Work in the Arur Lab is supported by grants from the National Institutes of Health, NIHGM98200; Cancer Prevention Research Institute of Texas, CPRIT RP160023; the American Cancer Society Research Scholar Award (ACS RSG014-044-DDC); and by the Anna Fuller Funds.

The protocol described here is primarily for the invertebrate model system C. elegans, and follows all the ethical guidelines set forth by the institution.
1. Animal Maintenance
<ol>
	<li>Maintain C. elegans working stock cultures on Nematode Growth Medium20,21 (NGM, see Materials Table) agar seeded with E. coli OP50.</li>
	<li>Maintain worm stocks at temperatures between 16 &#176;C and 25 &#176;C. 
	NOTE: Culturing worms at higher temperatures results in faster growth rate, often, aberrant germline development and lower brood sizes. Additionally, temperature sensitive genotypes will display different phenotypes at different temperatures.</li>
	<li>Transfer worms to new NGM plates every 2 - 3 days depending on their growth rate so as to maintain a constant supply of well-fed worms. 
	NOTE: For any given experiment involving dpERK levels, worms should NOT be obtained from a starved or crowded plate. Crowding or starvation impacts insulin signaling in worms22, and insulin signaling regulates ERK12 activity. Thus worms from starved or overcrowded plates will result in variable and difficult-to-reproduce staining patterns.</li>
</ol>
2. Dissection of Adult Worms for Obtaining Gonads
<ol>
	<li>Pick 100 - 150 WT (N2) or desired genotype of worms at the desired and specific developmental stage (L1, L2, L3, L4, adult, etc.) in 100 &#181;L of M9 buffer in a 1.5 mL microcentrifuge tube.</li>
	<li>Fill the microcentrifuge tube with 900 &#181;L of M9 buffer (see Materials Table). Centrifuge the tubes at 1,000 x g in a microcentrifuge for 1 min at ambient temperature.</li>
	<li>Gently remove 900 &#181;L of the M9 buffer and discard. Remove the buffer under a dissecting microscope to ensure that the worms are not accidently removed.</li>
	<li>Repeat steps 2.2 - 2.3 two more times with fresh M9 buffer each time. This ensures that any bacteria that were carried over with the worms are effectively washed away.</li>
	<li>After the final wash remove 900 &#181;L of M9 buffer from the tube and discard.</li>
	<li>Transfer the remaining 100 &#181;L of M9 buffer containing the 100 - 150 worms with a 200&#160;&#181;L-micropipette tip to a flat bottom glass watch dish. Ensure that the micropipette tip is cut at the 10&#160;&#181;L mark before use. Not cutting the tip will result in shearing of the worms.</li>
	<li>Add 1 - 3 &#181;L of 0.1 M levamisole to the glass watch dish containing the worms in M9 buffer. Gently swirl the levamisole and the M9 to enable even mixing until the worms stop moving. 
	NOTE: Levamisole stocks can be stored at -20 &#176;C for long-term storage. Here, use the higher concentration of 1 - 3 mM than what has been described in literature23 of 0.2 - 0.25 mM of levamisole in order to obtain rapid loss of mobility in the animals. If the animals flay around for too long in the liquid suspension, the resulting patterns of dpERK in the gonads can be variable (due to stress or lack of nutrition or both). More reproducible staining patterns have been achieved with 1 - 3 mM levamisole for dissection.</li>
	<li>Attach two 25 G needles to two 1 mL syringes (the size of the syringe does not matter, the gauge of the needle is critical). Position one syringe in each hand (Figure 2).</li>
	<li>Under a dissecting microscope, position each needle under and over each worm as shown in Figure 2 and place a fine cut on the worm near the second pharyngeal bulb in a scissor-like motion. After one cut in the worm move on to the next animal until all the animals in the dish have been cut at least once. 
	NOTE: Be mindful that steps 2.8 - 2.9 are time sensitive. Dissect all worms in the dish in under five min for a reproducible dpERK pattern. Longer dissection times will result in a turning off of the dpERK signal, especially in Zone 1. Adjust the number of worms per round of dissection (i.e., 50 worms vs.&#160;150) if necessary to stay in the allotted time frame.
	<ol>
		<li>In case an extruded gonad is not visible during the dissection, do not attempt another cut in the same animal. Usually that will only shear the animal and not result in extrusion of the gonad. Instead, move on to the next animal. Worms where the gonads do not extrude will appear as such on the slides that will be made in section 6 and can be ignored during imaging 
		NOTE: Through all of the dissection and processing steps, it is important to bear in mind that the gonad will remain attached to the body/carcass. Do not force the gonad and the body apart with the needle. Often times an aberrant tear in the gonadal tissue in an attempt to sever the gonad from the body can result in spurious antibody signal. The body/carcass can be ignored during the imaging steps (section 6), and only the gonad imaged. Additionally, sometimes the intestinal cells can also serve as internal controls/somatic controls for the antibody being assayed.</li>
	</ol>
	</li>
</ol>
3. Gonad Fixation
<ol>
	<li>After the dissection is complete, add 2 mL of 3% paraformaldehyde (PFA) directly to the glass watch dish containing the 100 &#956;L of M9 and the dissected animals and cover with Parafilm. Place the covered watch glass dish in a chemical fume hood. 
	Caution: PFA is a hazardous chemical. Thus, the fixation should be performed in the chemical fume hood.</li>
	<li>Incubate at RT for 10 min.</li>
	<li>Using a disposable 9&#34; glass Pasteur pipette add 3 mL of 1x PBS-T (see Materials Table) to the watch glass dish containing the dissected animals. Mix by drawing the PFA solution and the 1x PBS-T with the dissected animals into the Pasteur pipette and releasing gently back into the watch glass dish. Do not vortex the tubes during the washes.</li>
	<li>Using a fresh glass Pasteur pipette draw the 5 mL of liquid (containing dissected worms, PFA and 1x PBS-T) from the watch glass dish into the Pasteur pipette and transfer the contents into a fresh 5 mL glass conical tube.</li>
	<li>Centrifuge the conical tubes for 30 sec in a clinical centrifuge at 1,000 x g. Use glass Pasteur pipettes and conical tubes as dissected gonads stick to plastic.</li>
	<li>Remove and discard the supernatant taking care so as not to disturb the dissected animals. Remove the liquid after each wash from the conical tube under a dissecting microscope so as not to affect the dissected gonads, and minimize their loss.</li>
	<li>Using a fresh Pasteur pipette, add 5 mL of 1x PBS-T to the conical tubes. Centrifuge the conical tubes in a clinical centrifuge for 30 s at 1,000 x g. Remove and discard the supernatant taking care so as not to disturb the dissected animals.</li>
	<li>Repeat step 3.7 for a total of three times.</li>
	<li>Using a fresh Pasteur pipette, add 2 mL of 100% methanol to the tubes, and gently mix the gonads with the methanol by drawing up into the Pasteur pipette. Incubate the tube at -20 &#176;C for a minimum of 1 h. 
	NOTE: Dissected gonads can be stored in methanol for up to 2 days for dpERK staining. Storage for more than 2 days and up to 2 weeks is compatible with other antibody staining applications, such as anti-Lamin antibody, but not with the dpERK antibody.</li>
	<li>After the desired incubation time, repeat step 3.7 for a total of three times in order to wash the gonads. Do not vortex the tubes during the washes. After the final wash leave 500 &#956;L of 1x PBS-T in the 5 mL conical tube.</li>
	<li>Using a fresh Pasteur pipette transfer the contents of the conical glass tube into a fresh 1 mL glass tube (6 mm x 50 mm). Allow the dissected animals to settle by gravity for 5 - 10 min.</li>
	<li>Using a fresh Pasteur pipette and under a dissecting microscope, remove as much of the wash as possible from the 1 mL glass tubes without disturbing the dissected gonads.</li>
</ol>
4. Blocking and Primary Antibody Treatment
<ol>
	<li>Add 100 &#181;L of 30% Normal Goat Serum (NGS) diluted in 1x PBS-T (see Materials Table) to the dissected animals in the 1 mL glass tubes. 30% NGS serves as the blocking buffer. Cover the tube with Parafilm to avoid evaporation of the liquid. Incubate at RT for 1 h or at 4 &#176;C overnight.</li>
	<li>After the desired time of incubation, remove the blocking buffer using a fresh Pasteur pipette, removing as much liquid as possible. Use a dissecting microscope to ensure that gonads are not drawn up in the Pasteur pipette.</li>
	<li>Dilute the MAPKYT antibody (see&#160;Materials Table, referred to in this method as dpERK) at 1:400 in 30% NGS. Prepare enough antibody dilution to use 100 &#181;L per tube. Unused, diluted antibody can be stored at 4 &#176;C for a week.</li>
	<li>Add 100 &#181;L of the diluted anti-dpERK antibody solution to the 1 mL glass tubes containing the dissected animals. Seal the tubes with Parafilm. Incubate the tubes at 4 &#176;C overnight.</li>
	<li>After overnight incubation, add 800 &#181;L of 1x PBST to the tubes at RT.</li>
	<li>Draw the sample up in a fresh glass Pasteur pipette and release gently to ensure that the gonads are evenly suspended in the 1x PBS-T. Let the gonads settle to the bottom with gravity. Remove the supernatant. This ensures that the dissected gonads are washed but not damaged during the process.</li>
	<li>Repeat steps 4.5 - 4.6 for a total of three washes. Do not vortex the tubes during the washes. After the third wash be sure to remove and discard as much supernatant as possible without disturbing the dissected animals. Ensure that a fresh Pasteur pipette is used each time.</li>
</ol>
5. Secondary Antibody Treatment
<ol>
	<li>During the primary antibody washes prepare the dilution for the secondary antibody. Dilute the anti-mouse secondary antibody at 1:500 in 30% NGS. As with the primary antibody, use 100 &#181;L per tube. Unused antibody can be stored at 4 &#176;C for a week.</li>
	<li>Add 100 &#181;L secondary antibody solution to the 1 mL glass tubes containing the dissected animals. Cover each tube with Parafilm, and then wrap the tubes in aluminum foil to keep the contents of the tube in dark. Incubate the tubes at RT for 2 h, or alternatively at 4 &#176;C overnight.</li>
	<li>After the desired time of incubation, add 800 &#181;L of 1x PBS-T to the tubes and repeat steps 4.5 - 4.6 for a total of three times. After the final wash, remove and discard as much of the supernatant as possible under a dissecting microscope using a fresh Pasteur pipette.</li>
	<li>Prepare the DAPI solution during the washes for the secondary antibody. Dilute 1 &#181;L of DAPI (from a 1,000x stock of 1 mg/mL stored at -20 &#176;C) in- 1 mL of 1x PBST.</li>
	<li>Add 800 &#181;L of the diluted DAPI solution to the tubes containing the dissected animals. Cover the mouth of the tube with Parafilm, and wrap each tube in aluminum foil. Incubate at RT&#160;for 20 min.</li>
	<li>After the incubation with DAPI, remove as much of the DAPI solution as possible with a fresh Pasteur pipette, under a dissecting microscope so as not to disturb the dissected animals.</li>
	<li>Add 1 drop of mounting solution to the tubes. Wrap each tube in aluminum foil. 
	NOTE: For visualizing dpERK staining, stained gonads should be mounted for microscopy immediately. For other antibody applications such as phosphorylated serine 10 Histone H3, gonads can be kept in mounting media for up to 2 weeks at 4 &#176;C in the dark.</li>
</ol>
6. Assembling Slides and Imaging
<ol>
	<li>Place a layer of lab tape lengthwise, on top, of two glass microscope slides (25 mm x 75 mm x 1 mm) as seen in Figure 3. Place a fresh clean glass microscope slide in between the two taped slides. 
	NOTE: The thickness of the tape helps generate an agarose pad on the middle slide. Thickness of the agarose pad is almost equal to that of the tape, as the tape serves as a spacer for creating the agarose pad.</li>
	<li>Drop ~ 200 &#181;L of melted 2% agarose (made in distilled water) on the microscope slide in the middle. Quickly place a fresh clean slide, perpendicular to and on top, of the agarose.&#160;</li>
	<li>Let the agarose solidify (1 - 2 min).</li>
	<li>Remove the top microscope slide very gently, such that the agarose pad is not destroyed. The agarose pad can remain attached either to the top or the bottom slide. It does not matter which slide the agarose pad remains attached to, as long as it is a smooth surface of agarose.</li>
	<li>Using a Pasteur pipette transfer the dissected animals onto the agarose pad. Remove any excess liquid from the slide using a Pasteur pipette under a dissecting microscope.</li>
	<li>Using an eyelash hair or finely drawn capillary, push the gonads and intestines away from one another in order to spread the gonads out on the slide.</li>
	<li>Cover the microscope slide with a 24 mm x 50 mm size coverslip. Take care to ensure that the coverslip is lowered gently onto the slide with minimal air bubbles being formed between the coverslip and the slide.</li>
	<li>Store at 4 &#176;C overnight in a slide box (an opaque slide box with the slides lying flat, coverslip up) to allow excess liquid to evaporate and gonads to become somewhat flattened (due to the weight of the coverslip on the gonads). The slight flattening of the gonads allows for better imaging of an otherwise round gonadal tube.
	<ol>
		<li>Once the coverslip is mounted, do not to apply pressure to the top of the coverslip with fingers. Often this results in distortion of gonads and loss of the dpERK signal.</li>
		<li>Flatten the gonads overnight&#160;only for more effective imaging. The slides are ready to be viewed as soon as they are assembled. To take images immediately, gently absorb the excess liquid between the coverslip and the slide from the corners using a clean lab tissue.</li>
	</ol>
	</li>
	<li>After the overnight&#160;period, seal the slide and coverslip with transparent nail polish topcoat on the four corners of the coverslip and the slide boundary. The nail polish/coverslip seal ensures that the coverslip does not move while imaging and protects against destruction of the samples.</li>
	<li>Image the gonads with a compound microscope at 63X. However the objective lens and microscope can be varied based on the availability of microscopes and user application. Because the size of the entire gonad from the proliferative region to the oocytes is larger than can be captured in one image, the images are taken as a montage with overlapping boundaries as shown in Figure 4.11-14, 18.
	<ol>
		<li>In many cases, edit the carcass after the final imaging and assembly, as long as no major alterations are made to the germline image. Store the raw images along the assembled image to enable comparison of the &#39;before&#39; assembly/editing and &#39;after&#39; assembly/ editing of the images. 
		NOTE: The microscopy images must not be altered for signal strength or saturation during the assembly of the montage.</li>
	</ol>
	</li>
</ol>

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The authors declared that no competing interests exist.

Active ERK (dpERK) follows a stereotypical spatial and dynamic localization pattern in the C. elegans germline. This stereotyped dpERK spatial pattern (Figure 5), and amplitude in an adult C. elegans gonad, can be effectively correlated with the many biological processes that ERK regulates. For example, an inability to activate ERK in Zone 2 results in an arrest in oocytes in prophase of meiosis, a phenotype observed in female germlines that do not specify sperm, and thus do not activate ERK in mature oocytes (Figure 5). Mating with males&#39; results in effective accumulation of dpERK in Zone 2 in the female germlines11,13, coupled with onset of oocyte maturation and ovulation. Thus analysis of spatial patterns and temporal activation of dpERK upon certain genetic perturbations (such as RNA interference mediated gene inactivation) or chemical treatments allows for identification of factors that regulate ERK signaling and thus ERK-dependent biological processes. Such analyses also enable discovery of novel genetic interactions between signaling pathways.
A critical bottleneck for any imaging based analysis is the availability of functional reagents such as antibodies that are specific to the application. If such a reagent exists then methods such as the one described above can be easily adapted for the analysis of localization pattern for any protein of interest. The application is thus limited by the availability of a functioning antibody. Additionally, because the method describes dissection and staining at a given developmental time, it only enables the capturing of information in one time frame, and does not shed light on the dynamics or protein regulation in real time or rate of protein turnover.
The method described above is adapted from Francis et al.23, which is distinct from the Seydoux and Dunn method24 for gonad extrusion and antibody staining. The method based on Francis et al. will be called the &#34;suspension method&#34; and the Seydoux and Dunn method called the &#34;freeze cracking method&#34; for comparison. The suspension method has been very effective in our hands for obtaining reproducible localization patterns of signaling molecules such as dpERK, that are inherently more sensitive to harsh handling conditions. In the case of dpERK localization, in our experience, the signal in Zone 1 is virtually undetectable with the freeze cracking method. Additionally, sample drying, which can often occur with the freeze cracking method, results in spurious antibody signals. The suspension method minimizes sample drying, since the gonads are suspended in liquid throughout the process. We also find that the suspension method is useful for imaging multiple different genotypes on a single slide. For example, if two genotypes, such as hermaphrodites vs females are to be directly compared for dpERK localization, the two genotypes can be mixed together and processed. This is possible because the phenotypes are distinct and can be distinguished via DAPI morphologies. Staining in the same tube followed by imaging on the same slide allows for direct comparison between the gonads from different genotypes. Alternatively, if the DAPI morphologies are not distinguishable, then a two-step antibody staining can be adopted. First each individual genotype is treated with two distinct antibodies, Antibody A for WT and Antibody B for mutant (both antibodies need to be raised in a host distinct from the dpERK antibody). Once the two genotypes have been stained with the primary antibody, and washed, they can be mixed together in one tube and now stained with the dpERK antibody, followed by a secondary antibody treatment to visualize all the antibodies in the tube. An ability to compare different genotypes on a single slide, especially when deductions of activation patterns are being made ensures accurate interpretations not affected by distinct staining conditions.
While advantageous, there are multiple steps throughout the suspension method that require careful manipulation. It is critical that the dissections occur in a time efficient manner; importantly, the dissections should not take over 5 min. Longer dissection times result in variable dpERK signal, which can often be misinterpreted as regulated changes in ERK activation, rather than as technical failures. It is crucial to start with over 100 - 150 animals for each dissection because throughout the various wash steps the gonads can easily be lost from the tubes due to pipetting errors. Loss of gonads can be reduced either by dissecting in multiple small batches (such as three batches of 50 animals each) that can be combined, or by dissecting one large batch of 150. Another challenge is getting the gonads to lie in a well-spaced formation on the slide so that the entire distal gonad can be imaged. To enable this, use an eyelash on a matchstick or a pulled pipette to space out the gonads. However, care should be taken so as not to damage the gonads during this process. Often with these techniques it takes multiple attempts to master the dissections as well as arrangements of the gonads onto the slides.
Once this technique has been mastered, it serves as a powerful method for varied biological analyses, and can be used to study more than just dpERK levels. For example this method can be used to visualize active p38 levels, or active CDK levels, or any protein for which an antibody reagent exists. Additionally, the method can be coupled with a chemical inhibition screen in whole animals to assay for novel inhibitors or activators of the pathway, via assaying for dpERK levels. This will allow for an in vivo readout of both reaction and sensitivity to any inhibitors that may interact with the RTK-RAS-ERK signaling pathway. Moreover, this method can be used in antibody combinations with multiple signaling outputs that can all be used and visualized at one time. Using multiple antibodies allows correlation between multiple pathways, their activation status, and outcomes all within the same tissue, enabling better understanding of these interactions. These are just a few examples of further applications of this method, but this system can be modified to study multiple research interests.

The Receptor Tyrosine Kinase (RTK)-RAt Sarcoma (RAS)-Extracellular signal Regulated Kinase (ERK) pathway relays extracellular signals through a conserved kinase cascade that results in the phosphorylation and activation of ERK1-3. ERK proteins are members of the conserved proline-directed serine/threonine MAP (Mitogen Activated Protein) kinase family, and are directly activated by MEK via dual phosphorylation on the threonine (T) and tyrosine (Y) of the conserved TEY motif. Active ERK (referred to as diphosphorylated ERK, or dpERK) then regulates many cellular and developmental processes through its phosphorylation of a battery of downstream substrates1-3. Thus abnormal ERK activity leads to many cell and developmental defects4-7.
Stringent regulation of ERK activity is critical for normal development: in mammalian systems too much ERK activity leads to excessive cellular proliferation leading to oncogenic growth; too little activity leads to cell death4,6. Additionally, changes in the duration of ERK activity can also lead to distinct outcomes: in PC12 cells, ERK activation for 30 min or less induces cell proliferation, but ERK activation for 60 min or more induces neuronal differentiation8,9. Tight regulation of ERK activity is thus clearly essential for normal development and homeostasis.
C. elegans is a powerful, and genetically malleable model system to dissect the function and regulation of the RTK-RAS-ERK signaling pathway3,10-15. Relative to mammalian systems, which contain multiple genes for RAS and ERK, C. elegans contains one RAS gene (let-60) and one ERK gene (mpk-1), rendering it a genetically more facile system in which to dissect the function of this pathway3,10-14. The C. elegans germline is essentially a tube that consists of mitotic stem cells at its distal end and mature oocytes at its proximal end (Figure 1)16. Germ cells initiate meiosis just proximal to the distal mitotic zone, and progress through an extended meiotic prophase (pachytene), after which they begin to form oocytes in the loop region, finally undergoing oocyte maturation in the proximal region16.
Genetic studies from multiple labs, including our own, have shown that the RTK-RAS-ERK pathway is essential for germline development in C. elegans12,13,15,17-19. Specifically, we found that ERK controls and coordinates at least nine distinct biological processes during germline development, ranging from developmental switches such as germ cell apoptosis to cell biological processes such as oocyte growth11,18. Much like in mammalian systems, too much ERK activity in the C. elegans germline results in production of multiple small oocytes, while too little activity results in one large oocyte11. Thus tight regulation of dpERK is essential for normal germline development. The active form of ERK, as visualized by an antibody specific to dpERK, displays a stereotypical, dynamic, bimodal localization pattern: dpERK is high during mid-pachytene (Zone 1, Figure 1), low in the loop region and high again in mature oocytes (Zone 2, Figure 1). Recently, we found that nutrition acts through the Insulin-like Growth Factor receptor-1 (daf-2) to activate ERK in Zone 112; prior work showed that the sperm signal (via the Ephrin receptor tyrosine kinase) activates ERK in Zone 213.
Given that active ERK functions as a rheostat in the germline to regulate oocyte growth,&#160;spatial and temporal localization, as well as amplitude of dpERK, is key to understanding its normal signaling outcome. Using the method described here, changes in the stereotypical localization of dpERK can be easily monitored and predictions derived on the impact of the environmental or genetic perturbations on ERK activity and thus function. Thus, assaying for dpERK enables a comprehensive understanding of its role during germline development.

<table><tbody><tr><td>Agarose</td><td>Sigma Inc.</td><td>A9539</td><td>Dissolve 2 g in 100 mL to make the 2% solution</td></tr><tr><td>Flat bottom glass watch dish</td><td>Agar Scientific</td><td>AGL4161</td><td>We use glass because dissected gonads often stick to plastic</td></tr><tr><td>25 G needles</td><td>BD PrecisionGlide</td><td>305122</td><td></td></tr><tr><td>Syringes (could be 1 or 5 mL)</td><td>BD Syringes</td><td>1 mL: BD 309659</td><td></td></tr><tr><td>Microscope slides (25 x 75 x 1.0 mm)</td><td>Fisherbrand</td><td>12-550-343</td><td></td></tr><tr><td>Coverslips (24 x 50 mm)</td><td>Corning</td><td>2935-245</td><td></td></tr><tr><td>5 mL glass conical tube</td><td>Corning</td><td>8060-5</td><td></td></tr><tr><td>9&quot; disposable Pasteur pipette</td><td>Fisherbrand</td><td>13-678-20D</td><td></td></tr><tr><td>Clinical bench top centrifuge</td><td></td><td></td><td></td></tr><tr><td>Glass&amp;nbsp; tubes (6 x 50 mm)</td><td>Fisherbrand</td><td>14-958-A</td><td></td></tr><tr><td>*M9 solution</td><td></td><td></td><td></td></tr><tr><td>Levamisole</td><td>Sigma</td><td>L9756</td><td></td></tr><tr><td>3% Paraformaldehyde (PFA)</td><td>Electron microscopy services</td><td>17500</td><td>Obtained as 16% solution in ampoules, and diluted to 3% in Potassium Phosphate Buffer</td></tr><tr><td>1x PBS-T</td><td>1x PBS with 0.1% Tween 20</td><td></td><td></td></tr><tr><td>Methanol</td><td>Electron microscopy services</td><td>18510</td><td></td></tr><tr><td>Normal goat serum (NGS)</td><td>Cell Signaling</td><td>5425</td><td>Diluted to 30% in 1x PBS-T</td></tr><tr><td>MAPKYT (dpERK) antibody&amp;nbsp;</td><td>Sigma Inc.</td><td>M9692</td><td>Dilution 1:400 in 30% NGS</td></tr><tr><td>Secondary antibody</td><td>Invitrogen</td><td>A-11005</td><td>Dilution 1:500 in 30% NGS</td></tr><tr><td>DAPI</td><td>Sigma</td><td>D9542</td><td></td></tr><tr><td>Vectashield</td><td>Vector Labs</td><td>H-1000</td><td></td></tr><tr><td>*M9 Buffer Recipe</td><td></td><td></td><td>3 g KH&lt;sub&gt;2&lt;/sub&gt;PO&lt;sub&gt;4&lt;/sub&gt;, 6 g Na&lt;sub&gt;2&lt;/sub&gt;HPO&lt;sub&gt;4&lt;/sub&gt;, 5 g NaCl, 1 mL 1 M MgSO&lt;sub&gt;4&lt;/sub&gt;, H&lt;sub&gt;2&lt;/sub&gt;O to 1 L.&amp;nbsp;</td></tr><tr><td>**PBS (1x)</td><td></td><td></td><td>8 g NaCl, 0.2 g KCl, 1.44 g Na&lt;sub&gt;2&lt;/sub&gt;HPO&lt;sub&gt;4&lt;/sub&gt;, 0.24 g KH&lt;sub&gt;2&lt;/sub&gt;PO&lt;sub&gt;4&lt;/sub&gt;, 0.133 g CaCl&lt;sub&gt;2&lt;/sub&gt;, 0.10 g MgCl&lt;sub&gt;2&lt;/sub&gt;, H&lt;sub&gt;2&lt;/sub&gt;O to 1 L</td></tr><tr><td>Nematode Growth Medium (NGM)</td><td></td><td></td><td>3 g NaCl, 17 g Agar, 2.5 g Peptone, 975 mL of water in 2 L Erlenmayer Flask. Autoclave for 50 min. Cool, and add 1 mL of 1 M CaCl&lt;sub&gt;2&lt;/sub&gt;, 1 mL of 5 mg/mL cholesterol, 1 mL of 1 M MgSO&lt;sub&gt;4&lt;/sub&gt; and 25 mL of 1 M KPO&lt;sub&gt;4&lt;/sub&gt;.&amp;nbsp;</td></tr></tbody></table>

spatial and temporal analysis of active erk in the c. elegans germline

The evolutionarily conserved&#160;extracellular signal transducing RTK-RAS-ERK pathway is an important kinase-signaling cascade that controls multiple cellular and developmental processes principally via activation of ERK, the terminal kinase of the pathway. Tight regulation of ERK activity is essential for normal development and homeostasis; overly active ERK results in excessive cellular proliferation, while underactive ERK causes cell death. C. elegans is a powerful model system that has helped characterize the function and regulation of RTK-RAS-ERK signaling pathway during development. In particular, the RTK-RAS-ERK pathway is essential for C. elegans germline development, which is the focus of this method. Using antibodies specific to the active, diphosphorylated form of ERK (dpERK), the stereotypical localization pattern can be visualized within the germline. Because this pattern is both spatially and temporally controlled, the ability to reproducibly assay dpERK is useful to identify regulators of the pathway that affect dpERK signal duration and amplitude and thus germline development. Here we demonstrate how to successfully dissect, stain, and image dpERK within the C. elegans gonad. This method can be adapted for spatial localization of any signaling or structural protein in the C. elegans gonad, provided an antibody compatible with immunofluorescence is available.

In WT adult hermaphroditic animals, dpERK is typically visualized in mid-pachytene region, Zone 1, and in the most mature oocytes from -1 through -4 or -5 (Zone 2). Perturbations in this activation pattern reflect changes to the signaling pathway. Female germlines do not specify sperm, and thus do not display activation of ERK in Zone 2, with only weak activation in Zone 1. These are represented in Figure 5.
<img alt="Figure 1" src="/files/ftp_upload/54901/54901fig1.jpg" /> 
Figure 1: C. elegans&#160;germline Morphology. Differential Interference contrast (DIC) image of an adult hermaphrodite with one U-shaped gonad arm outlined with the white line. The adult hermaphroditic gonad contains mitotic cells at the distal tip. The mitotic cells enter meiosis and progress through meiotic prophase (pachytene) developing into mature oocytes in the proximal gonad. Zone 1 and Zone 2 mark stereotypical localization of dpERK in an adult hermaphroditic gonad. Scale: 20 &#181;m. <a href="https://cloudflare-jove-com.bdigitaluss.remotexs.co/files/ftp_upload/54901/54901fig1large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 2" src="/files/ftp_upload/54901/54901fig2.jpg" /> 
Figure 2: Demonstration of Needle Positions during dissections. Left: Photograph of a dissection in process. Right: Needle positions with reference to the worm. <a href="https://cloudflare-jove-com.bdigitaluss.remotexs.co/files/ftp_upload/54901/54901fig2large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 3" src="/files/ftp_upload/54901/54901fig3.jpg" /> 
Figure 3: Demonstration of Agarose Slide Preparation. (A) Place a tape lengthwise across 2 clean microscope slides. Place a fresh clean microscope slide between the two slides with the tape as shown. The three slides are next to each other lengthwise. (B) Add melted agarose to the slide in the center. (C) Place a fresh microscope slide perpendicular to, and on top of, the middle slide, carrying the drop of agarose. (D) Allow the agarose to solidify. (E) Remove the top slide leaving behind the solidified agarose pad. Use the bottom slide with the agarose pad for mounting dissected and stained germlines. <a href="https://cloudflare-jove-com.bdigitaluss.remotexs.co/files/ftp_upload/54901/54901fig3large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 4" src="/files/ftp_upload/54901/54901fig4.jpg" /> 
Figure 4: Imaging the Slides as a Montage. Montage of a wild type germline with DAPI (top) and dpERK (bottom) channel. Images taken at 63X with overlapping boundaries, white boxes for panel A and B and green boxes for panel B and C. The DAPI and dpERK images for each panel were taken simultaneously in two different channels. The assembled image is shown in Figure 5A. Scale bar: 20 &#956;m. <a href="https://cloudflare-jove-com.bdigitaluss.remotexs.co/files/ftp_upload/54901/54901fig4large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 5" src="/files/ftp_upload/54901/54901fig5.jpg" /> 
Figure 5: Dynamic Temporal/Spatial Activation of ERK. (A-C) WT&#160;(N2) adult (24 hours past mid-L4 stage of development) hermaphrodite germlines stained for DNA (B, DAPI, white) and dpERK (C, red). The dpERK signal in Zone 1, the pachytene region, and Zone 2, the mature oocytes is highlighted. (D-F) fog-2(oz40) female germlines (at 8 h&#160;past mid-L4 stage of development) stained for DNA (E, DAPI) and dpERK (F). Young female germlines display weak dpERK in Zone 1, and in a single oocyte in a sperm independent manner. Zone 2 is sperm dependent, and thus absent in the female germline. Scale bar: 20 &#956;m. <a href="https://cloudflare-jove-com.bdigitaluss.remotexs.co/files/ftp_upload/54901/54901fig5large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

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Spatial and Temporal Analysis of Active ERK in the C. elegans Germline | Developmental Biology | Video

Spatial and Temporal Analysis of Active ERK in the C. elegans Germline

We present an immunofluorescence&#160;imaging-based method for spatial and temporal localization of active ERK in the dissected C. elegans gonad. The protocol described here can be adapted for visualization of any signaling or structural protein in the C. elegans gonad, provided a suitable antibody reagent is available.

Spatial and Temporal Analysis of Active ERK in the C. elegans Germline

The evolutionarily conserved&#160;extracellular signal transducing RTK-RAS-ERK pathway is an important kinase-signaling cascade that controls multiple ...

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10.1K Views.  Baylor College of Medicine. We present an immunofluorescence imaging-based method for spatial and temporal localization of active ERK in the dissected C. elegans gonad. The protocol described here can be adapted for visualization of any signaling or structural protein in the C. elegans gonad, provided a suitable antibody reagent is available.

Video: Spatial and Temporal Analysis of Active ERK in the C. elegans Germline

Fourier-Based Diffraction Analysis of Live Caenorhabditis elegans

This manuscript describes how to classify nematodes using temporal far-field diffraction signatures. A single C. elegans is suspended in a water column inside an optical cuvette. A 632 nm continuous wave HeNe laser is directed through the cuvette using front surface mirrors. A significant distance of at least 20-30 cm traveled after the light passes through the cuvette ensures a useful far-field (Fraunhofer) diffraction pattern. The diffraction pattern changes in real time as the nematode swims within the laser beam. The photodiode is placed off-center in the diffraction pattern. The voltage signal from the photodiode is observed in real time and recorded using a digital oscilloscope. This process is repeated for 139 wild type and 108 &#34;roller&#34; C. elegans. Wild type worms exhibit a rapid oscillation pattern in solution. The &#34;roller&#34; worms have a mutation in a key component of the cuticle that interferes with smooth locomotion. Time intervals that are not free of saturation and inactivity are discarded. It is practical to divide each average by its maximum to compare relative intensities. The signal for each worm is Fourier transformed so that the frequency pattern for each worm emerges. The signal for each type of worm is averaged. The averaged Fourier spectra for the wild type and the &#34;roller&#34; C. elegans are distinctly different and reveal that the dynamic worm shapes of the two different worm strains can be distinguished using Fourier analysis. The Fourier spectra of each worm strain match an approximate model using two different binary worm shapes that correspond to locomotory moments. The envelope of the averaged frequency distribution for actual and modeled worms confirms the model matches the data. This method can serve as a baseline for Fourier analysis for many microscopic species, as every microorganism will have its unique Fourier spectrum.

Fourier-Based Diffraction Analysis of Live Caenorhabditis elegans

This manuscript describes how to distinguish different nematodes using far-field diffraction signatures. We compare the locomotion of 139 wild type and 108 &#34;Roller&#34; C. elegans by averaging frequencies associated with the temporal Fraunhofer diffraction signature at a single location using a continuous wave laser.

This manuscript describes how to classify nematodes using temporal far-field diffraction signatures. A single C. elegans is suspended in a water column ...

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Computational Analysis of the Caenorhabditis elegans Germline to Study the Distribution of Nuclei, Proteins, and the Cytoskeleton

The Caenorhabditis elegans (C. elegans) germline is used to study several biologically important processes including stem cell development, apoptosis, and chromosome dynamics. While the germline is an excellent model, the analysis is often two dimensional due to the time and labor required for three-dimensional analysis. Major readouts in such studies are the number/position of nuclei and protein distribution within the germline. Here, we present a method to perform automated analysis of the germline using confocal microscopy and computational approaches to determine the number and position of nuclei in each region of the germline. Our method also analyzes germline protein distribution that enables the three-dimensional examination of protein expression in different genetic backgrounds. Further, our study shows variations in cytoskeletal architecture in distinct regions of the germline that may accommodate specific spatial developmental requirements. Finally, our method enables automated counting of the sperm in the spermatheca of each germline. Taken together, our method enables rapid and reproducible phenotypic analysis of the C. elegans germline.

Computational Analysis of the Caenorhabditis elegans Germline to Study the Distribution of Nuclei, Proteins, and the Cytoskeleton

We present an automated method for three-dimensional reconstruction of the Caenorhabditis elegans germline. Our method determines the number and position of each nucleus within the germline and analyses germline protein distribution and cytoskeletal structure.

The Caenorhabditis elegans (C. elegans) germline is used to study several biologically important processes including stem cell development, apoptosis, and ...

Epigenetic Inheritance and ChIP in Caenorhabditis elegans

Analysis of Transgenerational Epigenetic Inheritance in C. elegans Using a Fluorescent Reporter and Chromatin Immunoprecipitation (ChIP)

Transgenerational epigenetic inheritance (TEI) allows the transmission of information through the germline without changing the genome sequence, through factors such as non-coding RNAs and chromatin modifications. The phenomenon of RNA interference (RNAi) inheritance in the nematode Caenorhabditis elegans is an effective model to investigate TEI that takes advantage of this model organism's short life cycle, self-propagation, and transparency. In RNAi inheritance, exposure of animals to RNAi leads to gene silencing and altered chromatin signatures at the target locus that persist for multiple generations in the absence of the initial trigger. This protocol describes the analysis of RNAi inheritance in C. elegans using a germline-expressed nuclear green fluorescent protein (GFP) reporter. Reporter silencing is initiated by feeding the animals bacteria expressing double-stranded RNA targeting GFP. At each generation, animals are passaged to maintain synchronized development, and reporter gene silencing is determined by microscopy. At select generations, populations are collected and processed for chromatin immunoprecipitation (ChIP)-quantitative polymerase chain reaction (qPCR) to measure histone modification enrichment at the GFP reporter locus. This protocol for studying RNAi inheritance can be easily modified and combined with other analyses to further investigate TEI factors in small RNA and chromatin pathways.

Author Spotlight: RNAi Inheritance and ChIP in C. elegans 

This protocol describes an RNA interference and ChIP assay to study the epigenetic inheritance of RNAi-induced silencing and associated chromatin modifications in C. elegans.

Transgenerational epigenetic inheritance (TEI) allows the transmission of information through the germline without changing the genome sequence, through ...

Cell Cycle Analysis in the C. elegans Germline with the Thymidine Analog EdU

Cell cycle analysis in eukaryotes frequently utilizes chromosome morphology, expression and/or localization of gene products required for various phases of the cell cycle, or the incorporation of nucleoside analogs. During S-phase, DNA polymerases incorporate thymidine analogs such as EdU or BrdU into chromosomal DNA, marking the cells for analysis. For C. elegans, the nucleoside analog EdU is fed to the worms during regular culture and is compatible with immunofluorescent techniques. The germline of C. elegans is a powerful model system for the studies of signaling pathways, stem cells, meiosis, and cell cycle because it is transparent, genetically facile, and meiotic prophase and cellular differentiation/gametogenesis occur in a linear assembly-like fashion. These features make EdU a great tool to study dynamic aspects of mitotically cycling cells and germline development. This protocol describes how to successfully prepare EdU bacteria, feed them to wild-type C. elegans hermaphrodites, dissect the hermaphrodite gonad, stain for EdU incorporation into DNA, stain with antibodies to detect various cell cycle and developmental markers, image the gonad and analyze the results. The protocol describes the variations in the method and analysis for the measurement of S-phase index, M-phase index, G2 duration, cell cycle duration, rate of meiotic entry, and rate of meiotic prophase progression. This method can be adapted to study the cell cycle or cell history in other tissues, stages, genetic backgrounds, and physiological conditions.

Cell Cycle Analysis in the C. elegans Germline with the Thymidine Analog EdU

An imaging-based method is described that can be used to identify S-phase and analyze cell cycle dynamics in the C. elegans hermaphrodite germline using the thymidine analog EdU. This method requires no transgenes and is compatible with immunofluorescent staining.

Cell cycle analysis in eukaryotes frequently utilizes chromosome morphology, expression and/or localization of gene products required for various phases ...

Isotropic Light-Sheet Microscopy and Automated Cell Lineage Analyses to Catalogue Caenorhabditis elegans Embryogenesis with Subcellular Resolution

Caenorhabditis elegans (C. elegans) stands out as the only organism in which the challenge of understanding the cellular origins of an entire nervous system can be observed, with single cell resolution, in vivo. Here, we present an integrated protocol for the examination of neurodevelopment in C. elegans embryos. Our protocol combines imaging, lineaging and neuroanatomical tracing of single cells in developing embryos. We achieve long-term, four-dimensional (4D) imaging of living C. elegans&#160;embryos with nearly isotropic spatial resolution through the use of Dual-view Inverted Selective Plane Illumination Microscopy (diSPIM). Nuclei and neuronal structures in the nematode embryos are imaged and isotropically fused to yield images with resolution of ~330 nm in all three dimensions. These minute-by-minute high-resolution 4D data sets are then analyzed to correlate definitive cell-lineage identities with gene expression and morphological dynamics at single-cell and subcellular levels of detail. Our protocol is structured to enable modular implementation of each of the described steps and enhance studies on embryogenesis, gene expression, or neurodevelopment.

Here, we present a combinatorial approach using high-resolution microscopy, computational tools, and single-cell labeling in living C. elegans embryos to understand single cell dynamics during neurodevelopment.

Caenorhabditis elegans (C. elegans) stands out as the only organism in which the challenge of understanding the cellular origins of an entire nervous ...

A Semi-high-throughput Imaging Method and Data Visualization Toolkit to Analyze C. elegans Embryonic Development

C. elegans is the premier system for the systematic analysis of cell fate specification and morphogenetic events during embryonic development. One challenge is that embryogenesis dynamically unfolds over a period of about 13 h; this half day-long timescale has constrained the scope of experiments by limiting the number of embryos that can be imaged. Here, we describe a semi-high-throughput protocol that allows for the simultaneous 3D time-lapse imaging of development in 80&#8211;100 embryos at moderate time resolution, from up to 14 different conditions, in a single overnight run. The protocol is straightforward and can be implemented by any laboratory with access to a microscope with point visiting capacity. The utility of this protocol is demonstrated by using it to image two custom-built strains expressing fluorescent markers optimized to visualize key aspects of germ-layer specification and morphogenesis. To analyze the data, a custom program that crops individual embryos out of a broader field of view in all channels, z-steps, and timepoints and saves the sequences for each embryo into a separate tiff stack was built. The program, which includes a user-friendly graphical user interface (GUI), streamlines data processing by isolating, pre-processing, and uniformly orienting individual embryos in preparation for visualization or automated analysis. Also supplied is an ImageJ macro that compiles individual embryo data into a multi-panel file that displays maximum intensity fluorescence projection and brightfield images for each embryo at each time point. The protocols and tools described herein were validated by using them to characterize embryonic development following knock-down of 40 previously described developmental genes; this analysis visualized previously annotated developmental phenotypes and revealed new ones. In summary, this work details a semi-high-throughput imaging method coupled with a cropping program and ImageJ visualization tool that, when combined with strains expressing informative fluorescent markers, greatly accelerates experiments to analyze embryonic development.

A Semi-high-throughput Imaging Method and Data Visualization Toolkit to Analyze C. elegans Embryonic Development

This work describes a semi-high-throughput protocol that allows simultaneous 3D time-lapse imaging of embryogenesis in 80&#8211;100 C. elegans embryos in a single overnight run. Additionally, image processing and visualization tools are included to streamline data analysis. The combination of these methods with custom reporter strains enables detailed monitoring of embryogenesis.

C. elegans is the premier system for the systematic analysis of cell fate specification and morphogenetic events during embryonic development. One ...

In Situ Detection of Ribonucleoprotein Complex Assembly in the C. elegans Germline using Proximity Ligation Assay

Understanding when and where protein-protein interactions (PPIs) occur is critical to understanding protein function in the cell and how broader processes such as development are affected. The Caenorhabditis elegans germline is a great model system for studying PPIs that are related to the regulation of stem cells, meiosis, and development. There are a variety of well-developed techniques that allow proteins of interest to be tagged for recognition by standard antibodies, making this system advantageous for proximity ligation assay (PLA) reactions. As a result, the PLA is able to show where PPIs occur in a spatial and temporal manner in germlines more effectively than alternative approaches. Described here is a protocol for the application and quantification of this technology to probe PPIs in the C. elegans germline.

In Situ Detection of Ribonucleoprotein Complex Assembly in the C. elegans Germline using Proximity Ligation Assay

This protocol demonstrates use of the proximity ligation assay to probe for protein-protein interactions in situ in the C. elegans germline.

Understanding when and where protein-protein interactions (PPIs) occur is critical to understanding protein function in the cell and how broader processes ...

4D Microscopy: Unraveling Caenorhabditis elegans Embryonic Development Using Nomarski Microscopy

4D microscopy is an invaluable tool for unraveling the embryonic developmental process in different animals. Over the last decades, Caenorhabditis elegans has emerged as one of the best models for studying development. From an optical point of view, its size and transparent body make this nematode an ideal specimen for DIC (Differential Interference Contrast or Nomarski) microscopy. This article illustrates a protocol for growing C. elegans nematodes, preparing and mounting their embryos, performing 4D microscopy and cell lineage tracing. The method is based on multifocal time-lapse records of Nomarski images and analysis with specific software. This technique reveals embryonic developmental dynamics at the cellular level. Any embryonic defect in mutants, such as problems in spindle orientation, cell migration, apoptosis or cell fate specification, can be efficiently detected and scored. Virtually every single cell of the embryo can be followed up to the moment the embryo begins to move. Tracing the complete cell lineage of a C. elegans embryo by 4D DIC microscopy is laborious, but the use of specific software greatly facilitates this task. In addition, this technique is easy to implement in the lab. 4D microscopy is a versatile tool and opens the possibility of performing an unparalleled analysis of embryonic development.

4D Microscopy: Unraveling Caenorhabditis elegans Embryonic Development Using Nomarski Microscopy

Here, we present a protocol for preparing and mounting Caenorhabditis elegans embryos, recording development under a 4D microscope and tracing cell lineage.

4D microscopy is an invaluable tool for unraveling the embryonic developmental process in different animals. Over the last decades, Caenorhabditis elegans ...

An Efficient Protocol to Assess ERK Activity Modulation in Early Zebrafish Noonan Syndrome Models via Live FRET Microscopy and Immunofluorescence

RASopathies are genetic syndromes caused by ERK hyperactivation and resulting in multisystemic diseases that can also lead to cancer predisposition. Despite a broad genetic heterogeneity, germline gain-of-function mutations in key regulators of the RAS-MAPK pathway underlie the majority of the cases, and, thanks to advanced sequencing techniques, potentially pathogenic variants affecting the RAS-MAPK pathway continue to be identified. Functional validation of the pathogenicity of these variants, essential for accurate diagnosis, requires fast and reliable protocols, preferably in vivo. Given the scarcity of effective treatments in early childhood, such protocols, especially if scalable in cost-effective animal models, can be instrumental in offering a preclinical ground for drug repositioning/repurposing. 
Here we describe step-by-step the protocol for rapid generation of transient RASopathy models in zebrafish embryos and direct inspection of live disease-associated ERK activity changes occurring already during gastrulation through real-time multispectral F&#246;rster resonance energy transfer (FRET) imaging. The protocol uses a transgenic ERK reporter recently established and integrated with the hardware of commercial microscopes. We provide an example application for Noonan syndrome (NS) zebrafish models obtained by expression of the Shp2D61G. We describe a straightforward method that enables registration of ERK signal change in the NS fish model before and after pharmacological signal modulation by available low-dose MEK inhibitors. We detail how to generate, retrieve, and assess ratiometric FRET signals from multispectral acquisitions before and after treatment and how to cross-validate the results via classical immunofluorescence on whole embryos at early stages. We then describe how, via examining standard morphometric parameters, to query late changes in embryo shape, indicative of a resulting impairment of gastrulation, in the same embryos whose ERK activity is assessed by live FRET at 6 h post fertilization.

RASopathies are multisystem genetic syndromes caused by RAS-MAPK pathway hyperactivation. Potentially pathogenic variants awaiting validation emerge continuously while poor preclinical evidence limits therapy. Here, we describe our in vivo protocol to test and cross-validate RASopathy-associated ERK activation levels and its pharmacological modulation during embryogenesis by live FRET imaging in Teen-reporter zebrafish.

RASopathies are genetic syndromes caused by ERK hyperactivation and resulting in multisystemic diseases that can also lead to cancer predisposition. ...

Biochemistry

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.

Spatial and Temporal Analysis of Active ERK in the C. elegans Germline

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Chapters in this video

Fourier-Based Diffraction Analysis of Live Caenorhabditis elegans

Computational Analysis of the Caenorhabditis elegans Germline to Study the Distribution of Nuclei, Proteins, and the Cytoskeleton

Analysis of Transgenerational Epigenetic Inheritance in C. elegans Using a Fluorescent Reporter and Chromatin Immunoprecipitation (ChIP)

Cell Cycle Analysis in the C. elegans Germline with the Thymidine Analog EdU

Isotropic Light-Sheet Microscopy and Automated Cell Lineage Analyses to Catalogue Caenorhabditis elegans Embryogenesis with Subcellular Resolution

A Semi-high-throughput Imaging Method and Data Visualization Toolkit to Analyze C. elegans Embryonic Development

In Situ Detection of Ribonucleoprotein Complex Assembly in the C. elegans Germline using Proximity Ligation Assay

4D Microscopy: Unraveling Caenorhabditis elegans Embryonic Development Using Nomarski Microscopy

An Efficient Protocol to Assess ERK Activity Modulation in Early Zebrafish Noonan Syndrome Models via Live FRET Microscopy and Immunofluorescence

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