This article was funded by NIH grants to JER. I would like to thank Dr. Alex Gottschalk for providing the transgenic worms expressing channel rhodopsin in cholinergic motor neurons used in one of the recording demonstrations.

A. Preparation of tools used for the dissection.
<ol>
	<li>The dissection/recording chamber: We typically construct our recording chamber out of a 1/16th inch magnetic sheet, with a circular hole drilled into the center that is large enough to accommodate a 22 mm diameter circular cover glass. The outer dimensions of the chamber will depend on the microscope stage used to do the recordings. On the reverse side of the magnetic sheet , we attach a 48 x 60 mm cover slip, which we adhere to the chamber by placing a pellet of low melt Paraplax tissue embedding wax in each corner of the cover slip sandwiched between the glass and the chamber. The wax is then melted on a hot plate, glass plate facing up, until it forms a continuous layer gluing the cover slip to the chamber. Do not leave this step unattended as the wax melts quickly. The chamber should be removed and cooled as soon as the wax has melted. Excess wax that has seeped along the inner edge of the 22mm circular cavity can be removed using a razor blade edge.</li>
	<li>Sylgard-coated cover slips: The dissection is performed on a 22 mm, Sylgard-coated cover slip that is placed within the circular cavity of the recording chamber. The Sylgard coat helps the cyanoacrylic glue adhere to the cover slip when gluing the worms down and also acts as a cushion on which to break back the glue pipette tip when it becomes plugged. Sylgard is made up fresh according to the manufactures instructions (10 parts silicone base to 1 part curing agent by weight). A small drop of Sylgard is then placed on each cover slip and smeared across using a razor blade edge to make an even surface. The coated cover slips are placed in large flat covered containers and left in a 65&#176; degree oven overnight to cure.</li>
	<li>Pipettes: Typically, we use the same 1 mm outer diameter filament borosilicate glass to generate the pipettes for gluing worms down, making cuticle incisions and recording from muscles, using an electrode puller. It is advisable to pull a jar of pipettes before attempting a dissection as this stage can require several rapid exchanges of pipettes.</li>
	<li>Glue applicator: Glue is loaded and applied from the tip of a glue pipette, by mouth-pressure. The gluing pipette is inserted into one end of a 2 foot piece of polyethylene tubing with an inner diameter that matches the outer diameter of the pipette glass (usually 1 mm). The other end of the tubing holds an eppendorf pipette tip that acts as a mouth piece.</li>
	<li>Extraction pipette: The worm viscera must be removed from the worm cavity once the cuticle incision has been made. This is achieved with a piece of tubing attached to an eppendorf pipette tip similar to the glue applicator but in this case the glass extraction pipette tip is broken back sufficiently to allow eggs and viscera to be easily vacuumed out of the worm cavity.</li>
</ol>
B. The dissection
<ol>
	<li>In the recording chamber a circular wax pen line is drawn close to the perimeter of the chamber to which a Sylgard-coated cover slip is pressed firmly in place (Sylgard-coat facing upwards). This wax line immobilizes the cover slip and prevents the recording solution from seeping under and dislodging the cover slip during dissection and recording. The chamber is then filled with extracellular recording solution and several worms are placed in the center using a worm pick. Worms will swim in solution, however with practice, these worms can be glued down without any further procedures such as cooling to prevent movement. During the training stage, it can help to practice gluing mutant worms in which swimming is impaired (such as unc-31 mutants).</li>
	<li>A glue container fashioned from a PCR cap held in wax or modeling clay, is used to hold a working stock of the cyanoacrylic glue. Glue pipettes with similar tip dimensions to the recording pipettes (normally ~4 megohms resistance) are used to apply glue. Using the glue applicator, a small amount of glue is sucked up into the glue pipette, using the dissection scope to visualize the pipette tip as the glue is sucked up.</li>
	<li>The cyanoacrylic glue (HistoacrylBlue, Aesulap) polymerizes on contact with extracellular recording solution, therefore it is important to maintain positive pressure on the glue pipette as it enters the chamber solution to prevent the glue from hardening and plugging the pipette. As soon as the glue pipette is in the chamber solution, a small but constant stream of glue needs to be applied onto the Sylgard surface under mouth-pressure control to prevent plugging. If the pipette becomes plugged it can sometimes be unblocked by gently tapping the tip on the sylgard-coated glass. It is recommended that you practice gluing before attempting to glue worms. If you can routinely write your name in glue on the Sylgard, with letters made of lines thinner than the worms width you are ready to proceed to gluing the worm cuticle.</li>
	<li>To glue a worm, use positive-pressure to attach a small amount of glue to either the head or tail of the worm and rapidly draw the worm down onto the Sylgard surface. Try to attach the worm near the center of the chamber as worms glued too near to the edge will be difficult to reach with recording pipettes. Then apply a constant stream of glue along the dorsal edge of the worm&#8217;s cuticle from the starting attachment, forcing the worm into a C-position with the vulva facing inward (for ventral muscle recordings). Any gaps in this glue line should be filled in so that the worm is strongly attached, to prevent the worm from detaching during the cuticle incision step.</li>
	<li>To make the cuticle incision, switch to a hand-held dissection pipette. This pipette needs to be sharper than the glue/recording pipettes and have a short, sturdy shaft. Using the highest-magnification on the dissecting scope, align the pipette parallel to the longitudinal axis of the worm and insert the tip about midpoint along the worm (close to the vulva) making the incision at the cuticle/glue interface. This incision releases the worms hydrostatic pressure forcing eggs and intestines out through the incision point. Continue cutting the cuticle towards the head of the worm with a slicing motion similar to opening a letter, until you reach the pharynx. Switch over to the extraction pipette to vacuum out the internal organs of the worm. After clearing the incision, the opened cuticle will retain it&#8217;s cylindrical shape. Use a fresh gluing pipette to spot weld the cut edge of the cuticle down onto the Sylgard surface. It is important to use minimal glue spots at this stage to avoid damaging or obscuring the NMJs. 
	The ventral nerve cord and body wall muscles are now exposed.</li>
	<li>To remove the basement membrane that covers the NMJs, suck off the extracellular recording solution from the chamber and apply collagenase (0.4mg/ml in extracellular recording solution) for about 10-20 seconds; remove and wash several times with fresh extracellular recording solution. Proceed to the recording stage.</li>
</ol>
C. Patch-clamp recording
<ol>
	<li>Position the recording chamber on an up-right microscope stage using a 10X objective to center the worm. Switch to a 40X water-emersion objective with DIC and check that the body wall muscles and nerve cord are intact. Position the worm so that the longitudinal axis of the worm is parallel with the front edge of the scope, allowing electrodes to be introduced from both the left and right sides.</li>
	<li>Patch the muscle using standard patch-clamping techniques. Recording electrodes of about 4 megohms resistance work well. Once the electrode touches the muscle membrane, increase negative-pressure until a gigohm seal is obtained. For the whole-cell voltage-clamp recording mode, apply more suction and zap the membrane until the gigohm seal is ruptured. In a good recording, body wall muscles of wild-type adult worms typically have a cell-membrane capacitance of ~70 pF and can be stably recorded for at least 10 minutes at a holding potential of -60 mV with minimal holding currents (~50 pA).</li>
	<li>The extracellular recording solution consists of (in mM): NaCl 150, KCl 5, CaCl2 5, MgCl2 4, glucose 10, sucrose 5, HEPES 15 (pH 7.3, ~ 330mOsm). The patch pipette contains (in mM): KCl 120, KOH 20, MgCl2 4, (N-tris[Hydroxymethyl] mthyl-2-aminoethane-sulfonic acid) TES 5, Dihydrate CaCl2 0.25, Na2ATP 4, sucrose 36, EGTA 5 (pH 7.2, ~312 mOsm). Note: These solutions artificially load muscle cells with chloride, so that the GABA receptor-mediated currents are inward due to chloride efflux at a holding potential of -60 mV. Variations on these solutions have been used (see publications for details).</li>
	<li>Evoked release. Typically we place a second electrode containing extracellular solution on the ventral nerve cord anterior to the patched muscle. This loose patch configuration requires large depolarizing stimuli (on the order of 10s of volts) of short duration (1-2ms) to achieve maximal evoked responses (typically 2.0-2.5 nA) in wild-type worms. The limitations of this approach are: 1) Only a few evoked responses can be delivered before the nerve cord is irreversibly damaged and 2) For unknown reasons, only cholinergic evoked responses can be elicited using this stimulation configuration.</li>
	<li>Light activated channel-rhodopsin stimulation. Recently, the Gottschalk lab has generated transgenic worms expressing algae-derived channel rhodopsins that depolarize the membrane upon light-stimulation in subsets of neurons, depending on the promoter used to drive expression 4. In the case of the C. elegans NMJ, integrated lines expressing channel rhodopsin in either the cholinergic motor neurons or GABAergic motor neurons are now available. Since channel rhodopsin only responds to light when all-trans retinal is associated with the opsin moiety, worms used for this application must be grown on bacteria laced with retinal. A green light pulse can then be used to repeatedly activate the channel rhodopsin causing evoked release of either ACh or GABA, depending on which transgenic worms are used. These transgenic worms can be crossed into mutant strains. The advantage of this technique over the previous electrical stimulation include; 1) Light-activation does not cause neuronal damage, allowing many repeat stimulations to be performed and 2) Evoked release from either GABAergic or cholinergic motor neurons can be independently studied.</li>
	<li>Hyperosmotic-induced release. To measure the size of the readily-releasable vesicle pool (reflecting the summed pool of primed vesicles from all NMJ&#8217;s onto the recorded muscle), a pressure-pipette containing hyperosmotic extracellular solution (adjusted to ~850 mOsm by adding additional sucrose) is used. Pressure-pulse applications of 1-3 seconds produce a barrage of asynchronous post-synaptic events using the standard solutions described above.</li>
</ol>
<img alt="" height="449" src="/files/ftp_upload/old/1165_2009_1_22_3/figure1.png" width="600" />
Figure 1: Schematic overview of the C. elegans dissection and neuromuscular recording configuration
A sylgard coated coverslip made by smearing a drop of Sylgard across the glass and curing overnight is placed in the circular cavity of the prefabricated recording chamber and covered with recording solution.&#160; A worm placed in the center is glued to the Sylgard surface starting from the head or tail, applying glue to the dorsal side of the worm, using a glue pipette,.&#160; Once gluing is complete, an incision adjacent to the vulva, at the interface between the cuticle and glue is made with a glass needle and extended toward the head (see red perforated line).&#160; Following extraction of viscera and eggs from the body cavity,&#160; the cut edge of the incision is spot-welded with glue to the Sylgard exposing the ventral nerve (green) and body wall muscles (red).&#160; The ventral-medial muscle anterior to the vulva is patch-clamped and the stimulating electrode is placed upstream on the nerve cord.
Representative results
<img alt="Represenative Figure" height="404" src="/files/ftp_upload/1165/Reprensentative.png" width="600" />
Whole-cell, voltage-clamped body wall muscle recordings are shown for each of the three stimulation protocols described above (electrically evoked, light-evoked and hyperosmotic responses).&#160; In all cases the ventral-medial body wall muscle is clamped at -60 mV.&#160; A. Wild-type muscle cells typically exhibit high rates of endogenous inward miniature synaptic currents in our standard recording solutions. B. A 2ms depolarization of the anterior ventral nerve cord causes synchronous release from several synapses resulting in a large evoked response of around 2-2.5 nA in wild-type worms.&#160; Representative traces to the right of the wild-type traces in A and B show synaptic release in a severely uncoordinated mutant (unc-18).&#160; Panel C shows a light-evoked response from the neuromuscular junction of a wild-type worm expressing channel rhodopsin in cholinergic neurons.&#160; A 2ms light pulse typically produces a response of 1.5-1.8 nA.&#160; D. Pressure-ejection of 850 mOsm hyperosmotic saline onto the muscle for 1s, produces an asynchronous barrage of miniature synaptic currents representing the readily-releasable pool of vesicles.

<ol>
	<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=.">.</a> Genetics. 77 (1), 71-71 (1974).</li><li>Richmond, J. E., Davis, W. S., Jorgensen, E. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=.">.</a> Nat Neurosci. 2 (11), 959-959 (1999).</li><li>Richmond, J. E. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=.">.</a> WormBook. 1, (2006).</li><li>Liewald, J. F., Brauner, M., Stephens, G. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=.">.</a> Nature methods. 5 (10), 895-895 (2008).</li><li>Lockery, S. R., Goodman, M. B. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=.">.</a> Methods in enzymology. 293, 201-201 (1998).</li><li>Brockie, P. J., Mellem, J. E., Hills, T. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=.">.</a> Neuron. 31 (4), 617-617 (2001).</li><li>Richmond, J. E., Jorgensen, E. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=.">.</a> Nat Neurosci. 2 (9), 791-791 (1999).</li></ol>

The genetic model organism C, elegans is ideally suited for the study of synaptic transmission, through the functional analysis of mutations in genes encoding synaptic proteins. Here we have described a dissection technique that renders the C. elegans NMJs accessible for electrophysiological analysis. The accompanying video depicts the critical steps in the C. elegans dissection and the typical recording configuration used to measure synaptic activity. The three steps that pose the greates...

dissecting and recording from the c. elegans neuromuscular junction

Neurotransmission is the process by which neurons transfer information via chemical signals to their post-synaptic targets, on a rapid time scale. This complex process requires the coordinated activity of many pre- and post-synaptic proteins to ensure appropriate synaptic connectivity, conduction of electrical signals, targeting and priming of secretory vesicles, calcium sensing, vesicle fusion, localization and function of postsynaptic receptors and finally, recycling mechanisms. As neuroscientists it is our goal to elucidate which proteins function in each of these steps and understand their mechanisms of action. Electrophysiological recordings from synapses provide a quantifiable read out of the underlying electrical events that occur during synaptic transmission. By combining this technique with the powerful array of molecular and genetic tools available to manipulate synaptic proteins in C. elegans, we can analyze the resulting functional changes in synaptic transmission. 
The C. elegans NMJs formed between motor neurons and body wall muscles control locomotion, therefore, mutants with uncoordinated locomotory phenotypes (known as unc s) often perturb synaptic transmission at these synapses 1. Since unc mutants are maintained on a rich supply of a bacterial food source, they remain viable as long as they retain some pharyngeal pumping ability to ingest food. This, together with the fact that C. elegans exist as hermaphrodites, allows them to pass on mutant progeny without the need for elaborate mating behaviors. These attributes, coupled with our recent ability to record from the worms NMJs 2,3,7 make this an excellent model organism in which to address precisely how unc mutants impact neurotransmission. 
The dissection method involves immobilizing adult worms using a cyanoacrylic glue in order to make an incision in the worm cuticle exposing the NMJs. Since C. elegans adults are only 1 mm in length the dissection is performed with the use of a dissecting microscope and requires excellent hand-eye coordination. NMJ recordings are made by whole-cell voltage clamping individual body wall muscle cells and neurotransmitter release can be evoked using a variety of stimulation protocols including electrical stimulation, light-activated channel-rhodopsin-mediated depolarization 4 and hyperosmotic saline, all of which will be briefly described.

Watch this Scientific Journal Video about Dissecting and Recording from The C. Elegans Neuromuscular Junction at JoVE.com

Dissecting and Recording from The C. Elegans Neuromuscular Junction

Application of electrophysiology to accessible synapses provides a quantifiable measure of synaptic activity, useful in analyzing synaptic mutants. This article describes a dissection method used to expose the neuromuscular junctions (NMJ) of Caenorhabditis elegans (C. elegans) and briefly discusses some of the uses to which this preparation can be applied.

Neurotransmission is the process by which neurons transfer information via chemical signals to their post-synaptic targets, on a rapid time scale.   This ...

dissecting-and-recording-from-the-c-elegans-neuromuscular-junction

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15.5K ビュー.  University of Illinois, Chicago. アクセス可能なシナプスの電気生理学のアプリケーションでは、シナプス変異体の分析に有用なシナプス活性の定量的な尺度を提供しています。この記事では、神経筋接合部（NMJ）のを公開するために使用される解剖の方法を説明します線虫Caenorhabditis elegans（線虫）と簡潔にこの製剤を適用することができる用途のいくつかについて説明しま?...

ビデオ: Dissecting and Recording from The C. Elegans Neuromuscular Junction

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 ...

Visualizing the Live Drosophila Glial-neuromuscular Junction with Fluorescent Dyes

Our project identified GFP labeled glial structures at the developing larval fly neuromuscular synapse. To look at development of live glial-nerve-muscle synapses, we developed a larval tissue preparation that had features of live intact larvae, but also had good optical properties.  This new preparation also allowed for access of perfusates to the synapse.  We used fly larvae, immersed them in artificial hemolymph, and relaxed their normal rhythmic body contractions by chilling them. Next we dissected off the posterior segments of each animal and with a blunt insect pin pushed the mouth parts backward through the body cavity.  This everted the larval body wall, like turning a sock inside-out.  We completed the dissection with ultra-fine dissection scissors and thus exposed the visceral side of the body wall muscles. The glial structures at the NMJ expressed membrane targeted GFP under the control of glial specific promoters. The post-synaptic membrane, the SSR (Subsynaptic Reticula) in muscle expressed synaptically targeted dsRed. We needed to acutely label the motor neuron terminals, the third part of the synapse. To do this we applied primary antibodies to HRP, conjugated to a far-red emitting flurophore.  To test for dye diffusion properties into the perisynaptic space between the motor neuron terminals and the SSR, we applied a solution of large Dextran molecules conjugated to far-red emitting flurophore and collected images.

Visualizing the Live Drosophila Glial-neuromuscular Junction with Fluorescent Dyes

We described structural features of the Glia-neuromuscular synapses in a novel Inside-out tissue preparation of live fly larvae using fluorescent dyes with confocal microscopy. We labeled live neuron terminals with fluorescent primary antibodies to HRP, and also visualized the perisynaptic space with fluorescent Dextrans.

Our project identified GFP labeled glial structures at the developing larval fly neuromuscular synapse. To look at development of live glial-nerve-muscle ...

The Production of C. elegans Transgenes via Recombineering with the galK Selectable Marker

The creation of transgenic animals is widely utilized in C. elegans research including the use of GFP fusion proteins to study the regulation and expression pattern of genes of interest or generation of tandem affinity purification (TAP) tagged versions of specific genes to facilitate their purification. Typically transgenes are generated by placing a promoter upstream of a GFP reporter gene or cDNA of interest, and this often produces a representative expression pattern. However, critical elements of gene regulation, such as control elements in the 3' untranslated region or alternative promoters, could be missed by this approach. Further only a single splice variant can be usually studied by this means. In contrast, the use of worm genomic DNA carried by fosmid DNA clones likely includes most if not all elements involved in gene regulation in vivo which permits the greater ability to capture the genuine expression pattern and timing. To facilitate the generation of transgenes using fosmid DNA, we describe an E. coli based recombineering procedure to insert GFP, a TAP-tag, or other sequences of interest into any location in the gene. The procedure uses the galK gene as the selection marker for both the positive and negative selection steps in recombineering which results in obtaining the desired modification with high efficiency. Further, plasmids containing the galK gene flanked by homology arms to commonly used GFP and TAP fusion genes are available which reduce the cost of oligos by 50% when generating a GFP or TAP fusion protein. These plasmids use the R6K replication origin which precludes the need for extensive PCR product purification. Finally, we also demonstrate a technique to integrate the unc-119 marker on to the fosmid backbone which allows the fosmid to be directly injected or bombarded into worms to generate transgenic animals. This video demonstrates the procedures involved in generating a transgene via recombineering using this method.

The Production of C. elegans Transgenes via Recombineering with the galK Selectable Marker

The ability to produce transgenes for Caenorhabditis elegans using genomic DNA carried by fosmids is particularly attractive as all of the native regulatory elements are retained. Described is a simple and robust procedure for the production of transgenes via recombineering with the galK selectable marker.

The creation of transgenic animals is widely utilized in C. elegans research including the use of GFP fusion proteins to study the regulation and ...

High-throughput Screening and Biosensing with Fluorescent C. elegans Strains

High-throughput screening (HTS) is a powerful approach for identifying chemical modulators of biological processes. However, many compounds identified in screens using cell culture models are often found to be toxic or pharmacologically inactive in vivo1-2. Screening in whole animal models can help avoid these pitfalls and streamline the path to drug development.
C. elegans is a multicellular model organism well suited for HTS. It is small (&lt;1 mm) and can be economically cultured and dispensed in liquids. C. elegans is also one of the most experimentally tractable animal models permitting rapid and detailed identification of drug mode-of-action3.
We describe a protocol for culturing and dispensing fluorescent strains of C. elegans for high-throughput screening of chemical libraries or detection of environmental contaminants that alter the expression of a specific gene. Large numbers of developmentally synchronized worms are grown in liquid culture, harvested, washed, and suspended at a defined density. Worms are then added to black, flat-bottomed 384-well plates using a peristaltic liquid dispenser. Small molecules from a chemical library or test samples (e.g., water, food, or soil) can be added to wells with worms. In vivo, real-time fluorescence intensity is measured with a fluorescence microplate reader. This method can be adapted to any inducible gene in C. elegans for which a suitable reporter is available. Many inducible stress and developmental transcriptional pathways are well defined in C. elegans and GFP transgenic reporter strains already exist for many of them4. When combined with the appropriate transgenic reporters, our method can be used to screen for pathway modulators or to develop robust biosensor assays for environmental contaminants. 
We demonstrate our C. elegans culture and dispensing protocol with an HTS assay we developed to monitor the C. elegans cap &#x2018;n&#x2019; collar transcription factor SKN-1. SKN-1 and its mammalian homologue Nrf2 activate cytoprotective genes during oxidative and xenobiotic stress5-10. Nrf2 protects mammals from numerous age-related disorders such as cancer, neurodegeneration, and chronic inflammation and has become a major chemotherapeutic target11-13.Our assay is based on a GFP transgenic reporter for the SKN-1 target gene gst-414, which encodes a glutathione-s transferase6. The gst-4 reporter is also a biosensor for xenobiotic and oxidative chemicals that activate SKN-1 and can be used to detect low levels of contaminants such as acrylamide and methyl-mercury15-16.

High-throughput Screening and Biosensing with Fluorescent C. elegans Strains

A procedure for liquid-based culturing and dispensing of C. elegans strains expressing fluorescent reporter proteins is described that does not require expensive sorting equipment. This approach can be applied to numerous inducible C. elegans genes for drug discovery or biosensing of contaminants.

High-throughput screening (HTS) is a powerful approach for identifying chemical modulators of biological processes.  However, many compounds identified in ...

RNAi Screening to Identify Postembryonic Phenotypes in C. elegans

C. elegans has proven to be a valuable model system for the discovery and functional characterization of many genes and gene pathways1. More sophisticated tools and resources for studies in this system are facilitating continued discovery of genes with more subtle phenotypes or roles. 
Here we present a generalized protocol we adapted for identifying C. elegans genes with postembryonic phenotypes of interest using RNAi2. This procedure is easily modified to assay the phenotype of choice, whether by light or fluorescence optics on a dissecting or compound microscope. This screening protocol capitalizes on the physical assets of the organism and molecular tools the C. elegans research community has produced. As an example, we demonstrate the use of an integrated transgene that expresses a fluorescent product in an RNAi screen to identify genes required for the normal localization of this product in late stage larvae and adults. First, we used a commercially available genomic RNAi library with full-length cDNA inserts. This library facilitates the rapid identification of multiple candidates by RNAi reduction of the candidate gene product. Second, we generated an integrated transgene that expresses our fluorecently tagged protein of interest in an RNAi-sensitive background. Third, by exposing hatched animals to RNAi, this screen permits identification of gene products that have a vital embryonic role that would otherwise mask a post-embryonic role in regulating the protein of interest. Lastly, this screen uses a compound microscope equipped for single cell resolution.

RNAi Screening to Identify Postembryonic Phenotypes in C. elegans

We describe a sensitized method to identify postembryonic regulators of protein expression and localization in C. elegans using an RNAi-based genomic screen and an integrated transgene that expresses a functional, fluorescently tagged protein.

C. elegans has proven to be a valuable model system for the discovery and functional characterization of many genes and gene pathways1. More sophisticated ...

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 ...

Creating Defined Gaseous Environments to Study the Effects of Hypoxia on C. elegans

Oxygen is essential for all metazoans to survive, with one known exception1. Decreased O2 availability (hypoxia) can arise during states of disease, normal development or changes in environmental conditions2-5. Understanding the cellular signaling pathways that are involved in the response to hypoxia could provide new insight into treatment strategies for diverse human pathologies, from stroke to cancer. This goal has been impeded, at least in part, by technical difficulties associated with controlled hypoxic exposure in genetically amenable model organisms.
The nematode Caenorhabditis elegans is ideally suited as a model organism for the study of hypoxic response, as it is easy to culture and genetically manipulate. Moreover, it is possible to study cellular responses to specific hypoxic O2 concentrations without confounding effects since C. elegans obtain O2 (and other gasses) by diffusion, as opposed to a facilitated respiratory system6. Factors known to be involved in the response to hypoxia are conserved in C. elegans. The actual response to hypoxia depends on the specific concentration of O2 that is available. In C. elegans, exposure to moderate hypoxia elicits a transcriptional response mediated largely by hif-1, the highly-conserved hypoxia-inducible transcription factor6-9. C .elegans embryos require hif-1 to survive in 5,000-20,000 ppm O27,10. Hypoxia is a general term for "less than normal O2". Normoxia (normal O2) can also be difficult to define. We generally consider room air, which is 210,000 ppm O2 to be normoxia. However, it has been shown that C. elegans has a behavioral preference for O2 concentrations from 5-12% (50,000-120,000 ppm O2)11. In larvae and adults, hif-1 acts to prevent hypoxia-induced diapause in 5,000 ppm O212. However, hif-1 does not play a role in the response to lower concentrations of O2 (anoxia, operational definition &lt;10 ppm O2)13. In anoxia, C. elegans enters into a reversible state of suspended animation in which all microscopically observable activity ceases10. The fact that different physiological responses occur in different conditions highlights the importance of having experimental control over the hypoxic concentration of O2.
Here, we present a method for the construction and implementation of environmental chambers that produce reliable and reproducible hypoxic conditions with defined concentrations of O2. The continual flow method ensures rapid equilibration of the chamber and increases the stability of the system. Additionally, the transparency and accessibility of the chambers allow for direct visualization of animals being exposed to hypoxia. We further demonstrate an effective method of harvesting C. elegans samples rapidly after exposure to hypoxia, which is necessary to observe many of the rapidly-reversed changes that occur in hypoxia10,14. This method provides a basic foundation that can be easily modified for individual laboratory needs, including different model systems and a variety of gasses.

Creating Defined Gaseous Environments to Study the Effects of Hypoxia on C. elegans

This paper details how to use continuous-flow hypoxia chambers to generate atmospheres with defined concentrations of O2 to understand biological responses to decreased O2. This system is easy to setup and maintain, and flexible enough to suit a wide range of O2 concentrations and model systems

Oxygen is essential for all metazoans to survive, with one known exception1. Decreased O2 availability (hypoxia) can arise during states of disease, ...

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 ...

A Method for Culturing Embryonic C. elegans Cells

C. elegans is a powerful model system, in which genetic and molecular techniques are easily applicable. Until recently though, techniques that require direct access to cells and isolation of specific cell types, could not be applied in C. elegans. This limitation was due to the fact that tissues are confined within a pressurized cuticle which is not easily digested by treatment with enzymes and/or detergents. Based on early pioneer work by Laird Bloom, Christensen and colleagues 1 developed a robust method for culturing C. elegans embryonic cells in large scale. Eggs are isolated from gravid adults by treatment with bleach/NaOH and subsequently treated with chitinase to remove the eggshells. Embryonic cells are then dissociated by manual pipetting and plated onto substrate-covered glass in serum-enriched media. Within 24 hr of isolation cells begin to differentiate by changing morphology and by expressing cell specific markers. C. elegans cells cultured using this method survive for up 2 weeks in vitro and have been used for electrophysiological, immunochemical, and imaging analyses as well as they have been sorted and used for microarray profiling.

A Method for Culturing Embryonic C. elegans Cells

We describe here a revised protocol for large-scale culture of embryonic C. elegans cells. Embryonic C. elegans cells cultured in vitro using this method, appear to differentiate and recapitulate the expression of genes in a cell specific manner. Techniques that require direct access to the cells or isolation of specific cell types from the other tissues can be applied on C. elegans cultured cells.

C. elegans is a powerful model system, in which genetic and molecular techniques are easily applicable. Until recently though, techniques that require ...

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 ...