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

1. Preparation of EdU-labeled Bacteria

1. Grow a starter culture of MG1693. Escherichia coli (E. coli) MG1693 carries a mutation in thyA.
   1. Streak out E. coli MG1693 from a frozen glycerol stock onto a 120 mm lysogeny broth (LB) agar Petri dish. Culture at 37 °C overnight.
   2. Inoculate from two individual E. coli MG1693 colonies into two duplicate 4 mL tubes of liquid LB. Culture at 37 °C for ~16 h.
      NOTE: MG1693 grows fine in LB without supplementing with thymine or thymidine.
2. Grow E. coli MG1693 in minimal media supplemented with EdU.
   1. Autoclave a 500 mL conical flask.
   2. Use sterile technique to add 5 mL of 20% glucose, 50 μL of 10 mg/mL of thiamine, 120 μL of 5 mM thymidine, 100 μL of 1 M MgSO₄, 200 μL of 10 mM EdU, 100 mL of M9 buffer, and 4 mL of freshly-grown overnight MG1693 culture.
      NOTE: The final concentration of EdU is 20 μM in this culture^1,7. This concentration leads to DNA damage and cell cycle arrest if applied directly to mammalian cells in culture²⁰. However, only a fraction of this EdU is incorporated into the E. coli and thus available to C. elegans. There is no evidence of cell cycle arrest and no change in the size of the progenitor zone or M-phase index after EdU treatment of young adult hermaphrodites.
   3. Culture overnight, but no longer than 24 h at 37 °C with shaking at 200 rpm.
3. Concentrate EdU-labeled E. coli and apply to M9 agar Petri dishes.
   1. Pre-cool a tabletop centrifuge to 4 °C.
   2. Use sterile technique to transfer the culture into 2–4 sterile 50 mL conical tubes.
   3. Centrifuge the cultures at 3,000 x g at 4 °C for 30 min to pellet the EdU-labeled E. coli.
      NOTE: Dispose of EdU-containing supernatant according to local and institutional guidelines.
   4. Resuspend the pellets with 4 mL of fresh M9. Use a sterile 1 mL pipet tip or a sterile 5 mL serological pipet. Resuspending the pellets may take several minutes.
   5. Use the same pipet to apply and spread ~8 drops of resuspended EdU-labeled E. coli MG1693 to the center of room-temperature 60 mm M9 agar Petri dishes. One batch yields ~16 dishes.
   6. Allow the dishes to dry for several hours or overnight at room temperature, then seal each dish with a strip of laboratory film. Dishes can be stored at 15 °C for ~2 weeks and at 4 °C for ~2 months. Use the same batch of EdU dishes for each set of experiments.

2. Feeding EdU to C. elegans

1. Synchronize population by timed egg lay, alkaline hypochlorite treatment followed by hatching into S-medium with cholesterol, or by picking the appropriate stage²¹. Grow the animals to desired stage (here, 24 h post-L4) on Nematode Growth Medium seeded with E. coli OP50 at 20 °C.
   NOTE: Prepare 50–100 animals per experiment, as some may not dissect well, and others will be lost in the process of washing and transferring.
2. Allow the EdU dishes to warm to 20 °C (or the temperature required for the experiment).
3. Wash the animals from NGM dishes using M9 or phosphate-buffered saline (PBS) into a 1.5 mL tube. Allow the animals to settle briefly by gravity or a brief spin in a microcentrifuge.
4. Remove the supernatant and wash the animals 1–2 times with ~1 mL of M9 or PBS. Remove the supernatant.
5. Use a glass Pasteur pipette to transfer the animals in a tiny drop of M9 or PBS onto the center of the EdU lawn. Wait a few minutes for the liquid to be absorbed, then incubate at 20 °C for 30 min (for direct S-phase measurement) or longer (to measure history of S-phase), as needed.
   NOTE: EdU signal is detectable in germline nuclei after as little as 15 min of EdU feeding¹.
6. Wash the worms off the EdU dish with ~2 mL of M9 or PBS into a glass dissecting dish.

3. Dissection and Fixation of C. elegans Germline

NOTE: This protocol for the dissection, fixation, and antibody staining of the C. elegans hermaphrodite germline is nearly identical to that published by Gervaise and Arur (2016)²², except that the 1 mL glass tubes can be centrifuged to speed up washing steps and a drawn-out glass Pasteur pipette can be used to remove the liquid from 1 mL glass tubes more effectively.

1. Wash and dissect C. elegans germlines.
   1. Allow the animals to settle to the bottom of the dissecting dish, swirl to collect the animals in the center, and use a long Pasteur pipette to remove PBS. Wash 1–2 times with ~2 mL of PBS.
   2. Add 2 mL of PBS and 4 μL of 100 mM levamisole to immobilize the animals. Swirl the dish again to collect the animals in the center of the dish.
      NOTE: Immobilization can take between a few seconds and a few minutes. Complete immobilization is not necessary for successful dissection. Some people have better success when dissecting incompletely immobilized animals.
   3. Dissect the animals with a pair of 25G 5/8” needles by cutting at the head (approximately between the two pharyngeal bulbs) or the tail. Take care not to cut the loop of the germline. Intestine and germline should “pop out” of the body cavity due to internal pressure, but remain attached. This protocol is similar to previously published Gervaise and Arur (2016)²².
      NOTE: Keep the dissection time to ~5 min, certainly no more than 15 min. Longer dissection times may result in the loss of antibody staining signal (Sudhir Nayak, personal communication) and starvation in PBS may affect the animals’ physiology. Learning to dissect quickly and accurately may take some practice.
   4. If needed, swirl to collect the dissected animals in the center, and use a long Pasteur pipette to remove as much PBS as possible.
2. Fix and dehydrate tissues
   1. Add 2 mL of 3% paraformaldehyde (PFA) in PBS solution. Cover the dish loosely with laboratory film and store on a bench or in a drawer for 10 min.
      NOTE: Thaw PFA solution in a 37 °C water bath and then cool to room temperature prior to dissecting germlines.
      CAUTION: PFA solution is moderately toxic and a probable carcinogen and teratogen. Vapors emitting from paraformaldehyde solutions are flammable. Wear nitrile gloves. Dilute PFA from 16% to 3% in a chemical fume hood. When working outside of fume hood, keep all containers covered.
   2. Transfer the gonads carefully to a clean 5 mL glass centrifuge tube.
   3. Add ~3 mL of PBSTw (PBS with 0.1% Tween-20) to the dish that contained the gonads to help retrieve remaining gonads and to dilute the PFA solution.
   4. Spin down in a clinical centrifuge at 870 x g for ~1 min. Younger or smaller animals require longer spin times than older or larger animals.
   5. Using a long glass pipette, transfer the supernatant to unwanted material beaker for eventual discard in unwanted material bottle in the chemical hood.
   6. Add 2 mL of high-grade methanol pre-chilled to -20 °C. Cover the centrifuge tube tightly with laboratory film.
      NOTE: Use of fresh high-grade “gold label” methanol is essential for proper morphology with certain antibodies.
      CAUTION: Methanol is a highly flammable liquid and vapor, that is toxic if swallowed, inhaled, or allowed to contact skin. Wear gloves and appropriate personal protective equipment. Use a freezer appropriate for small volumes of flammables.
   7. Store in -20 °C freezer for 1 h, even overnight or even several months.
      NOTE: The protocol can be paused here.

4. Rehydrate Germlines

1. Fill the glass centrifuge tube to top with PBSTw to dilute the methanol and rehydrate the gonads. Spin down in a clinical centrifuge at 870 x g for ~1 min.
2. Using a long glass Pasteur pipette, transfer the supernatant to unwanted material beaker for eventual discard in unwanted material bottle in the chemical hood.
3. Wash the gonads 3 times using ~5 mL of PBSTw, spinning down in a clinical centrifuge at 870 x g for ~1 min each time. Remove the supernatant.
4. Rinse a small 1 mL borosilicate glass tube and a long glass Pasteur pipette with PBSTw.
5. Add ~700 μL of PBSTw and use the long glass Pasteur pipette to transfer the gonads to the small tube. Use a few additional drops of PBSTw to ensure that all gonads are transferred.
6. Spin down in a clinical centrifuge at 870 x g for ~1 min. Using a drawn-out long glass Pasteur pipette, remove as much liquid as possible without disturbing the gonads. Leave no more than 50 μL.
   NOTE: If antibody detection is not necessary, skip to Step 6.

5. Detect Antigens with Antibodies

1. Dilute the primary antibodies in 30% serum in PBS. In the present example, anti-WAPL-1 antibody is diluted 1:2,000 and anti-pH3 antibody is diluted 1:500. Centrifuge freshly thawed serum for 10 min in a microfuge at 20,000 x g at 4 °C to remove the particulates. Use the supernatant, which can be stored at 4 °C for several days. Use the appropriate serum to match the host organism of secondary antibodies (goat serum is used here). An optional blocking step may be added prior to the addition of primary antibodies.
2. Apply 100 μL of diluted primary antibody to each small glass tube. Incubate at room temperature for 4 h.
   NOTE: Incubation times vary by antibody. For some antibodies, 2 h at room temperature is sufficient. Longer incubations (e.g., overnight) are possible, but may increase background.
3. Fill the tubes to top with PBSTw and spin down in a clinical centrifuge at 870 x g for ~1 min.
4. Wash the gonads 3 times using ~1 mL of PBSTw. Incubate for ~5 min per wash to allow excess primary antibody to diffuse into wash. Using a drawn-out long glass Pasteur pipette, remove as much liquid as possible without disturbing the gonads. Leave no more than 50 μL.
5. Dilute the secondary antibodies in 30% goat serum in PBS. In the present example, goat-anti-rabbit-594 and goat-anti-mouse-647 are each diluted at 1:400.
   NOTE: Select secondary antibodies carefully to make sure dyes are distinct from the dye in the EdU kit. In the present example, the EdU kit contained a 488 nm excitation dye.
6. Apply 100 μL of diluted secondary antibody to each small glass tube. Incubate in the dark at room temperature 2 h.
   NOTE: Incubation times vary by antibody. For some secondary antibodies, 1 h at room temperature is sufficient. Longer incubations (e.g., overnight) are possible, but may increase background.
7. Wash the gonads 3 times using ~1 mL of PBSTw. Incubate for ~5 min per wash to allow excess secondary antibody to diffuse into wash. Using a drawn-out long glass Pasteur pipette, remove as much liquid as possible without disturbing the gonads. Leave no more than 50 μL.
   NOTE: Gonads can be stored in 100 μL of PBS after this step, if necessary, although this may reduce signal. Remove PBS prior to proceeding.

6. Perform the EdU Click Reaction to Detect EdU

NOTE: Performing the EdU click reaction before the antibody staining steps (perform Step 6 before Step 5) is possible, depending on the antibodies used⁷. However, the click reagents may interfere with certain antigens (e.g., REC-8 antibody is sensitive to fixation and permeabilization). The order presented here yields bright antibody staining with the REC-8, WAPL-1, HIM-3, pH3, FLAG, and CYE-1 antibodies used, among others.

1. Prepare the click EdU cocktail⁸ fresh by adding the following to a clean 1.5 mL tube. The order of additions is important. Protect from light and maintain all reagents on ice. This recipe yields enough for one sample (100 μL); multiply the recipe as needed.
   1. Add 2 mL of ultrapure water to the buffer additive. This makes 10x buffer additive, which must be diluted to 1x immediately prior to use.
   2. Add 8.5 μL of 10x buffer to 76.5 μL of ultrapure water. Mix well.
   3. Add 4 μL of 100 mM CuSO₄ (may be labeled as Component E). Mix well.
   4. Add 0.25 μL of the 488 nm dye Azide. It must be thawed at room temperature, as its solvent, dimethyl sulfoxide, is solid at 4 °C. Mix well and protect from light.
   5. Mix 9 μL of ultrapure water with 1 μL of the buffer additive in the cap of the tube. Pipet from the cap to add to the remaining cocktail, and mix well by pipetting up and down.
2. Perform the EdU click reaction.
   1. Add ~100 μL of click EdU cocktail to the gonads in the small tube. Cover with laboratory film and incubate for 30-60 min at room temperature.
   2. Wash once with 100 μL of reaction rinse buffer.
   3. Wash the gonads 4 times using ~1 mL of PBSTw. Incubate for ~15 min per wash to allow excess EdU cocktail components to diffuse into wash. Using a drawn-out long glass Pasteur pipette, remove as much liquid as possible without disturbing the gonads. Leave no more than 50 μL.

7. Stain DNA and Prepare Slides

1. Add 1 drop (~25 μL) of antifade mounting medium with 4’,6-diamidino-2-phenylindole (DAPI, used to visualize DNA) to the gonads. Wait a few minutes so it can settle and mix with the gonads.
   NOTE: Alternately, a 1:1,000 dilution of DAPI (from a 0.1 mg/mL stock) in PBS may be applied for 5 min, followed by 20 μL of 1,4-diazabicyclo[2.2.2]octane (DABCO) in 90% glycerol, or another antifade mounting medium.
2. Prepare a large 2.5% agarose pad on a standard glass microscope slide.
3. Use a new clean dust-free long glass Pasteur pipette to transfer the gonads to the agarose pad. Keep all liquid and gonads in the narrow bottom of the pipette to minimize the loss of gonads.
   NOTE: Gonads stuck to small glass tube or in long glass Pasteur pipette can be “rescued” by rinsing with PBSTw, collecting the liquid in a dissecting dish, and picking individual animals onto the slide with an eyelash.
4. Use an eyelash (or a loop of thin hair) glued to a toothpick to distribute the gonads over the agarose pad and remove the dust particles.
5. Apply a rectangular glass coverslip. Lower slowly from one side to avoid air bubbles. Use a tissue to remove excess solution thus preventing the coverslip from moving freely.
   NOTE: Choose coverslips that match to the microscope that will be used. #1 and #1.5 coverslips work well.
6. Allow the slides to settle and dry slightly overnight at room temperature or 4 °C. This helps to slightly flatten the gonads. Slides should be stored at 4 °C.
7. Optional: Seal the edges of the slide with nail polish, or another slide sealer. Sealing the corners first, then the sides, prevents the coverslip from shifting.

8. Confocal Imaging and Analysis

1. Image the distal gonad with a spinning disc confocal fluorescent microscope equipped with a high energy light source, plan-apochromatic objectives, and a high efficiency microscope camera. Capture the images with 1 μm or tighter spacing between z-stacks. Take note of laser power, sensitivity, and exposure time for all channels.
   1. Use the following: 405 nm laser line excitation with a 485 nm (W60) emission filter for DAPI, 488 nm laser line excitation with a 527 nm (W55) emission filter for EdU, 561 nm laser line excitation with a 615 nm (W70) emission filter for WAPL-1, and a 640 nm laser line excitation with a 705 nm (W90) emission filter for pH3.
      NOTE: Signal intensity and background intensity will vary. Likewise, the required exposure times will vary, possibly up to 10-fold.
2. Use the Cell Counter plug-in²³ in Fiji^24,25 to manually count each nucleus. Label each individual nucleus according to the presence and absence of pH3, EdU, and WAPL-1. Use the classes of nuclei described in Table 1 and Figure 2, as these will facilitate all of the calculations outlined below.
   NOTE: Skilled experimentalists can accurately count all nuclei in a 3D image without double-counting or missing any nuclei. Alternately, one may count every nucleus in every z-plane, and the Marks-to-Cells R script³ may be used to remove multiply-counted nuclei.
3. Calculate cell numbers and frequencies from the above counts depending on the type of cell cycle measurement needed. The types of nuclei are defined in Table 1 and Figure 2. The calculations are summarized in Table 2.
   1. (Variation I) To identify the nuclei in S-phase, feed the animals EdU for 30 min. Any nuclei displaying EdU label are S-phase nuclei. To calculate, take the sum A and C nuclei, see Table 1 and Figure 2.
      NOTE: In a 30 min EdU pulse in wild-type adult hermaphrodites, all EdU labeled nuclei are co-labeled with progenitor zone markers¹.
   2. To measure the progenitor zone, stain with a progenitor zone marker such as REC-8 or WAPL-1 antibody. The progenitor zone is defined here as all nucleoplasmic REC-8¹⁸ or WAPL-1 immunoreactive germline nuclei. To calculate, sum all WAPL-1 immunoreactive nuclei (A+B+C+D, see Table 1 and Figure 2).
      NOTE: WAPL-1 also labels the DTC and somatic gonad nuclei which should not be counted. Somatic nuclei are easy to identify by extremely intense WAPL-1 signal, position slightly outside the germline, and a “fried-egg” morphology of the nuclei.
   3. To measure the S-phase index, perform a 30 min EdU experiment and co-label with REC-8 or WAPL-1 antibody. The S-phase index is defined as the proportion of the progenitor zone that is in S-phase. To calculate, count all S-phase nuclei, and then divide by the total number of progenitor zone nuclei (A+C / A+B+C+D, see Table 1 and Figure 2).
   4. (Variation II) To identify the nuclei in M-phase, stain with pH3 antibody. Any pH3 immunoreactive nuclei are M-phase nuclei. This works regardless of the duration of the EdU feed. To calculate, take the sum of A and B nuclei, see Table 1 and Figure 2.
      NOTE: WAPL-1 also labels the DTC and somatic gonad nuclei which should not be counted. Somatic nuclei are easy to identify by extremely intense WAPL-1 signal, position slightly outside the germline, and a “fried-egg” morphology of the nuclei.
   5. To measure the M-phase index, co-label with pH3 and REC-8 or WAPL-1 antibodies. The M-phase index is defined as the proportion of the progenitor zone that is in M-phase. To calculate, count all M-phase nuclei, and then divide by the total number of progenitor zone nuclei (A+B / A+B+C+D, see Table 1 and Figure 2).
   6. (Variation III) Nuclei in mitotic and meiotic S-phase both label with EdU. To tell the two populations apart, determine whether the S-phase was followed by mitosis or by meiosis. To determine whether nuclei are in mitotic or meiotic S-phase, feed EdU for 4 h and co-label for pH3 (an M-phase marker) and HIM-3 (a meiotic chromosome axis protein) by antibody staining. Record the nuclei that display both EdU and pH3 (type A, see Table 1 and Figure 2) as mitotic S-phase while nuclei that display both EdU and HIM-3 (type E, see Table 1 and Figure 2) as meiotic S-phase.
   7. (Variation IV) Calculate the duration of G2 phase.
      NOTE: G2-phase separates S-phase from M-phase. Although no marker has been reported to label G2 in the C. elegans germline, one can calculate the duration of G2 phase by combining data from several experiments that label M-phase (at the time of dissection) and S-phase (starting at several h before dissection). A cell that displays both M-phase and S-phase markers completed G2-phase during the course of the experiment. A cell that displays only the M-phase marker and not the S-phase marker was not in S-phase during the experiment.
      1. To calculate the duration of G2-phase, feed EdU for 2 h and co-label with pH3 antibody. Examine only nuclei that label with pH3 (these are in M-phase at the time of dissection) for the presence of EdU (these were in S-phase during the 2 h EdU label prior to dissection). Calculate the fraction of M-phase nuclei that completed G2-phase (A / A+B, see Table 1 and Figure 2).
      2. Repeat this experiment with a 3 h EdU label, and again with a 4 h EdU label (and optionally a 5 h EdU label). Plot the percent of pH3 positive nuclei that are EdU positive on the y-axis and the duration of EdU label on the x-axis, as shown in Figure 3A.
      3. Calculate the median duration of G2-phase by connecting the points on the graph and determining where the line crosses 50%, as shown in Figure 3A.
      4. Calculate the maximum duration of G2-phase by connecting the points on the graph and determining where the line crosses 99%, as shown in Figure 3A.
   8. (Variation V) Calculate the duraion of G2+M+G1.
      NOTE: In the C. elegans germline, G1 phase is unusually short. Although no marker has been reported to label G1 in the C. elegans germline, one can estimate the sum duration of G2, M, and G1 phase, and then compare this time with the G2-phase time described above. The maximum duration of G2+M+G1 is estimated from the percentage of all progenitor zone nuclei (WAPL-1 immunoreactive) that remain EdU negative (did not undergo S-phase) after EdU labeling for several hours.
      1. To calculate the duration of G2+M+G1, feed EdU for 2 h and co-label with REC-8 or WAPL-1 antibody. Determine the fraction of the progenitor zone that underwent S-phase during this time (A+C / A+B+C+D, see Table 1 and Figure 2).
      2. Repeat this experiment with a 3 h EdU label, and again with a 4 h EdU label (and optionally a 5 h EdU label). Plot the percent of REC-8 or WAPL-1 positive nuclei that are EdU positive on the y-axis and the duration of EdU label on the x-axis, as shown in Figure 3B.
      3. Calculate the maximum duration of G2+M+G1 by connecting the points on the graph and finding where the line crosses 99%, as shown in Figure 3B.
         NOTE: It is possible to perform the experiments to determine the duration of G2 and G2+M+G1 as a single set of 2, 3, 4, and 5 h EdU experiments by co-labeling with both rabbit-anti-WAPL-1 and mouse-anti-pH3 antibodies.
   9. (Variation VI) To identify the nuclei that replicated in the progenitor zone but have since entered meiosis, feed the animals EdU for 10 h and co-label with REC-8 or WAPL-1 antibodies. Any nuclei displaying EdU label were in S-phase during those 10 h. Any nuclei that do not display nucleoplasmic REC-8 or WAPL-1 staining were in meiosis. Simply count the nuclei with EdU labeling that do not display labeling with the progenitor zone marker (E, see Table 1).
      NOTE: Conversely, if gonads are stained for the meiotic prophase marker HIM-3 with anti-HIM-3 antibodies, count the number of nuclei with EdU labeling that are also positive for HIM-3.
   10. To calculate the rate of meiotic entry, perform the above experiment with a 5 h, 10 h, and 15 h label of EdU. Plot the number of nuclei that entered meiosis on the y-axis and the duration of the EdU label on the x-axis, as shown in Figure 3C. Then use a simple linear regression to calculate the slope (nuclei entered meiosis per h) from y=mx+b.
       NOTE: It is critical to use a linear regression to calculate the rate of meiotic entry. It would be incorrect to simply divide the number of nuclei that entered meiosis by the duration of the EdU label, because the y-intercept is not zero.
   11. (Variation VII) Measure the rate of meiotic progression.
       NOTE: Since EdU is covalently incorporated into DNA, it can be used to track a population of cells through differentiation. The cells that underwent S-phase in the progenitor zone retain the EdU label as they enter into meiosis, progress through meiosis, and undergo oogenesis. A pulse-chase experiment with EdU can be used to measure the rate of meiotic progression.
       1. Feed EdU-labeled bacteria to the animals for 4 h (the “pulse”). Transfer the animals to unlabeled OP50 bacteria for 48 h (the “chase”), then dissect and co-label with a progenitor zone marker such as REC-8 or WAPL-1 (or a meiotic prophase marker such as HIM-3) if desired.
       2. When imaging, look for the position of the most proximal EdU-labeled nucleus. The rate of meiotic progression is the distance (in cell diameters from the end of the progenitor zone) traveled by the most proximal EdU labeled nucleus during the 48 h chase.