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><tbody><tr><td>&lt;em&gt;&lt;strong&gt;Rotifer Culture Infrastructure&lt;/strong&gt;&lt;/em&gt;</td><td></td><td></td><td></td></tr><tr><td>100 L Culture Vessel</td><td>Aquaneering</td><td>Custom</td><td>Polycarbonate culture vessel, conical bottomed, with drain valve</td></tr><tr><td>5 Gallon Culture Bucket Kit</td><td>Reed Mariculture</td><td>CCS Starter Kit</td><td>Small volume culture vessel for small facilities</td></tr><tr><td>Rigid Clear Tubing 1/2&quot; O.D., 36&amp;rdquo;</td><td>Pentair Aquatic Ecosystems</td><td>16025</td><td>Rigid clear tubing for air delivery</td></tr><tr><td>Mesh tube</td><td>Pentair Aquatic Ecosystems</td><td>RT444X</td><td>Mesh tube support for floss filter</td></tr><tr><td>Rotifer Floss</td><td>Reed Mariculture</td><td>Rotifer floss 12&amp;rdquo; x 42&amp;rdquo;</td><td>Particulate waste trap</td></tr><tr><td>Peristaltic Metering Timer Pump, 5 GPD</td><td>Grainger</td><td>38M003&amp;nbsp;</td><td>Metering pump with timer for dosing feed to rotifers</td></tr><tr><td>Peristaltic Metering Timer Pump, 1-100 mL/hr (for smaller-scale culture)</td><td>Coral Vue</td><td>SKU: IC-LQD-DSR</td><td>Metering pump with timer for dosing feed to rotifers</td></tr><tr><td>Silicone Tubing&amp;nbsp;</td><td>Cole Parmer</td><td></td><td>Tubing for algae delivery to rotifer vessel</td></tr><tr><td>Rigid Clear Tubing &quot; O.D.,36&amp;rdquo;</td><td>Pentair Aquatic Ecosystems</td><td>16025</td><td>Rigid clear tubing for air delivery to algae paste</td></tr><tr><td>Rigid Clear Tubing O.D., 36&amp;rdquo;</td><td>Pentair Aquatic Ecosystems</td><td>16025</td><td>Rigid clear tubing for algae delivery</td></tr><tr><td>&lt;em&gt;&lt;strong&gt;Rotifers&lt;/strong&gt;&lt;/em&gt;</td><td></td><td></td><td></td></tr><tr><td>Live Rotifers Brachionus plicatilis Type L</td><td>Reed Mariculture</td><td>Type L 5 million</td><td>Rotifer stock culture for system startup</td></tr><tr><td>&lt;em&gt;&lt;strong&gt;Rotifer Feed&lt;/strong&gt;&lt;/em&gt;</td><td></td><td></td><td></td></tr><tr><td>Sodium hydroxymethylsulfonate</td><td>Reed Mariculture</td><td>ClorAm-X&lt;sup&gt;&amp;reg;&lt;/sup&gt; 1lb tub</td><td>Ammonia reducer for algae feed mix</td></tr><tr><td>Sodium Bicarbonate</td><td>Fisher Scientific</td><td>S25533B</td><td>pH buffer for algae feed mix</td></tr><tr><td>Microalgae concentrate</td><td>Reed Mariculture</td><td>Rotigrow Plus&lt;sup&gt;&amp;reg;&lt;/sup&gt; 1 liter bag</td><td>Nutritionally optimized rotifer feed</td></tr><tr><td>RG Complete</td><td>Reed Mariculture</td><td>RG Complete 6 oz bottle</td><td>All in one microalgae based feed for small scale cultures</td></tr><tr><td>&lt;em&gt;&lt;strong&gt;Water Preparation&lt;/strong&gt;&lt;/em&gt;</td><td></td><td></td><td></td></tr><tr><td>&amp;nbsp;Reef Crystals Reef Salt</td><td>That Fish Place</td><td>198210</td><td>Salt for making culture water (NOTE: this item is an example only; any contaminant free salt formulations may be used).&amp;nbsp;</td></tr><tr><td>Refractometer</td><td>Pentair Aquatic Ecosystems</td><td>SR6</td><td>measuring salinity</td></tr><tr><td>&lt;em&gt;&lt;strong&gt;Rotifer Culture Equipment&lt;/strong&gt;&lt;/em&gt;</td><td></td><td></td><td></td></tr><tr><td>Plankton Collectors 12&quot; Dia, 53 microns</td><td>Pentair Aquatic Ecosystems</td><td>BBPC20</td><td>Mesh screen for collecting rotifers</td></tr><tr><td>Scrub Pads</td><td>Pentair Aquatic Ecosystems</td><td>SCR-58</td><td>Scrub pad for cleaning inside of culturing vessels</td></tr><tr><td>Scrub Brush</td><td></td><td></td><td></td></tr><tr><td>Bucket</td><td>Grainger Supply&amp;nbsp;</td><td>43Y530</td><td>Graduated bucket for mixing culture water</td></tr><tr><td>Hatching Jar</td><td>Pentair Aquatic Ecosystems</td><td>J30</td><td>Storage of algae feed mix</td></tr><tr><td>Lugol&amp;rsquo;s Solution, Dilute</td><td>Fisher Scientific</td><td>S99481</td><td>Agent used to immobilize live rotifers for counting</td></tr><tr><td>Sedgewick-Rafter plankton counting slide with grid&amp;nbsp;</td><td>Pentair Aquatic Eco-Systems</td><td>M415</td><td>Counting rotifers</td></tr><tr><td>&lt;em&gt;&lt;strong&gt;Miscelleneous&lt;/strong&gt;&lt;/em&gt;</td><td></td><td></td><td></td></tr><tr><td>Tea Strainer</td><td>Kitchenworks</td><td>971972</td><td>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 ...

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Universidad San Sebastian

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

Profiling DNA Replication Timing Using Zebrafish as an In Vivo Model System

DNA replication timing is an important cellular characteristic, exhibiting significant relationships with chromatin structure, transcription, and DNA mutation rates. Changes in replication timing occur during development and in cancer, but the role replication timing plays in development and disease is not known. Zebrafish were recently established as an in vivo model system to study replication timing. Here is detailed the protocols for using the zebrafish to determine DNA replication timing. After sorting cells from embryos and adult zebrafish, high-resolution genome-wide DNA replication timing patterns can be constructed by determining changes in DNA copy number through analysis of next generation sequencing data. The zebrafish model system allows for evaluation of the replication timing changes that occur in vivo throughout development, and can also be used to assess changes in individual cell types, disease models, or mutant lines. These methods will enable studies investigating the mechanisms and determinants of replication timing establishment and maintenance during development, the role replication timing plays in mutations and tumorigenesis, and the effects of perturbing replication timing on development and disease.

Profiling DNA Replication Timing Using Zebrafish as an In Vivo Model System

Zebrafish were recently used as an in vivo model system to study DNA replication timing during development. Here is detailed the protocols for using zebrafish embryos to profile replication timing. This protocol can be easily adapted to study replication timing in mutants, individual cell types, disease models, and other species.

DNA replication timing is an important cellular characteristic, exhibiting significant relationships with chromatin structure, transcription, and DNA ...

Genetics

Development of an Ethanol-induced Fibrotic Liver Model in Zebrafish to Study Progenitor Cell-mediated Hepatocyte Regeneration

Sustained liver fibrosis with continuation of extracellular matrix (ECM) protein build-up results in the loss of cellular competency of the liver, leading to cirrhosis with hepatocellular dysfunction. Among multiple hepatic insults, alcohol abuse can lead to significant health problems including liver failure and hepatocellular carcinoma. Nonetheless, the identity of endogenous cellular sources that regenerate hepatocytes in response to alcohol has not been properly investigated. Moreover, few studies have effectively modeled hepatocyte regeneration upon alcohol-induced injury. We recently reported on establishing an ethanol (EtOH)-induced fibrotic liver model in zebrafish in which hepatic progenitor cells (HPCs) gave rise to hepatocytes upon near-complete hepatocyte loss in the presence of fibrogenic stimulus. Furthermore, through chemical screens using this model, we identified multiple small molecules that enhance hepatocyte regeneration. Here we describe in detail the procedures to develop an EtOH-induced fibrotic liver model and to perform chemical screens using this model in zebrafish. This protocol will be a critical tool to delineate the molecular and cellular mechanisms of how hepatocyte regenerates in the fibrotic liver. Furthermore, these methods will facilitate potential discovery of novel therapeutic strategies for chronic liver disease in vivo.

Sustained fibrosis with deposition of excessive extracellular matrix proteins leads to cirrhosis. Alcohol abuse is one of the main causes of severe liver disease. We established an ethanol-induced zebrafish fibrotic liver model to study the mechanisms and strategies of promoting hepatocyte regeneration upon alcohol-induced injury.

Sustained liver fibrosis with continuation of extracellular matrix (ECM) protein build-up results in the loss of cellular competency of the liver, leading ...

Culture and Transfection of Zebrafish Primary Cells

Zebrafish embryos are transparent and develop rapidly outside the mother, thus allowing for excellent in vivo imaging of dynamic biological processes in an intact and developing vertebrate. However, the detailed imaging of the morphologies of distinct cell types and subcellular structures is limited in whole mounts. Therefore, we established an efficient and easy-to-use protocol to culture live primary cells from zebrafish embryos and adult tissue.
In brief, 2 dpf zebrafish embryos are dechorionated, deyolked, sterilized, and dissociated to single cells with collagenase. After a filtration step, primary cells are plated onto glass bottom dishes and cultivated for several days. Fresh cultures, as much as long term differenciated ones, can be used for high resolution confocal imaging studies. The culture contains different cell types, with striated myocytes and neurons being prominent on poly-L-lysine coating. To specifically label subcellular structures by fluorescent marker proteins, we also established an electroporation protocol which allows the transfection of plasmid DNA into different cell types, including neurons. Thus, in the presence of operator defined stimuli, complex cell behavior, and intracellular dynamics of primary zebrafish cells can be assessed with high spatial and temporal resolution. In addition, by using adult zebrafish brain, we demonstrate that the described dissociation technique, as well as the basic culturing conditions, also work for adult zebrafish tissue.

We present an efficient and easy-to-use protocol for preparing primary cell cultures of zebrafish embryos for transfection and live cell imaging as well as a protocol to prepare primary cells from adult zebrafish brain.

Zebrafish embryos are transparent and develop rapidly outside the mother, thus allowing for excellent in vivo imaging of dynamic biological processes in ...

Optimizing GF Zebrafish Models: Generation, Maintenance, and Microbial Studies

Generation, Maintenance, and Identification of Germ-Free Zebrafish Models from Larvae to Juvenile Stages

Zebrafish serve as valuable models for research on growth, immunity, and gut microbiota due to their genomic similarities with mammals, transparent embryos developed in a relatively clean chorion environment, and extremely rapid development of larvae compared to rodent models. Germ-free (GF) zebrafish (Danio rerio) are crucial for evaluating pollutant toxicity and establishing human-like disease models related to microbial functions. In comparison to conventionally raised (CR) models (fish in common husbandry), GF zebrafish allow for more accurate manipulation of the host microbiota, aiding in determining the causal relationship between microorganisms and hosts. Consequently, they play a critical role in advancing our understanding of these relationships. However, GF zebrafish models are typically generated and researched during the early life stages (from embryos to larvae) due to limitations in immune function and nutrient absorption. This study optimizes the generation, maintenance, and identification of early GF zebrafish models without feeding and with long-term feeding using GF food (such as Artemia sp., brine shrimp). Throughout the process, daily sampling and culture were performed and identified through multiple detections, including plates and 16S rRNA sequencing. The aseptic rate, survival, and developmental indexes of GF zebrafish were recorded to ensure the quality and quantity of the generated models. Importantly, this study provides details on bacterial isolation and infection techniques for GF fish, enabling the efficient creation of GF fish models from larvae to juvenile stages with GF food support. By applying these procedures in biomedical research, scientists can better understand the relationships between intestinal bacterial functions and host health.

Author Spotlight: Establishing Germ-Free Zebrafish for Microbiota-Host Relationship Studies

This protocol outlines the primary steps for obtaining germ-free (GF) fish embryos and maintaining them from larvae until the juvenile stage, including sampling and detecting their sterile status. The use of GF models with infection is important for understanding the role of microbes in host health.

Zebrafish serve as valuable models for research on growth, immunity, and gut microbiota due to their genomic similarities with mammals, transparent ...

Regular Care and Maintenance of a Zebrafish (Danio rerio) Laboratory: An Introduction

This protocol describes regular care and maintenance of a zebrafish laboratory. Zebrafish are now gaining popularity in genetics, pharmacological and behavioural research. As a vertebrate, zebrafish share considerable genetic sequence similarity with humans and are being used as an animal model for various human disease conditions. The advantages of zebrafish in comparison to other common vertebrate models include high fecundity, low maintenance cost, transparent embryos, and rapid development. Due to the spur of interest in zebrafish research, the need to establish and maintain a productive zebrafish housing facility is also increasing. Although literature is available for the maintenance of a zebrafish laboratory, a concise video protocol is lacking. This video illustrates the protocol for regular housing, feeding, breeding and raising of zebrafish larvae. This process will help researchers to understand the natural behaviour and optimal conditions of zebrafish husbandry and hence troubleshoot experimental issues that originate from the fish husbandry conditions. This protocol will be of immense help to researchers planning to establish a zebrafish laboratory, and also to graduate students who are intending to use zebrafish as an animal model.

Regular Care and Maintenance of a Zebrafish Danio rerio Laboratory: An Introduction

This protocol outlines regular maintenance and care to maintain optimal conditions for zebrafish husbandry. The video illustrates the protocol for system maintenance, regular housing, feeding, breeding, and raising of zebrafish larvae.

This protocol describes regular care and maintenance of a zebrafish laboratory. Zebrafish are now gaining popularity in genetics, pharmacological and ...

Biology

Flat Mount Preparation for Observation and Analysis of Zebrafish Embryo Specimens Stained by Whole Mount In situ Hybridization

The zebrafish embryo is now commonly used for basic and biomedical research to investigate the genetic control of developmental processes and to model congenital abnormalities. During the first day of life, the zebrafish embryo progresses through many developmental stages including fertilization, cleavage, gastrulation, segmentation, and the organogenesis of structures such as the kidney, heart, and central nervous system. The anatomy of a young zebrafish embryo presents several challenges for the visualization and analysis of the tissues involved in many of these events because the embryo develops in association with a round yolk mass. Thus, for accurate analysis and imaging of experimental phenotypes in fixed embryonic specimens between the tailbud and 20 somite stage (10 and 19 hours post fertilization (hpf), respectively), such as those stained using whole mount in situ hybridization (WISH), it is often desirable to remove the embryo from the yolk ball and to position it flat on a glass slide. However, performing a flat mount procedure can be tedious. Therefore, successful and efficient flat mount preparation is greatly facilitated through the visual demonstration of the dissection technique, and also helped by using reagents that assist in optimal tissue handling. Here, we provide our WISH protocol for one or two-color detection of gene expression in the zebrafish embryo, and demonstrate how the flat mounting procedure can be performed on this example of a stained fixed specimen. This flat mounting protocol is broadly applicable to the study of many embryonic structures that emerge during early zebrafish development, and can be implemented in conjunction with other staining methods performed on fixed embryo samples.

Flat Mount Preparation for Observation and Analysis of Zebrafish Embryo Specimens Stained by Whole Mount In situ Hybridization

The zebrafish embryo is an excellent model for developmental biology research. During embryogenesis, zebrafish develop with a yolk mass, which presents three-dimensional challenges for sample observation and analysis. This protocol describes how to create two-dimensional flat mount preparations of whole mount in situ (WISH) stained zebrafish embryo specimens.

The zebrafish embryo is now commonly used for basic and biomedical research to investigate the genetic control of developmental processes and to model ...

Construction of an Affordable and Easy-to-Build Zebrafish Facility

In vivo biomedical research is pivotal to translate in vitro findings into clinical advances. Small academic institutions with limited resources find it virtually impossible to build and maintain typical rodent facilities for research. Zebrafish research has been demonstrated to be a valuable alternative for in vivo research in pharmacology, physiology, development and genetic studies. This article demonstrates that a functional zebrafish facility can be built in an easy and affordable manner. We demonstrate that such a facility could be built in about one working day with minimal tools and expertise. The cost of the 27 1.8 L fish tank zebrafish facility constructed in this study was approximately $1,500. We estimate that the maintenance of an initial working 150 fish colony for 3 months is $1,000. This project involved students, who were introduced to aquaculturing of zebrafish for research proposes.

A Zebrafish facility suitable for breeding and in vivo research is built in an easy and affordable manner. Maintenance is minimal. This facility is ideal for student involvement and independent research.

In vivo biomedical research is pivotal to translate in vitro findings into clinical advances. Small academic institutions with limited resources find it ...

High-throughput Measurement of Gut Transit Time Using Larval Zebrafish

Zebrafish are used as alternative model organisms for drug safety testing. The gastrointestinal (GI) tract of zebrafish has genetic, neuronal, and pharmacological similarities to that of the mammals. GI intolerance during clinical testing of drug candidates is common and may pose a serious threat to human health. Testing for GI toxicity in preclinical mammalian models can be expensive in terms of time, test compound, and labor. The high-throughput method presented here may be used to predict GI safety issues. Compared to mammalian models, this method allows for more expedient assessment of test compound effects on GI transit while using low quantities of compound. In this method, larval zebrafish (7 days post fertilization) are fed food containing a fluorescent label. After feeding, each larval fish is placed into a well of a 96-conical-bottom-well plate and dosed with test compound (dissolved in water) or the vehicle. As gut transit occurs, fecal matter accumulates on the bottom of the wells, and the rate at which this happens is monitored by measuring fluorescence from the bottom of the well repeatedly over time using a plate spectrophotometer. The fluorescence from larvae in a given treatment group are averaged and these values are graphed along with standard error, for each measurement time, yielding a curve representing average transit of food over time. Effects on gut transit time are identified by comparing the area under the curve for each treatment group to that of the vehicle-treated group. This method detected changes in zebrafish GI transit time induced by drugs with known clinical GI effects; it can be employed to interrogate dozens of treatments for GI effects per day. As such, safer compounds can be quickly prioritized for mammalian testing, which expedites discovery and proffers 3Rs advancement.

The goal of this protocol is to measure the transit time of fluorescently labeled food through the gut of larval zebrafish in a high throughput fashion.

Zebrafish are used as alternative model organisms for drug safety testing. The gastrointestinal (GI) tract of zebrafish has genetic, neuronal, and ...

Small Molecule Screening and Toxicity Testing in Early-stage Zebrafish Larvae

Zebrafish have become a prominent model organism for human translational research, toxicity testing of small molecules and environmental pollutants, therapeutic drug discovery, and other biomedical research applications. The presented protocol aims to provide a clear guide for drug screening and toxicity testing in early-stage zebrafish larvae (aged 3-7 days post fertilization, dpf). Several methodologies describing toxicity testing in zebrafish larvae have been published, albeit lacking a standardized protocol for exposure durations, rate of media changes, and morphology scoring, amongst others. We experimentally determined the ideal larvae ages for conducting the experiments, the plate sizes that work best for those ages, the rate of media changes that provide reliable results while limiting wastage&#160;of test compound, and a morphology scoring system that is easily quantifiable in a statistical manner. The manuscript describes in detail a protocol for toxicity testing in zebrafish larvae aged between&#160;3 to 7 dpf, aiming to target the gap in drug toxicity testing and taking the first step towards a standardized method similar to the Fish Embryo Acute Toxicity test.

This protocol describes a drug screening procedure in zebrafish larvae 3-7 days post fertilization followed by morphological assessment.

Zebrafish have become a prominent model organism for human translational research, toxicity testing of small molecules and environmental pollutants, ...

Zebrafish Maintenance and Husbandry

The zebrafish (Danio rerio) is a powerful vertebrate model system for studying development, modeling disease, and screening for novel therapeutics. Due to their small size, large numbers of zebrafish can be housed in the laboratory at low cost. Although zebrafish are relatively easy to maintain, special consideration must be given to both diet and water quality to in order to optimize fish health and reproductive success.
This video will provide an overview of zebrafish husbandry and maintenance in the lab. After a brief review of the natural zebrafish habitat, techniques essential to recreating this environment in the lab will be discussed, including key elements of fish facility water recirculation systems and the preparation of brine shrimp as part of the zebrafish diet. Additionally, the presentation will include information on how specific zebrafish strains are tracked in a laboratory setting, with specific reference to the collection of tail fin samples for DNA extraction and genotyping. Finally, experimental modifications of the zebrafish environment will be discussed as a means to further our understanding of these fish, and in turn, ourselves.

Maintenance and Husbandry of Danio rerio Zebrafish

The zebrafish (Danio rerio) is a powerful vertebrate model system for studying development, modeling disease, and screening for novel therapeutics. Due to ...

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

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Chapters in this video

Profiling DNA Replication Timing Using Zebrafish as an In Vivo Model System

Development of an Ethanol-induced Fibrotic Liver Model in Zebrafish to Study Progenitor Cell-mediated Hepatocyte Regeneration

Culture and Transfection of Zebrafish Primary Cells

Generation, Maintenance, and Identification of Germ-Free Zebrafish Models from Larvae to Juvenile Stages

Regular Care and Maintenance of a Zebrafish (Danio rerio) Laboratory: An Introduction

Flat Mount Preparation for Observation and Analysis of Zebrafish Embryo Specimens Stained by Whole Mount In situ Hybridization

Construction of an Affordable and Easy-to-Build Zebrafish Facility

High-throughput Measurement of Gut Transit Time Using Larval Zebrafish

Small Molecule Screening and Toxicity Testing in Early-stage Zebrafish Larvae

Zebrafish Maintenance and Husbandry