In Vivo Visualization of Calcium Transients during Fertilization and Early Development in C. elegans

1. C. elegans maintenance

1. Nematode Growth Medium (NGM) plate preparation
   1. Combine 3 g of sodium chloride (NaCl), 2.5 g of Bacto-Peptone, 20 g of Agar, and 1 L of deionized water in a 4 L flask (see Table of Materials). Cover the top of the flask with aluminum foil before autoclaving.
   2. Autoclave to sterilize using liquid sterilization cycle settings. A sterilization cycle set at 121 °C for 30 min is recommended.
   3. Allow the mixture to cool to ~50 °C, approximately 20 min at room temperature (RT).
   4. Add 25 mL of sterile 1 M KH₂PO₄ (dissolved in water, pH = 6), 1 mL of sterile 1 M CaCl₂ (dissolved in water), 1 mL of sterile 1 M MgSO₄ (dissolved in water), and 1 mL of 5 mg/mL cholesterol (dissolved in 100% ethanol), gently swirling after each addition (see Table of Materials).
   5. Using a sterile technique, transfer a portion of the mixture into a sterile 300 mL beaker for easier pouring. Pour the warm media to cover the bottom of the plate.
      NOTE: Approximately 10 mL per plate in 60 mm x 15 mm sterile Petri plates (see Table of Materials) will produce 100 plates. Alternatively, 35 mm x 10 mm sterile Petri dishes may be used, where ~4 mL per plate will produce ~250 plates.
   6. Allow the NGM plates to dry at RT for at least 1 day before seeding with bacteria. Store plates upside down at 4 °C for future use.
2. OP50 bacterial lawn preparation
   1. Make Luria Broth (LB) by combining 5 g of yeast extract, 10 g of Bacto-Tryptone, 5 g of NaCl, and 1 L of deionized water in a 2 L flask (see Table of Materials). Using a stir plate and stir bar, mix until dissolved.
   2. Aliquot LB into bottles and autoclave using liquid sterilization cycle settings. A sterilization cycle set at 121 °C for 30 min is recommended. If using a container with a screw cap, loosen the screw cap a quarter of a turn for proper ventilation during the autoclave cycle.
   3. Allow the broth to cool to RT before using.
   4. Using sterile technique, inoculate 50 mL of LB in a 250 mL Erlenmeyer flask with OP50 bacteria and grow overnight at 37 °C with shaking.
   5. Using sterile technique, transfer 300 μL of saturated culture onto the center of a 60 mm x 15 mm NGM plate. For 35 mm x 10 mm plates, 100 μL of saturated culture is recommended.
   6. Allow the plates to dry at RT for 1-3 days until the OP50 bacterial spot is no longer liquid. Ensure the plates are dry and are free of contamination before use. Store the plates at 4 °C for future use. Storing plates upside down reduces condensation on the plate surface.
3. Worm pick construction
   1. Loosen the silver collar towards the head of the worm pick handle (see Table of Materials) by gently unscrewing so the four sections come apart.
   2. Cut 1.5 inches (about the length of the thumb) of platinum wire (see Table of Materials) using scissors and place it in the middle of the four sections; about an inch will stick out of the handle.
   3. Hold the wire in the center while tightening the collar; the four sections will come together in the middle and hold the platinum wire in place.
   4. Carefully shape the wire using a pair of flat-nose pliers. A rainbow or arc shape tends to work best for picking.
   5. Create about a 2-3 mm "foot" on the end of the wire by repeatedly clamping down with the flat-nose pliers. Use this "foot" to pick up the worms and keep the "foot" flat to the surface of the plate.
      NOTE: Clamping too hard will cut the platinum wire. Avoid creating sharp edges using the flat-nose pliers (or equivalent) to prevent puncturing the worms or the plates.
4. Worm handling
   1. Hold the worm pick similar to a pencil and sterilize the "foot" by placing the wire in the flame of a 95% ethanol lamp until it is bright orange.
   2. Place an OP50 seeded plate onto the stage of a dissecting scope and remove the lid. Gently touch the "foot" of the pick to an area of the bacterial lawn and use a gentle swiping motion to gather bacteria onto the bottom. The bacteria that are stuck to the pick (referred to as "goo") will act as a glue to pick up worms.
   3. Move a plate of worms to the dissecting scope stage and remove the lid while keeping the pick from touching other surfaces. Quickly and gently touch the "foot" of the pick onto a worm to pick it up. This technique enables multiple worms to be moved between plates simultaneously.
   4. Gently touch the "foot" of the pick to the unseeded portion of a new plate to transfer the worms; typically, this is the same plate that had the bacteria taken from it in section 1.4.2.
   5. Sterilize the pick between each plate and/or worm strain to avoid contamination and allow the pick to cool after sterilizing.
      NOTE: Touching a hot pick on a worm will stress or kill the animal. The worm pick will cool after touching the bacterial lawn to pick up "goo" for worm transfer.
5. Maintain worms at 20 °C on NGM plates (step 1.1) seeded with OP50 Escherichia coli bacteria (step 1.2). Passage worms 2 – 3 times per week by transferring several adults to fresh plates (60 mm x 15 mm) using a worm pick (steps 1.3-1.4) to avoid starvation, which can alter physiology.
   NOTE: C. elegans strains and OP50 bacteria are available through the Caenorhabditis Genetics Center (CGC).

2. C. elegans developmental synchronization

1. Age-synchronize worms using a timed egg-lay (described here). If a large number of worms are needed for a classroom or other purpose, perform bleach synchronization as described in Golden et al.¹⁰.
2. Transfer adult worms to a new seeded OP50 plate using a platinum wire pick (described in section 1). The number of worms is dictated by the experimental need. Day 1 worms lay approximately 3 eggs per hour. Therefore, 10 worms will generate approximately 30 eggs in 1 h. Preferably, use worms on the first day of adulthood (Day 1).
3. After approximately 1 h, remove the adult worms from the plate, leaving only the eggs.
   NOTE: Inadvertently leaving a worm behind is a common error. Worms that remain will continue to lay eggs and disrupt synchronization. Worms mature into Day 1 adults approximately 72 h from being laid when incubated at 20°C.

3. Mounting worms for imaging

1. When imaging fertilization events, use Day 1 adults to maximize visualization opportunities.
   NOTE: In Day 1 adults, an oocyte is fertilized every ~23 min. However, ovulation slows with age, and a typical hermaphrodite will stop laying eggs after Day 4.
2. Prepare a solution of 3% agarose in water by mixing 3 g of agarose in 100 mL of water. Heat the agarose solution using a microwave until the agarose is fully dissolved. To reuse a solidified solution, heat until fully dissolved. After several uses, generate a fresh solution.
3. Place a blank microscope slide for imaging in between two other microscope slides that have a single layer of lab tape across the long axis of the slide (see Table of Materials). The tape creates a spacer for the agarose pad, allowing for even distribution and a uniform width. Ensure that the three slides are in a row with the long sides touching: two slides with tape that flank a blank microscope slide.
   1. Drop the heated 3% agarose solution (~100-150 μL) onto the center of the microscope slide that is between the taped slides using a 1 mL pipette.
   2. Quickly place a new microscope slide onto the agarose droplet, ensuring the new slide is perpendicular to the slide with the agarose droplet and resting across the adjacent taped slides.
   3. Gently separate the slides. Avoid tearing the pad.
   4. Prepare the agarose pads fresh each day of imaging. It is recommended to make multiple pads when preparing for imaging, but refrain from exposing more than one pad at the same time. Exposed pads will dry out rapidly.
4. Using a 20 μL pipette, add a small drop (~7 μL) of 1 mM levamisole in M9 buffer to the center of the agarose pad to paralyze the worms. Use a filter-sterilized 100 mM stock solution of levamisole to make the 1 mM working solution.
   1. To make M9 buffer, combine 6 g of Na₂HPO₄, 3 g of KH₂PO₄, 0.5 g of NaCl, 1 g of NH₄Cl, and 1 L of deionized water in a suitable container (see Table of Materials).
   2. Autoclave the solution using liquid sterilization cycle settings and let the solution cool to RT before use. If using a container with a screw cap, loosen the screw cap about a quarter of a turn for proper ventilation. A sterilization cycle set at 121 °C for 30 min is recommended.
5. Pick at least 5 synchronized worms and gently touch the "foot" to the levamisole droplet to transfer all worms. Use at least 5 worms per slide, as the orientation of the body can cause the intestine to obscure the germline, blocking visualization of the calcium wave in the oocyte.
   NOTE: Ensure that the worms are transferred quickly as the levamisole will evaporate if left sitting on the agarose pad for longer than 5 min.
6. Carefully cover the agarose pad with a coverslip. Check the location of the worms, as the coverslip may cause the dispersal of the worms from the initial placement. Label the microscope slide with appropriate details, such as the name of the worm strain (see Table of Materials).
   NOTE: Ensure imaging is done as soon as possible after mounting worms, as prolonged exposure to levamisole is toxic.

4. Imaging calcium during fertilization in  C. elegans

1. Locate the worms using a low-powered objective, such as 4x, on the microscope of choice. Use brightfield illumination.
2. Use a minimum of a 20x air objective for clearly visualizing the calcium wave in oocytes. Use an immersion objective of 40x or greater.
3. Depending on the system one uses, the excitation intensity typically is a fixed value. For optimal visualization of the fluorescent reporters, adjust the gain or exposure time.
   NOTE: Continuous high-intensity light will damage the sample and is deleterious to worm health. If desired, turn off the laser while waiting for a fertilization event to avoid continuous GFP illumination. See the hallmarks of a fertilization event in sections 4.8 – 4.9.
4. Ensure the 488 nm GFP channel is set to a laser power where the CaFE reporter signal is easily visible.
   1. For a Nikon ECLIPSE Ti2 laser scanning confocal, detect the CaFE reporter at a laser power of 30% for the GFP channel and a gain of 75. The pinhole size for the Nikon ECLIPSE Ti2 laser scanning confocal is 30 μm.
   2. For the Andor Dragonfly spinning disk confocal using the Zyla camera, detect the CaFE reporter at a laser power of 30% for the GFP channel. The pinhole size for the Andor Dragonfly spinning disk confocal is 40 μm.
   3. Adjust the laser settings and exposure time for every sample examined.
5. If using the EAG28 (CaFE; PH-mCherry) or EAG25 (CaFE; H2B-mCherry) strain, ensure the red channel is set to a laser power where the mCherry signal is easily visible in addition to the GFP channel.
   1. For the Nikon ECLIPSE Ti2 laser scanning confocal, detect the mCherry reporter at a laser power of 15 for the RFP channel and a gain of 130.
   2. For the Andor Dragonfly spinning disk confocal using the Zyla camera, detect the mCherry reporter at a laser power of 30% for the RFP channel.
   3. Adjust the laser settings and exposure time for every sample examined.
6. Set the frame rate or frames per second (fps) as fast as the system allows.
   NOTE: The CaFE reporter is typically observed at approximately 1 fps for the Nikon ECLIPSE Ti2 laser scanning confocal and 10 fps for the Andor Dragonfly spinning disk confocal with the Zyla camera.
7. Select time lapse or time series protocol and adjust to desired settings. Acquire time-lapse images for 30 min or less, with the repeats option set to '1'.
   NOTE: If the fertilization event occurs before the time interval is complete, then stop the video to preserve file size.
8. Locate the worms on the slide and select a worm where the proximal gonad is visible. The intestine and the gonad twist around each other inside the worm. As such, a gonad arm is often obscured by the intestine, preventing visualization of fertilization. If the -1 oocyte is obscured, select another worm for visualization. To increase the probability of the selected worm having a visible gonad arm, mount a minimum of 5 worms per slide. Use early Day 1 adults to visualize fertilization.
9. Examine each -1 oocyte in the gonad arms of the worm to determine if a fertilization event is imminent.
   NOTE: Prior to fertilization, during meiotic maturation, the nucleus in the -1 oocyte will migrate to the back of the oocyte, and the nuclear envelope will dissolve. These events are visible in both Nomarksi and GFP channels using the CaFE reporter, and such events indicate that ovulation will occur within the next ~10 min.
10. After nuclear envelope breakdown, the cytoskeleton of the most proximal, -1 oocyte will rearrange, take on a rounded appearance, and separate from the -2 oocyte. At this point, ovulation will occur imminently. Start recording the video immediately if these events are observed. Take videos in a widefield setting.
    NOTE: If movement of the nucleus or rounding of the oocyte is not visible after 20 min, an ovulation event is unlikely to occur. Focus on a different worm and/or prepare a fresh slide.

5. Quantitation of calcium transients in C. elegans fertilization

1. Quantitate calcium transients from fertilization videos using the microscope system software like Nikon NIS Elements (described here). ImageJ/FIJI are also suitable for image analysis¹¹.
2. To quantitate the change in fluorescence of fertilization videos, use the following signal values: the signal at time 0, the signal from the frame of interest, and the baseline signal.
3. Identify the time and frame that captures the first burst of fluorescence occurs in the -1 oocyte. Define this frame as time 0.
4. To obtain the signal from the frame of interest, draw the region of interest (ROI) to encompass the following areas: the -1 oocyte prior to fertilization, the spermatheca, and the newly fertilized oocyte in the uterus. The signal displayed in the ROI is termed F₁.
   NOTE: Ensure the ROI covers the entire area trafficked by the oocyte. The ROI should be identical in every frame of the time-lapse recording being analyzed.
5. Define a blank area of the image with no fluorescence using the same dimensions as the ROI defined in step 5.4 in order to calculate the background signal in each frame. Subtract the background signal from every frame.
6. To obtain the baseline signal (F₀), locate the frame in a minimum of 15 frames before time 0. Using the same ROI as in step 5.4, note the signal at each frame until time 0. Average the signals (with background subtraction) from the starting frame until time 0 to calculate the baseline signal.
7. Calculate the change in fluorescence (ΔF) using the following formula: . F₁ is defined in each frame as the signal in the ROI without the background signal. he F0 is the averaged baseline fluorescent signal of ≥15 frames before time 0, as calculated in section 5.6. 
8. Determine the rate of change between frames by subtracting the change in fluorescence between 2 frames and dividing by the time.

6. RNAi in C. elegans

1. Prepare RNAi plates
   1. Prepare plates according to step 1.1 and add the following additional reagents to allow for gene knockdown via RNAi at step 6.1.4.
   2. Add 1 mL of 1 M sterile-filtered isopropyl β-d-1-thiogalactopyranoside (IPTG) (dissolved in deionized water) for a final concentration of 1 mM IPTG in the media (see Table of Materials).
   3. Add 1 mL of 100 mg/mL sterile-filtered ampicillin (dissolved in deionized water) for a final concentration of 100 μg/mL ampicillin in the media (see Table of Materials). Alternatively, use 40 μg/mL carbenicillin¹².
   4. Swirl gently but thoroughly to mix and pour plates as described in steps 1.1.4 – 1.1.5.
2. Preparation of RNAi-saturated culture
   1. Calculate the amount of saturated culture by multiplying the number of seeded RNAi plates needed by 300 μL when using 60 mm x 15 mm plates or by 100 μL when using 35 mm x 10 mm plates.
   2. Using a sterile technique, grow desired RNAi bacteria in LB (described in steps 1.2.1-1.2.3) containing 100 μg/mL ampicillin overnight at 37 °C until saturated. Include a bacterial strain containing the L4440 empty vector as a negative control and a bacterial strain containing an RNAi positive control (e.g., egg-5) to verify the RNAi is working. Additionally, include a "blank" condition containing only LB to verify there is no contamination in the stock LB.
   3. Add sterile-filtered IPTG to a final concentration of 1 mM to the saturated overnight culture for induction of dsRNA expression and incubate for at least 2 h 30 min at 37 °C before seeding. Alternatively, dilute back the saturated overnight culture out of the log-phase prior to inducing with IPTG.
      NOTE: IPTG is included in both the plates and the saturated culture for optimal dsRNA expression.
   4. Using sterile technique, transfer 300 μL of the induced culture onto the center of a 60 mm x 15 mm RNAi plate (solidified). Alternatively, use 100 μL of saturated culture for a 35 mm x 10 mm plate.
   5. Allow the plates to dry at RT for 1-3 days until the RNAi bacterial spot is no longer liquid. Ensure plates are dry and are free of contamination before use. Store plates at 4 °C upside down for future use.
      NOTE: Use RNAi plates as soon as possible for optimal knockdown, as RNAi efficacy decreases over time. Discard if >1 month old.
3. As described in section 2, synchronize the worms onto RNAi plates.
   1. Check worms 1-2 days after the egg-lay synchronization for developmental delays.
   2. Some RNAi may cause developmental defects when the knockdown begins at egg-lay. If needed, place worms onto the RNAi plates at a later stage, such as L3.
4. As described in sections 3 and 4, mount the worms and prepare for imaging using any microscope system equipped to detect GFP fluorescence.

7. Imaging calcium during early embryogenesis in C. elegans

1. Synchronize worms as described in section 2. Use the EAG28 strain, which combines the CaFE reporter and a pleckstrin-homology domain-mCherry marker highlighting the plasma membrane boundary. Alternatively, to visualize fertilized embryos ex utero, gently cut open the worm along the uterus using a sterile syringe needle to release the embryos.
2. Prepare worms or embryos for imaging as described in section 3, and follow the same instructions described in section 4 for setting imaging parameters.
3. Center the area of interest on the uterus of the worm to visualize embryos or a laid egg.
   1. For embryos in utero, ensure that the uterus is not behind the intestine, the uterus is not crowded, and that the individual cells in the embryo are clear in the focal plane.
   2. For eggs ex utero, ensure that the stages of the cells are not past the comma stage of the worm life cycle.
4. Select time lapse or time series protocol and adjust to desired settings.
   1. For optimal visualization of embryo calcium transients, increase the time settings for prolonged movies. Keep in mind that constant exposure to the laser will harm the embryo over long periods of time.
5. Capture videos or images by focusing on cells in embryos in the desired cell stage.
6. Use a 40x immersion objective or greater for imaging, although the transients are readily detected using a 20x air objective.