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

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

1. Maintain C. elegans working stock cultures on Nematode Growth Medium^20,21 (NGM, see Materials Table) agar seeded with E. coli OP50.
2. Maintain worm stocks at temperatures between 16 °C and 25 °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.
3. 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.

2. Dissection of Adult Worms for Obtaining Gonads

1. 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 µL of M9 buffer in a 1.5 mL microcentrifuge tube.
2. Fill the microcentrifuge tube with 900 µL of M9 buffer (see Materials Table). Centrifuge the tubes at 1,000 x g in a microcentrifuge for 1 min at ambient temperature.
3. Gently remove 900 µL of the M9 buffer and discard. Remove the buffer under a dissecting microscope to ensure that the worms are not accidently removed.
4. 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.
5. After the final wash remove 900 µL of M9 buffer from the tube and discard.
6. Transfer the remaining 100 µL of M9 buffer containing the 100 – 150 worms with a 200 µL-micropipette tip to a flat bottom glass watch dish. Ensure that the micropipette tip is cut at the 10 µL mark before use. Not cutting the tip will result in shearing of the worms.
7. Add 1 – 3 µL 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 °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.
8. 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).
9. 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. 150) if necessary to stay in the allotted time frame.
   1. 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.

3. Gonad Fixation

1. After the dissection is complete, add 2 mL of 3% paraformaldehyde (PFA) directly to the glass watch dish containing the 100 μL 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.
2. Incubate at RT for 10 min.
3. Using a disposable 9" 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.
4. 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.
5. 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.
6. 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.
7. 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.
8. Repeat step 3.7 for a total of three times.
9. 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 °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.
10. 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 μL of 1x PBS-T in the 5 mL conical tube.
11. 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.
12. 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.

4. Blocking and Primary Antibody Treatment

1. Add 100 µL 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 °C overnight.
2. 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.
3. Dilute the MAPKYT antibody (see Materials Table, referred to in this method as dpERK) at 1:400 in 30% NGS. Prepare enough antibody dilution to use 100 µL per tube. Unused, diluted antibody can be stored at 4 °C for a week.
4. Add 100 µL 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 °C overnight.
5. After overnight incubation, add 800 µL of 1x PBST to the tubes at RT.
6. 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.
7. 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.

5. Secondary Antibody Treatment

1. 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 µL per tube. Unused antibody can be stored at 4 °C for a week.
2. Add 100 µL 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 °C overnight.
3. After the desired time of incubation, add 800 µL 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.
4. Prepare the DAPI solution during the washes for the secondary antibody. Dilute 1 µL of DAPI (from a 1,000x stock of 1 mg/mL stored at -20 °C) in- 1 mL of 1x PBST.
5. Add 800 µL 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 for 20 min.
6. 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.
7. 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 °C in the dark.

6. Assembling Slides and Imaging

1. 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.
2. Drop ~ 200 µL 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. 
3. Let the agarose solidify (1 – 2 min).
4. 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.
5. 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.
6. 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.
7. 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.
8. Store at 4 °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.
   1. 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.
   2. Flatten the gonads overnight 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.
9. After the overnight 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.
10. 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}.
    1. 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 'before' assembly/editing and 'after' assembly/ editing of the images.
       NOTE: The microscopy images must not be altered for signal strength or saturation during the assembly of the montage.