting, with Fluorescence</td>
<td>Microscope</td>
<td>Nikon</td>
<td>SMZ800</td>
<td>&nbsp;</td>
<tr>
<td>Dissecting</td>
<td>Microscope</td>
<td>Nikon</td>
<td>SMZ645</td>
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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.

Watch this Scientific Journal Video about Generation of Stable Transgenic C. elegans Using Microinjection at JoVE.com

Generation of Stable Transgenic C. elegans Using Microinjection

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 ...

generation-of-stable-transgenic-c-elegans-using-microinjection

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52.7K Views.  University of Alabama. This video demonstrates the technique of microinjection into the gonad of C. elegans to create transgenic animals.

Video: Generation of Stable Transgenic C. elegans Using Microinjection

Microinjection of A. aegypti Embryos to Obtain Transgenic Mosquitoes

In this video, Nijole Jasinskiene demonstrates the methodology employed to generate transgenic Aedes aegypti mosquitoes, which are vectors for dengue fever.  The techniques for correctly preparing microinjection needles, dessicating embryos, and performing microinjection are demonstrated.

In this video, Nijole Jasinskiene demonstrates the methodology employed to generate transgenic Aedes aegypti mosquitoes, which are vectors for dengue fever. The techniques for correctly preparing microinjection needles, dessicating embryos, and performing microinjection are demonstrated.

In this video, Nijole Jasinskiene demonstrates the methodology employed to generate transgenic Aedes aegypti mosquitoes, which are vectors for dengue ...

Generation of Transgenic C. elegans by Biolistic Transformation

The number of laboratories using the free living nematode C. elegans is rapidly growing. The popularity of this biological model is attributed to a rapid generation time and short life span, easy and inexpensive maintenance, fully sequenced genome, and array of RNAi resources and mutant animals. Additionally, analysis of the C. elegans genome revealed a great similarity between worms and higher vertebrates, which suggests that research in worms could be an important adjunct to studies performed in whole mice or cultured cells. A powerful and important part of worm research is the ability to use transgenic animals to study gene localization and function. Transgenic animals can be created either via microinjection of the worm germline or through the use of biolistic bombardment. Bombardment is a newer technique and is less familiar to a number of labs. Here we describe a simple protocol to generate transgenic worms by biolistic bombardment with gold particles using the Bio-Rad PDS-1000 system. Compared with DNA microinjection into hermaphrodite germline, this protocol has the advantage of not requiring special skills from the operator with regards to identifying worm anatomy or performing microinjection. Further multiple transgenic lines are usually obtained from a single bombardment. Also in contrast to microinjection, biolistic bombardment produces transgenic animals with both extrachromosomal arrays and integrated transgenes. The ability to obtain integrated transgenic lines can avoid the use of mutagenic protocols to integrate foreign DNA. In conclusion, biolistic bombardment can be an attractive method for the generation of transgenic animals, especially for investigators not interested in investing the time and effort needed to become skilled at microinjection.

Generation of Transgenic C. elegans by Biolistic Transformation

Transgenic worms are commonly used in C. elegans research. Described is a simple, yet effective, protocol to introduce transgenes into worms using biolistic bombardment with DNA-coated gold particles. The effort involved and results of bombardment compare favorably with microinjection for the generation of transgenic animals.

The number of laboratories using the free living nematode C. elegans is rapidly growing. The popularity of this biological model is attributed to a rapid ...

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.

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.

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

Activation of Apoptosis by Cytoplasmic Microinjection of Cytochrome c

Apoptosis, or programmed cell death, is a conserved and highly regulated pathway by which cells die1. Apoptosis can be triggered when cells encounter a wide range of cytotoxic stresses. These insults initiate signaling cascades that ultimately cause the release of cytochrome c from the mitochondrial intermembrane space to the cytoplasm2. The release of cytochrome c from mitochondria is a key event that triggers the rapid activation of caspases, the key cellular proteases which ultimately execute cell death3-4.
The pathway of apoptosis is regulated at points upstream and downstream of cytochrome c release from mitochondria5. In order to study the post-mitochondrial regulation of caspase activation, many investigators have turned to direct cytoplasmic microinjection of holocytochrome c (heme-attached) protein into cells6-9. Cytochrome c is normally localized to the mitochondria where attachment of a heme group is necessary to enable it to activate apoptosis10-11. Therefore, to directly activate caspases, it is necessary to inject the holocytochrome c protein instead of its cDNA, because while the expression of cytochrome c from cDNA constructs will result in mitochondrial targeting and heme attachment, it will be sequestered from cytosolic caspases. Thus, the direct cytosolic microinjection of purified heme-attached cytochrome c protein is a useful tool to mimic mitochondrial cytochrome c release and apoptosis without the use of toxic insults which cause cellular and mitochondrial damage.
In this article, we describe a method for the microinjection of cytochrome c protein into cells, using mouse embryonic fibroblasts (MEFs) and primary sympathetic neurons as examples. While this protocol focuses on the injection of cytochrome c for investigations of apoptosis, the techniques shown here can also be easily adapted for microinjection of other proteins of interest.

Activation of Apoptosis by Cytoplasmic Microinjection of Cytochrome c

In this protocol, we describe the direct cytoplasmic microinjection of cytochrome c protein into fibroblasts and primary sympathetic neurons. This technique allows for the introduction of cytochrome c protein into the cytoplasm of cells and mimics the release of cytochrome c from mitochondria, which occurs during apoptosis.

Apoptosis, or programmed cell death, is a conserved and highly regulated pathway by which cells die1.  Apoptosis can be triggered when cells encounter a ...

Determining Genetic Expression Profiles in C. elegans Using Microarray and Real-time PCR

Synapses are composed of a presynaptic active zone in the signaling cell and a postsynaptic terminal in the target cell. In the case of chemical synapses, messages are carried by neurotransmitters released from presynaptic terminals and received by receptors on postsynaptic cells. Our previous research in Caenorhabditis elegans has shown that VSM-1 negatively regulates exocytosis. Additionally, analysis of synapses in vsm-1 mutants showed that animals lacking a fully functional VSM-1 have increased synaptic connectivity. Based on these preliminary findings, we hypothesized that C. elegans VSM-1 may play a crucial role in synaptogenesis. To test this hypothesis, double-labeled microarray analysis was performed, and gene expression profiles were determined. First, total RNA was isolated, reversely transcribed to cDNA, and hybridized to the DNA microarrays. Then, in-silico analysis of fluorescent probe hybridization revealed significant induction of many genes coding for members of the major sperm protein family (MSP) in mutants with enhanced synaptogenesis. MSPs are the major component of sperm in C. elegans and appear to signal nematode oocyte maturation and ovulation . In fruit flies, Chai and colleagues 1 demonstrated that MSP-like molecules regulate presynaptic bouton number and size at the neuromuscular junction. Moreover, analysis performed by Tsuda and coworkers 2 suggested that MSPs may act as ligands for Eph receptors and trigger receptor tyrosine kinase signaling cascades. Lastly, real time PCR analysis corroborated that the gene coding for MSP-32 is induced in vsm-1(ok1468) mutants. Taken together, research performed by our laboratory has shown that vsm-1 mutants have a significant increase in synaptic density, which could be mediated by MSP-32 signaling.

Determining Genetic Expression Profiles in C. elegans Using Microarray and Real-time PCR

Microarray analysis was conducted to determine genetic expression profiles in C. elegans, and real-time PCR was used to validate and quantify microarray data.

Synapses are composed of a presynaptic active zone in the signaling cell and a postsynaptic terminal in the target cell. In the case of chemical synapses, ...

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 ...

Generation of Transgenic Hydra by Embryo Microinjection

As a member of the phylum Cnidaria, the sister group to all bilaterians, Hydra can shed light on fundamental biological processes shared among multicellular animals. Hydra is used as a model for the study of regeneration, pattern formation, and stem cells. However, research efforts have been hampered by lack of a reliable method for gene perturbations to study molecular function. The development of transgenic methods has revitalized the study of Hydra biology1. Transgenic Hydra allow for the tracking of live cells, sorting to yield pure cell populations for biochemical analysis, manipulation of gene function by knockdown and over-expression, and analysis of promoter function. Plasmid DNA injected into early stage embryos randomly integrates into the genome early in development. This results in hatchlings that express transgenes in patches of tissue in one or more of the three lineages (ectodermal epithelial, endodermal epithelial, or interstitial). The success rate of obtaining a hatchling with transgenic tissue is between 10% and 20%. Asexual propagation of the transgenic hatchling is used to establish a uniformly transgenic line in a particular lineage. Generating transgenic Hydra is surprisingly simple and robust, and here we describe a protocol that can be easily implemented at low cost.

Generation of Transgenic Hydra by Embryo Microinjection

Stably transgenic Hydra are made by microinjection of plasmid DNA into embryos followed by random genomic integration and asexual propagation to establish a uniform line. Transgenic Hydra are used to track cell movements, overexpress genes, study promoter function, or knock down gene expression using RNAi.

As a member of the phylum Cnidaria, the sister group to all bilaterians, Hydra can shed light on fundamental biological processes shared among ...

Investigating the Spreading and Toxicity of Prion-like Proteins Using the Metazoan Model Organism C. elegans

Prions are unconventional self-propagating proteinaceous particles, devoid of any coding nucleic acid. These proteinaceous seeds serve as templates for the conversion and replication of their benign cellular isoform. Accumulating evidence suggests that many protein aggregates can act as self-propagating templates and corrupt the folding of cognate proteins. Although aggregates can be functional under certain circumstances, this process often leads to the disruption of the cellular protein homeostasis (proteostasis), eventually leading to devastating diseases such as Alzheimer&#8217;s disease (AD), Parkinson&#8217;s disease (PD), Amyotrophic lateral sclerosis (ALS), or transmissible spongiform encephalopathies (TSEs). The exact mechanisms of prion propagation and cell-to-cell spreading of protein aggregates are still subjects of intense investigation. To further this knowledge, recently a new metazoan model in Caenorhabditis elegans, for expression of the prion domain of the cytosolic yeast prion protein Sup35 has been established. This prion model offers several advantages, as it allows direct monitoring of the fluorescently tagged prion domain in living animals and ease of genetic approaches. Described here are methods to study prion-like behavior of protein aggregates and to identify modifiers of prion-induced toxicity using C. elegans.

Investigating the Spreading and Toxicity of Prion-like Proteins Using the Metazoan Model Organism C. elegans

Prion-like propagation of protein aggregates has recently emerged as being implicated in many neurodegenerative diseases. The goal of this protocol is to describe, how to use the nematode C. elegans as a model system to monitor protein spreading and to investigate prion-like phenomena.

Prions are unconventional self-propagating proteinaceous particles, devoid of any coding nucleic acid. These proteinaceous seeds serve as templates for ...