Funding was provided by NSF CCLI#0633618 and Ohio University. Some strains were provided by the Caenorhabditis Genetics Center. Graduate student Reetobrata Basu appears in the video.

NOTE: Multiple protocol steps are presented with options for set-up and preparation depending on the specific conditions of the experiment. For these cases, alternative steps are presented as A, B, C, etc. The protocol with alternative steps is outlined in Figure 1.
1. Preparation of Polylysine Coated Slides
Three different types of slides may be used, depending upon the desired trade-off between ease of preparation, relative adhesion, and non-specific binding of the antibody. Details of slide preparation alternatives are given below in steps 1A-1C in order of increasing adhesion and complexity. Slides prepared by these different methods can be used together for a single experiment.
1A. No slide preparation
Commercially available polylysine coated slides provide low adhesion and low background.
1B. Slide preparation optimum for methanol-acetone fixation
Medium adhesion and low background.
<ol>
	<li>Prepare a polylysine solution for dipping the slides: Add 400 mg of very high molecular weight (150,000 - 300,000 D) poly-L-lysine to 200 ml double distilled water. Add 2 ml 10% sodium azide stock (final concentration 0.1%) as a preservative. CAUTION: dry sodium azide is reactive and all forms are toxic. Wear mask and gloves while weighing powder to make 10% stock in double distilled water.</li>
	<li>Wipe single end frosted glass slides with lint-free wipes, removing smudges and particles. This should be done even on slides purchased with the designation &#34;precleaned.&#34;</li>
	<li>Place slides back-to-back in a slide holder, coplin jar or other slide staining container, keeping frosted edges up and not quite perfectly aligned (to make later separation easier). Place slice holder in beaker with 70% ethanol (reagent grade alcohol not needed) with 1% HCl in distilled water or fill staining jar to level of frosting. Rock slides in solution for a minimum of 5 min. CAUTION: This solution is corrosive, wear gloves. NOTE: This solution can be reused several times (up to several months).</li>
	<li>Rinse slides in running distilled water for 1 min.</li>
	<li>Dry slides completely in a dust free environment either by placing in a 40-60 &#176;C oven for 30-60 min or by keeping overnight at room temperature.</li>
	<li>Incubate slides in polylysine solution (covering unfrosted parts) on rocker for 15 min.</li>
	<li>Repeat drying of slides.</li>
	<li>Repeat polylysine solution incubation and drying steps.</li>
	<li>Separate the slides using a slim metal object such as a thin weighing spatula. If they are properly adhered, they are difficult to separate. Wear appropriate safety gear (safety glasses) and work on bench top paper (to be discarded after use), since small bits of glass may fly loose when slides are separated.</li>
	<li>Store slides in clean (dust-free) slide box at 4 &#176;C until needed.</li>
</ol>
1C. Slide preparation optimum for formaldehyde fixation
Highest adhesion but high background.
<ol>
	<li>Place slides prepared with method 1B flat in a dust-free, oven-safe dish (for drying in oven) or on a flat dust free surface (for air drying).</li>
	<li>Place a 20-60 &#956;l drop of the polylysine solution on the middle of each slide.</li>
	<li>Dry slides completely in oven or at room temperature. The polylysine will leave behind pale ovals as it evaporates. These ovals are very adhesive. Unfortunately, they also bind to the primary and secondary antibodies, even after blocking.</li>
	<li>Store slides in clean (dust-free) slide box at 4 &#176;C until needed.</li>
</ol>
2. Preparation of Frozen Nematode Slides
Prepare nematodes for staining by completely rinsing free of bacteria using method 2A (for plates of nematodes) or 2B (for individual nematodes).
2A. Preparation of slides from plates of nematodes
<ol>
	<li>Place a flat piece of metal on dry ice to prechill.</li>
	<li>Use distilled water and a glass pipette (decreases loss of worms) or micropipette to rinse worms from NGM plate with bacteria into a 1.5 ml microfuge tube. Plates can be synchronized or mixed ages; do not use plates with many dauers (difficult to crack) or starved worms (increased autofluorescence) unless necessary.</li>
	<li>Rinse the worms 3-4x&#160;with distilled water until free of bacteria. (To decrease loss of worms use the same glass pipette.) To pellet worms, either let them sit on the bench for 3-5 min (to collect mostly adults at bottom of tube) or spin 30 sec at ~1,000 rpm (~500 g) in a tabletop centrifuge (to collect adults, larva, and embryos).</li>
	<li>Remove most of water from tube with worms, leaving about twice the volume of water as worms (but at least 25 &#956;l total volume).</li>
	<li>Label frosted side of the &#34;bottom&#34; (treated in step 1) polylysine slides with indelible ink and place slides on counter next to container with dry ice, label side up.</li>
	<li>Have an equal number of regular (wiped clean but not polylysine treated) &#34;top&#34; slides available.</li>
	<li>Place ~25 &#956;l worms in water on the &#34;bottom&#34; adherent slide and spread the liquid containing the worms over the central portion of the slide using the side of pipette tip.</li>
	<li>Let the worms settle for 30 sec to 2 min. If the slide is appropriately sticky, the ends of the worms will stick as they contact the slide.</li>
	<li>Hold the bottom slide in one hand and place the top slide over the bottom slide so that all but the frosted parts are overlapping. The liquid should keep the slides slightly apart, so that the worms still move slightly. Do not let the slides slip over one another - this twists the worms.</li>
	<li>Carefully press straight down slightly on the top slide, using the thumbs and forefingers of each hand. With the ideal amount of pressure, a few of the largest hermaphrodites on the edge of the slides will rupture, while most of the adults and larvae will contact each slide but will stay intact.</li>
	<li>Immediately and carefully (without slipping) put the slides on the piece of metal on dry ice. The slides should appear frozen within one minute. Keep on dry ice for 5 min&#160;until thoroughly frozen.</li>
	<li>At this point, slides may be stored in a labeled box at -80 &#176;C for several days. (If they are kept weeks at -80 &#176;C, the water will start to sublime away and the worms will not stain as well). If slides are stored at -80 &#176;C, let them &#39;warm up&#39; on dry ice for 10 min before cracking.</li>
	<li>Proceed to step 3 (fixation).</li>
</ol>
2B. Preparation of slides of individual nematodes
<ol>
	<li>Place a flat piece of metal on dry ice to prechill.</li>
	<li>Use a worm pick to transfer individual nematodes to a drop of water on an NGM plate to free them from bacteria. Nematode(s) should be well-fed to decrease autofluorescence, if possible. Repeat transfer to new spots of water as needed until worm(s) are free of bacteria.</li>
	<li>Label frosted side of the &#34;bottom&#34; polylysine slides with indelible ink and place slides on counter next to container with dry ice, label side up. Have an equal number of less adherent &#34;top&#34; (untreated) slides available.</li>
	<li>Place a 5-10 &#956;l drop of water on the region of the bottom slide double-treated with polylysine. Use the larger volume if transferring more worms, to prevent the water from evaporating before completion.</li>
	<li>Transfer the worm(s) to the drop of water with the worm pick, using the microscope to watch as the worms sink down to touch the sticky slide surface. Transfer as few as one worm to as many as 20+ worms per drop.</li>
	<li>Hold the bottom slide in one hand and place the top slide over the bottom slide so that all but the frosted parts are overlapping.</li>
	<li>Immediately and carefully (without slipping or compressing) put the slides on the piece of metal on dry ice. Keep on dry ice for 5-30 min. (Do not store these slides long term, as the slides easily dry out.)</li>
	<li>Proceed to step 3 (fixation).</li>
</ol>
3. Fixation
Fixatives are necessary to &#39;fix&#39; the antigen in place in the cell, by either precipitating (&#34;light fixation&#34;) or cross-linking (&#34;hard fixation&#34;) the antigen. Fixation may disrupt antigenicity; most known antibodies work best with a particular fixation condition. For new antibodies, a range of fixation conditions should be tested.
For any fixation, the first step is to make phosphate buffered saline solution. Next fix slides with nematodes for antibody staining using method A (light fix using methanol and acetone) or B (harder fix using formaldehyde or glutaraldehyde).
<ol>
	<li>Prepare a sterile 10x&#160;PBS (phosphate buffered saline) stock solution by adding 80 g NaCl, 2.0 g KCl, 27.2 g Na2HPO4&bull;H20, 2.4 g KH2PO4, 20 ml 10% sodium azide stock. To make this stock solution, weigh out sodium azide powder to make 10% solution in double distilled water. Stir saline with gentle heat until dissolved. CAUTION: dry sodium azide is reactive and all forms are toxic. Wear mask and gloves when working with dry sodium azide or solutions.</li>
	<li>Let saline cool (Tris pH is temperature sensitive), then pH to 7.2 with 1-10 M NaOH. Top to 1 L with double distilled water and autoclave to sterilize. Store at room temperature and dilute to 1x&#160;as needed with sterile double distilled water. CAUTION: NaOH is caustic - wear gloves during use.</li>
</ol>
3A. Light fix using methanol-acetone
<ol>
	<li>Put 100% methanol, 100% acetone, and 1x&#160;phosphate buffered saline (PBS) in staining jars on ice to prechill for at least 10 min. CAUTION: Methanol and acetone are toxic, wear gloves.</li>
	<li>Take a pair of bottom and top slides from the dry ice, swiftly twist them apart, and immediately immerse the &#34;bottom&#34; slide in ice cold methanol for 2 min. The &#34;top&#34; slide may be discarded.</li>
	<li>Transfer slides from methanol to ice cold acetone for 4 min. If the slides had many worms, most (90%) will fall off in the fix, even with the best prepared slides. If single worm slides were properly prepared, the worms should stay adhered.</li>
	<li>Place or dip the slides in the jar with PBS (to rinse off fixative) and proceed to step 4 (staining).</li>
</ol>
3B. Hard fix using formaldehyde or glutaraldehyde
<ol>
	<li>Dilute reagent grade formaldehyde or glutaraldehyde solution in 1x&#160;PBS to 0.5-4% final concentration. Aliquots (1.5 ml) may be stored tightly capped at -20 to -30 &#176;C; defrost aliquots on ice just before using. NOTE: formaldehyde solutions degrade over time and with exposure to air. CAUTION: formaldehyde and glutaraldehyde are toxic, wear gloves and work in the hood.</li>
	<li>Take a pair of bottom and top slides from the dry ice, swiftly twist them apart, and immediately place the &#34;bottom&#34; slide with worms in the flat dish with lid. (The top slide can be discarded).</li>
	<li>Immediately pipette 200 &#956;l toxic fixative over the center of the area of the slide with worms. The fixative will partially freeze in place, then melt to cover a larger region of the slide.</li>
	<li>If the worms are not totally covered with fixative, immediately and carefully add more fixative using a micropipette. Do not allow fixative to flow over the edge of the slide.</li>
	<li>Cover the dish with a lid and leave for 10 min to overnight at room temperature. Be sure the dish is humidified if longer incubations are used - this can be done by adding folded wet lint-free wipes to the dish. Do not let the wipes touch the slides.</li>
	<li>After fixation is complete, carefully pipette the fixative with loose worms into a 1.5 ml tube and dip the slide in a staining jar with PBS.</li>
	<li>Spin the tube with worms that came loose at high speed for 30 sec to pellet. Rinse the worms twice with PBS, then process in parallel with the worms on the slides (doing steps in microfuge tubes, rather than on slides).</li>
	<li>Proceed to step 4 (staining).</li>
</ol>
4. Antibody Staining
The antibody staining protocol is similar to standard protocols for any tissue on slides. This protocol is the same for all slides regardless of the preparation in steps 1-3.
<ol>
	<li>Prepare Antibody Buffer by mixing 700 ml double distilled water, 100 ml 10x&#160;PBS, 5 ml Triton X-100 (0.5% final), 2 ml 0.5 M EDTA pH 8 (1 mM final), 1 g BSA (0.1% final), 5 ml 10% sodium azide solution (0.05% final). pH to 7.2 with HCl, then add double distilled water to 1 L; store at 4 &#176;C.</li>
	<li>Prepare Block by reconstituting donkey serum with double distilled water, then adding 5 ml serum to 45 ml antibody buffer (final 10% serum).</li>
	<li>Prepare primary and secondary Antibody Solutions by diluting antibody in antibody buffer plus 1% BSA. Recommended dilutions are 1:50-1:2,000 for primary antibodies and 1:1,000-1:5,000 for secondary antibodies (Cy3or OG488 conjugated donkey antibodies against species used to raise primary antibody).</li>
	<li>Prepare DAPI stock by mixing 2.5 mg/ml in double distilled water; store in 8 &#956;l aliquots frozen at -30 &#176;C. CAUTION: DAPI is a toxic DNA binding dye, wear gloves while weighing and dispensing into aliquots.</li>
	<li>Prepare 10 ml Mounting Medium by mixing 200 mg n-propyl gallate, 0.3 ml 1 M Tris pH 9, 2.7 ml double distilled water in a 15 ml conical tube. Heat at 65 &#176;C for about 10 min until dissolved. Add 7 ml glycerol and an 8 &#956;l aliquot of DAPI and vortex well. Aliquot medium into 1.5 ml tubes, wrap in foil, and store in dark at 4 &#176;C (both air and light degrade the medium - if it becomes more yellow, discard in Biohazard waste). CAUTION: n-propyl gallate is a reactive anti-oxidant, wear gloves while weighing.</li>
	<li>Transfer slides to a staining jar with Block for 1 hr at room temperature or overnight at 4 &#176;C (overnight preferable for hard fix). Avoid shaking to avoid losing worms.</li>
	<li>Transfer slides to staining jar with primary antibody or place slides flat in a humidified dish and top each with 100-500 &#956;l primary antibody. Incubate slides overnight at 4 &#176;C without shaking.</li>
	<li>After primary antibody incubation, transfer slides to staining jar of PBS.</li>
	<li>Filter Block through funnel lined with filter paper and store at 4 &#176;C; if sterile filtered every 1-2 weeks, Block can be reused for 1 or more months. Similarly, filter and save primary antibody at 4 &#176;C for reuse (for up to 10 or more rounds of staining).</li>
	<li>Rinse slides three times (~20 min each) in antibody buffer.</li>
	<li>Place slides in secondary antibody in a light-tight staining container and incubate 4 hr at room temperature or overnight at 4 &#176;C.</li>
	<li>Transfer slides to staining jar of PBS. Filter and save secondary antibody at 4 &#176;C for reuse (for up to 10 or more rounds of staining).</li>
	<li>Rinse slides three times (~20 min each) in antibody buffer.</li>
	<li>Transfer slides to PBS and leave in PBS (minutes to overnight at 4 &#176;C) until ready to mount.</li>
	<li>Work rapidly on individual slides, so that the worms on front of slide stay wet. Remove one slide from the PBS, dry back of the slide with a lint-free wipe, and place on paper towel.</li>
	<li>Place ~20 &#956;l of mounting medium over worms so that there are approximately equal parts medium and PBS on slide and tilt slide gently to mix the two solutions. CAUTION: Mounting medium is toxic.</li>
	<li>Tilt slide so frosted edge is lower and solution gathers near frosting, then place one edge of 24 x 60 mm coverslip on solution.</li>
	<li>Gently lower the coverslip to minimize air bubbles (which increase bleaching and disrupt viewing). If bubbles form, lift the edge of the coverslip slowly, then relower it without sliding the coverslip across the worms.</li>
	<li>Carefully dry off the edge of the slide without moving the coverslip, by compressing the edges with a lint-free wipe. The relatively dry edge of the slide must be visible all around the edge of the coverslip</li>
	<li>Seal the coverslip with 2-3 layers of nail polish. Slides can be placed in a flat slide holder during preparation and storage to protect from light. Once slide is well sealed and dry, clean the coverslip and back of the slide.</li>
	<li>Well-sealed slides can be kept in the dark for days at 4 &#176;C or weeks at -20 &#176;C. Over longer periods, DAPI staining of DNA and most green dyes (e.g.&#160;488 nm excitation) fade. Reseal the coverslip with more nail polish as needed, e.g.&#160;after using any oil immersion lens.</li>
</ol>

<ol>
	<li>Harlow, E., Lane, D. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=.">.</a> Antibodies: A Laboratory Manual. , (1988).</li><li>Mello, C., 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=DNA+Transformation.">DNA Transformation.</a> Methods Cell Biol. 48, 451-482 (1995).</li><li>Boulin, T., Etchberger, J. F., Hobert, O. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Reporter+gene+fusions.">Reporter gene fusions.</a> WormBook. , (2006).</li><li>Duerr, J. S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Immunohistochemistry.">Immunohistochemistry.</a> WormBook. , (2006).</li><li>Finney, M., Ruvkun, G. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+unc-86+gene+product+couples+cell+lineage+and+cell+identity+in+C.+elegans.">The unc-86 gene product couples cell lineage and cell identity in C. elegans.</a> Cell. 63, 895-905 (1990).</li><li>Garriga, G., Desai, C., Horvitz, H. R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Cell+interactions+control+the+direction+of+outgrowth,+branching+and+fasciculation+of+the+HSN+axons+of+Caenorhabditis+elegans.">Cell interactions control the direction of outgrowth, branching and fasciculation of the HSN axons of Caenorhabditis elegans.</a> Development. 117, 1071-1087 (1993).</li><li>Miller, D. M., Shakes, D. C., Epstein, H. F., Shakes, D. C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Immunofluorescence+microscopy.">Immunofluorescence microscopy.</a> Caenorhabditis elegans: Modern Biological Analysis of an Organism. , 365-394 (1995).</li><li>Crittenden, S. L., Troemel, E. R., Evans, T. C., Kimble, J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=GLP-1+is+localized+to+the+mitotic+region+of+the+C.+elegans+germ+line.">GLP-1 is localized to the mitotic region of the C. elegans germ line.</a> Development. 120, 2901-2911 (1994).</li><li>Strome, S., Wood, W. B. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Immunofluorescence+visualization+of+germ-line-specific+cytoplasmic+granules+in+embryos,+larvae,+and+adults+of+Caenorhabditis+elegans.">Immunofluorescence visualization of germ-line-specific cytoplasmic granules in embryos, larvae, and adults of Caenorhabditis elegans.</a> Proc. Natl. Acad. Sci. U.S.A. 79, 1558-1562 (1982).</li><li>Albertson, D. G. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Formation+of+the+first+cleavage+spindle+in+nematode+embryos.">Formation of the first cleavage spindle in nematode embryos.</a> Dev. Biol. 101, 61-72 (1984).</li><li>Harlow, 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> Using Antibodies: A Laboratory Manual. , (1999).</li><li>Ahringer, J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Reverse+Genetics.">Reverse Genetics.</a> WormBook. , (2006).</li></ol>

The author declares that she has no competing financial interests.

The freeze-cracking protocol is one of several methods for antibody staining in C. elegans. It provides a relatively simple way to stain worms, but does require specific reagents and practice for optimum results. Critical steps (outlined in Figure 1) include: 1) slide preparation (see steps 1 and&#160;2) manual compression of the nematodes (see steps 2 and 3) rapid fixation (see step 3). First, to maximize adhesion of the nematodes to the slides, it is best to prepare slides with high molecular ...

To determine the cellular and subcellular localization of proteins, scientists have traditionally labeled tissues with antibodies selected to specifically recognize particular proteins1. In some model organisms such as C. elegans, antibody staining has often been replaced by molecular genetic techniques, which yield results more quickly. These include transforming organisms with constructs consisting of gene fusions between the promoter and coding regions of the gene of interest and green fluorescent protein. However, molecular techniques are subject to a number of artifacts, including problems with knowing the true promoter and changes in expression of constructs at high copy number (the usual techniques in C. elegans)2,3. Therefore, staining with antibodies remains a goal for many scientists studying protein function in vivo.
Staining tissues with antibodies can be difficult, since antibodies may only recognize their antigen in a particular conformation1. For example, antibodies may recognize only denatured or intact antigen fixed in a particular way and may not recognize the antigen in situ. The problem of antibody staining in nematodes is exacerbated by the fact that the cuticle of the nematode forms a relatively impermeable barrier, blocking access of antibodies to tissues.
There are several different methods used for antibody staining in C. elegans (reviewed in our previous work4). To stain intact &#39;worms,&#39; methods were developed to freeze and thaw in a relatively hard fixative (formaldehyde or glutaraldehyde); the freeze-thaw cycles help to crack the cuticle to allow rapid penetration of the fixative5. After fixation, the cuticle was permeabilized to allow penetration of the antibody; methods included treatment with reducing agents, collagenase, or both5-7. These treatments preserved morphology, but often reduced or destroyed antibody recognition. Alternative methods include dissection8 to allow antibody penetration.
&#34;Freeze-cracking&#34; is one way to gain access to the interior of the semi-intact worm to allow antibody staining9-10. Freeze-cracking can be performed with a variety of fixation conditions while avoiding collagenase and reduction treatments. The nematodes are placed between two adhesives slides, frozen, and then the slides are separated, leaving most of the nematode on the bottom slide and much of the cuticle on the top slide. The bottom slide can be placed in fixative and the entire slide with adhered nematodes is transferred through the antibody staining procedure. The method does present two major difficulties. First, it is difficult to apply the correct amount of pressure necessary to split the nematodes without seriously deforming them. Second, many of the worms will not stick to either slide, and will be lost in the fixative or rinses. However, with the proper slides and practice, the method will rapidly yield nematodes with reasonable morphology which can be used with a wide variety of fixatives and antibodies.
The method may be varied slightly, depending upon the goal of the experimenter. If staining of single nematodes is desired (e.g.&#160;to determine whether a single transformant has altered antibody staining), then slides with maximal adhesion (but higher background staining) can be used, such as laboratory-prepared slides with extra polylysine (see below). For formaldehyde or glutaraldehyde fixation, higher adhesion slides (laboratory-prepared polylysine slides) should be used, since nematodes adhere more poorly to polylysine after these fixations. If wild-type nematodes are being fixed with methanol and/or acetone, then slides with lower adhesion and lower background staining should be used (commercial or laboratory-prepared slides). If a variety of antibodies and fixatives are being used in the lab, a selection of slides may be prepared ahead of time and stored until needed.
After access to the nematode tissue has been gained, antibody staining procedures follow standard methods (with longer incubations characteristic of tissues rather than cells1,11).

<table border="1">
		<tbody>
		 <tr>
			<td>Name of Reagent/Material</td>
			<td>Company</td>
			<td>Catalog Number</td>
			<td>Comments</td>
		 </tr>
		 <tr>
			<td>poly-L-lysine slide (ESCO brand)</td>
			<td>Fisher</td>
			<td>12-545-78</td>
			<td>Sigma brand slides also suitable</td>
		 </tr>
		 <tr>
			<td>poly-L-lysine hydrobromide</td>
			<td>Sigma</td>
			<td>P1524</td>
			<td>IMPORTANT: Brand and high molecular weight of polylysine critical</td>
		 </tr>
		 <tr>
			<td>single frosted edge slides</td>
			<td>Fisher</td>
			<td>48312-002</td>
			<td>Other brands are suitable</td>
		 </tr>
		 <tr>
			<td>sodium azide</td>
			<td>Sigma</td>
			<td>S2002-25G</td>
			<td>TOXIC, wear gloves and mask while weighing out powder to make 10% stock. Wear gloves when pipeting stock solution</td>
		 </tr>
		 <tr>
			<td>coplin staining jar</td>
			<td>VWR</td>
			<td>47751-792</td>
			<td>Other brands are suitable</td>
		 </tr>
		 <tr>
			<td>5-slide mailer</td>
			<td>Electron Microscopy Science</td>
			<td>71549-01</td>
			<td>One of many different brands of small slide holder, suitable for staining</td>
		 </tr>
		 <tr>
			<td>Paraformaldehyde</td>
			<td>VWR</td>
			<td>MK262159</td>
			<td>IMPORTANT: Regular formalin solutions (37% formaldehyde) are NOT suitable. Either 1) Dissolve crysatlaline paraformaldehyde in phosphate buffer and store in small aliquots frozen. Or 2) buy electron microscopy grade paraformaldehyde in aliquots. TOXIC -wear gloves; dispose of excess down sink with water.</td>
		 </tr>
		 <tr>
			<td>Glutaraldehyde solution, 25% in water</td>
			<td>Sigma</td>
			<td>G 5882</td>
			<td>IMPORTANT: purchase electron microscopy grade glutaraldehyde in ampules; TOXIC -wear gloves; dispose of excess down sink with water.</td>
		 </tr>
		 <tr>
			<td>Triton X-100</td>
			<td>VWR</td>
			<td>EM-9410</td>
			<td>Other brands are suitable</td>
		 </tr>
		 <tr>
			<td>Bovine Serum Albumin</td>
			<td>Fisher</td>
			<td>ICN820451</td>
			<td>Other brands are suitable</td>
		 </tr>
		 <tr>
			<td>Donkey Serum, lyophilized</td>
			<td>Jackson Immunoresearch</td>
			<td>017-000-121</td>
			<td>Other brands are suitable</td>
		 </tr>
		 <tr>
			<td>Cy3 Donkey anti-Mouse IgG multi-labeling</td>
			<td>Jackson Immunoresearch</td>
			<td>715-165-150</td>
			<td>This company is recommended for high quality multi-label (affinity depleted) antibodies. Select appropriate dye and antigen (animal used to raise primary antibody). </td>
		 </tr>
		 <tr>
			<td>n-propyl gallate</td>
			<td>Fisher</td>
			<td>ICN10274750</td>
			<td>Other brands are suitable</td>
		 </tr>
		 <tr>
			<td>DAPI, 4',6'-diamidino-2-phenylindole dihydrochloride</td>
			<td>Sigma</td>
			<td>D 9542</td>
			<td>TOXIC. Wear gloves and dispense into small aliquots and feeze to minimize exposure</td>
		 </tr>
		 <tr>
			<td>Whatman #1 15 cm diameter filters</td>
			<td>Fisher</td>
			<td>09805G</td>
			<td></td>
		 </tr>
		 <tr>
			<td>Coverslip #1 1/2 rectangle 24 x 60 mm</td>
			<td>VWR</td>
			<td>48393-252</td>
			<td>#1 1/2 thickness is optically best</td>
		 </tr>
		 <tr>
			<td>Microscope slide folder</td>
			<td>VWR</td>
			<td>48429-092</td>
			<td>Other brands are suitable</td>
		 </tr>
		</tbody>
 </table>

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.

When worms are properly compressed, cracked, and lightly fixed, virtually the entire worm can be accessible to antibody staining (see Figure 2). The location of the nuclei as indicated by DAPI staining indicates which parts of the worm are intact When the worms are subjected to a harder fixative, such as formaldehyde, the morphology of the worm may become distorted (as seen by the unnaturally wavy appearance of the muscles in Figures 3A-D). Another common problem is uneven fixation or pe...

Watch this Scientific Journal Video about Antibody Staining in C. Elegans Using "Freeze-Cracking" at JoVE.com

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.

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 ...

antibody-staining-in-c-elegans-using-freeze-cracking

Research

JoVE Journal

Biology

22.5K Views.  Ohio University. "Freeze-cracking," a method for exposing the inner tissues of the nematode C. elegans to antibodies for protein localization, is demonstrated.

Video: Antibody Staining in C. Elegans Using "Freeze-Cracking"

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 ...

Method for Whole Mount Antibody Staining in Chick

The chick embryo is a valuable tool in the study of early embryonic development. Its transparency, accessibility and ease of manipulation, make it an ideal tool for studying antibody expression in developing brain, neural tube and somite. This video demonstrates the different steps in whole-mount antibody staining using HRP conjugated secondary antibodies; First, the embryo is dissected from the egg and fixed in paraformaldehyde. Second, endogenous peroxidase is inactivated; The embryo is then exposed to primary antibody. After several washes, the embryo is incubated with secondary antibody conjugated to HRP. Peroxidase activity is revealed using reaction with diaminobenzidine substrate. Finally, the embryo is fixed and processed for photography and sectioning. The advantage of this method over the use of fluorescent antibodies is that embryos can be processed for wax sectioning, thus enabling the study of antigen sites in cross section. This method was originally introduced by Jane Dodd and Tom Jessell 1.

This video demonstrates whole mount immunohistochemistry, a method by which the spatial and temporal expression pattern of an antigen can be visualized in young chick embryos. This method was originally introduced by Jane Dodd and Tom Jessell.

The chick embryo is a valuable tool in the study of early embryonic development. Its transparency, accessibility and ease of manipulation, make it an ...

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 ...

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, ...

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 ...

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 ...

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 ...

Methods to Assess Subcellular Compartments of Muscle in C. elegans

Muscle is a dynamic tissue that responds to changes in nutrition, exercise, and disease state. The loss of muscle mass and function with disease and age are significant public health burdens. We currently understand little about the genetic regulation of muscle health with disease or age. The nematode C. elegans is an established model for understanding the genomic regulation of biological processes of interest. This worm&#8217;s body wall muscles display a large degree of homology with the muscles of higher metazoan species. Since C. elegans is a transparent organism, the localization of GFP to mitochondria and sarcomeres allows visualization of these structures in vivo. Similarly, feeding animals cationic dyes, which accumulate based on the existence of a mitochondrial membrane potential, allows the assessment of mitochondrial function in vivo. These methods, as well as assessment of muscle protein homeostasis, are combined with assessment of whole animal muscle function, in the form of movement assays, to allow correlation of sub-cellular defects with functional measures of muscle performance. Thus, C. elegans provides a powerful platform with which to assess the impact of mutations, gene knockdown, and/or chemical compounds upon muscle structure and function. Lastly, as GFP, cationic dyes, and movement assays are assessed non-invasively, prospective studies of muscle structure and function can be conducted across the whole life course and this at present cannot be easily investigated in vivo in any other organism.

Methods to Assess Subcellular Compartments of Muscle in C. elegans

Skeletal muscle is essential for locomotion and is the bodies&#8217; main protein store. Muscle health measurements within C. elegans are described. Prospective changes to muscle structure and function are assessed using localized GFP and cationic dyes.

Muscle is a dynamic tissue that responds to changes in nutrition, exercise, and disease state. The loss of muscle mass and function with disease and age ...

Tracking and Quantifying Developmental Processes in C. elegans Using Open-source Tools

Quantitatively capturing developmental processes is crucial to derive mechanistic models and key to identify and describe mutant phenotypes. Here protocols are presented for preparing embryos and adult C. elegans animals for short- and long-term time-lapse microscopy and methods for tracking and quantification of developmental processes. The methods presented are all based on C. elegans strains available from the Caenorhabditis Genetics Center and on open-source software that can be easily implemented in any laboratory independently of the microscopy system used. A reconstruction of a 3D cell-shape model using the modelling software IMOD, manual tracking of fluorescently-labeled subcellular structures using the multi-purpose image analysis program Endrov, and an analysis of cortical contractile flow using PIVlab (Time-Resolved Digital Particle Image Velocimetry Tool for MATLAB) are shown. It is discussed how these methods can also be deployed to quantitatively capture other developmental processes in different models, e.g., cell tracking and lineage tracing, tracking of vesicle flow.

Tracking and Quantifying Developmental Processes in C. elegans Using Open-source Tools

Here it is shown how to track and quantify developmental processes in C. elegans. The methods presented are based on open-source tools that can be easily implemented. It is demonstrated how to reconstruct 3D cell-shape models, how to manually track subcellular structures, and how to analyze cortical contractile flow.

Quantitatively capturing developmental processes is crucial to derive mechanistic models and key to identify and describe mutant phenotypes. Here ...

Measurements of Physiological Stress Responses in C. Elegans

Organisms are often exposed to fluctuating environments and changes in intracellular homeostasis, which can have detrimental effects on their proteome and physiology. Thus, organisms have evolved targeted and specific stress responses dedicated to repair damage and maintain homeostasis. These mechanisms include the unfolded protein response of the endoplasmic reticulum (UPRER), the unfolded protein response of the mitochondria (UPRMT), the heat shock response (HSR), and the oxidative stress response (OxSR). The protocols presented here describe methods to detect and characterize the activation of these pathways and their physiological consequences in the nematode, C. elegans. First, the use of pathway-specific fluorescent transcriptional reporters is described for rapid cellular characterization, drug screening, or large-scale genetic screening (e.g., RNAi or mutant libraries). In addition, complementary, robust physiological assays are described, which can be used to directly assess sensitivity of animals to specific stressors, serving as functional validation of the transcriptional reporters. Together, these methods allow for rapid characterization of the cellular and physiological effects of internal and external proteotoxic perturbations.

Measurements of Physiological Stress Responses in C. Elegans

Here, we characterize cellular proteotoxic stress responses in the nematode C. elegans by measuring the activation of fluorescent transcriptional reporters and assaying sensitivity to physiological stress.

Organisms are often exposed to fluctuating environments and changes in intracellular homeostasis, which can have detrimental effects on their proteome and ...