This research was funded by the Ministry of Science and Technology (Taiwan), grant numbers MOST 111-2611-M-291-005 and MOST 111-2811-M-291-001.

1. Hanging coral colonies in&#160;ex situ aquaculture tanks
<ol>
	<li>Position a notched bar (length x width x height: 75 cm x 1 cm x 3 cm), hereafter referred to as a &#34;hanging bar&#34;, across the culture tank in preparation for hanging the coral colonies. 
	NOTE: The hanging bar used in this experiment was custom-made, but a simple PVC pipe with protruding screws (i.e., to act as the notches) would be sufficient as long as it can be positioned in a stable manner across the top of the culture tank and is strong enough to hold the corals.</li>
	<li>Measure a piece of fishing line (see the Table of Materials) to ~1.5 m in length, and then fold it in half twice. 
	NOTE: The initial length of the fishing line should be chosen based on the desired final position of the coral colony in the culture tank.</li>
	<li>Make a small overhand knot at the end of the folded fishing line that has the initial ends of the fishing line. 
	NOTE: After making the knot, there should be two large loops at the bottom and one small loop at the top.</li>
	<li>Place the coral colony in the middle of the two large loops such that the loops are positioned around the colony and can securely hold the coral when it is hung in the water.</li>
	<li>Hook the small top loop of the fishing line into a notch on the hanging bar (Figure 1B).</li>
</ol>
2. Coral feeding
<ol>
	<li>Making the feeding container
	<ol>
		<li>Construct a rectangular frame using acrylic pipe (length x width x height: 25 cm x 60 cm x 25 cm). Make two separate compartments in the frame where the fed and unfed corals can be placed, respectively (Figure 1C). 
		NOTE: Acrylic pipe was used because it is lightweight (i.e., as opposed to heavier PVC pipe) and, therefore, could facilitate easier movement of the feeding container in/out of the culture tanks.</li>
		<li>Use a hot glue gun to adhere 100 &#181;m of plankton mesh to the bottom and sides of the frame.</li>
		<li>Drill a total of ~10 small holes (0.5 cm in diameter) into the pipes (especially along the sides and the bottom of the frame) to prevent the feeding container from floating when placed in the culture tank.</li>
		<li>Drill holes (~0.5 cm in diameter) through the plankton mesh at each corner of the feeding container.</li>
		<li>Place an 8 cm length of 0.5 cm diameter tubing through the corner holes, and use a hot glue gun to fix it into position. 
		NOTE: These pieces of tubing will be connected to an air pump and bubble stones during feeding (see step 2.3.2 for more detail).</li>
	</ol>
	</li>
	<li>Artemia cultivation
	<ol>
		<li>Collect 2 L of seawater from an independent feeding tank, and pour the seawater into an Artemia hatching container (Figure 1D). 
		NOTE: In the present experiment used to demonstrate the protocols, two independent treatment-specific feeding tanks were used, which necessitated the preparation of two hatching containers for Artemia cultivation.</li>
		<li>Connect an air pump to tubing connected to the bottom of the hatching container for approximately 10 min prior to adding Artemia cysts.</li>
		<li>While waiting, use a balance to measure 8 g of Artemia cysts (see the Table of Materials). 
		NOTE: To obtain a mean density of 35 individual Artemia nauplii/mL, as suggested by Huang et al.19, use a ratio of 4 g of Artemia cysts to 1 L of seawater.</li>
		<li>After 10 min, pour the 8 g of Artemia cysts into the hatching container.</li>
		<li>Incubate the Artemia cysts for 48 h.</li>
	</ol>
	</li>
	<li>Preparing the feeding tank
	<ol>
		<li>Place the feeding container into the feeding tank such that the top of the container is above the surface of the water.</li>
		<li>Connect the outer portion of the feeding container&#39;s corner tubing to an air pump, which will supply air to bubble stones for facilitating water circulation during feeding.</li>
		<li>Turn on the air pump ~5 min prior to the start of feeding.</li>
	</ol>
	</li>
	<li>Artemia nauplii enrichment and collection
	<ol>
		<li>Add 1.5 mL of enrichment diet (see the Table of Materials) to the hatching container 2 h before the desired feeding time. 
		NOTE: A ratio of 0.75 mL of enrichment diet to 1 L of seawater is recommended by Huang et al.19.</li>
		<li>After 2 h, turn off the valve supplying air to the hatching container.</li>
		<li>Cover the hatching container with a cardboard box to exclude ambient light, and place a light source (a cell phone flashlight is sufficient) at the base of the hatching container for 5 min to attract Artemia nauplii to the bottom of the container and thereby facilitate the separation of live Artemia nauplii from empty shells.</li>
		<li>After 5 min, remove the box and the light source.</li>
		<li>Place a 3 L measuring jug below the hatching container.</li>
		<li>Detach the tubing from the hatching container to allow the Artemia nauplii and seawater solution to flow into the measuring jug; collect 1 L of the Artemia nauplii and seawater solution. 
		NOTE: Collect only half of the volume in the hatching container to exclude unwanted empty shells.</li>
		<li>While standing in close proximity to the feeding tank, pour the Artemia nauplii and seawater solution through a 100 &#181;m strainer to separate the Artemia nauplii (which will remain in the strainer) from the seawater.</li>
		<li>Rinse the Artemia nauplii held within the strainer twice with water from the feeding tank.</li>
		<li>The Artemia nauplii are now ready to be used.</li>
	</ol>
	</li>
	<li>Feeding the coral colonies
	<ol>
		<li>Unload the Artemia nauplii by placing the strainer from step 2.4.8 into the feeding tank.</li>
		<li>Stir the water in the tank by hand to evenly distribute the Artemia nauplii. 
		NOTE: Collect samples for the &#34;pre-feeding&#34; quantification of the Artemia nauplii density after this step (see step 3.1 for more detail).</li>
		<li>Move each hanging bar (with the coral colonies still hanging from the bar) from the culture tank to the feeding tank, and position the bar so that it is securely resting across the top of the feeding tank. The duration for which the corals are exposed to the air should be kept as short as possible. 
		NOTE: Make sure colonies are not touching each other and have enough space to capture food (e.g., ~5 cm apart).</li>
		<li>Turn off the lights in the feeding tanks, or use a non-airtight lid to cover the feeding tank to avoid light disturbance during feeding.</li>
		<li>Allow the colonies to feed undisturbed for 4 h.</li>
		<li>After 4 h, collect the samples for the &#34;post-feeding&#34; quantification of the Artemia nauplii density (see step 3.1 for more detail).</li>
	</ol>
	</li>
	<li>Post-feeding clean up
	<ol>
		<li>After the feeding session is complete, remove the coral colonies. Take the hanging bars out of the feeding tank individually, and thoroughly rinse each coral with seawater from its respective culture tank to remove any residual Artemia nauplii. 
		NOTE: Rinse the colonies on a stable surface rather than while hanging to reduce the risk of damage that could occur if the colonies were to swing back and forth during rinsing. As per the initial transfer, keep the duration for which the corals are exposed to the air as short as possible.</li>
		<li>Place the hanging bars (with corals hung) back into the culture tanks.</li>
		<li>Detach the tubes connecting the feeding container to the air pump, and remove the feeding container from the feeding tank.</li>
		<li>Rinse the feeding container thoroughly with fresh water to remove all the remaining Artemia nauplii.</li>
	</ol>
	</li>
</ol>
3. Quantifying Artemia nauplii density pre- and post-feeding
<ol>
	<li>Collecting the samples
	<ol>
		<li>Collect samples at two time points: first, when&#160;the Artemia nauplii have been unloaded and distributed evenly in the feeding container (step 2.5.2), and again after the feeding session has been completed (step 2.5.6).</li>
		<li>For each time point, use three syringes to draw 20 mL of water from the surface, the middle layer, and the bottom layer of the feeding container, respectively.</li>
	</ol>
	</li>
	<li>Sample dilution
	<ol>
		<li>For each syringe, transfer the 20 mL of water sample into an independent 500 mL beaker.</li>
		<li>Add 180 mL of hot water (~60 &#176;C) to the beaker (1:10 dilution). 
		NOTE: The hot water is used to immobilize the Artemia nauplii to increase the accuracy of enumeration.</li>
		<li>Add 2 mL of the water sample from the beaker into each well of a 9-well plate. 
		NOTE: Mix the sample in the beaker to distribute the Artemia nauplii evenly in the water column before drawing the 2 mL of sample.</li>
		<li>Count the number of Artemia nauplii in each well under a stereo microscope using 6.5x magnification (see the Table of Materials).</li>
	</ol>
	</li>
	<li>Calculating the density of Artemia nauplii
	<ol>
		<li>Divide the number of Artemia nauplii in each well by 2 to obtain the number of Artemia nauplii per mL. Then, multiply that number by 10 (to account for dilution) to calculate the Artemia nauplii density.</li>
		<li>Calculate the mean density of Artemia nauplii (i.e., average density across the 27 well replicates before vs. after feeding) to compare the Artemia nauplii density between pre- and post-feeding.</li>
	</ol>
	</li>
</ol>
4. Coral larvae collection
<ol>
	<li>Making the larvae collection container (Figure 1E)
	<ol>
		<li>Select a 6 L plastic water bottle, and cut the bottom of the bottle off completely. 
		NOTE: This opening will be used for transferring the colonies in and out of the larvae collection container.</li>
		<li>Create two windows by cutting out a ~15 cm x 20 cm rectangle from each side of the bottle. 
		NOTE: A 6 L plastic water bottle is appropriate for corals that are ~15 cm in diameter; modify the size of the bottle based on the size of the corals being studied.</li>
		<li>Use a hot glue gun and then epoxy to adhere a 100 &#181;m plankton mesh onto each of the windows.</li>
		<li>Create two small holes (~0.5 cm in diameter) on each side of the bottom of the bottle.</li>
		<li>Put a string through the two small holes, and tie both ends to create a handle for hooking the larvae collection container onto the hanging bar.</li>
		<li>Before initial use, place the bottles into a flow-through tank (with no corals) for at least 24 h to remove any glue residue.</li>
	</ol>
	</li>
	<li>Preparing for coral collection
	<ol>
		<li>Immerse the larvae collection container completely into the culture tank.</li>
		<li>Place the colony into the larvae collection container while keeping both the colony and the container submerged in water.</li>
		<li>Hook the handle of the larvae collection container onto the hanging bar. 
		NOTE: After hanging, ensure that the top of the collection container is ~3 cm above the water.</li>
		<li>Repeat steps 4.2.1-4.2.3 until all the colonies are in their larvae collection containers.</li>
	</ol>
	</li>
	<li>Collecting and enumerating the coral larvae
	<ol>
		<li>Prepare a 3 L measuring jug, a bowl, a 3 mL pipette, and 50 mL tubes.</li>
		<li>Unhook the fishing line from the hanging bar, and remove one colony from its larvae collection container. Place the colony back into the culture tank immediately. 
		NOTE: Ensure that the duration of air exposure is as short as possible.</li>
		<li>Place one hand on the cap end of the larvae collection container. 
		NOTE: When the larvae collection container is filled with water, it can be heavy. Without proper support, the container can break when it is being removed from the water.</li>
		<li>Unhook the larvae collection container &#34;handle&#34; from the hanging bar.</li>
		<li>Slowly lift the larvae collection container out of the water.</li>
		<li>Hold the collection container at an approximately 45&#176; angle above the culture tank for a few seconds to allow excess water to flow back into the tank via the larvae collection container windows. 
		NOTE: Do not angle the container past 45&#176; to mitigate the chance of pouring out larvae from the top of the container.</li>
		<li>Remove the larvae collection container from the tank, and position it on top of the measuring jug.</li>
		<li>Before unscrewing the cap, use one finger to apply a moderate amount of pressure against the cap, and then unscrew the cap. 
		NOTE: Water inside of the collection container can be released quickly when the cap is removed if it is not first supported by one&#39;s finger (i.e., potentially resulting in a loss of larvae).</li>
		<li>Transfer some of the water inside of the measuring jug into a bowl.</li>
		<li>Manually count the number of larvae in the bowl by using a 3 mL pipette to move the larvae into a 50 mL tube. 
		NOTE: Be aware that some of the larvae may get stuck inside the pipette. If this happens, draw some seawater into the pipette, and shake gently while sealing the pipette with one finger to loosen the larvae.</li>
		<li>Continue step 4.3.9 and step 4.3.10 until all the larvae have been counted. At this stage, the larvae can be used in subsequent experiments.</li>
		<li>Repeat steps 4.3.2-4.3.10 for all the other coral colonies. 
		NOTE: The measuring jug and bowl should be rinsed between colonies.</li>
		<li>After the counting is finished, rinse each collection container thoroughly with fresh water, especially the windows.</li>
	</ol>
	</li>
</ol>

Ex Situ Culture of Pocillopora acuta and Effective Feeding Techniques

The authors have no competing financial interests or other conflicts of interest.

This preliminary assessment of the effect of temperature and feeding on coral reproduction revealed differences in reproductive output and timing among colonies cultured under distinct treatment conditions. Further, it was found that feeding Artemia nauplii to coral colonies appeared to be effective at relatively cool (24&#176;C) as well as warm temperatures (28 &#176;C). These combined findings highlight the applicability of these straightforward techniques for the feeding and culture of reproducing scleractini...

Many coral reef ecosystems globally are being lost and degraded as a result of high-temperature stress driven by climate change1,2. Coral bleaching (i.e., the breakdown of the coral-algal symbiosis3) was considered relatively rare in the past4 but is now occurring more frequently5, with annual bleaching expected to occur in many regions by&#160;mid to late century6,7. This shortening of the interim period between bleaching events can limit the capacity for reef resilience8. The direct impacts of high-temperature stress on coral colonies (e.g., tissue damage9; energy depletion10) are intrinsically linked to indirect impacts at the reef-scale level, of which a reduction in reproductive/recruitment capacity is of particular concern11. This has spurred a range of applied research exploring, for example, the active in situ enhancement of recruitment (e.g., reef seeding12), new technologies for scaling-up coral restoration13, and the simulation of reproductive cues to induce reproduction in ex situ systems14. Complementary to these active interventions are the recent recognition of the advantages of heterotrophic feeding in corals under high-temperature stress15 and the exploration of the role that food provision may play in reproduction16.
Heterotrophic feeding is known to influence the performance of corals17 and has been specifically linked to increased coral growth18,19, as well as thermal resistance and resilience20,21. Yet, the benefits of heterotrophy are not ubiquitous among coral species22 and can differ based on the type of food being consumed23, as well as the level of light exposure24. In the context of coral reproduction, heterotrophic feeding has shown variable results, with observations of higher25 as well as lower26 reproductive capacity following heterotrophic feeding being reported. The influence of heterotrophic feeding on coral reproduction across a spectrum of temperatures is rarely assessed, yet in the temperate coral Cladocora caespitosa, heterotrophy was found to be more important for reproduction under lower temperature conditions27. A better understanding of the role of temperature and feeding on reproductive output is likely needed to determine whether specific reefs (e.g., reefs associated with high food availability28) possess a higher capacity for recruitment under climate change.
Similar to reproductive output, the effect of temperature and feeding on reproductive timing in corals remains relatively understudied, despite the synchronization of reproduction with abiotic/biotic conditions being an important consideration for recruitment success in a warming ocean29. Warmer temperatures have been shown to result in earlier reproduction in coral thermal conditioning studies conducted in the lab30, and this has also been observed in corals collected from natural reefs across seasons31. Yet, interestingly the opposite trend was recently observed in fed corals cultured over the course of 1 year in an ex situ flow-through system (i.e., reproduction occurred earlier in the lunar cycle at cooler winter temperatures and later in the lunar cycle at warmer summer temperatures)32. This contrasting result suggests that reproductive timing may stray from typical patterns under conditions associated with abundant energetic resources.
Long-term controlled experiments under different temperature scenarios could contribute to a better understanding of the influence of heterotrophy on reproduction in scleractinian corals. Maintaining reproducing coral colonies under ex situ conditions for multiple reproductive cycles, however, can be challenging (but see previous research32,33). Herein, straightforward and effective techniques for the active feeding (food source: Artemia nauplii) and long-term culture of a brooding coral (Pocillopora acuta) in a flow-through aquaculture system are described; yet, it should be noted that all the techniques described can also be used in recirculating aquaculture systems. To demonstrate these techniques, a preliminary comparison of the reproductive output and timing of coral colonies held at 24 &#176;C and 28 &#176;C under &#34;fed&#34; and &#34;unfed&#34; treatments was conducted. These temperatures were chosen to approximate seawater temperatures in winter and summer, respectively, in southern Taiwan30,34; a higher temperature was not chosen because the promotion of long-term ex situ culture, rather than testing coral response to thermal stress, was a primary goal of this experiment. Further, the density of Artemia nauplii before and after the feeding sessions was quantified&#160;to compare the feasibility of heterotrophic feeding at both temperature treatments.
Specifically, 24 colonies of P. acuta (mean total linear extension &#177; standard deviation: 21.3 cm &#177; 2.8 cm) were obtained from flow-through tanks at the research facilities of the National Museum of Marine Biology &#38; Aquarium, southern Taiwan. Pocillopora&#8203;&#160;acuta is a common coral species that possesses both a broadcast spawning, but&#160;typically brooding reproductive strategy35,36. The parent colonies of these corals were originally collected from the Outlet reef (21.931&#176;E, 120.745&#176;N) approximately 2 years earlier for another experiment32. Consequently, the coral colonies used in the present experiment had been reared for their entire lives under ex situ culture conditions; specifically, the colonies were exposed to ambient temperature and a 12 h:12 h light: dark cycle at 250 &#181;mol quanta m&#8722;2&#183;s&#8722;1 and were fed Artemia nauplii twice per week. We recognize that this long-term ex situ culture could have affected how the colonies responded to the treatment conditions in this experiment. We, therefore, would like to emphasize that the primary aim here is to illustrate how the described techniques can be effectively used to culture corals ex situ by demonstrating an applied example wherein the effects of temperature and feeding on coral reproduction were assessed.
Coral colonies were evenly distributed across six flow-through system culture tanks (tank interior length x width x height: 175 cm x 62 cm x 72 cm; tank light regime: 12 h:12h light:dark cycle at 250 &#181;mol quanta m&#8722;2&#183;s&#8722;1) (Figure 1A). The temperature in three of the tanks was set at 28 &#176;C, and the temperature in the other three tanks was set at 24 &#176;C; each tank had a logger that recorded the temperature every 10 min (see the Table of Materials). The temperature was independently controlled in each tank using chillers and heaters, and water circulation was maintained using flow motors (see the Table of Materials). Half of the colonies in each tank (n = 2 colonies/tank) were fed Artemia nauplii twice per week, while the other colonies were not fed. Each feeding session was 4 h in duration and was conducted in two independent temperature-specific feeding tanks. During feeding, all the colonies were moved into the feeding tanks, including the unfed colonies, to standardize the potential stress effect of moving the colonies between the tanks. The colonies in the fed and unfed treatments were positioned in their own compartment using a meshed frame within the temperature-specific feeding tanks so that only the colonies in the fed condition received food. The coral reproductive output and timing were assessed for each colony daily at 09:00 A.M. by counting the number of larvae that had been released into the larval collection containers overnight.

<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>Artemia cysts&#160;</td>
			<td>Supreme plus</td>
			<td>NA</td>
			<td>Food source&#160;</td>
		</tr>
		<tr>
			<td>Chiller</td>
			<td>Resun</td>
			<td>CL650</td>
			<td>To cool down water temperature if needed</td>
		</tr>
		<tr>
			<td>Conductivity portable meter</td>
			<td>WTW</td>
			<td>Cond 3110</td>
			<td>To measure salinity</td>
		</tr>
		<tr>
			<td>Enrichment diets</td>
			<td>Omega</td>
			<td>NA</td>
			<td>Used in Artemia cultivation</td>
		</tr>
		<tr>
			<td>Fishing line</td>
			<td>Super</td>
			<td>Nylon monofilament</td>
			<td>To hang the coral colonies</td>
		</tr>
		<tr>
			<td>Flow motors</td>
			<td>Maxspect</td>
			<td>GP03</td>
			<td>To create water flow</td>
		</tr>
		<tr>
			<td>Heater 350 W</td>
			<td>ISTA</td>
			<td>NA</td>
			<td>Heaters used in tanks</td>
		</tr>
		<tr>
			<td>HOBO pendant temperature logger</td>
			<td>Onset Computer</td>
			<td>UA-002-08</td>
			<td>To record water temperature</td>
		</tr>
		<tr>
			<td>LED lights</td>
			<td>Mean Well</td>
			<td>FTS: HLG-185H-36B</td>
			<td>NA</td>
		</tr>
		<tr>
			<td>Light portable meter</td>
			<td>LI-COR</td>
			<td>LI-250A</td>
			<td>Device used with light sensor to measure light intensity in PAR</td>
		</tr>
		<tr>
			<td>Light sensor</td>
			<td>LI-COR</td>
			<td>LI-193SA</td>
			<td>NA</td>
		</tr>
		<tr>
			<td>Plankton net 100 &#181;m mesh size</td>
			<td>Omega</td>
			<td>NA</td>
			<td>To collect larvae and artemia&#160;</td>
		</tr>
		<tr>
			<td>Primary pump 6000 L/H</td>
			<td>Mr. Aqua</td>
			<td>BP6000</td>
			<td>To draw water from tanks into chiller</td>
		</tr>
		<tr>
			<td>Propeller-type current meter</td>
			<td>KENEK</td>
			<td>GR20</td>
			<td>Device used with propeller-type detector to measure flow rate</td>
		</tr>
		<tr>
			<td>Propeller-type detector</td>
			<td>KENEK</td>
			<td>GR3T-2-20N</td>
			<td>NA</td>
		</tr>
		<tr>
			<td>Stereo microscope</td>
			<td>Zeiss</td>
			<td>Stemi 2000-C&#160;</td>
			<td>To count the number of artemia&#160;</td>
		</tr>
		<tr>
			<td>Temperature controller 1000 W</td>
			<td>Rep Park</td>
			<td>O-RP-SDP-1</td>
			<td>To set and maintain water temperature</td>
		</tr>
	</tbody>
</table>

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.

The described protocols allowed for (1) the comparison of the reproductive output and timing of individual coral colonies among distinct feeding and temperature treatments and (2) an assessment of the feasibility of Artemia nauplii feeding at different temperatures. Herein, a brief overview of the findings is given, but caution should be exercised with regard to the broad interpretation of the reported effects of temperature and feeding on coral reproduction due to the short-term nature of this experiment (i.e.,...

Watch this Scientific Journal Video about Advancing Coral Research by Exploring Climate Change Resistance, Ex Situ Aquaculture, and Reproduction Strategies at JoVE.com

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.

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 ...

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1.2K Views.  National Museum of Marine Biology & Aquarium. 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.

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

Method for Culture of Early Chick Embryos ex vivo (New Culture)

The chick embryo is a valuable tool in the study of early embryonic development. Its transparency, accessibility and ease of manipulation, make it an ideal tool for studying  the formation and patterning of brain, neural tube, somite and heart primordia. Applications of chick embryo culture include electroporation of DNA or RNA constructs in order to analyze gene function, grafts of growth factor coated beads such as FGFs and BMPs , as well as whole mount in situ hybridization and immunohistochemistry. This video demonstrates the different steps in chick embryo culture; First, the embryo is explanted in saline. Then, the embryo is centered on a glass ring. The membranes surrounding the embryo are lifted along the walls of the ring. The ring is then placed in a culture dish containing a pool of albumine. The culture dish is sealed and placed in a humid chamber, where the embryo is cultured for up to 24 hrs. Finally, the embryo is removed from the ring, fixed and processed for further applications. A troubleshooting guide is also presented.

Method for Culture of Early Chick Embryos ex vivo New Culture

This video demonstrates New culture, a method by which chick embryos are cultured outside the egg for up to 24 hr. This method enables one to study early development (primitive streak to 14 som.), a period corresponding to E7-9 in mouse. Applications of this technique include electroporation, in situ hybridization and immunohistochemistry.

The chick embryo is a valuable tool in the study of early embryonic development. Its transparency, accessibility and ease of manipulation, make it an ...

Chick ex ovo Culture and ex ovo CAM Assay: How it Really Works

Chicken eggs in the early phase of breeding are between in vitro and in vivo systems and provide a vascular test environment not only to study angiogenesis but also to study tumorigenesis. After the chick chorioallantoic membrane (CAM) has developed, its blood vessel network can be easily accessed, manipulated and observed and therefore provides an optimal setting for angiogenesis assays. Since the lymphoid system is not fully developed until late stages of incubation, the chick embryo serves as a naturally immunodeficient host capable of sustaining grafted tissues and cells without species-specific restrictions. In addition to nurturing developing allo- and xenografts, the CAM blood vessel network provides a uniquely supportive environment for tumor cell intravasation, dissemination, and vascular arrest and a repository where arrested cells extravasate to form micro metastatic foci.
For experimental purposes, in most of the recent studies the CAM was exposed by cutting a window through the egg shell and experiments were carried out in ovo, resulting in significant limitations in the accessibility of the CAM and possibilities for observation and photo documentation of effects. When shell-less cultures of the chick embryo were used1-4, no experimental details were provided and, if published at all, the survival rates of these cultures were low. We refined the method of ex ovo culture of chick embryos significantly by introducing a rationally controlled extrusion of the egg content. These ex ovo cultures enhance the accessibility of the CAM and chick embryo, enabling easy in vivo documentation of effects and facilitating experimental manipulation of the embryo. This allows the successful application to a large number of scientific questions: (1) As an improved angiogenesis assay5,6, (2) an experimental set up for facilitated injections in the vitreous of the chick embryo eye7-9, (3) as a test environment for dissemination and intravasation of dispersed tumor cells from established cell lines inoculated on the CAM10-12, (4) as an improved sustaining system for successful transplantation and culture of limb buds of chicken and mice13 as well as (5) for grafting, propagation, and re-grafting of solid primary tumor tissue obtained from biopsies on the surface of the CAM14.
In this video article we describe the establishment of a refined chick ex ovo culture and CAM assay with survival rates over 50%. Besides we provide a step by step demonstration of the successful application of the ex ovo culture for a large number of scientific applications. 
Daniel S. Dohle, Susanne D. Pasa, and Sebastian Gustmann contributed equally to this study.

Chick ex ovo Culture and ex ovo CAM Assay: How it Really Works

The chick chorioallantoic membrane (CAM) is a unique, naturally immunodeficient supportive culture environment to study angiogenesis and tumorigenesis. This video article demonstrates the different steps in chick ex ovo culture, application of potentially angiogenic substances and successful inoculation of tumor cells and tissues on the surface of the CAM.

Chicken eggs in the early phase of breeding are between in vitro and in vivo systems and provide a vascular test environment not only to study ...

An ex-ovo Chicken Embryo Culture System Suitable for Imaging and Microsurgery Applications

Understanding the relationships between genetic and microenvironmental factors that drive normal and malformed embryonic development is fundamental for discovering new therapeutic strategies. Advancements in imaging technology have enabled quantitative investigation of the organization and maturing of the body plan, but later stage embryonic morphogenesis is less clear. Chicken embryos are an attractive vertebrate animal model system for this application because of its ease of culture and surgical manipulation. Early embryos can be cultured for a short time on filter paper rings, which enables complete optical access for cell patterning and fate studies1,2. Studying advanced developmental processes such as cardiac morphogenesis are traditionally performed through a window of the eggshell3-5, but this technique limits optical access due to window size. We previously developed a simple method to culture whole embryos ex-ovo on hexagonal weigh boats for up to 10 days, which enabled high resolution imaging via ultrasonography6,7. These cultures were difficult to transport, limiting the types of imaging tools available for live experiments. We here present an improved shell-less culture system with a cost-effective, portable environmental chamber. Eggs were cracked onto a hammock created by a polyurethane membrane (cling wrap) affixed circumferentially to a plastic cup partially filled with sterile water. The dimensions of the circumference and depth of the hammock were both critical to maintain surface tension, while the mechanics of the hammock and water beneath helped dampen vibrations induced by transportation. A small footprint circulating water bath was also developed to enable continuous temperature control during experimentation. We demonstrate the ability to culture embryos in this way for at least 14 days without morphogenic defect or delay and employ this system in several microsurgical and imaging applications.

An ex-ovo Chicken Embryo Culture System Suitable for Imaging and Microsurgery Applications

In this article, we present a simple methodology to enable long-term ex-ovo avian embryo culture. This technique is ideal for longitudinal experimentation requiring complete optical accessibility and/or sterile transportation in avian embryos.

Understanding the relationships between genetic and microenvironmental factors that drive normal and malformed embryonic development is fundamental for ...

A Novel Ex vivo Culture Method for the Embryonic Mouse Heart

Developmental studies in the mouse are hampered by the inaccessibility of the embryo during gestation. Thus, protocols to isolate and culture individual organs of interest are essential to provide a method of both visualizing changes in development and allowing novel treatment strategies. To promote the long-term culture of the embryonic heart at late stages of gestation, we developed a protocol in which the excised heart is cultured in a semi-solid, dilute Matrigel. This substrate provides enough support to maintain the three-dimensional structure but is flexible enough to allow continued contraction. In brief, hearts are excised from the embryo and placed in a mixture of cold Matrigel diluted 1:1 with growth medium. After the diluted Matrigel solidifies, growth medium is added to the culture dish. Hearts excised as late as embryonic day 16.5 were viable for four days post-dissection. Analysis of the coronary plexus shows that this method does not disrupt coronary vascular development. Thus, we present a novel method for long-term culture of embryonic hearts.

A Novel Ex vivo Culture Method for the Embryonic Mouse Heart

Developmental studies in the mouse are hampered by the inaccessibility of the embryo during gestation. To promote the long-term culture of the embryonic heart at late stages of gestation, we developed a protocol in which the excised heart is cultured in a semi-solid, dilute Matrigel.

Developmental studies in the mouse are hampered  by the inaccessibility of the embryo during gestation. Thus, protocols to  isolate and culture individual ...

The Slice Culture Method for Following Development of Tooth Germs In Explant Culture

Explant culture allows manipulation of developing organs at specific time points&#160;and is therefore an important method for the developmental biologist. For many organs it is difficult to access developing tissue to allow monitoring during ex vivo culture. The slice culture method allows access to tissue so that morphogenetic movements can be followed and specific cell populations can be targeted for manipulation or lineage tracing.
In this paper we describe a method of slice culture that has been very successful for culture of tooth germs in a range of species. The method provides excellent access to the tooth germs, which develop at a similar rate to that observed in vivo, surrounded by the other jaw tissues. This allows tissue interactions between the tooth and surrounding tissue to be monitored. Although this paper concentrates on tooth germs, the same protocol can be applied to follow development of a number of other organs, such as salivary glands, Meckel&#39;s cartilage, nasal glands, tongue, and ear.

Here we detail a method to culture tooth germs in mandible slices using a tissue chopper. This method allows unique access to the tooth during development, providing excellent opportunity for manipulation and lineage tracing, not available using more traditional culture methods.

Explant culture allows manipulation of developing organs at specific time points&#160;and is therefore an important method for the developmental ...

High-fat Feeding Paradigm for Larval Zebrafish: Feeding, Live Imaging, and Quantification of Food Intake

Zebrafish are emerging as a model of dietary lipid processing and metabolic disease. This protocol describes how to feed larval zebrafish a lipid-rich meal, which consists of an emulsion of chicken egg yolk liposomes created by sonicating egg yolk in embryo media. Detailed instructions are provided to screen larvae for egg yolk consumption so that larvae that fail to feed will not confound experimental results. The chicken egg yolk liposomes can be spiked with fluorescent lipid analogs, including fatty acids and cholesterol, enabling both systemic and subcellular visualization of dietary lipid processing. Several methods are described to mount larvae that are conducive to short- and long-term live imaging with both upright and inverted objectives at high and low magnification. Additionally presented is an assay to quantify larval food intake by extracting the lipids of larvae fed fluorescent lipid analogs, spotting the lipids on a thin layer chromatography plate, and quantifying the fluorescence. Finally, critical aspects of the procedures, important controls, options for modifying the protocols to address specific experimental questions, and potential limitations are discussed. These techniques can be applied not only to focused, hypothesis driven inquiries, but also to a variety of screens and live imaging techniques to study dietary lipid metabolism and the control of food intake.

Zebrafish are emerging as a valuable model of dietary lipid processing and metabolic disease. Described are protocols of lipid-rich larval feeds, live imaging of dietary fluorescent lipid analogs, and quantification of food intake. These techniques can be applied to a variety of screening, imaging, and hypothesis driven inquiry techniques.

Zebrafish are emerging as a model of dietary lipid processing and metabolic disease. This protocol describes how to feed larval zebrafish a lipid-rich ...

Simple and Effective Administration and Visualization of Microparticles in the Circulatory System of Small Fishes Using Kidney Injection

The systemic administration of micro-size particles into a living organism can be applied for vasculature visualization, drug and vaccine delivery, implantation of transgenic cells and tiny optical sensors. However, intravenous microinjections into small animals, which are mostly used in biological and veterinary laboratories, are very difficult and require trained personnel. Herein, we demonstrate a robust and efficient method for the introduction of microparticles into the circulatory system of adult zebrafish (Danio rerio) by injection into the fish kidney. To visualize the introduced microparticles in the vasculature, we propose a simple intravital imaging technique in fish gills. In vivo monitoring of the zebrafish blood pH was accomplished using an injected microencapsulated fluorescent probe, SNARF-1, to demonstrate one of the possible applications of the described technique. This article provides a detailed description of the encapsulation of pH-sensitive dye and demonstrates the principles of the quick injection and visualization of the obtained microcapsules for in vivo recording of the fluorescent signal. The proposed method of injection is characterized by a low mortality rate (0-20%) and high efficiency (70-90% success), and it is easy to institute using commonly available equipment. All described procedures can be performed on other small fish species, such as guppies and medaka.

This article demonstrates the principles of a quick, minimally invasive injection of fluorescent microparticles into the circulatory system of small fishes and the in vivo visualization of the microparticles in fish blood.

The systemic administration of micro-size particles into a living organism can be applied for vasculature visualization, drug and vaccine delivery, ...

Applications of Spatio-temporal Mapping and Particle Analysis Techniques to Quantify Intracellular Ca2+ Signaling In Situ

Ca2+ imaging of isolated cells or specific types of cells within intact tissues often reveals complex patterns of Ca2+ signaling. This activity requires careful and in-depth analyses and quantification to capture as much information about the underlying events as possible. Spatial, temporal and intensity parameters intrinsic to Ca2+ signals such as frequency, duration, propagation, velocity and amplitude may provide some biological information required for intracellular signalling. High-resolution Ca2+ imaging typically results in the acquisition of large data files that are time consuming to process in terms of translating the imaging information into quantifiable data, and this process can be susceptible to human error and bias. Analysis of Ca2+ signals from cells in situ typically relies on simple intensity measurements from arbitrarily selected regions of interest (ROI) within a field of view (FOV). This approach ignores much of the important signaling information contained in the FOV. Thus, in order to maximize recovery of information from such high-resolution recordings obtained with Ca2+dyes or optogenetic Ca2+ imaging, appropriate spatial and temporal analysis of the Ca2+ signals is required. The protocols outlined in this paper will describe how a high volume of data can be obtained from Ca2+ imaging recordings to facilitate more complete analysis and quantification of Ca2+ signals recorded from cells using a combination of spatiotemporal map (STM)-based analysis and particle-based analysis. The protocols also describe how different patterns of Ca2+ signaling observed in different cell populations in situ can be analyzed appropriately. For illustration, the method will examine Ca2+ signaling in a specialized population of cells in the small intestine, interstitial cells of Cajal (ICC), using GECIs.

Applications of Spatio-temporal Mapping and Particle Analysis Techniques to Quantify Intracellular Ca2+ Signaling In Situ

Genetically encoded Ca2+ indicators (GECIs) have radically changed how in situ Ca2+ imaging is performed. To maximize data recovery from such recordings, appropriate analysis of Ca2+ signals is required. The protocols in this paper facilitate the quantification of Ca2+ signals recorded in situ using spatiotemporal mapping and particle-based analysis.

Ca2+ imaging of isolated cells or specific types of cells within intact tissues often reveals complex patterns of Ca2+ signaling. This activity requires ...

Fluorescence-Activated Cell Sorting for the Isolation of Scleractinian Cell Populations

Coral reefs are under threat due to anthropogenic stressors. The biological response of coral to these stressors may occur at a cellular level, but the mechanisms are not well understood. To investigate coral response to stressors, we need tools for analyzing cellular responses. In particular, we need tools that facilitate the application of functional assays to better understand how cell populations are reacting to stress. In the current study, we use fluorescence-activated cell sorting (FACS) to isolate and separate different cell populations in stony corals. This protocol includes: (1) the separation of coral tissues from the skeleton, (2) creation of a single cell suspension, (3) labeling the coral cells using various markers for flow cytometry, and (4) gating and cell sorting strategies. This method will enable researchers to work on corals at the cellular level for analysis, functional assays, and gene expression studies of different cell populations.

Corals create biodiverse ecosystems important for both humans and marine organisms. However, we still do not understand the full potential and function of many coral cells. Here, we present a protocol developed for the isolation, labeling, and separation of stony coral cell populations.

Coral reefs are under threat due to anthropogenic stressors. The biological response of coral to these stressors may occur at a cellular level, but the ...

At-Risk Butterfly Captive Propagation Programs to Enhance Life History Knowledge and Effective Ex Situ Conservation Techniques

Improving knowledge of ex situ best practices for at-risk butterflies is important for generating successful conservation and recovery program outcomes. Research on such captive populations can also yield valuable data to address key information gaps about the behavior, life history, and ecology of the target taxa. We describe a protocol for captive propagation of the federally endangered Cyclargus thomasi bethunebakeri that can be used as a model for other at-risk butterfly ex situ programs, especially those in the family Lycaenidae. We further provide a simple and straightforward protocol for recording various life history metrics that can be useful for informing ex situ methodologies as well as adapted for laboratory studies of other lepidoptera.

Here, we present protocols for 1) the laboratory captive propagation of the federally endangered Miami blue butterfly (Cyclargus thomasi bethunebakeri), and 2) assessing basic life history information such as immature development time and number of larval stadia. Both methods can be adapted for use with other ex situ conservation programs.

Improving knowledge of ex situ best practices for at-risk butterflies is important for generating successful conservation and recovery program outcomes. ...