Individual Culturing of Tigriopus Copepods and Quantitative Analysis of Their Mate-guarding Behavior

1. Preparation of Virgin Animals for Behavioral Observation

1. Obtain fertilized eggs and allow them to hatch.
   1. Collect gravid females carrying clear orange (i.e., fertilized and in advanced stages of development) eggs (Figure 3C) from a stock culture with a Pasteur pipette. Rinse the females by gently pipetting them in clean culture medium to avoid transfer of other animals, including nauplial larvae that may have hatched in the culture.
      Note: In this protocol, artificial seawater with a salinity of 35% is used as culture medium. Use different medium (e.g. artificial seawater with a higher salinity or an additional solute) depending on the purpose of an experiment.
      Note: See Barreto et al. 2015¹² for a method for establishment of laboratory culture stocks and Pereira et al. 2016¹³ for examples of collection sites of natural populations of T. californicus. Peterson et al. also reported their collection sites of T. californicus, T. fulvus, and T. japonicus with latitude and longitude information⁹. Use plastic pipettes with a capacity of approximately 30-50 mL to collect Tigriopus copepods from rock pools.
   2. Place each female individually in a well of a 6-well cell culture plate with clean culture medium (Figure 4, left). Make sure that no other animal (including nauplial larva) is contaminating the well.
   3. Maintain the plates in an incubator set at 20 °C with a 12-hour light-dark cycle to culture the females until they release egg sacs. This step generally takes one to ten days depending on species and developmental stages of embryos. Meanwhile, feed each gravid female twice a week with two grains of finely ground (approximately < 0.5 mm in diameter) fish food (see Table of Materials for details).
      Note: Use different temperatures and light cycles depending on the purpose of an experiment. Higher temperatures can facilitate more rapid development of the embryos.
      Note: Avoid leaving excess food decaying in wells. If food debris starts to decay, pipette it out of wells with a Pasteur pipette.
   4. After egg sacs are released, remove the females from the culture wells with a Pasteur pipette.
      Note: Inseminated females are capable of spawning multiple clutches of offspring^2,3. If needed, transfer the females into other wells to collect further clutches.
2. Collect copepodids for individual culture.
   1. Keep the plates with hatched nauplii in the incubator and culture the nauplii until they develop to the first copepodid (CI) stage (Figure 4, middle). This step generally takes from one to two weeks. Feed each well with several grains of the finely ground fish food while searching for CI animals once every two days. Refill evaporated rearing water with distilled water.
      Note: If floating debris obstructs the view, skim it with a small piece of paper towel.
   2. As soon as CI animals start to emerge, collect them from the wells with a P-10 micropipette with its pipetting volume set at approximately 8 µL, under a stereomicroscope at 10X to 40X magnification (Figure 4, right). Wash each CI animal by gently pipetting it in clean artificial seawater and place it in a well of 24-well cell culture plate containing 2-3 mm depth (approximately 400-600 µL) of artificial seawater.
      Note: Avoid carrying over nauplial exuviae or other animals into the wells for individual culture.
3. Track development of the individuals by counting molted exuviae.
   1. Examine inside of each well for molted exuviae every two to three days (adjust frequency if needed) by stereomicroscopic observation under a dark-field illumination at 10X to 40X magnification. Exuviae of copepodids are transparent and recognizable with bristled legs, a pair of caudal rami (i.e., thin protruding structures at the caudal end), and/or segmentation of prosome and urosome (Figure 5). Change focus and illumination to detect exuviae at different depths in a well (Figure 6A).
      Note: Molted exuviae are smaller than the animal that has molted them. Start examination from lower magnification for an animal and its relatively recent exuviae and shift to higher magnification for older (i.e., smaller) exuviae.
      Note: If exuviae in a well are damaged, eliminate the well from further developmental tracking to avoid misjudgment of developmental stages.
      Note: If floating debris obstructs the view (Figure 6B), skim it with a small piece of paper towel.
   2. Record a total number of copepodid exuviae in each well with a tally mark on the plate lid (Figure 7). Add a line to the mark as a new exuviae is found in the well.
      Note: If there are nauplial exuviae contaminated in the well, eliminate them from the count.
   3. Feed each individual with two grains of the finely ground fish food. Refill evaporated rearing water with distilled water to maintain the salinity.
      Note: Feed copepodids every two to three days to prevent them from consuming and damaging molted exuviae, which could potentially influence estimation of developmental stage (step 1.3.4).
   4. Estimate developmental stage of the animal based on the number of the exuviae. If there is no exuviae, the individual is estimated to be at CI stage. If there are one to four exuviae, the individual is estimated to be at CII to CV stages. If there are five exuviae, the individual is estimated to be an adult.
4. Identify sex of adults based on morphology.
   1. Sex animals by examining their morphology. Adult Tigriopus males possess geniculate first antennae with hooked and globular structures at the distal end (Figure 3A). Adult females possess smoother and relatively thinner first antennae (Figure 3B) than those of males. Some females also exhibit dark green color from gonadal lipid in their bodies (Figure 3B, 3D).
   2. Mark the sex on the lid of the well (Figure 7B).

2. Behavioral Test and Video Recording of Mate-guarding Behavior

1. On a testing day, select animals of desired stage and sex for a behavioral test.
   1. Feed the selected individuals in culture wells with finely ground fish food and allow them to eat it for 30 minutes. Meanwhile, proceed to step 2.1.2 to prepare testing chambers.
   2. Prepare two 48-well flat bottom cell culture plates as testing chambers; one plate (hereafter called “plate A”) is for males and the other (“plate B”) is for targets (e.g. copepodids, adult females, adult males). Add 400 µL of clean artificial seawater with a salinity of 35% in wells of the plates. Place the plates on an LED light pad covered with a translucent acrylic board as a light diffuser (Figure 8).
      Note: Use different medium depending on a purpose of an experiment.
      Note: To avoid excess heat, do not use incandescent or fluorescent lamps as a backlight.
   3. After the 30-min feeding time described in 2.1.1, rinse each animal by gently pipetting it into a succession of four wells of a 24-well plate filled with clean artificial seawater (Figure 9) to prevent carry-over of debris and exuviae from culture wells.
   4. Place the rinsed individuals in the wells of plates A and B on the LED light pad. Allow 30 minutes for adjustment of the animals to the wells.
   5. Set a video camera above plate A during the adjustment period (Figure 8). Use a tripod or a stand to hold the camera.
   6. Focus the camera on the males inside of plate A to allow an observation of antennae.
      Note: To reduce flicker of the backlight in movies, set a frame rate equal to or half of an electrical frequency and use a longer exposure time. Also, keep the LED light pad covered with a translucent acrylic board as described in step 2.1.2.
2. After the adjustment period described in step 2.1.4, start video recording and the behavioral test.
   1. Transfer the targets from plate B to plate A with a Pasteur pipette. If the experiment is sensitive to chemical components in medium, change pipettes for each testing pair.
      Note: When pipetting an animal from plate B, do not chase it in the water with a Pasteur pipette as water disturbance can stimulate the animal and elicit its excess movement. Hold a tip of the pipette slightly above the water surface and wait for the individual to come beneath the tip. Gently pipette it up with a small amount of artificial seawater and then eject it into a well on plate A.
   2. According to an experimental plan, allow a male and a target to interact in each well for a desired period of observation time (e.g. 10 minutes) after the transfer.
   3. Stop video recording after the observation time. If necessary, download the recorded movies from the camera to a computer.

3. Manual Analysis of Behavioral Properties

1. Examine the movie for the timing when each target was transferred into the well on plate A.
2. Examine initiation and termination timing of events of interest and calculate duration, latency, and frequency of the events.
   1. Define initiation of guarding attempt by a male as a contact of the male’s antenna with any body part of the target (Figure 10, top left). The antennal contact is often preceded by a swift (<0.5 s) chase or pounce.
   2. Define termination of guarding attempt as a point when both antennae of the male detach from the body of a target (Figure 10, top right). Calculate the frequency of guarding attempts as:
      
   3. Define initiation of copulation by a dorsal body bend of a male that is followed by a repetitive press of an urosome against that of the female (Figure 10, bottom right). Frequency of the push is several times per second.
      Note: A guarding male crawls down to the caudal end of a female’s body prior to copulation (Figure 10, bottom left).
   4. Define termination of copulation by detachment of the urosomes after the events described in 3.2.4.
      Note: Copulation generally takes place for several minutes in cases of T. californicus and T. japonicus.

4. Two-dimensional Tracking of Individuals and pairs

1. Generate a video clip featuring an episode of interest (e.g. the first 3 s of a guarding attempt) by trimming the movie recorded in step 2 with movie editing software.
2. Install ImageJ¹⁴ from: https://imagej.nih.gov/ij/download.html. Then download MTrackJ, a motion-tracking plugin for ImageJ¹⁵, by following instructions on: https://imagescience.org/meijering/software/mtrackj/.
3. Open ImageJ and import a movie of interest (e.g., File > Import > Using QuickTime; choose “Convert 8-bit grayscale” to reduce memory required for image processing).
4. Perform spatial and time calibration.
   1. Select the Straight Line Selection tool in the ImageJ window for spatial calibration.
   2. Draw a selection line along an object with a known length (e.g. diameter of a well) in the movie window.
   3. Open the “Set Scale” menu (Analyze > Set Scale). Enter the known length in the “Known Distance” box and its unit (e.g. mm) in the “Unit of Length” box. Click the “Global” checkbox to use the scale for other video clips.
   4. Open the “Properties” menu (Image > Properties) for time calibration. Enter frame interval (reciprocal of the frame rate) in the “Frame interval” box.
5. Configure tracking with MTrackJ.
   1. Open the MTrackJ plugin from Plugins > MtrackJ on ImageJ. Click the “Tracking” button in the MTrackJ dialog window to open a tracking configuration menu.
   2. Set a tracking interval by checking “Move to next time index after adding point” and entering a number in “Time step size” box in the configuration dialog window. To perform frame-by-frame tracking, enter “1” in the “Time step size” box.
   3. Check “Apply local cursor snapping during tracking” and choose “Dark centroid” as a snap feature. This option allows an automatic detection of the centroid of a dark object (i.e., an individual or a pair) within a detection square (“snap range”) around a cursor. Choose a size of the square to cover a whole pair or animal in the movie (e.g. 31 x 31 pixels).
   4. Click “OK” to apply the chosen options.
6. Perform two-dimensional tracking.
   1. Click the “Add” button in the MTrackJ dialog window to start tracking.
   2. Place a cursor (detection square) to cover a pair or an individual to be tracked. Click to detect the dark centroid of the object within the square.
   3. Repeat step 4.6.2 for a desired number of frames.
   4. To generate data tables, click “Measure” button in the MTrackJ dialog window. The data is displayed in two windows: “MTrackJ: Tracks” window shows a summary of the tracked two-dimensional trajectory and “MTrackJ: Points” shows detailed data for each time point. To save each table, choose File > Save as… menu on ImageJ.
      Note: Refer to the online manual provided by the developer (https://imagescience.org/meijering/software/mtrackj/manual/) for descriptions of each data column.
   5. To generate a movie with a tracked trajectory drawn as a colored line, click the “Movie” button in the MTrackJ dialog window. To save the generated movie, choose File > Save as… menu on ImageJ.
7. Save the whole result. Click the “Save” button in the MTrackJ dialog window to save the results including the data (step 4.6.4) and the tracked two-dimensional trajectory (steps 4.6.2 and 4.6.3).