Name: individual culturing of tigriopus copepods and quantitative analysis of their mate-guarding behavior
Uploaded: 2018-09-26T14:30:00.000+00:00
Duration: 6 min 24 s
Description: Watch this Scientific Journal Video about Individual Culturing of Tigriopus Copepods and Quantitative Analysis of Their Mate-guarding Behavior at JoVE.com

This work is supported by grants from the Sumitomo Foundation, Japan (Grant for Basic Science Research Projects, grant number: 150932) and Research Institute of Marine Invertebrates, Japan (2018 individual research grant) to STI and RSB, and a grant from the US National Science Foundation (DEB-1556466) to RSB. We thank Ms. Kiana Michelle Woodward for feedback on the culturing and staging method.

1. Preparation of Virgin Animals for Behavioral Observation
<ol>
	<li>Obtain fertilized eggs and allow them to hatch.
	<ol>
		<li>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.</li>
		<li>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.</li>
		<li>Maintain the plates in an incubator set at 20&#160;&#176;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 &#60; 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.</li>
		<li>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 offspring2,3. If needed, transfer the females into other wells to collect further clutches.</li>
	</ol>
	</li>
	<li>Collect copepodids for individual culture.
	<ol>
		<li>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.</li>
		<li>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 &#181;L, 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 &#181;L) of artificial seawater. 
		Note: Avoid carrying over nauplial exuviae or other animals into the wells for individual culture.</li>
	</ol>
	</li>
	<li>Track development of the individuals by counting molted exuviae.
	<ol>
		<li>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.</li>
		<li>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.</li>
		<li>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).</li>
		<li>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.</li>
	</ol>
	</li>
	<li>Identify sex of adults based on morphology.
	<ol>
		<li>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).</li>
		<li>Mark the sex on the lid of the well (Figure 7B).</li>
	</ol>
	</li>
</ol>
2. Behavioral Test and Video Recording of Mate-guarding Behavior
<ol>
	<li>On a testing day, select animals of desired stage and sex for a behavioral test.
	<ol>
		<li>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.</li>
		<li>Prepare two 48-well flat bottom cell culture plates as testing chambers; one plate (hereafter called &#8220;plate A&#8221;) is for males and the other (&#8220;plate B&#8221;) is for targets (e.g. copepodids, adult females, adult males). Add 400 &#181;L 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.</li>
		<li>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.</li>
		<li>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.</li>
		<li>Set a video camera above plate A during the adjustment period (Figure 8). Use a tripod or a stand to hold the camera.</li>
		<li>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.</li>
	</ol>
	</li>
	<li>After the adjustment period described in step 2.1.4, start video recording and the behavioral test.
	<ol>
		<li>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.</li>
		<li>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.</li>
		<li>Stop video recording after the observation time. If necessary, download the recorded movies from the camera to a computer.</li>
	</ol>
	</li>
</ol>
3. Manual Analysis of Behavioral Properties
<ol>
	<li>Examine the movie for the timing when each target was transferred into the well on plate A.</li>
	<li>Examine initiation and termination timing of events of interest and calculate duration, latency, and frequency of the events.
	<ol>
		<li>Define initiation of guarding attempt by a male as a contact of the male&#8217;s antenna with any body part of the target (Figure 10, top left). The antennal contact is often preceded by a swift (&#60;0.5 s) chase or pounce.</li>
		<li>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: 
		<img alt="Equation" src="/files/ftp_upload/58378/58378eq1.jpg" /></li>
		<li>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&#8217;s body prior to copulation (Figure 10, bottom left).</li>
		<li>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.</li>
	</ol>
	</li>
</ol>
4. Two-dimensional Tracking of Individuals and pairs
<ol>
	<li>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.</li>
	<li>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/.</li>
	<li>Open ImageJ and import a movie of interest (e.g., File &#62; Import &#62; Using QuickTime; choose &#8220;Convert 8-bit grayscale&#8221; to reduce memory required for image processing).</li>
	<li>Perform spatial and time calibration.
	<ol>
		<li>Select the Straight Line Selection tool in the ImageJ window for spatial calibration.</li>
		<li>Draw a selection line along an object with a known length (e.g. diameter of a well) in the movie window.</li>
		<li>Open the &#8220;Set Scale&#8221; menu (Analyze &#62; Set Scale). Enter the known length in the &#8220;Known Distance&#8221; box and its unit (e.g. mm) in the &#8220;Unit of Length&#8221; box. Click the &#8220;Global&#8221; checkbox to use the scale for other video clips.</li>
		<li>Open the &#8220;Properties&#8221; menu (Image &#62; Properties) for time calibration. Enter frame interval (reciprocal of the frame rate) in the &#8220;Frame interval&#8221; box.</li>
	</ol>
	</li>
	<li>Configure tracking with MTrackJ.
	<ol>
		<li>Open the MTrackJ plugin from Plugins &#62; MtrackJ on ImageJ. Click the &#8220;Tracking&#8221; button in the MTrackJ dialog window to open a tracking configuration menu.</li>
		<li>Set a tracking interval by checking &#8220;Move to next time index after adding point&#8221; and entering a number in &#8220;Time step size&#8221; box in the configuration dialog window. To perform frame-by-frame tracking, enter &#8220;1&#8221; in the &#8220;Time step size&#8221; box.</li>
		<li>Check &#8220;Apply local cursor snapping during tracking&#8221; and choose &#8220;Dark centroid&#8221; 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 (&#8220;snap range&#8221;) 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).</li>
		<li>Click &#8220;OK&#8221; to apply the chosen options.</li>
	</ol>
	</li>
	<li>Perform two-dimensional tracking.
	<ol>
		<li>Click the &#8220;Add&#8221; button in the MTrackJ dialog window to start tracking.</li>
		<li>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.</li>
		<li>Repeat step 4.6.2 for a desired number of frames.</li>
		<li>To generate data tables, click &#8220;Measure&#8221; button in the MTrackJ dialog window. The data is displayed in two windows: &#8220;MTrackJ: Tracks&#8221; window shows a summary of the tracked two-dimensional trajectory and &#8220;MTrackJ: Points&#8221; shows detailed data for each time point. To save each table, choose File &#62; Save as&#8230; 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.</li>
		<li>To generate a movie with a tracked trajectory drawn as a colored line, click the &#8220;Movie&#8221; button in the MTrackJ dialog window. To save the generated movie, choose File &#62; Save as&#8230; menu on ImageJ.</li>
	</ol>
	</li>
	<li>Save the whole result. Click the &#8220;Save&#8221; 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).</li>
</ol>

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J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Seasonal+abundance+and+population+flux+of+Tigriopus+californicus+(Copepoda+:+Harpacticoida)+in+Barkley+Sound,+British+Columbia.">Seasonal abundance and population flux of Tigriopus californicus (Copepoda : Harpacticoida) in Barkley Sound, British Columbia.</a> Journal of the Marine Biological Association of the United Kingdom. 78 (2), 467-481 (1998).</li><li>Lard, M., Backman, J., Yakovleva, M., Danielsson, B., Hansson, L. A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Tracking+the+small+with+the+smallest+-+Using+nanotechnology+in+tracking+zooplankton.">Tracking the small with the smallest - Using nanotechnology in tracking zooplankton.</a> PLoS ONE. 5 (10), e13516 (2010).</li></ol>

The authors have nothing to disclose.

Individual culturing and determination of stage and sex
Here we described the method used in our previous study5 to prepare virgin Tigriopus animals with their pairing experience controlled while tracking their development (Figure 4 and Figure 7). As Tigriopus species are utilized as model animals in various biological fields such as toxicology16,18, ecological physiology19,20,21, and evolutionary genetics13,22,23,24, this method has a potential to provide a valuable means to assess influence of environmental and genetic factors on the life cycle of these copepods.
To achieve successful staging, regular search and collection of CI copepodids from a mass culture (step 1.2) is critical, as collection at later stages may result in mis-staging of the animals. In addition, a thorough search for exuviae (step 1.3) is also essential for accurate staging. Increase the frequency of collection and staging if necessary, as the interval between molts varies from one to several days depending on species and rearing condition2,25,26. Differences in antennae morphology between copepodids and adult females are not visibly significant in some species and populations of Tigriopus16,25. Hence, staging prior to sexing is helpful to distinguish adult females from advanced copepodids of either sex.
Behavioral tests and manual analysis of behavioral properties
In general, one of the most critical parts of ethological studies is definition and description of events of interest. The methods introduced in this paper were first developed for our recent study5 and supplemented with the description of copulation and the visual aids (Figure 10). In addition to that, consistency in animal handling also plays an important role in behavioral experiments. For example, rinsing of copepods may potentially facilitate some aspects of their behavior (Supplemental Figure 1) and therefore is desirable to be performed in a consistent manner among samples, such as standardized in step 2.1.3. We expect the materials provided in this paper will assist controlled and reproducible studies on the mate-guarding behavior of Tigriopus, promoting reproductive and ecological studies of this abundant inhabitant of high tide pools.
One possible limitation with this method is low magnification of obtained images. Although movies recorded with our system allow identification of prominent body structures including urosomes and male first antennae, one may not be able to observe more subtle structures such as legs and genitalia with our method since it does not employ microscopic magnification for video recording. While Kelly et al. reported that they were able to observe spermatophore transfer from males to females of T. japonicus under a microscopic observation at 100X magnification (no video recorded)7, we have not been able to observe a spermatophore in our movies, perhaps due to the limitation of the image resolution.
Two-dimensional tracking of pairs
Although this method does not allow three-dimensional tracking of animals, it enables two-dimensional trajectory analysis of zooplankton without chemically labeling animals (cf. Lard et al. 201027) by utilizing programs distributed for free. If the size of a movie file is too large to be processed in ImageJ, one can reduce resolution of the file and convert it into a gray scale movie. While the described method was originally developed for adult pairs of T. californicus (Figure 12 and Figure 13), it is also available for adult-juvenile pairs and single individuals (Supplemental Figure 1) as well as other Tigriopus species in principle. We further expect the method to be applicable to short-term (from the millisecond to the second time scales) trajectory analysis of zooplankton of other taxa with appropriate adjustments.

Intertidal copepods of the genus Tigriopus are widely distributed in rocky high intertidal pools across several continents1. These copepods exhibit mate-guarding behavior as a part of their reproduction, where an adult male captures a potential mate (juvenile or adult) utilizing its hooked first antennae prior to copulation (Figure 1 and Figure 2)2,3,4,5. Although this phenomenon has been a subject of ethological and biochemical studies for decades2,3,6,7, detailed procedures for studies of this behavior, including individual culture of virgin animals and criteria of behavioral events seen in a mate-guarding attempt, have not been well described. Thus, here we establish methods to enable studies of the behavior under a controlled experimental environment.
Individual culturing and staging of animals
Previous studies of reproduction of Tigriopus conventionally employed a pair detaching method to prepare juveniles (copepodids) and adult females for behavioral tests and breeding experiments3,8,9,10. However, this method allows animals to form pairs and potentially to copulate prior to tests (discussed in Ito 198811), which may alter behavioral properties of animals5. In addition, there is also a potential to misjudge developmental stages of copepods with the conventional protocols as they rely on apparent body size for staging. In this paper, we describe an individual culturing method used in our recent study with Tigriopus californicus5, which was designed to address these limitations by controlling pairing experience of animals and tracking their development from copepodid to adult stages.
Quantitative analysis of mate-guarding behavior
The mate-guarding behavior of Tigriopus species has been studied not only in the field of ethology2,6 but also in other fields such as ecotoxicology and evolutionary genetics3,4,7,8,9,10. However, the previous studies have explained the course of this behavior mainly in written forms without sufficient visual illustrations to outline both the behavior and the methods to study it, which creates technical obstacles for the replication and advancement of the studies. Here, we provide detailed descriptions of some of the key events in the mate-guarding behavior of Tigriopus copepods supported by visual materials. We also demonstrate equipment and methods for quantitative analysis of the behavior. These methods allow evaluation of behavioral properties of animals during mate-guarding attempts in precisely replicated experiments.
With these methods we aim to provide a methodological basis of controlled and reproducible studies on the mate-guarding behavior of the genus Tigriopus.

<table><tbody><tr><td>Instant Ocean Sea Salt</td><td>Spectrum Brands. Inc.</td><td>SS1-160P</td><td>For preparation of culture medium</td></tr><tr><td>PRO PlecoWafers</td><td>Tetra</td><td>16447</td><td>Food for copepods (used after being ground in a mortar)</td></tr><tr><td>Flat bottom 6-well tissue culture plate with lid</td><td>Corning Co., Ltd.</td><td>353224</td><td>Container for culture of gravid females and hatched nauplii</td></tr><tr><td>Flat bottom 24-well cell culture plate with lid</td><td>Corning Co., Ltd.</td><td>353226</td><td>Container for individual culturing</td></tr><tr><td>Flat bottom 48-well cell culture plate with lid</td><td>Nest Biotechnology Co., Ltd.</td><td>748001</td><td>Behavioral observation chambers</td></tr><tr><td>LED light pad</td><td>Shenzhen Huion Animation Technology Co., Ltd.</td><td>Litup LP4</td><td>Backlight for behavioral observation</td></tr><tr><td>Camera</td><td>Canon</td><td>0591C003 (model: Rebel T6i)</td><td>For recording of behavior</td></tr><tr><td>Pasteur pipette</td><td>Fisher Scientific</td><td>13-678-6A</td><td>For transfer of copepods</td></tr><tr><td>P10 micropipette tips</td><td>VWR</td><td>613-0735</td><td>For transfer of C1 stage copepodids</td></tr><tr><td>ImageJ</td><td>NIH</td><td>Version 1.49t</td><td>For semi-automatic analysis of movies</td></tr><tr><td>MTrackJ</td><td></td><td>Version 1.5.1</td><td>ImageJ plugin for tracking developed by Dr. Erik Meijering (Biomedical Imaging Group Rotterdam, Erasmus University Medical Center, The Netherlands)</td></tr></tbody></table>

individual culturing of tigriopus copepods and quantitative analysis of their mate-guarding behavior

Copepods of the genus Tigriopus, which are common zooplankton in rocky tide pools, show precopulatory mate-guarding behavior where a male clasps a potential mate to form a pair. While this phenomenon has attracted interest of researchers, methods for its analysis have not been well described. Here we describe procedures for: 1) individual culturing and staging of Tigriopus juveniles and adults, and 2) video-based analysis of their mate-guarding behavior. The culturing method enables experimental control of paring experience of animals as well as the ability to track their development before behavioral tests. The analysis method allows quantitative evaluation of several aspects of the mate-guarding behavior, including capturing attempts by males and swimming trajectory of mate-guarding pairs. Although these methods were originally established for ethological studies on Tigriopus, with proper modifications they can also be applied to studies of other zooplankton in different research fields, such as physiology, toxicology, and ecological genetics.

The individual culturing method described in step 1 allows preparation and staging of virgin animals with no prior experience of pairing.
The behavioral test described in step 2 allows video recording and observation of the mate-guarding behavior of Tigriopus copepods. The following examination of the recorded video with the methods described in steps 3 and 4 enables quantitative analysis of several aspects of the behavior shown in Figure 1.
Figure 11 shows a difference in average duration of guarding attempts between male-female pairs and male-male pairs of T. californicus. A manual analysis demonstrated that male-male pairs showed relatively shorter duration of pairing than male-female pairs.
Examples of trajectories tracked with the analysis method are presented in Figure 12 and a representative result of the velocity analysis is shown in Figure 13. Two-dimensional spatial tracking of newly formed guarding pairs of T. californicus revealed that male-male pairs tended to show higher velocity than male-female pairs in the first 3 s of guarding attempts.
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/58378/58378fig1.jpg" /> 
Figure 1: Mate-guarding behavior in Tigriopus. (A) An adult male clasping a juvenile (copepodid) with the first antennae (indicated by blue arrows). Bar = 1 mm. (B) An outline of mate-guarding behavior. A male tries to capture a target individual (left) and forms a pair with it (middle). In male-male pairs and some male-female pairs, a guarding attempt terminates without copulation. This figure has been modified from Tsuboko-Ishii and Burton 20175. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/58378/58378fig1large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 2" class="xfigimg" src="/files/ftp_upload/58378/58378fig2.jpg" /> 
Figure 2: Developmental stages of Tigriopus. Tigriopus species generally undergo six nauplius stages (from NI to NVI), five copepodid stages (from CI to CV), and an adult stage (CVI)16. Males make guarding attempts to juveniles from an early stage of copepodids (CI in T. japonicus and T. fulvus6,17 and CII in T. californicus3), as well as to adults of both sexes5. This figure has been modified from Tsuboko-Ishii and Burton 20175. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/58378/58378fig2large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 3" class="xfigimg" src="/files/ftp_upload/58378/58378fig3.jpg" /> 
Figure 3: Morphology of adults (T. californicus). (A) Adult male. (B) Adult female. (C) Gravid adult female with an egg sac with fertilized and developed (clear orange) eggs. (D) Gravid adult female with an egg sac with unfertilized or undeveloped (dark green) eggs. Arrows indicate egg sacs. Bar = 1 mm. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/58378/58378fig3large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 4" class="xfigimg" src="/files/ftp_upload/58378/58378fig4.jpg" /> 
Figure 4: Outline of preparation for individual culture. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/58378/58378fig4large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 5" class="xfigimg" src="/files/ftp_upload/58378/58378fig5.jpg" /> 
Figure 5: Molted exuviae. (A) Exuviae from CI to CV stages of a single animal of T. californicus. Bar = 1 mm. (B) Exuviae from CI to CV stages in an individual culture well. White debris is excrement of an animal cultured in the well (not shown in the image). <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/58378/58378fig5large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 6" class="xfigimg" src="/files/ftp_upload/58378/58378fig6.jpg" /> 
Figure 6: Search for exuviae under a stereomicroscope. Bars = 1 mm. (A) Focal change of a stereomicroscope for detection of exuviae at different depths. Magenta arrows indicate focused exuviae and gray arrows indicate unfocused exuviae. Top image focuses on the left exuviae, which is sunken at the bottom of the well. Bottom image focuses on the right exuviae, which is floating beneath the medium surface. (B) Top image shows an example of obstruction of examination by debris floating on the medium surface. A green arrow indicates an exuviae hidden beneath the debris. Bottom image shows a result of surface cleaning with a small piece of paper towel. An exuviae (indicated by a green arrow) is visible after the cleaning. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/58378/58378fig6large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 7" class="xfigimg" src="/files/ftp_upload/58378/58378fig7.jpg" /> 
Figure 7: Staging and sexing of animals. Examples of how to mark number of exuviae and sex of animals on a lid of a culture plate. (A) An example for a well containing five exuviae and an adult animal. The number of the lines in the tally mark on the lid represents the number of the exuviae found in the well. When the number of exuviae reaches five, the sex of the animal can be determined based on antennae morphology (see also Figure 3). (B) An example of a marked lid of a culture plate. Top rows contain older (CIV to adult) animals and bottom rows contain younger (i.e., newly collected) animals (CI to CIII). <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/58378/58378fig7large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 8" class="xfigimg" src="/files/ftp_upload/58378/58378fig8.jpg" /> 
Figure 8: Behavioral test scheme. Setup for video recording of mate-guarding behavior (left) and an outline of behavioral test (right). This figure has been modified from Tsuboko-Ishii and Burton 20175. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/58378/58378fig8large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 9" class="xfigimg" src="/files/ftp_upload/58378/58378fig9.jpg" /> 
Figure 9: Rinse of animals prior to a behavioral test. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/58378/58378fig9large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 10" class="xfigimg" src="/files/ftp_upload/58378/58378fig10.jpg" /> 
Figure 10: Definition of events examined in the manual analysis. Illustrations of events observed in relation to mate-guarding attempt. Names of the events defined and analyzed in step 3 are highlighted. Events enclosed in a dotted line may not be observed in some mate-guarding attempts. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/58378/58378fig10large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 11" class="xfigimg" src="/files/ftp_upload/58378/58378fig11.jpg" /> 
Figure 11: Difference in guarding duration between male-female pairs and male-male pairs of T. californicus. Each triangle symbol represents data from one tested pair. Bars and whiskers represent medians and interquartile range respectively. Average duration of capture was greater for male-female pairs (male-female (n=22), male-male (n=29); **p &#60; 0.01 by Mann-Whitney U test). <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/58378/58378fig11large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 12" class="xfigimg" src="/files/ftp_upload/58378/58378fig12.jpg" /> 
Figure 12: Trajectories of pairs in the first three seconds of guarding attempts. Examples of tracked two-dimensional trajectories of male-female pairs (left) and male-male pairs (right) of T. californicus. Dots on the trajectories represent time points (30 frames per second). Bars = 10 mm. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/58378/58378fig12large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 13" class="xfigimg" src="/files/ftp_upload/58378/58378fig13.jpg" /> 
Figure 13: Difference in mean velocity after initiation of guarding between male-female pairs and male-male pairs of T. californicus. Each triangle symbol represents data from one tested pair. Bars and whiskers represent medians and interquartile range respectively. Average velocity of the pair in the first 3 s of guarding attempts was greater for male-male pairs (male-female (n=13), male-male (n=35). ***p &#60; 0.001 by Mann-Whitney U test). <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/58378/58378fig13large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Suppl Figure 1" src="/files/ftp_upload/58378/58378sfig1.jpg" /> 
Supplemental Figure 1: Effect of rinsing treatment on behavior of Tigriopus. Each circle or triangle symbol represents data from one tested individual or pair. Bars and whiskers represent medians and interquartile range respectively. Individuals in &#34;rinsed&#34; and &#34;not rinsed&#34; groups were handled in the same manner, except that the &#34;not rinsed&#34; group did not experience the rinsing treatment (step 2.1.3) before the 30-minute adjustment time (step 2.1.4). (A) Average velocity of rinsed males tended to be greater than that of not rinsed males (rinsed (n=6), not rinsed (n=6), n.s.: no significant difference was detected by Mann-Whitney U test). The velocity was measured for 30 s based on videos recorded after the adjustment time. (B) Average velocity of rinsed females tended to be greater than that of not rinsed females (rinsed (n=6), not rinsed (n=6), n.s.: no significant difference was detected by Mann-Whitney U test). The velocity was measured for 30 s based on videos recorded after the adjustment time, following step 4 (tracking interval = 0.5 s). (C) Frequency of guarding attempts was greater for pairs of rinsed individuals (rinsed (n=6), not rinsed (n=6). ** p &#60; 0.01 by Mann-Whitney U test). (D) Duration of guarding attempts tended to be greater for pairs of rinsed individuals (rinsed (n=6), not rinsed (n=6), n.s.: no significant difference was detected by Mann-Whitney U test). <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/58378/58378sfig1large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

Watch this Scientific Journal Video about Individual Culturing of Tigriopus Copepods and Quantitative Analysis of Their Mate-guarding Behavior at JoVE.com

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

Mate-guarding behavior plays an important role in reproduction of intertidal copepods of the genus Tigriopus. However, methods for studying this behavior have not been well described. Here we describe methods for: 1) individual culture of virgin Tigriopus animals, and 2) quantitative analysis of their mate-guarding behavior.

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

Copepods of the genus Tigriopus, which are common zooplankton in rocky tide pools, show precopulatory mate-guarding behavior where a male clasps a ...

individual-culturing-tigriopus-copepods-quantitative-analysis-their

Nanyang Technological University

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Environment

9.5K Views.  University of California, San Diego. Het belangrijkste voordeel van deze kweektechniek is dat je maagdelijke roeipootkreeften verkrijgen terwijl je hun koppelingservaring controleert en hun ontwikkeling volgt. De volgende gedragsanalyse methode kan helpen gecontroleerde en reproduceerbare experimenten op de mate-guarding gedrag van de Tigriopus roeipootkreeften die gemeenschappelijk zoöplankton in rotsachtige getijdenpoelen. Hoewel deze methoden inzicht kunnen geven in het voortplantingsgedrag van Tigriopus, kunnen ze ook worde...

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

Assaying Predatory Feeding Behaviors in Pristionchus and Other Nematodes

This protocol provides multiple methods for the analysis and quantification of predatory feeding behaviors in nematodes. Many nematode species including Pristionchus pacificus display complex behaviors, the most striking of which is the predation of other nematode larvae. However, as these behaviors are absent in the model organism Caenorhabditis elegans, they have thus far only recently been described in detail along with the development of reliable behavioral assays 1. These predatory behaviors are dependent upon phenotypically plastic but fixed mouth morphs making the correct identification and categorization of these animals essential. In P. pacificus there are two mouth types, the stenostomatous and eurystomatous morphs 2, with only the wide mouthed eurystomatous containing an extra tooth and being capable of killing other nematode larvae. Through the isolation of an abundance of size matched prey larvae and subsequent exposure to predatory nematodes, assays including both "corpse assays" and "bite assays" on correctly identified mouth morph nematodes are possible. These assays provide a means to rapidly quantify predation success rates and provide a detailed behavioral analysis of individual nematodes engaged in predatory feeding activities. In addition, with the use of a high-speed camera, visualization of changes in pharyngeal activity including tooth and pumping dynamics are also possible.

Assaying Predatory Feeding Behaviors in Pristionchus and Other Nematodes

Behavioral assays provide powerful tools for understanding neuronal function. Here we present several protocols for quantifying predatory feeding behavior found in the model nematode Pristionchus pacificus and its relatives. Additionally, we provide methods for analyzing predatory feeding adaptations including mouth structures and teeth.

This protocol provides multiple methods for the analysis and quantification of predatory feeding behaviors in nematodes. Many nematode species including ...

Fatty Acid 13C Isotopologue Profiling Provides Insight into Trophic Carbon Transfer and Lipid Metabolism of Invertebrate Consumers

Fatty acids (FAs) are useful biomarkers in food web ecology because they are typically assimilated as a complete molecule and transferred into consumer tissue with minor or no modification, allowing the dietary routing between different trophic levels. However, the FA trophic marker approach is still hampered by the limited knowledge in lipid metabolism of the soil fauna. This study used entirely labelled palmitic acid (13C16:0, 99 atom%) as a tracer in fatty acid metabolism pathways of two widespread soil Collembola, Protaphorura fimata and Heteromurus nitidus. In order to investigate the fate and metabolic modifications of this precursor, a method of isotopologue profiling is presented, performed by mass spectrometry using single ion monitoring. Moreover, the upstream laboratory feeding experiment is described, as well as the extraction and methylation of dominant lipid fractions (neutral lipids, phospholipids) and the related formula and calculations. Isotopologue profiling does not only yield the overall 13C enrichment in fatty acids derived from the 13C labeled precursor but also produces the pattern of isotopologues exceeding the mass of the parent ion (i.e., the FA molecular ion M+) of each labeled FA by one or more mass units (M+1, M+2, M+3, etc.). This knowledge allows conclusions on the ratio of dietary routing of an entirely consumed FA in comparison to de novo biosynthesis. The isotopologue profiling is suggested as a useful tool for evaluation of fatty acid metabolism in soil animals to disentangle trophic interactions.

Fatty Acid 13C Isotopologue Profiling Provides Insight into Trophic Carbon Transfer and Lipid Metabolism of Invertebrate Consumers

The fatty acid trophic marker approach, i.e., the assimilation of fatty acids as entire molecule and transfer into consumer tissue with no or minor modification, is hampered by knowledge gaps in fatty acid metabolism of small soil invertebrates. Isotopologue profiling is provided as a valuable tool to disentangle trophic interactions.

Fatty acids (FAs) are useful biomarkers in food web ecology because they are typically assimilated as a complete molecule and transferred into consumer ...

Biochemistry

Colonization of Euprymna scolopes Squid by Vibrio fischeri

Specific bacteria are found in association with animal tissue1-5. Such host-bacterial associations (symbioses) can be detrimental (pathogenic), have no fitness consequence (commensal), or be beneficial (mutualistic). While much attention has been given to pathogenic interactions, little is known about the processes that dictate the reproducible acquisition of beneficial/commensal bacteria from the environment. The light-organ mutualism between the marine Gram-negative bacterium V. fischeri and the Hawaiian bobtail squid, E. scolopes, represents a highly specific interaction in which one host (E. scolopes) establishes a symbiotic relationship with only one bacterial species (V. fischeri) throughout the course of its lifetime6,7. Bioluminescence produced by V. fischeri during this interaction provides an anti-predatory benefit to E. scolopes during nocturnal activities8,9, while the nutrient-rich host tissue provides V. fischeri with a protected niche10. During each host generation, this relationship is recapitulated, thus representing a predictable process that can be assessed in detail at various stages of symbiotic development. In the laboratory, the juvenile squid hatch aposymbiotically (uncolonized), and, if collected within the first 30-60 minutes and transferred to symbiont-free water, cannot be colonized except by the experimental inoculum6. This interaction thus provides a useful model system in which to assess the individual steps that lead to specific acquisition of a symbiotic microbe from the environment11,12.
Here we describe a method to assess the degree of colonization that occurs when newly hatched aposymbiotic E. scolopes are exposed to (artificial) seawater containing V. fischeri. This simple assay describes inoculation, natural infection, and recovery of the bacterial symbiont from the nascent light organ of E. scolopes. Care is taken to provide a consistent environment for the animals during symbiotic development, especially with regard to water quality and light cues. Methods to characterize the symbiotic population described include (1) measurement of bacterially-derived bioluminescence, and (2) direct colony counting of recovered symbionts.

Colonization of Euprymna scolopes Squid by Vibrio fischeri

The method outlines the procedure by which the Hawaiian bobtail squid, Euprymna scolopes and its bacterial symbiont, Vibrio fischeri, are raised separately and then introduced to allow for specific colonization of the squid light organ by the bacteria. Colonization detection by bacterially-derived luminescence and by direct colony counting are described.

Specific bacteria are found in association with animal tissue1-5. Such host-bacterial associations (symbioses) can be detrimental (pathogenic), have no ...

Immunology and Infection

Automated Analysis of a Nematode Population-based Chemosensory Preference Assay

The nematode, Caenorhabditis elegans' compact nervous system of only 302 neurons underlies a diverse repertoire of behaviors. To facilitate the dissection of the neural circuits underlying these behaviors, the development of robust and reproducible behavioral assays is necessary. Previous C. elegans behavioral studies have used variations of a "drop test", a "chemotaxis assay", and a "retention assay" to investigate the response of C. elegans to soluble compounds. The method described in this article seeks to combine the complementary strengths of the three aforementioned assays. Briefly, a small circle in the middle of each assay plate is divided into four quadrants with the control and experimental solutions alternately placed. After the addition of the worms, the assay plates are loaded into a behavior chamber where microscope cameras record the worms' encounters with the treated regions. Automated video analysis is then performed and a preference index (PI) value for each video is generated. The video acquisition and automated analysis features of this method minimizes the experimenter's involvement and any associated errors. Furthermore, minute amounts of the experimental compound are used per assay and the behavior chamber's multi-camera setup increases experimental throughput. This method is particularly useful for conducting behavioral screens of genetic mutants and novel chemical compounds. However, this method is not appropriate for studying stimulus gradient navigation due to the close proximity of the control and experimental solution regions. It should also not be used when only a small population of worms is available. While suitable for assaying responses only to soluble compounds in its current form, this method can be easily modified to accommodate multimodal sensory interaction and optogenetic studies. This method can also be adapted to assay the chemosensory responses of other nematode species.

We present a behavior recording setup and protocol that enables automated analysis of the nematode, Caenorhabditis elegans' preference for soluble compounds in a population-based assay. This article describes the construction of a behavior chamber, the behavioral assay protocol, and video analysis software usage.

The nematode, Caenorhabditis elegans' compact nervous system of only 302 neurons underlies a diverse repertoire of behaviors. To facilitate the dissection ...

Behavior

Cultivation of the Marine Pelagic Tunicate Dolioletta gegenbauri (Uljanin 1884) for Experimental Studies

Gelatinous zooplanktons play a crucial role in ocean ecosystems. However, it is generally difficult to investigate their physiology, growth, fecundity, and trophic interactions primarily due to methodological challenges, including the ability to culture them. This is particularly true for the doliolid, Dolioletta gegenbauri. D. gegenbauri commonly occurs in productive subtropical continental shelf systems worldwide, often at bloom concentrations capable of consuming a large fraction of daily primary production. In this study, we describe cultivation approaches for collecting, rearing, and maintaining D. gegenbauri for the purpose of conducting laboratory-based studies. D. gegenbauri and other doliolid species can be captured live using obliquely towed conical 202 &micro;m mesh plankton nets from a drifting ship. Cultures are most reliably established when water temperatures are below 21 &deg;C and are started from immature gonozooids, maturing phorozooids, and large nurses. Cultures can be maintained in rounded culture vessels on a slowly rotating plankton wheel and sustained on a diet of cultured algae in natural seawater for many generations. In addition to the ability to establish laboratory cultures of D. gegenbauri, we demonstrate that the collection condition, algae concentration, temperature, and exposure to naturally conditioned seawater are all critical to the culture establishment, growth, survival, and reproduction of D. gegenbauri.

Cultivation of the Marine Pelagic Tunicate Dolioletta gegenbauri Uljanin 1884 for Experimental Studies

Doliolids, including the species Dolioletta gegenbauri, are small gelatinous marine zooplankton of ecological significance found on productive subcontinental shelf systems worldwide. The difficulty of culturing these delicate organisms limits their investigation. In this study, we describe cultivation approaches for collecting, rearing, and maintaining the doliolid Dolioletta gegenbauri.

Gelatinous zooplanktons play a crucial role in ocean ecosystems. However, it is generally difficult to investigate their physiology, growth, fecundity, ...

Culture and Assay of Large-Scale Mixed-Stage Caenorhabditis elegans Populations

Caenorhabditis elegans (C. elegans) has been and remains a valuable model organism to study developmental biology, aging, neurobiology, and genetics. The large body of work on C. elegans makes it an ideal candidate to integrate into large-population, whole-animal studies to dissect the complex biological components and their relationships with another organism. In order to use C. elegans in collaborative -omics research, a method is needed to generate large populations of animals where a single sample can be split and assayed across diverse platforms for comparative analyses.
Here, a method to culture and collect an abundant mixed-stage C. elegans population on a large-scale culture plate (LSCP) and subsequent phenotypic data is presented. This pipeline yields sufficient numbers of animals to collect phenotypic and population data, along with any data needed for -omics experiments (i.e., genomics, transcriptomics, proteomics, and metabolomics). In addition, the LSCP method requires minimal manipulation to the animals themselves, less user preparation time, provides tight environmental control, and ensures that handling of each sample is consistent throughout the study for overall reproducibility. Lastly, methods to document population size and population distribution of C. elegans life stages in a given LSCP are presented.

Culture and Assay of Large-Scale Mixed-Stage Caenorhabditis elegans Populations

To use Caenorhabditis elegans (C. elegans) in omics research, a method is needed to generate large populations of worms where a single sample can be measured across platforms for comparative analyses. Here, a method to culture C. elegans populations on large-scale culture plates (LSCPs) and to document population growth is presented.

Caenorhabditis elegans (C. elegans) has been and remains a valuable model organism to study developmental biology, aging, neurobiology, and genetics. The ...

Genetics

Caenorhabditis Sieve: A Low-tech Instrument and Methodology for Sorting Small Multicellular Organisms

Caenorhabditis elegans (C. elegans) is a well-established model organism used across a range of basic and biomedical research. Within the nematode research community, there is a need for an affordable and effective way to maintain large, age-matched populations of C. elegans. Here, we present a methodology for mechanically sorting and cleaning C. elegans. Our aim is to provide a cost-effective, efficient, fast, and simple process to obtain animals of uniform sizes and life stages for their use in experiments. This tool, the Caenorhabditis Sieve, uses a custom-built lid system that threads onto common conical lab tubes and sorts C. elegans based on body size. We also demonstrate that the Caenorhabditis Sieve effectively transfers animals from one culture plate to another allowing for a rapid sorting, synchronizing, and cleaning without impacting markers of health, including motility and stress-inducible gene reporters. This accessible and innovative tool is a fast, efficient, and non-stressful option for maintaining C. elegans populations.

Caenorhabditis Sieve: A Low-tech Instrument and Methodology for Sorting Small Multicellular Organisms

The current protocol includes a methodology for the sorting and cleaning of age-matched populations of Caenorhabditis elegans. It uses a simple, inexpensive, and efficient custom-made tool to obtain a large experimental population of nematodes for research.

Caenorhabditis elegans (C. elegans) is a well-established model organism used across a range of basic and biomedical research. Within the nematode ...

Developmental Biology

Using Coculture to Detect Chemically Mediated Interspecies Interactions

In nature, bacteria rarely exist in isolation; they are instead surrounded by a diverse array of other microorganisms that alter the local environment by secreting metabolites. These metabolites have the potential to modulate the physiology and differentiation of their microbial neighbors and are likely important factors in the establishment and maintenance of complex microbial communities. We have developed a fluorescence-based coculture screen to identify such chemically mediated microbial interactions. The screen involves combining a fluorescent transcriptional reporter strain with environmental microbes on solid media and allowing the colonies to grow in coculture. The fluorescent transcriptional reporter is designed so that the chosen bacterial strain fluoresces when it is expressing a particular phenotype of interest (i.e. biofilm formation, sporulation, virulence factor production, etc.) Screening is performed under growth conditions where this phenotype is not expressed (and therefore the reporter strain is typically nonfluorescent). When an environmental microbe secretes a metabolite that activates this phenotype, it diffuses through the agar and activates the fluorescent reporter construct. This allows the inducing-metabolite-producing microbe to be detected: they are the nonfluorescent colonies most proximal to the fluorescent colonies. Thus, this screen allows the identification of environmental microbes that produce diffusible metabolites that activate a particular physiological response in a reporter strain. This publication discusses how to: a) select appropriate coculture screening conditions, b) prepare the reporter and environmental microbes for screening, c) perform the coculture screen, d) isolate putative inducing organisms, and e) confirm their activity in a secondary screen. We developed this method to screen for soil organisms that activate biofilm matrix-production in Bacillus subtilis; however, we also discuss considerations for applying this approach to other genetically tractable bacteria.

Bacteria produce secreted compounds that have the potential to affect the physiology of their microbial neighbors. Here we describe a coculture screen that allows detection of such chemically mediated interspecies interactions by mixing soil microbes with fluorescent transcriptional reporter strains of Bacillus subtilis on solid media.

In nature, bacteria rarely exist in isolation; they are instead surrounded by a diverse array of other microorganisms that alter the local environment by ...

Biology

Maintaining Laboratory Cultures of Gryllus bimaculatus, a Versatile Orthopteran Model for Insect Agriculture and Invertebrate Physiology

Gryllus bimaculatus (De Geer)&#160;is a large-bodied cricket distributed throughout Africa and Southern Eurasia where it is often wild-harvested as human food. Outside its native range, culturing G. bimaculatus is feasible due to its dietary plasticity, rapid reproductive cycle, lack of diapause requirement, tolerance for high-density rearing, and robustness against pathogens. Thus, G. bimaculatus can be&#160;a versatile model for studies of insect physiology, behavior, embryology, or genetics.
Cultural parameters, such as stocking density, within-cage refugia, photoperiod, temperature, relative humidity, and diet, all impact cricket growth, behavior, and gene expression and should be standardized. In the burgeoning literature on farming insects for human consumption, these crickets are frequently employed to evaluate candidate feed admixtures derived from crop residues, food-processing byproducts, and other low-cost waste streams.
To support ongoing experiments evaluating G. bimaculatus growth performance and nutritional quality in response to variable feed substrates, a comprehensive set of standard protocols for breeding, upkeep, handling, measurement, and euthanasia in the laboratory was developed and is presented here. An industry-standard cricket feed has proven nutritionally adequate and functionally appropriate for the long-term maintenance of cricket breeding stocks, as well as for use as an experimental control feed. Rearing these crickets at a density of 0.005 crickets/cm3 in screen-topped 29.3 L polyethylene cages at an average temperature of 27 &#176;C on a 12 light (L)/12 dark (D) photoperiod, with moistened coconut coir serving both as hydration source and oviposition medium has successfully sustained healthy crickets over a 2-year span. Following these methods, crickets in a controlled experiment yielded an average mass of 0.724 g 0.190 g at harvest, with 89% survivorship and 68.2% sexual maturation between stocking (22 days) and harvest (65 days).

Maintaining Laboratory Cultures of Gryllus bimaculatus, a Versatile Orthopteran Model for Insect Agriculture and Invertebrate Physiology

This paper outlines basic methods to standardize important factors such as density, feed availability, hydration source, and environmental controls for the long-term rearing of laboratory cultures of the edible cricket, Gryllus bimaculatus.

Gryllus bimaculatus (De Geer)&#160;is a large-bodied cricket distributed throughout Africa and Southern Eurasia where it is often wild-harvested as human ...

Ex Situ Culture of Pocillopora acuta and Effective Feeding Techniques

Effective Techniques for the Feeding and Ex Situ Culture of a Brooding Scleractinian Coral, Pocillopora acuta

Climate change is affecting the survival, growth, and recruitment of corals globally, with large-scale shifts in abundance and community composition expected in reef ecosystems over the next several decades. Recognition of this reef degradation has prompted a range of novel research- and restoration-based active interventions. Ex situ aquaculture can play a supporting role through the establishment of robust coral culture protocols (e.g., to improve health and reproduction in long-term experiments) and through the provision of a consistent broodstock supply (e.g., for use in restoration projects). Here, simple techniques for the feeding and ex situ culture of brooding scleractinian corals are outlined using the common and well-studied coral, Pocillopora acuta, as an example. To demonstrate this approach, coral colonies were exposed to different temperatures (24 &#176;C vs. 28 &#176;C) and feeding treatments (fed vs. unfed) and the reproductive output and timing, as well as the feasibility of feeding Artemia nauplii to corals at both temperatures, was compared. Reproductive output showed high variation across colonies, with differing trends observed between the temperature treatments; at 24 &#176;C, fed colonies produced more larvae than unfed colonies, but the opposite was found in colonies cultured at 28 &#176;C. All colonies reproduced before the full moon, and differences in reproductive timing were only found between unfed colonies in the 28 &#176;C treatment and fed colonies in the 24 &#176;C treatment (mean lunar day of reproduction &#177; standard deviation: 6.5 &#177; 2.5 and 11.1 &#177; 2.6, respectively). The coral colonies fed efficiently on Artemia nauplii at both treatment temperatures. These proposed feeding and culture techniques focus on the reduction of coral stress and the promotion of reproductive longevity in a cost-effective and customizable manner, with versatile applicability in both flow-through and recirculating aquaculture systems.

Author Spotlight: Advancing Coral Research by Exploring Climate Change Resistance, Ex Situ Aquaculture, and Reproduction Strategies 

Climate change is impacting coral reef ecosystems globally. Corals sourced from ex situ aquaculture systems can help support restoration and research efforts. Herein, feeding and coral culture techniques that can be used to promote the long-term maintenance of brooding scleractinian corals ex situ are outlined.

Climate change is affecting the survival, growth, and recruitment of corals globally, with large-scale shifts in abundance and community composition ...

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

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