The care and usage of fish generated for representative results described in this protocol was performed in full accordance with the guidelines set forth by the Institutional Animal Care and Use Committee at Boston Children&#39;s Hospital, protocol # 14-05-2673R.

1. Rotifer Culture
<ol>
	<li>Basic Components of a Culture System using a 100 L Culture Vessel
	<ol>
		<li>Gather all the components for the rotifer culture setup . The rotifer culture setup consists of a culture vessel (CV) to grow the rotifers; a similar vessel to maintain feedout rotifers (feedout culture vessel, FCV); a round-bottomed hatching jar (Feed Reservoir, FR) for storage of the algae feed mixture (AFM); an air supply (AS) to aerate the CV, FCV and the FR; a peristaltic pump with a metering timer (PMT) to control delivery of algae feed into the CV and FCV; and a floss particle filter (FPF) that sits inside the CV. 
		NOTE: A complete list of supplies and components is provided in the Materials List.</li>
	</ol>
	</li>
	<li>Configuration
	<ol>
		<li>Elevate the CV and FCV on a stand or table so that the cultures may be easily harvested via a drain fitting into a collection container (Figure 1). Use flexible air supply tubing to connect the AS to a length of rigid tubing in each culture vessel. Make sure that the tubing is long enough to deliver air to the bottom of the CV or FCV.</li>
		<li>Use a small-capacity air line to connect the AS to a length of rigid tubing that extends to the bottom of the FR that contains the AFM. Install a valve into each air line to regulate air flow. Connect the FR to the PMT with feed delivery tubing, and run the tubing from the PMT into a hole drilled into the side of the CV/FCV, near the top. Figure 1.</li>
	</ol>
	</li>
	<li>Startup
	<ol>
		<li>Fill the culture vessel to 90% of the available volume with Reverse Osmosis water (RO). If RO is not available, use clean, dechlorinated municipal water; however, a biosecurity risk assessment should be performed to ensure that no potentially pathogenic organisms are present in the source water. NOTE: Such an analysis can be performed by any qualified water testing laboratory.</li>
		<li>Dose the culture vessel water with aquarium salt to reach a salinity of 15 g/L. Set the air flow into the vessel so that it maintains a &#34;rolling boil&#34;, and then slowly add the measured amount of salt to the culture vessel until it is completely dissolved by the aeration. Continue aerating the water for &#62;1 hr to ensure that it is fully oxygenated.</li>
		<li>Make the algae feed mixture. To 3 L of clean, dechlorinated fresh (0 ppm) water add 100 g of NaHCO3 and 100 g of ammonia neutralizer (sodium hydroxymethylsulfonate). This last reagent provides the additional benefit of neutralizing any residual chlorine from tap water or bleach residues from sanitizing of culture equipment. It is critical to ensure that these compounds are fully dissolved. Then add 1 L of algae concentrate (biomass dry weight ~15%). Add the feed mixture to the FR and store at 4 &#176;C.</li>
		<li>Add a starter culture of 5 - 10 million Brachionus plicatilis rotifers to the CV containing the aerated 15 g/L salinity water. If the rotifers have been chilled during shipping or storage, they should be gradually (during 30 min or longer) acclimated to the temperature of the water in the culture vessel (25 - 27 &#176;C).</li>
		<li>Turn on the PMT and begin pumping the algae feed into the rotifer culture vessel. Using the timer feature of the PMT, set the delivery rate of algae feed mixture so that ~1.6 ml of algae feed mixture is delivered per million rotifers in the culture, per day. Distribute the feedings in small portions at regular intervals over the course of a 24 hr period; the more frequent the feedings, the better.</li>
		<li>Calibrate the delivery rate of the pump by manually turning on the PMT for a set period (e.g., 1 min) and collect the algae that it pumps during this interval into a graduated cylinder or beaker. For example, if the PMT doses 5 ml of algae in 1 min, then the dose rate would be 5 ml algae/min.</li>
		<li>Calculate the required daily feeding rate by multiplying the number of rotifers present, in millions, by 1.6 ml. For example, a rotifer culture with a population size of 100 million rotifers would require ~160 ml of feed per day (100 x 1.6 ml).</li>
		<li>Set the PMT to dose the total daily feed requirement at regular intervals throughout a 24 hr period. For example, delivery of a total daily feed amount of 160 ml could be delivered in portions once every 3 hr over a 24 hr period using a PMT set with a dosing pump rate of 5 ml/min for 4 min, 8 times daily (5 ml/min x 4 min = 20 ml x 8 feedings = 160 ml).</li>
		<li>Allow the culture to grow until it generates the required population, typically for 48 - 72 hr, before harvesting. At 24 h post-start up, add the floss particle filters to the culture vessel and begin normal maintenance.</li>
	</ol>
	</li>
	<li>Maintenance 
	NOTE: The culture operates on a continuous basis and requires routine maintenance that ideally should be performed at the same time each day, in the following sequence.
	<ol>
		<li>Fill the FCV to 90% of available volume with clean, dechlorinated fresh water, dosed with 10 g/L aquarium salts. Ensure that the water is well mixed, and that all of the salt is fully dissolved. Set the air flow into the vessel so that it maintains a &#34;rolling boil&#34;. Measure the salinity with a refractometer and ensure that the salinity is 10 g/L. It is critical to meet this target and not to exceed it.</li>
		<li>Sample the rotifers in the CV: Ensure that the culture is well-mixed, then collect 3 samples of a 2 - 3 ml each using a transfer pipette or autopipettor, from different parts of the culture. Combine these samples in a tube or vial of convenient size (e.g., 10 ml).</li>
		<li>Transfer 1 - 2 ml of the combined sample onto a petri dish so that it can be visualized under a dissecting microscope. Check the quality of the culture (swimming behavior of the rotifers, presence of detached eggs, contaminating protozoa).</li>
		<li>Immobilize the rotifers in the remaining combined sample by adding 100 &#181;l of 50% Lugols iodine solution to the sample. Within seconds after addition of the Lugols, observe the rotifers to stop swimming. Now, easily count the rotifers . 
		NOTE: Ethanol, diluted bleach, or vinegar can be used in place of Lugols. Vinegar (2 drops/10 ml) has the advantages of being non-hazardous, not losing strength in storage as bleach and iodine solutions can, and not making the rotifers contract, so the corona of cilia and the &#34;foot&#34; remain extended and the animals look more natural.</li>
		<li>Ensure that the sample is well mixed (immobilized rotifers will settle rapidly), then quickly take a ~2 ml subsample in a plastic pipet and dispense 1 ml into a Sedgewick-Rafter counting slide (20 x 50 1-mm squares) (Figure 2). Using a dissecting or compound microscope, count the intact rotifers and the total number of eggs attached to these rotifers (Figure 2). Count as much of the slide area as necessary to count ~100 rotifers. Calculate the number of rotifers per ml, and record this in a spreadsheet or logbook.</li>
		<li>Harvest ~30% of the volume of the rotifers in the CV: Remove the air supply and floss filter, slowly open the valve at the bottom of the CV and allow the water to flow into a plankton collector with a 53 &#181;m mesh screen. Collect the water flowing out of the bottom of the collector after filtering into a bucket or drain. Use a gentle to moderate flow to avoid damaging the rotifers. Do not allow the rotifers to dry on the screen.</li>
		<li>For consistency reasons, it is advisable to establish the FCV with a standard number of rotifers each day.&#160; Therefore, based on the known CV rotifer density and harvest volume, adjust the FCV total volume to achieve a consistent final density (e.g., 1,500 rotifers/ml).&#160;Add the harvested rotifers to the FCV: Gently transfer the rotifers from the collection screen using a wash bottle filled with clean salt water (10 - 15 g/L). Invert the screen over the FCV and wash the rotifers into the FCV with a gentle stream of salt water. Start the PMT to deliver feed (~1.57 ml per million rotifers per day) to the FCV.</li>
		<li>Scrub the entire inside of the CV with a clean, soft nylon brush or scrub pad.</li>
		<li>Make a new mix of 15 g/L water by adding the appropriate amount of salt to a measured amount of clean RO water in a 5 gallon bucket to replace the volume of water lost to harvest. Add the salt to the water in the bucket and stir vigorously until it is completely dissolved, and then add to the CV.</li>
		<li>Using a high-pressure spray in a sink, rinse the floss filter until it is free of debris, and then return it to the CV.</li>
		<li>Adjust feed rates of algae delivered to the CV by changing the duration of each dosing event, according to the daily count of rotifers/ml. Use the calculations provided in step 1.3.8, above to determine the appropriate amount of feed to be delivered.</li>
		<li>Approximately 24 hr later, repeat the process. Start by harvesting the rotifers remaining in the FCV (that were not needed for the previous day) in the same manner as described above (steps 1.4.2 - 1.4.10). Concentrate them in 2 L of fresh, clean dechlorinated water (5 g/L salt). These can be stored at 4 &#176;C as a backup supply, or used to feed later stages of fish, beyond what is described in this protocol. 
		NOTE: This protocol permits up to 2 - 3 days of reduced culture maintenance (with normal automatic feeding), because the rotifers in the CV can tolerate omission of harvests without serious consequences.</li>
	</ol>
	</li>
</ol>
2. Polyculture
<ol>
	<li>Setup
	<ol>
		<li>Collect zebrafish embryos from a spawning event by pouring spawned embryos through a tea-strainer and then gently rinsing with sterile fish water (or any other non-contaminated source of appropriately conditioned solution; e.g., embryo medium, E3, etc.) from a wash bottle into petri dishes.</li>
		<li>Incubate the embryos at 25 - 28 &#176;C in petri dishes at a density of 40 - 50 embryos per dish for 5 days.</li>
		<li>Begin the polyculture phase on day 5 post-fertilization, or when greater than 90% of the hatched larvae are actively swimming up in the water column.</li>
	</ol>
	</li>
	<li>Inoculation
	<ol>
		<li>Add 500 ml of rotifer culture directly from the FCV to a 3.5 L nursery tank; inclusion of rotifer culture water provides algal feed that maintains the nutritional content of the rotifers during polyculture.</li>
		<li>Gently pour the larvae from one petri dish into the nursery tank. Ensure that no larvae remain in the dish.</li>
		<li>Add 500 ml of clean, conditioned fish water from a recirculating system or dedicated water source to the tank to reach a final volume of 1 L and a final salinity of 5 g/L. 
		NOTE: This salinity is critical because zebrafish larvae survival will be negatively impacted if salinity is &#62;7 g/L and rotifer survival will be negatively impacted if salinity &#60;2 g/L.</li>
	</ol>
	</li>
	<li>Polyculture phase 
	NOTE: The polyculture phase should last for up to 4 days post-inoculation (a total of 5 days, corresponding to days 5-9 post-fertilization).
	<ol>
		<li>Observe the polyculture tank at least once per day during this period to ensure that rotifers and fish are present and growing. Ensure that the rotifers are visible throughout the water column. Ensure that the the fishs are also visible within the water column, swimming among the rotifers.</li>
		<li>Start normal water flow through the tank. Place a screen or baffle over the drain port to ensure that larvae are not flushed out of the tank. 
		NOTE: At the end of this phase, the fish will be large enough to consume larger prey items such as Artemia nauplii or processed feed items in the size range of 75 - 125 &#181;m. 
		NOTE: The rotifer population dynamics within a representative polyculture tank were measured by sampling/counting rotifers from the tank in the same manner as described in steps 1.4.2 - 1.4.5. This was done once per day from the beginning of the polyculture phase until it was completed.</li>
	</ol>
	</li>
</ol>

<ol>
	<li>Ribas, L., Piferrer, F. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+zebrafish+(Danio+rerio)+as+a+model+organism,+with+emphasis+on+applications+for+finfish+aquaculture+research.">The zebrafish (Danio rerio) as a model organism, with emphasis on applications for finfish aquaculture research.</a> Reviews in Aquaculture. 6, 209-240 (2014).</li><li>Poss, K. D. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Advances+in+understanding+tissue+regenerative+capacity+and+mechanisms+in+animals.">Advances in understanding tissue regenerative capacity and mechanisms in animals.</a> Nature reviews. Genetics. 11, 710-722 (2010).</li><li>Gemberling, M., Bailey, T. J., Hyde, D. R., Poss, K. D. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+zebrafish+as+a+model+for+complex+tissue+regeneration.">The zebrafish as a model for complex tissue regeneration.</a> Trends in genetics TIG. 29, 611-620 (2013).</li><li>Santoriello, C., Zon, L. I. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Hooked!+modeling+human+disease+in+zebrafish.">Hooked! modeling human disease in zebrafish.</a> Journal of Clinical Investigation. 122, 2337-2343 (2012).</li><li>Selderslaghs, I. W. T., Blust, R., Witters, H. E. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Feasibility+study+of+the+zebrafish+assay+as+an+alternative+method+to+screen+for+developmental+toxicity+and+embryotoxicity+using+a+training+set+of+27+compounds.">Feasibility study of the zebrafish assay as an alternative method to screen for developmental toxicity and embryotoxicity using a training set of 27 compounds.</a> Reproductive Toxicology. 33 (2), 142-154 (2012).</li><li>Lawrence, C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+husbandry+of+zebrafish+(Danio+rerio):+A+review.">The husbandry of zebrafish (Danio rerio): A review.</a> Aquaculture. 269, 1-20 (2007).</li><li>Harper, C., Lawrence, C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=.">.</a> The Laboratory Zebrafish (Laboratory Animal Pocket Reference). , (2010).</li><li>Nusslein-Volhard, C., Dahm, R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=.">.</a> Zebrafish, A Practical Approach. , (2002).</li><li>Westerfield, M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=.">.</a> The Zebrafish Book. A Guide for the Laboratory Use of Zebrafish. , (2007).</li><li>Carvalho, P., Arau, L. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Rearing+zebrafish+(Danio+rerio)+larvae+without+live+food:+evaluation+of+a+commercial,+a+practical+and+a+purified+starter+diet+on+larval+performance.">Rearing zebrafish (Danio rerio) larvae without live food: evaluation of a commercial, a practical and a purified starter diet on larval performance.</a> Aquaculture Research. 37, 1107-1111 (2006).</li><li>Spence, R., Gerlach, G., Lawrence, C., Smith, C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+behaviour+and+ecology+of+the+zebrafish,+Danio+rerio.">The behaviour and ecology of the zebrafish, Danio rerio.</a> Biol Rev Camb Philos Soc. 83 (1), 13-34 (2008).</li><li>Best, J., Adatto, I., Cockington, J., James, A., Lawrence, C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+novel+method+for+rearing+first-feeding+larval+zebrafish:+polyculture+with+Type+L+saltwater+rotifers+(Brachionus+plicatilis).">A novel method for rearing first-feeding larval zebrafish: polyculture with Type L saltwater rotifers (Brachionus plicatilis).</a> Zebrafish. 7 (3), 289-295 (2010).</li><li>Lawrence, C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Advances+in+zebrafish+husbandry+and+management.">Advances in zebrafish husbandry and management.</a> Methods in Cell Biology. 104, 429-451 (2011).</li><li>Lawrence, C., Sanders, E., Henry, E. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Methods+for+culturing+saltwater+rotifers+(Brachionus+plicatilis)+for+rearing+larval+zebrafish.">Methods for culturing saltwater rotifers (Brachionus plicatilis) for rearing larval zebrafish.</a> Zebrafish. 9, 140-146 (2012).</li><li>Tucker, C. S., Hargreaves, J. A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=.">.</a> Biology and Culture of Channel Catfish. 34, 634-657 (2004).</li></ol>

E.C. Henry is employed by Reed Mariculture, Inc., a company that provides rotifers, algae concentrates, and other supplies to the aquaculture and fish hobbyist markets.

Successful implementation of the rotifer polyculture method for feeding early larval zebrafish requires effective protocols for two tasks: the establishment and maintenance of a continuous rotifer culture system to feed the fish, and culturing first-feeding zebrafish larvae along with rotifers in the same tank.
The setup for a continuous saltwater rotifer production system for zebrafish laboratories first described by Lawrence and co-authors14 has been modified and enhanced in a num...

The zebrafish (Danio rerio) is a pre-eminent laboratory animal utilized in a growing number of scientific disciplines, including but not limited to developmental genetics, toxicology, behavior, aquaculture, regenerative biology, and the modeling of many human disorders1-5. Although the species is relatively easy to maintain in the laboratory, there are a number of management challenges associated with their culture6. The most prominent of these is larval rearing, particularly when the fish first begin to feed subsequent to gas bladder inflation7. Under normal, controlled conditions, this developmental event occurs at ~5 days post-fertilization (dpf), with the following 3 - 5 days of growth being particularly critical7. The central technical difficulty during this stage is to adequately meet the nutritional demands of the first feeding larvae - feed items must be appropriately sized, digestible, attractive, and available on a nearly continuous basis, without creating excessive waste in culturing tanks. Historically this has been achieved typically by delivering numerous small amounts of feed to the fish in tanks, along with routine water exchange8,9. While these methods are to some degree successful, they are inefficient, require high labor inputs, and return only variable and limited rates of growth and survival10.
In nature, zebrafish larvae presumably feed on abundant small zooplankton present in the water column11. For this reason, larviculture protocols that incorporate live feeds such as Paramecium, rotifers, and Artemia are typically most efficient7. In 2010, Best and co-authors demonstrated that it was possible to grow larval zebrafish in static, brackish water along with saltwater rotifers for the first 5 days of exogenous feeding12. This approach, which harnesses the natural high productivity of rotifer cultures to provide ample, highly nutritious prey without polluting the water, yields very high rates of larval growth and survival with low labor input12,13. In recent years, an increasing number of laboratories around the world have adopted variations of this protocol, and many are now culturing rotifers in a continuous fashion to support nursery systems14.
Over the past several years, methods for both rotifer/zebrafish polyculture and rotifer production have been refined and improved to become more standardized and readily scalable. This article provides step-by-step instructions for 1) continuous and robust rotifer production and 2) the establishment of the rotifer/zebrafish polyculture system used to support robust growth of fish for the first 5 days of exogenous feeding.

<table border="0" cellpadding="0" cellspacing="0" width="823">
	<tbody>
		<tr height="22">
			<td height="22" style="height:22px;width:305px;">Rotifer Culture Infrastructure</td>
			<td style="width:135px;"></td>
			<td style="width:119px;"></td>
			<td style="width:264px;"></td>
		</tr>
		<tr height="41">
			<td height="41" style="height:41px;width:305px;">100 Liter Culture Vessel</td>
			<td style="width:135px;">Aquaneering</td>
			<td style="width:119px;">Custom</td>
			<td style="width:264px;">Polycarbonate culture vessel, conical bottomed, with drain valve</td>
		</tr>
		<tr height="41">
			<td height="41" style="height:41px;width:305px;">5 Gallon Culture Bucket Kit</td>
			<td style="width:135px;">Reed Mariculture</td>
			<td style="width:119px;">CCS Starter Kit</td>
			<td style="width:264px;">Small volume culture vessel for small facilities</td>
		</tr>
		<tr height="41">
			<td height="41" style="height:41px;width:305px;">Rigid Clear Tubing 1/2&#34; O.D., 36&#8221;</td>
			<td style="width:135px;">Pentair Aquatic Ecosystems</td>
			<td style="width:119px;">16025</td>
			<td style="width:264px;">Rigid clear tubing for air delivery</td>
		</tr>
		<tr height="41">
			<td height="41" style="height:41px;width:305px;">Mesh tube</td>
			<td style="width:135px;">Pentair Aquatic Ecosystems</td>
			<td style="width:119px;">RT444X</td>
			<td style="width:264px;">Mesh tube support for floss filter</td>
		</tr>
		<tr height="41">
			<td height="41" style="height:41px;width:305px;">Rotifer Floss</td>
			<td style="width:135px;">Reed Mariculture</td>
			<td style="width:119px;">Rotifer floss 12&#8221; x 42&#8221;</td>
			<td style="width:264px;">Particulate waste trap</td>
		</tr>
		<tr height="41">
			<td height="41" style="height:41px;width:305px;">Peristaltic Metering Timer Pump, 5 GPD</td>
			<td style="width:135px;">Grainger</td>
			<td style="width:119px;">38M003&#160;</td>
			<td style="width:264px;">Metering pump with timer for dosing feed to rotifers</td>
		</tr>
		<tr height="41">
			<td height="41" style="height:41px;width:305px;">Peristaltic Metering Timer Pump, 1-100 mL/h (for smaller-scale culture)</td>
			<td style="width:135px;">Coral Vue</td>
			<td style="width:119px;">SKU: IC-LQD-DSR</td>
			<td style="width:264px;">Metering pump with timer for dosing feed to rotifers</td>
		</tr>
		<tr height="41">
			<td height="41" style="height:41px;width:305px;">Silicone Tubing&#160;</td>
			<td style="width:135px;">Cole Parmer</td>
			<td style="width:119px;"></td>
			<td style="width:264px;">Tubing for algae delivery to rotifer vessel</td>
		</tr>
		<tr height="41">
			<td height="41" style="height:41px;width:305px;">Rigid Clear Tubing &#34; O.D.,36&#8221;</td>
			<td style="width:135px;">Pentair Aquatic Ecosystems</td>
			<td style="width:119px;">16025</td>
			<td style="width:264px;">Rigid clear tubing for air delivery to algae paste</td>
		</tr>
		<tr height="41">
			<td height="41" style="height:41px;width:305px;">Rigid Clear Tubing O.D., 36&#8221;</td>
			<td style="width:135px;">Pentair Aquatic Ecosystems</td>
			<td style="width:119px;">16025</td>
			<td style="width:264px;">Rigid clear tubing for algae delivery</td>
		</tr>
		<tr height="22">
			<td height="22" style="height:22px;width:305px;">Rotifers</td>
			<td style="width:135px;"></td>
			<td style="width:119px;"></td>
			<td style="width:264px;"></td>
		</tr>
		<tr height="41">
			<td height="41" style="height:41px;width:305px;">Live Rotifers Brachionus plicatilis Type L</td>
			<td style="width:135px;">Reed Mariculture</td>
			<td style="width:119px;">Type L 5 million</td>
			<td style="width:264px;">Rotifer stock culture for system startup</td>
		</tr>
		<tr height="22">
			<td height="22" style="height:22px;width:305px;">Rotifer Feed</td>
			<td style="width:135px;"></td>
			<td style="width:119px;"></td>
			<td style="width:264px;"></td>
		</tr>
		<tr height="45">
			<td height="45" style="height:45px;width:305px;">Sodium hydroxymethylsulfonate</td>
			<td style="width:135px;">Reed Mariculture</td>
			<td style="width:119px;">ClorAm-X&#174; 1lb tub</td>
			<td style="width:264px;">Ammonia reducer for algae feed mix</td>
		</tr>
		<tr height="22">
			<td height="22" style="height:22px;width:305px;">Sodium Bicarbonate</td>
			<td style="width:135px;">Fisher Scientific</td>
			<td style="width:119px;">S25533B</td>
			<td style="width:264px;">pH buffer for algae feed mix</td>
		</tr>
		<tr height="45">
			<td height="45" style="height:45px;width:305px;">Microalgae concentrate</td>
			<td style="width:135px;">Reed Mariculture</td>
			<td style="width:119px;">Rotigrow Plus&#174; 1 liter bag</td>
			<td style="width:264px;">Nutritionally optimized rotifer feed</td>
		</tr>
		<tr height="22">
			<td height="22" style="height:22px;width:305px;">Water Preparation</td>
			<td style="width:135px;"></td>
			<td style="width:119px;"></td>
			<td style="width:264px;"></td>
		</tr>
		<tr height="96">
			<td height="96" style="height:96px;width:305px;">&#160;Reef Crystals Reef Salt</td>
			<td style="width:135px;">That Fish Place</td>
			<td style="width:119px;">198210</td>
			<td style="width:264px;">Salt for making culture water (NOTE: this item is an example only; any contaminant free salt formulations may be used).&#160;</td>
		</tr>
		<tr height="41">
			<td height="41" style="height:41px;width:305px;">Refractometer</td>
			<td style="width:135px;">Pentair Aquatic Ecosystems</td>
			<td style="width:119px;">SR6</td>
			<td style="width:264px;">measuring salinity</td>
		</tr>
		<tr height="22">
			<td height="22" style="height:22px;width:305px;">Rotifer Culture Equipment</td>
			<td style="width:135px;"></td>
			<td style="width:119px;"></td>
			<td style="width:264px;"></td>
		</tr>
		<tr height="41">
			<td height="41" style="height:41px;width:305px;">Plankton Collectors 12&#34; Dia, 53 microns</td>
			<td style="width:135px;">Pentair Aquatic Ecosystems</td>
			<td style="width:119px;">BBPC20</td>
			<td style="width:264px;">Mesh screen for collecting rotifers</td>
		</tr>
		<tr height="41">
			<td height="41" style="height:41px;width:305px;">Scrub Pads</td>
			<td style="width:135px;">Pentair Aquatic Ecosystems</td>
			<td style="width:119px;">SCR-58</td>
			<td style="width:264px;">Scrub pad for cleaning inside of culturing vessels</td>
		</tr>
		<tr height="22">
			<td height="22" style="height:22px;width:305px;">Scrub Brush</td>
			<td style="width:135px;"></td>
			<td style="width:119px;"></td>
			<td style="width:264px;"></td>
		</tr>
		<tr height="41">
			<td height="41" style="height:41px;width:305px;">Bucket</td>
			<td style="width:135px;">Grainger Supply&#160;</td>
			<td style="width:119px;">43Y530</td>
			<td style="width:264px;">Graduated bucket for mixing culture water</td>
		</tr>
		<tr height="41">
			<td height="41" style="height:41px;width:305px;">Hatching Jar</td>
			<td style="width:135px;">Pentair Aquatic Ecosystems</td>
			<td style="width:119px;">J30</td>
			<td style="width:264px;">Storage of algae feed mix</td>
		</tr>
		<tr height="41">
			<td height="41" style="height:41px;width:305px;">Lugol&#8217;s Solution, Dilute</td>
			<td style="width:135px;">Fisher Scientific</td>
			<td style="width:119px;">S99481</td>
			<td style="width:264px;">Agent used to immobilize live rotifers for counting</td>
		</tr>
		<tr height="41">
			<td height="41" style="height:41px;width:305px;">Sedgewick-Rafter plankton counting slide with grid&#160;</td>
			<td style="width:135px;">Pentair Aquatic Eco-Systems</td>
			<td style="width:119px;">M415</td>
			<td style="width:264px;">Counting rotifers</td>
		</tr>
		<tr height="22">
			<td height="22" style="height:22px;width:305px;">Miscelleneous</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr height="41">
			<td height="41" style="height:41px;width:305px;">Tea Strainer</td>
			<td style="width:135px;">Kitchenworks</td>
			<td style="width:119px;">971972</td>
			<td style="width:264px;">Used for collecting zebrafish embryos after spawning</td>
		</tr>
	</tbody>
</table>

the complete and updated "rotifer polyculture method" for rearing first feeding zebrafish

The zebrafish (Danio rerio) is a model organism of increasing importance in many fields of science. One of the most demanding technical aspects of culture of this species in the laboratory is rearing first-feeding larvae to the juvenile stage with high rates of growth and survival. The central management challenge of this developmental period revolves around delivering highly nutritious feed items to the fish on a nearly continuous basis without compromising water quality. Because larval zebrafish are well-adapted to feed on small zooplankton in the water column, live prey items such as brachionid rotifers, Artemia, and Paramecium are widely recognized as the feeds of choice, at least until the fish reach the juvenile stage and are able to efficiently feed on processed diets. This protocol describes a method whereby newly hatched zebrafish larvae are cultured together with live saltwater rotifers (Brachionus plicatilis) in the same system. This polyculture approach provides fish with an "on-demand", nutrient-rich live food source without producing chemical waste at levels that would otherwise limit performance. Importantly, because the system harnesses both the natural high productivity of the rotifers and the behavioral preferences of the fish, the labor involved with maintenance is low. The following protocol details an updated, step-by-step procedure that incorporates rotifer production (scalable to any desired level) for use in a polyculture of zebrafish larvae and rotifers that promotes maximal performance during the first 5 days of exogenous feeding.

The continuous rotifer culture system described here is dynamic, and it is normal for rotifer numbers to fluctuate to a small extent over time if there are variations in daily feeding and harvest rates. The population of the rotifers in one of the active cultures in the aquaculture facilities at Boston Children&#39;s Hospital, maintained in the manner described above, was monitored for 30 days (Figure 3). The mean culture density during this period was 932 rotifers/mL, wi...

Watch this Scientific Journal Video about The Complete and Updated "Rotifer Polyculture Method" for Rearing First Feeding Zebrafish at JoVE.com

The Complete and Updated "Rotifer Polyculture Method" for Rearing First Feeding Zebrafish

Larval zebrafish are adapted to feed on zooplankton. It is possible to capitalize on this natural feature in the laboratory by growing first feeding fish together in the same system with live saltwater rotifers. This "polyculture" strategy promotes high growth and survival with minimal labor and disturbance to the larvae.

The zebrafish (Danio rerio) is a model organism of increasing importance in many fields of science.  One of the most demanding technical aspects of ...

the-complete-updated-rotifer-polyculture-method-for-rearing-first

Research

JoVE Journal

Developmental Biology

15.3K Views.  Boston Children's Hospital. Larval zebrafish are adapted to feed on zooplankton. It is possible to capitalize on this natural feature in the laboratory by growing first feeding fish together in the same system with live saltwater rotifers. This "polyculture" strategy promotes high growth and survival with minimal labor and disturbance to the larvae.

Video: The Complete and Updated "Rotifer Polyculture Method" for Rearing First Feeding Zebrafish

Live Imaging of Innate Immune and Preneoplastic Cell Interactions Using an Inducible Gal4/UAS Expression System in Larval Zebrafish Skin

Here we describe a method to conditionally induce epithelial cell transformation by the use of the 4-Hydroxytamoxifen (4-OHT) inducible KalTA4-ERT2/UAS expression system1 in zebrafish larvae, and the subsequent live imaging of innate immune cell interaction with HRASG12V expressing skin cells. The KalTA4-ERT2/UAS system is both inducible and reversible which allows us to induce cell transformation with precise temporal/spatial resolution in vivo. This provides us with a unique opportunity to live image how individual preneoplastic cells interact with host tissues as soon as they emerge, then follow their progression as well as regression. Recent studies in zebrafish larvae have shown a trophic function of innate immunity in the earliest stages of tumorigenesis2,3. Our inducible system would allow us to live image the onset of cellular transformation and the subsequent host response, which may lead to important insights on the underlying mechanisms for the regulation of oncogenic trophic inflammatory responses. We also discuss how one might adapt our protocol to achieve temporal and spatial control of ectopic gene expression in any tissue of interest.

Studying the earliest events of preneoplastic cell progression and innate immune cell interaction is pivotal to understand and treat cancer. Here we describe a method to conditionally induce epithelial cell transformations and the subsequent live imaging of innate immune cell interaction with HRASG12V expressing skin cells in zebrafish larvae.

Here we describe a method to conditionally induce epithelial cell transformation by the use of the 4-Hydroxytamoxifen (4-OHT) inducible KalTA4-ERT2/UAS ...

Use of the TetON System to Study Molecular Mechanisms of Zebrafish Regeneration

The zebrafish has become a very important model organism for studying vertebrate development, physiology, disease, and tissue regeneration. A thorough understanding of the molecular and cellular mechanisms involved requires experimental tools that allow for inducible, tissue-specific manipulation of gene expression or signaling pathways. Therefore, we and others have recently adapted the TetON system for use in zebrafish. The TetON system facilitates temporally and spatially-controlled gene expression and we have recently used this tool to probe for tissue-specific functions of Wnt/beta&#8211;catenin signaling during zebrafish tail fin regeneration. Here we describe the workflow for using the TetON system to achieve inducible, tissue-specific gene expression in the adult regenerating zebrafish tail fin. This includes the generation of stable transgenic TetActivator and TetResponder lines, transgene induction and techniques for verification of tissue-specific gene expression in the fin regenerate. Thus, this protocol serves as blueprint for setting up a functional TetON system in zebrafish and its subsequent use, in particular for studying fin regeneration.

Here we outline the workflow for using the TetON system to achieve tissue-specific gene expression in the adult regenerating zebrafish tail fin.

The zebrafish has become a very important model organism for studying vertebrate development, physiology, disease, and tissue regeneration. A thorough ...

Analysis of Zebrafish Larvae Skeletal Muscle Integrity with Evans Blue Dye

The zebrafish model is an emerging system for the study of neuromuscular disorders. In the study of neuromuscular diseases, the integrity of the muscle membrane is a critical disease determinant. To date, numerous neuromuscular conditions display degenerating muscle fibers with abnormal membrane integrity; this is most commonly observed in muscular dystrophies. Evans Blue Dye (EBD) is a vital, cell permeable dye that is rapidly taken into degenerating, damaged, or apoptotic cells; in contrast, it is not taken up by cells with an intact membrane. EBD injection is commonly employed to ascertain muscle integrity in mouse models of neuromuscular diseases. However, such EBD experiments require muscle dissection and/or sectioning prior to analysis. In contrast, EBD uptake in zebrafish is visualized in live, intact preparations. Here, we demonstrate a simple and straightforward methodology for performing EBD injections and analysis in live zebrafish. In addition, we demonstrate a co-injection strategy to increase efficacy of EBD analysis. Overall, this video article provides an outline to perform EBD injection and characterization in zebrafish models of neuromuscular disease.

In this study, we describe a straightforward method to perform Evans Blue Dye (EBD) analysis on zebrafish larvae. This technique is a powerful tool for the characterization of skeletal muscle integrity and delineation of zebrafish models of muscular dystrophy, and is a valuable method for the development of novel therapeutics.

The zebrafish model is an emerging system for the study of neuromuscular disorders.  In the study of neuromuscular diseases, the integrity of the muscle ...

Imaging Subcellular Structures in the Living Zebrafish Embryo

In vivo imaging provides unprecedented access to the dynamic behavior of cellular and subcellular structures in their natural context. Performing such imaging experiments in higher vertebrates such as mammals generally requires surgical access to the system under study. The optical accessibility of embryonic and larval zebrafish allows such invasive procedures to be circumvented and permits imaging in the intact organism. Indeed the zebrafish is now a well-established model to visualize dynamic cellular behaviors using in vivo microscopy in a wide range of developmental contexts from proliferation to migration and differentiation. A more recent development is the increasing use of zebrafish to study subcellular events including mitochondrial trafficking and centrosome dynamics. The relative ease with which these subcellular structures can be genetically labeled by fluorescent proteins and the use of light microscopy techniques to image them is transforming the zebrafish into an in vivo model of cell biology. Here we describe methods to generate genetic constructs that fluorescently label organelles, highlighting mitochondria and centrosomes as specific examples. We use the bipartite Gal4-UAS system in multiple configurations to restrict expression to specific cell-types and provide protocols to generate transiently expressing and stable transgenic fish. Finally, we provide guidelines for choosing light microscopy methods that are most suitable for imaging subcellular dynamics.

Imaging the dynamic behavior of organelles and other subcellular structures in vivo can shed light on their function in physiological and disease conditions. Here, we present methods for genetically tagging two organelles, centrosomes and mitochondria, and imaging their dynamics in living zebrafish embryos using wide-field and confocal microscopy.

In vivo imaging provides unprecedented access to the dynamic behavior of cellular and subcellular structures in their natural context. Performing such ...

Rearing the Fruit Fly Drosophila melanogaster Under Axenic and Gnotobiotic Conditions

The influence of microbes on myriad animal traits and behaviors has been increasingly recognized in recent years. The fruit fly Drosophila melanogaster is a model for understanding microbial interactions with animal hosts, facilitated by approaches to rear large sample sizes of Drosophila under microorganism-free (axenic) conditions, or with defined microbial communities (gnotobiotic). This work outlines a method for collection of Drosophila embryos, hypochlorite dechorionation and sterilization, and transfer to sterile diet. Sterilized embryos are transferred to sterile diet in 50 ml centrifuge tubes, and developing larvae and adults remain free of any exogenous microbes until the vials are opened. Alternatively, flies with a defined microbiota can be reared by inoculating sterile diet and embryos with microbial species of interest. We describe the introduction of 4 bacterial species to establish a representative gnotobiotic microbiota in Drosophila. Finally, we describe approaches for confirming bacterial community composition, including testing if axenic Drosophila remain bacteria-free into adulthood.

Rearing the Fruit Fly Drosophila melanogaster Under Axenic and Gnotobiotic Conditions

A method for rearing Drosophila melanogaster under axenic and gnotobiotic conditions is presented. Fly embryos are dechorionated in sodium hypochlorite, transferred aseptically to sterile diet, and reared in closed containers. Inoculating diet and embryos with bacteria leads to gnotobiotic associations, and bacterial presence is confirmed by plating whole-body Drosophila homogenates.

The influence of microbes on myriad animal traits and behaviors has been increasingly recognized in recent years. The fruit fly Drosophila melanogaster is ...

Inducing Complete Polyp Regeneration from the Aboral Physa of the Starlet Sea Anemone Nematostella vectensis

Cnidarians, and specifically Hydra, were the first animals shown to regenerate damaged or severed structures, and indeed such studies arguably launched modern biological inquiry through the work of Trembley more than 250 years ago. Presently the study of regeneration has seen a resurgence using both &#34;classic&#34; regenerative organisms, such as the Hydra, planaria and Urodeles, as well as a widening spectrum of species spanning the range of metazoa, from sponges through mammals. Besides its intrinsic interest as a biological phenomenon, understanding how regeneration works in a variety of species will inform us about whether regenerative processes share common features and/or species or context-specific cellular and molecular mechanisms. The starlet sea anemone, Nematostella vectensis, is an emerging model organism for regeneration. Like Hydra, Nematostella is a member of the ancient phylum, cnidaria, but within the class anthozoa, a sister clade to the hydrozoa that is evolutionarily more basal. Thus aspects of regeneration in Nematostella will be interesting to compare and contrast with those of Hydra and other cnidarians. In this article, we present a method to bisect, observe and classify regeneration of the aboral end of the Nematostella adult, which is called the physa. The physa naturally undergoes fission as a means of asexual reproduction, and either natural fission or manual amputation of the physa triggers re-growth and reformation of complex morphologies. Here we have codified these simple morphological changes in a Nematostella Regeneration Staging System (the NRSS). We use the NRSS to test the effects of chloroquine, an inhibitor of lysosomal function that blocks autophagy. The results show that the regeneration of polyp structures, particularly the mesenteries, is abnormal when autophagy is inhibited.

Inducing Complete Polyp Regeneration from the Aboral Physa of the Starlet Sea Anemone Nematostella vectensis

Here we demonstrate how to induce and monitor regeneration in the Starlet Sea Anemone Nematostella vectensis, a model cnidarian anthozoan. We demonstrate how to amputate and categorize regeneration using a morphological staging system, and we use this system to reveal a requirement for autophagy in regenerating polyp structures.

Cnidarians, and specifically Hydra, were the first animals shown to regenerate damaged or severed structures, and indeed such studies arguably launched ...

AFM and Microrheology in the Zebrafish Embryo Yolk Cell

Elucidating the factors that direct the spatio-temporal organization of evolving tissues is one of the primary purposes in the study of development. Various propositions claim to have been important contributions to the understanding of the mechanical properties of cells and tissues in their spatiotemporal organization in different developmental and morphogenetic processes. However, due to the lack of reliable and accessible tools to measure material properties and tensional parameters in vivo, validating these hypotheses has been difficult. Here we present methods employing atomic force microscopy (AFM) and particle tracking with the aim of quantifying the mechanical properties of the intact zebrafish embryo yolk cell during epiboly. Epiboly is an early conserved developmental process whose study is facilitated by the transparency of the embryo. These methods are simple to implement, reliable, and widely applicable since they overcome intrusive interventions that could affect tissue mechanics. A simple strategy was applied for the mounting of specimens, AFM recording, and nanoparticle injections and tracking. This approach makes these methods easily adaptable to other developmental times or organisms.

The lack of tools to measure material properties and tensional parameters in vivo prevents validating their roles during development. We employed atomic force microscopy (AFM) and nanoparticle-tracking to quantify mechanical features on the intact zebrafish embryo yolk cell during epiboly. These methods are reliable and widely applicable avoiding intrusive interventions.

Elucidating the factors that direct the spatio-temporal organization of evolving tissues is one of the primary purposes in the study of development. ...

Visualization of Cellular Electrical Activity in Zebrafish Early Embryos and Tumors

Bioelectricity, endogenous electrical signaling mediated by ion channels and pumps located on the cell membrane, plays important roles in signaling processes of excitable neuronal and muscular cells and many other biological processes, such as embryonic developmental patterning. However, there is a need for in vivo electrical activity monitoring in vertebrate embryogenesis. The advances of genetically encoded fluorescent voltage indicators (GEVIs) have made it possible to provide a solution for this challenge. Here, we describe how to create a transgenic voltage indicator zebrafish using the established voltage indicator, ASAP1 (Accelerated Sensor of Action Potentials 1), as an example. The Tol2 kit and a ubiquitous zebrafish promoter, ubi, were chosen in this study. We also explain&#160;the processes of Gateway site-specific cloning, Tol2 transposon-based zebrafish transgenesis, and the imaging process for early-stage fish embryos and fish tumors using regular epifluorescent microscopes. Using this fish line, we found that there are cellular electric voltage changes during zebrafish embryogenesis, and fish larval movement. Furthermore, it was observed that in a few zebrafish malignant peripheral nerve sheath tumors, the tumor cells were generally polarized compared to the surrounding normal tissues.

Here, we show the process of creating a cellular electric voltage reporter zebrafish line to visualize embryonic development, movement, and fish tumor cells in vivo.

Bioelectricity, endogenous electrical signaling mediated by ion channels and pumps located on the cell membrane, plays important roles in signaling ...

An In Vivo Method to Study Mouse Blood-Testis Barrier Integrity

Spermatogenesis is the development of spermatogonia into mature spermatozoa in the seminiferous tubules of the testis. This process is supported by Sertoli cell junctions at the blood-testis barrier (BTB), which is the tightest tissue barrier in the mammalian body and segregates the seminiferous epithelium into two compartments, a basal and an adluminal. The BTB creates a unique microenvironment for germ cells in meiosis I/II and for the development of postmeiotic spermatids into spermatozoa via spermiogenesis. Here, we describe a reliable assay to monitor BTB integrity of mouse testis in vivo. An intact BTB blocks the diffusion of FITC-conjugated inulin from the basal to the apical compartment of the seminiferous tubules. This technique is suitable for studying gene candidates, viruses, or environmental toxicants that may affect BTB function or integrity, with an easy procedure and a minimal requirement of surgical skills compared to alternative methods.

An In Vivo Method to Study Mouse Blood-Testis Barrier Integrity

Here, we present a protocol to assess the blood-testis barrier integrity by injecting inulin-FITC into testes. This is an efficient in vivo method to study blood-testis barrier integrity that can be compromised by genetic and environmental elements.

Spermatogenesis is the development of spermatogonia into mature spermatozoa in the seminiferous tubules of the testis. This process is supported by ...

In Vitro Assays to Evaluate the Migration, Invasion, and Proliferation of Immortalized Human First-trimester Trophoblast Cell Lines

Cell movement is a critical property of trophoblasts during placental development and early pregnancy. The significance of proper trophoblast migration and invasion is demonstrated by pregnancy disorders such as pre-eclampsia and intrauterine growth restriction, which are associated with inadequate trophoblast invasion of the maternal vasculature. Unfortunately, our understanding of the mechanisms by which the placenta develops from migrating trophoblasts is limited. In vitro analysis of cell migration via the scratch assay is a useful tool in identifying factors that regulate trophoblast migratory capacity. However, this assay alone does not define the cellular changes that can result in altered cell migration. This protocol describes three different in vitro assays that are used collectively to evaluate trophoblast cell movement: the scratch assay, the invasion assay, and the proliferation assay. The protocols described here may also be modified for use in other cell lines to quantify cell movement in response to stimuli. These methods allow investigators to identify individual factors that contribute to the cell movement and provide a thorough examination of potential mechanisms underlying apparent changes in cell migration.

Here, we present a highly accessible protocol for evaluating the cell movement in human trophoblast cells using three in vitro assays: the scratch assay, the transwell invasion assay, and the cell proliferation assay.

Cell movement is a critical property of trophoblasts during placental development and early pregnancy. The significance of proper trophoblast migration ...

The Complete and Updated "Rotifer Polyculture Method" for Rearing First Feeding Zebrafish

Podsumowanie

Przeglądaj więcej filmów

Rozdziały w tym wideo

Live Imaging of Innate Immune and Preneoplastic Cell Interactions Using an Inducible Gal4/UAS Expression System in Larval Zebrafish Skin

Use of the TetON System to Study Molecular Mechanisms of Zebrafish Regeneration

Analysis of Zebrafish Larvae Skeletal Muscle Integrity with Evans Blue Dye

Imaging Subcellular Structures in the Living Zebrafish Embryo

Rearing the Fruit Fly Drosophila melanogaster Under Axenic and Gnotobiotic Conditions

Inducing Complete Polyp Regeneration from the Aboral Physa of the Starlet Sea Anemone Nematostella vectensis

AFM and Microrheology in the Zebrafish Embryo Yolk Cell

Visualization of Cellular Electrical Activity in Zebrafish Early Embryos and Tumors

An In Vivo Method to Study Mouse Blood-Testis Barrier Integrity

In Vitro Assays to Evaluate the Migration, Invasion, and Proliferation of Immortalized Human First-trimester Trophoblast Cell Lines