The authors would like to thank Tammy Esmaili for her assistance in the management of laboratory consumables. AA is the recipient of a PhD Scholarship from the Centre of Excellence for Alzheimer's Disease Research and Care, School of Medical Sciences, Edith Cowan University. MC receives PhD funding support from the Rotary Club, Perth. AM, GV, KT, and RNM are funded by the McCusker Alzheimer's Research foundation.

1. System Maintenance
<ol>
 <li>Zebrafish are kept in a circulating system that continuously filters and aerates the system water to maintain the water quality required for a healthy aquatic environment. The circulating system also helps to filter excess food and fish excreta. Different companies provide zebrafish systems but we use systems from Aquatic Habitats, USA in our laboratory. The room temperature or the tank temperature is generally maintained between 26-28.5 &deg;C and the lighting conditions are 14:10 hr (light: dark). A zebrafish system from Aquatic Habitats (e.g., Benchtop system) costs ~9,000 USD. This benchtop system with two shelves can hold six 10-liter, twelve 3-liter, or twenty 1.5-liter tanks on each shelf. Multiple lines of fish (e.g., transgenic, mutant, wild type) can also be housed on the same system. </li>
 <li>A set of different kinds of filters are used in the system. In our system, water from all the tanks passes through a 120-micron filter pad, 50-micron canister filter, biological filter , active carbon absorption filter and UV disinfection filter before being circulated back into the tank. De-chlorinated/aged water is used in the zebrafish system. Water can be de-chlorinated by ageing for at least 48 hr. Under ideal conditions, water should be kept in a reservoir with a pump circulating the water to keep it warm, and expedite the de-chlorination.</li>
 <li>The pH of the system water should be checked daily and maintained between 6.8 and 7.5. When necessary, sodium bicarbonate should be used to increase the pH. </li>
 <li>Fish tanks should be cleaned regularly. To clean a fish tank, close the water flow to this tank, drain excess water by tilting the tank backwards and remove the tank carefully from the system. Dirt and algae growth will be apparent on the bottom and sides of the tank. </li>
 <li>Place the baffle into a clean tank, and fill with de-chlorinated water (a.k.a. System water). Carefully transfer the fish into this tank with a fish net. Close the lid and transfer the name tag of the tank. Carefully place the clean tank into the system and switch on the water supply. </li>
 <li>To decontaminate the fish net, spray with 70% ethanol, rinse in water, and let it dry before re-using. Remove the baffle from the dirty tank and spray both parts with 70% ethanol. Rinse thoroughly with tap water and allow the tank and baffle to dry fully before re-using. </li>
 <li>The circulating system filters have to be checked and changed regularly to ensure their proper function. These filters should be changed regularly to ensure proper and clean water supply to all the fish tanks.</li>
 <li>The 120-micron filter pad is usually repositioned or replaced daily; reposition towards the water flow to use it completely before replacing with a new one. </li>
 <li>The Canister filter should be changed weekly. To change the canister filter, remove the filter unit by twisting anticlockwise with a wrench or hands. Place a plastic sheet or towel below to prevent or absorb water spillage. Replace the canister filter with a new one and fit the filter unit back into the system, carefully tighten with hands and a wrench if necessary. </li>
 <li>The carbon filter should be changed fortnightly (every two weeks). To change the carbon filter, remove the carbon filter unit carefully with a wrench. Discard the used activated carbon and replace it with new activated carbon. Re-fit the carbon holder and place it back into the filter unit. Fit the filter unit back into the system. Switch on the system and check that water is flowing to the filter. If necessary, flush some water through the pipes to clean dirt in the pipes before fitting the filter into the system. </li>
 <li>The biological filter should be cleaned every six months. A biological filter is generally located between the canister and carbon filter in the circulating system. To change the biological filter, remove the filter unit from the system. Release the pressure from the filter by pressing the pressure release button. Unscrew the filter with a wrench. Generally two people are needed for this step. Remove the lid of the filter unit. Empty the contents of the filter into a sieved container to separate the siporax from the water. Note: Siporax is a fine pore bio-filter media that has the ability to perform both nitrification and de-nitrification. </li>
 <li>If the siporax is very dirty replace it with new. Fill the filter unit with system water (de-chlorinated water), close the lid, return the filter unit to the system and switch on the water supply. 
 Note: In an acclimated aquarium, the siporax will be home to several nitrifying bacteria. These microorganisms are critical for maintaining the nitrogen cycle within the aquarium, and removing the primary housing for the biological filter (the dirty siporax) could result in a serious ammonia spike, followed by a nitrite spike while the new biological filter (with new siporax) is re-establishing. Both of these intermediate states of the nitrogen cycle may be toxic to aquatic organisms, and may kill zebrafish if not responded to appropriately. Therefore, it is important to have a secondary biological filter elsewhere in the zebrafish housing system (e.g., in the collection tank in Aquatic Habitats system) to allow rapid repopulation of these important microorganisms on the new siporax. </li>
 <li>UV filters are used to control system biological contaminants (such as bacteria) and should be replaced every nine-ten months. It should be noted that UV filter disinfection dose rate is ~110 mJ/cm2 at the beginning of the lamp life and the dose rate decreases over the course of time, hence it is necessary to replace the globe even when it appears to still be functional. </li>
</ol>
2. Feeding
<ol>
 <li>Zebrafish can be fed with dry food (food size from 100 microns for larvae to 300/400 microns for adult fish) or live food (brine shrimps). Brine shrimp (Artemia sp.) eggs are available from local pet shops and can be hatched in the laboratory by following the simple steps described below. 
 <ol class="lalpha">
 <li> Dissolve red sea salt in aged water by placing a beaker with salt on a magnetic stirrer for better solubility. Alternatively, salt can be dissolved in water by aerating it with the aeration tube. Instant ocean salt can also be used as a hatching solution if red sea salt is not available. Brine shrimps can tolerate a wide range of salinity; however, we hatch our brine shrimp (10-15 g) in 30-35 g/L red sea salt water. Note: In case brine shrimp eggs are not readily available in the local pet stores then encapsulated brine shrimps can also be used after "decapping" them in batches before adding them to the shrimp hatchery. This might also avoid frequent trips to the pet store.</li>
 <li> Fill the brine shrimp hatcher with salt water and add the shrimp eggs at a concentration of 1.2 tablespoons/liter. Aerate the hatcher vigorously with an air pump and leave the brine shrimp eggs to hatch for ~ 48 hr. 
 Note: It should be noted that un-enriched brine shrimp lose nutritional value every hour post-hatching. Therefore, it may be appropriate to either hatch over 24 hr or enrich over 48 hr in order to obtain the maximum nutritional value from the brine shrimp.</li>
 <li> Waste water from the hatching system is added to quarantine waste to be disinfected with bleach and disposed later on. </li>
 </ol>
 </li>
 <li>To collect the brine shrimps, remove the air pipe and allow the culture to settle for 4-5 min but not for more than 10 min. The hatched brine shrimps gather at the bottom of the Hatcher.
 <ol class="lalpha">
 <li> Collect brine shrimps using the tap at the bottom of the hatcher. Discard the initial flow which consists of un-hatched brine shrimp eggs. </li>
 <li> After discarding the un-hatched brine shrimps, collect the hatched brine shrimps which will be used as zebrafish food. </li>
 <li> Separate the brine shrimps from the salty water, using a brine shrimp collection net (~350 micron nylon mesh). Rinse the brine shrimps from the net into a container using system water. </li>
 </ol>
 </li>
 <li>The collected brine shrimps are generally present in high concentration at the bottom of the container giving it a rather orange colour. </li>
 <li>Feed the brine shrimps to the zebrafish using a pipette or a dropper/squeezy bottle. The amount of food dispensed depends on the population size of individual tanks. The commonly accepted ratio for zebrafish is to receive 4% of body weight in food per day. Zebrafish should never be overfed as this may increase the nitrate level in the water, possibly affecting their breeding, or viability, as some fish may die due to overeating.
 <ol class="lalpha">
 <li>Upon injecting food into the water, hungry fish swim to catch the brine shrimps. </li>
 </ol>
 </li>
 <li>Dry feeding can be performed using an Aquatic Eco-System's/Aquatic Habitats simple spring-based fish food dispenser. Alternatively, dry feeding can also be performed using a simple spoon or by cutting a plastic dropper diagonally with scissors to give it an appearance of a small spoon. </li>
</ol>
3. Breeding
<ol>
 <li>Zebrafish initiate breeding at the onset of light. Fertilized eggs can be obtained either through in-tank breeding or pairwise breeding. While in-tank breeding is more labour-efficient and is implemented for regular embryo collection in our laboratory, pairwise breeding is preferred when genes or mutations are to be screened from individual fish. </li>
 <li>For in-tank breeding, assemble the in-tank breeder and drop slowly into the fish tank after the onset of light. Alternatively, in-tank breeding set-up can be left overnight in the fish tank. </li>
 <li>Leave the in-tank breeder for around 15 min to allow the fish to mate before removing the breeder from the tank and collecting the eggs. </li>
 <li>Pairwise breeding is usually set up late in the afternoon after feeding.</li>
 <li>Assemble the breeding tank and fill it with aged system water. </li>
 <li>Transfer one female and one male to opposite sides of the breeding tank. Females can be distinguished from males because of their bigger underbelly. Males can also be distinguished from females because they are more slender and darker in colour than females. Moreover, males have more yellow colouration in the anal fin compared to females (see Figure 1). When in doubt look for the ovipositor in female zebrafish (see Figure 4).</li>
 <li>Remove the divider the next morning shortly after the onset of light. Allow mating to occur undisturbed for 20 min or until sufficient numbers of embryos are laid at the bottom of the tank. </li>
 <li>After breeding, return the fish to their tanks. Collect the eggs using a strainer. </li>
 <li>Wash the embryos thoroughly with system water. </li>
 <li>Transfer the embryos to a Petri dish by rinsing the strainer with embryo medium; a.k.a. EM3 (NaCl, 13.7 mM; KCl, 0.54 mM; MgSO4, 1.0 mM; CaCl2, 1.3 mM; Na2HPO4, 0.025 mM; KH2PO4, 0.044 mM; NaHCO3, 4.2 mM). </li>
 <li>Embryos can be observed under a microscope. Fertilized eggs are then separated from the unfertilized eggs using a needle and a pipette (see Figure 3). </li>
</ol>
4. Raising of Larvae
<ol>
 <li>Fertilized eggs are kept in an incubator (~28.5 &deg;C) for 72 hr until the larvae are hatched.</li>
 <li>Now the larvae are out of the chorion and swimming freely they are ready to transfer to a main fish tank. Larvae need to be fed from 5 days post-fertilization (dpf) and are kept in embryo medium (composition described in Part 3, 10) or system water. Larvae can be kept in round dishes with ~50% or more of water therein changed on a daily basis. The water change should include removing dead or diseased larvae and any other debris. </li>
 <li>Transfer the larvae gently into a tank containing a small sized baffle (of around 300-400 microns). Dead and diseased larvae should be removed and a few milliliters of water should be added slowly on a daily basis. </li>
 <li>After 14 days, larvae tanks can be shelved into the system, and supplied with a small stream of cycling water (1-2 drops per second). As the larvae grow, water flow can be increased. Different sizes of baffles can be used depending on the size of the larvae (e.g. baffle size 300-400, 500, 700-750, and 1000 microns) and a normal plastic baffle should be used for the adult fish. </li>
 <li>It usually takes 3 months for the embryos to develop into sexually mature adult. </li>
</ol>

<ol>
	<li>Kimmel, C. B., Ballard, W. W., Kimmel, S. R., Ullmann, B., Schilling, T. F. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Stages+of+embryonic+development+of+the+zebrafish.">Stages of embryonic development of the zebrafish.</a> Dev. Dyn. 203, 253-310 (1995).</li><li>Avdesh, A., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Evaluation+of+color+preference+in+zebrafish+for+learning+and+memory.">Evaluation of color preference in zebrafish for learning and memory.</a> J. Alzheimers Dis. 28, 459-469 (2012).</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, 13-34 (2008).</li><li>Best, J. D., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Non-associative+learning+in+larval+zebrafish.">Non-associative learning in larval zebrafish.</a> Neuropsychopharmacology. 33, 1206-1215 (2008).</li><li>Wolman, M. A., Jain, R. A., Liss, L., Granato, M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Chemical+modulation+of+memory+formation+in+larval+zebrafish.">Chemical modulation of memory formation in larval zebrafish.</a> Proc. Natl. Acad. Sci. U.S.A. 108, 15468-15473 (2011).</li><li>Rosen, J. N., Sweeney, M. F., Mably, J. D. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Microinjection+of+Zebrafish+Embryos+to+Analyze+Gene+Function.">Microinjection of Zebrafish Embryos to Analyze Gene Function.</a> J. Vis. Exp. (25), e1115 (2009).</li><li>Gerlai, R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Zebra+fish:+an+uncharted+behavior+genetic+model.">Zebra fish: an uncharted behavior genetic model.</a> Behav. Genet. 33, 461-468 (2003).</li><li>Eddins, D., Petro, A., Williams, P., Cerutti, D. T., Levin, E. D. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Nicotine+effects+on+learning+in+zebrafish:+the+role+of+dopaminergic+systems.">Nicotine effects on learning in zebrafish: the role of dopaminergic systems.</a> Psychopharmacology. (Berl). 202, 103-109 (2009).</li><li>Colwill, R. M., Raymond, M. P., Ferreira, L., Escudero, H. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Visual+discrimination+learning+in+zebrafish+(Danio+rerio.">Visual discrimination learning in zebrafish (Danio rerio.</a> Behav. Processes. 70, 19-31 (2005).</li><li>Blaser, R. E., Chadwick, L., McGinnis, G. C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Behavioral+measures+of+anxiety+in+zebrafish+(Danio+rerio).">Behavioral measures of anxiety in zebrafish (Danio rerio).</a> Behav. Brain Res. 208, 56-62 (2010).</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+guide+for+the+laboratory+use+of+zebrafish+(Danio+rerio).">A guide for the laboratory use of zebrafish (Danio rerio).</a> THE ZEBRAFISH BOOK. , (2007).</li><li>Delaney, M., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Social+interaction+and+distribution+of+female+zebrafish+(Danio+rerio)+in+a+large+aquarium.">Social interaction and distribution of female zebrafish (Danio rerio) in a large aquarium.</a> Biol. Bull. 203, 240-241 (2002).</li></ol>

No conflicts of interest declared.

Zebrafish originate from the Ganges river in northern India and are becoming popular in research in both their adult and larval stages2, reviewed by Spence et al.3. Zebrafish possess several advantages over other animal models such as high fecundity, ease of maintenance, optical clearance of embryos, rapid embryonic development, and low maintenance cost. They are amenable to genetic manipulation6 and suitable for high-throughput drug screening4,5. Their fertilization is external which is advantageous for their use by developmental biologists. Due to these favourable characteristics, zebrafish are gaining popularity in genetics7, pharmacological8, and behavioural research9,10. There are a number of challenges to maintaining a zebrafish facility and zebrafish husbandry to obtain embryos. Herein, we describe our experiences and recommendations in addressing these challenges and outline a protocol for system maintenance, feeding, breeding and raising of the larvae. 
System maintenance
To maintain zebrafish in a healthy condition, it is important to provide them with a clean environment in a properly functioning aquarium system. An important part of this is changing system filters regularly so that all the tanks receive proper water flow and clean water. It is vital to avoid failure of the cycling water supply to each tank due to blocked system pipes. The pipes can be cleaned using a higher than normal water pressure and flow if blockage does occur. Ideally, around 10% of the system's water should be replaced daily to maintain good water quality. Alternatively, water can be replaced while changing the Canister or Carbon filter. This ensures that dirt deposited in the pipes connecting these filters is removed. The quality of water should be checked on a regular basis. Parameters such as alkalinity, pH, temperature, hardness, ammonia, dissolved oxygen, salinity, and conductivity should be considered as important factors in representing the quality of the system water (see Table 2 for details). At the very least nitrate, pH, and temperature should be monitored on a regular basis to ensure good water quality for housing zebrafish. Ideal nitrate (NO3-) levels are &lt;50 mg/L11; if high these levels can be reduced by replacing the water in the circulating system with the fresh system water. Occasionally, filters do not fit well and leak so it is recommended to check for any leaks after a filter change. If the water flow from the main reservoir is blocked either after changing the water pump or changing a filter, the water flow can be restored by either loosening or removing the filter for a few seconds to release any vacuum being generated in the pipes. The time required to change filters can vary depending on various factors such as total biological load on the system, cleanliness of other filters, and dirt deposited in the pipes. Hence, filters should immediately be changed if they appear dirty or if all the tanks are not receiving the correct water supply. It is also recommended that the fish net be cleaned with 70% ethanol, and rinsed in water to decontaminate it, and allowed to dry before being re-used. Drying ensures evaporation of ethanol which otherwise is toxic to fish. 
Most of the zebrafish systems use de-chlorinated tap water; however, some systems use deionized water. It is important to keep the conductivity of the system water between 300 and ~1,500 &mu;S as this reduces the energy the fish needs to maintain body salts. Therefore, zebrafish cannot be kept in deionised water unless salts are added to maintain the optimal conductivity levels. There is a risk of possible high copper concentrations in the system water if tap water is used because the carbon filter does not remove copper. Therefore, users should check for copper concentrations and avoid copper piping where possible.
Feeding
Zebrafish should never be overfed as this may increase the nitrate level in the water, possibly affecting their breeding11, or viability, as some fish may die due to overeating. We recommend providing no more food during any one feeding than a tank of fish can finish within 10 min. It is very important to remove salt from the brine shrimps before feeding them to the zebrafish as excess salt concentration causes death. If more zebrafish eggs are required, fish can be fed three times a day. Cleaning the breeder fish tanks daily also improves levels of egg production. 
When feeding on the Aquatic Habitats systems we usually turn off the water pump and air pump to allow the fish to eat the food for 10 min. This decreases the amount of food that is washed into the filters. However, users must be careful to remember to turn on these pumps again afterwards.
Breeding
Zebrafish are usually at optimal breeding condition between ~3 and 18 months of age. Pairwise breeding should not be performed for two consecutive days11; however, in-tank breeding can be performed daily as a tank can hold many fish which reduces the chance of the same pair of fish being bred for two days in a row. Breeding should be undertaken at regular intervals even if eggs are not required. This process will ensure the breeding cycle of the fish is maintained. It is recommended that there are more females than males in a breeding set-up. Male zebrafish change their female partners on a daily basis12 which further supports this recommendation. Furthermore, within our laboratory we initially experienced problems with breeding, however, using more females than males in a breeding set-up helped solve the problem. Moreover, feeding with a high protein content diet and brine shrimp two-three times a day, mixing fish from different tanks (from different parents), maintaining the temperature of the breeding set-up between 27 and 28 &deg;C, and squeezing the bellies of females with blocked ovary tubes using gentle massage further improved egg production. We recommend keeping a record of fish lines/ origins to avoid in-breeding between siblings. This further improves embryo production. Keeping a record of the number of embryos laid by fish from each tank is also recommended. This assists with keeping a track of the best breeding fish tanks and taking measures to improve breeding in the fish not laying eggs. 
Raising of larvae
Feeding of larvae should commence from 5 dpf (days post fertilization). Young larvae can be fed with dry food of ~100 microns in size (e.g., ZM100) or live food such as paramecium and rotifers (which stimulates growth). The food size can slowly be increased to 200 microns (e.g. ZM200) or 300/400 microns (e.g. ZM300). A population of adult fish should be around 6-7 fish per liter of water. This practice is recommended for better maintenance of BOD (Biological Oxygen Demand) to the tanks.

<table><tbody><tr><td>Zebrafish circulating system</td><td>Aquatic Habitats, USA</td><td>AHAB stand-alone, bench-top systems</td><td></td></tr><tr><td>Sodium Bicarbonate</td><td>Sigma Aldrich</td><td>S6297</td><td></td></tr><tr><td>Fish food dispenser/Dry food feeder</td><td>Aquatic Habitats, USA</td><td>AH19</td><td></td></tr><tr><td>Microscope </td><td>Olympus</td><td>SZX12</td><td></td></tr><tr><td>Dry Food </td><td>ZM Fish food, UK</td><td>ZM100, ZM200, ZM300</td><td></td></tr><tr><td>Brine shrimp eggs</td><td>Salt Creek, Inc., USA</td><td>Premium Grade</td><td></td></tr><tr><td>In-tank breeders</td><td>Aquatic Habitats, USA</td><td>ITSTS-A</td><td></td></tr><tr><td>Activated carbon</td><td>Penn-Plax Pro-crab, USA</td><td>PBC3MF</td><td></td></tr><tr><td>Breeding tanks</td><td>Aquatic Habitats, USA</td><td>SBTANK (1L)&lt;br/&gt; Breeder Tank-2 (2L)</td><td></td></tr><tr><td>Red sea salt</td><td>Red sea salt, USA</td><td>Local pet store/&lt;br/&gt; &lt;a href=&quot;http://www.redseafish.com&quot; target=&quot;_blank&quot;&gt;www.redseafish.com&lt;/a&gt;</td><td></td></tr><tr><td>Filters (Canister, filter pads, and UV) </td><td>Aquatic Habitats, USA</td><td>&lt;a href=&quot;http://www.aquatichabitats.com&quot; target=&quot;_blank&quot;&gt;aquatichabitats.com&lt;/a&gt;</td><td></td></tr><tr><td>Siporax Media</td><td>Aquatic Habitats, USA</td><td>BF 820</td><td></td></tr><tr><td>Brine shrimp net </td><td>Aquatic Habitats, USA</td><td>BSN 1</td><td></td></tr><tr><td>Brine shrimp hatcher</td><td>Aquatic Habitats, USA</td><td>BS252</td><td></td></tr><tr><td>Baffles</td><td>Aquatic Habitats, USA</td><td>&lt;a href=&quot;http://www.aquatichabitats.com&quot; target=&quot;_blank&quot;&gt;aquatichabitats.com&lt;/a&gt;</td><td></td></tr><tr><td colspan="4">&lt;span class=&quot;jove_content&quot;&gt;&lt;strong&gt;Table 1. &lt;/strong&gt;Table of specific reagents and equipment used in this protocol.&lt;/span&gt;</td></tr></tbody></table>

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.

Zebrafish housing and maintenance is easier and cheaper than traditional rodent models. Several thousand zebrafish can be housed in a small laboratory. As a result of this protocol, researchers will be able to manage a zebrafish facility which will provide healthy conditions to the zebrafish. In addition, the following illustrations will help identify fertilized eggs, adult zebrafish, and their food. An illustration of a male zebrafish (Figure 1A) and female zebrafish (Figure 1B &amp; 1C) is shown to help researchers distinguish between a male and a female zebrafish for breeding purpose. Figure 2 depicts a microscopic view of; brine shrimps at 12X (Figure 2A), a single brine shrimp at 90X (Figure 2B), and an unfertilized brine shrimp egg at 90X (Figure 2C). This will help in understanding the difference between unfertilized and fertilized brine shrimps for proper feeding purposes. Fertilized and unfertilized eggs are shown in Figure 3. Figure 3A illustrates a microscopic view of fertilized and unfertilized embryos. Unfertilized embryos are generally opaque and/or with ruptured cell(s) inside the chorion (black arrow) whilst fertilized embryos appears intact and growing to the next cell division state (for detailed reading of different stages of zebrafish embryos see1). Higher magnification view of a fertilized and an unfertilized egg is shown in Figure 3B and 3C respectively. Figure 4 illustrates a female zebrafish's ovipositor to help researchers distinguish between a male and a female zebrafish. 
Several critical problems that might occur in a zebrafish laboratory include blockage of water supply to individual/all tanks in the housing system, poor water quality, and leakage in pipes or reservoir of the circulating system. In addition, problems in obtaining embryos from breeding could be another concern. Troubleshooting of these issues is discussed below.
<img src="/files/ftp_upload/4196/4196fig1.jpg" alt="Zebrafish phenotype comparison diagram for genetic study; three different specimens labeled A, B, C." data-altupdatedat="1747911167000" data-alt="Figure 1"> 
Figure 1. An illustration of a male zebrafish (A) and female zebrafish (B, C).
<img src="/files/ftp_upload/4196/4196fig2.jpg" alt="Microscope images of plankton sample; includes A) multiple specimens, B) close-up detail, C) single droplet." data-altupdatedat="1747911167000" data-alt="Figure 2"> 
Figure 2. A microscopic view of brine shrimps at 12X (A), a single brine shrimp at 90X (B), and an unfertilized brine shrimp egg at 90X (C). 
<img src="/files/ftp_upload/4196/4196fig3.jpg" alt="Embryonic development stages diagram; cell division, early embryo microscopy, biology study." data-altupdatedat="1747911167000" data-alt="Figure 3"> 
Figure 3. Microscopic view (16X) of fertilized and unfertilized eggs, where only two eggs are unfertilized, the unfertilized eggs are indicated with black arrows (A). Higher magnification view (90X) of a fertilized (B) and an unfertilized egg (C). 
<img src="/files/ftp_upload/4196/4196fig4.jpg" alt="Zebrafish anatomy diagram showing ovipositor location, relevant for embryonic development studies." data-altupdatedat="1747911167000" data-alt="Figure 4"> 
Figure 4. A female zebrafish's ovipositor (indicated with black arrow) illustration. 
 <table border="1">
 <tbody>
 <tr>
 <td fo:width="2in">Parameter</td>
 <td fo:width="3in">Optimum range</td>
 </tr>
 <tr>
 <td>Alkalinity</td>
 <td>50-150 mg/L CaCO3</td>
 </tr>
 <tr>
 <td>pH</td>
 <td>6.8-7.5 
(6.0-8.5 tolerated)</td>
 </tr>
 <tr>
 <td>Temperature</td>
 <td>26-28.5 °C</td>
 </tr>
 <tr>
 <td>Hardness</td>
 <td>50-100 mg/L CaCO3</td>
 </tr>
 <tr>
 <td>Un-ionized Ammonia</td>
 <td>&lt;0.02 mg/L</td>
 </tr>
 <tr>
 <td>Nitrate (NO3-)</td>
 <td>&lt;50 mg/L</td>
 </tr>
 <tr>
 <td>Nitrite (NO2-)</td>
 <td>&lt;0.1 mg/L</td>
 </tr>
 <tr>
 <td>Dissolved oxygen</td>
 <td>&gt;6.0 mg/L</td>
 </tr>
 <tr>
 <td>Salinity</td>
 <td>0.5-1 g/L</td>
 </tr>
 <tr>
 <td>Conductivity</td>
 <td>300 -1,500 μS</td>
 </tr>
 </tbody>
 </table>
Table 2. Water quality parameters. Water quality parameters. Optimum range of environmental parameters in the zebrafish aquarium11.

Watch this Scientific Journal Video about Regular Care and Maintenance of a Zebrafish Danio rerio Laboratory: An Introduction at JoVE.com

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.

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 ...

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Nanyang Technological University

Research

JoVE Journal

Biology

89.1K Views.  Edith Cowan University. Dit protocol beschrijft regelmatig onderhoud en zorg aan optimale omstandigheden voor zebravis veeteelt te behouden. De video illustreert het protocol dat voor systeem onderhoud, regelmatige huisvesting, voeding, fokkerij, en het verhogen van de zebravis larven.

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

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.

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 ...

Developmental Biology

Behavioral Approaches to Studying Innate Stress in Zebrafish

Responding appropriately to stressful stimuli is essential for survival of an organism. Extensive research has been done on a wide spectrum of stress-related diseases and psychiatric disorders, yet further studies into the genetic and neuronal regulation of stress are still required to develop better therapeutics. The zebrafish provides a powerful genetic model to investigate the neural underpinnings of stress, as there exists a large collection of mutant and transgenic lines. Moreover, pharmacology can easily be applied to zebrafish, as most drugs can be added directly to water. We describe here the use of the 'novel tank test' as a method to study innate stress responses in zebrafish, and demonstrate how potential anxiolytic drugs can be validated using the assay. The method can easily be coupled with zebrafish lines harboring genetic mutations, or those in which transgenic approaches for manipulating precise neural circuits are used. The assay can also be used in other fish models. Together, the described protocol should facilitate the adoption of this simple assay to other laboratories.

This manuscript describes a simple method to measure stress behaviorally in adult zebrafish. The approach takes advantage of the innate tendency that zebrafish prefer the bottom half of a tank when in a stressful state. We also describe methods for coupling the assay with pharmacology.

Responding appropriately to stressful stimuli is essential for survival of an organism. Extensive research has been done on a wide spectrum of ...

Behavior

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 ...

Boldness, Aggression, and Shoaling Assays for Zebrafish Behavioral Syndromes

A behavioral syndrome exists when specific behaviors interact under different contexts. Zebrafish have been test subjects in recent studies and it is important to standardize protocols to ensure proper analyses and interpretations. In our previous studies, we have measured boldness by monitoring a series of behaviors (time near surface, latency in transitions, number of transitions, and darts) in a 1.5 L trapezoidal tank. Likewise, we quantified aggression by observing bites, lateral displays, darts, and time near an inclined mirror in a rectangular 19 L tank. By dividing a 76 L tank into thirds, we also examined shoaling preferences. The shoaling assay is a highly customizable assay and can be tailored for specific hypotheses. However, protocols for this assay also must be standardized, yet flexible enough for customization. In previous studies, end chambers were either empty, contained 5 or 10 zebrafish, or 5 pearl danios (D. albolineatus). In the following manuscript, we present a detailed protocol and representative data that accompany successful applications of the protocol, which will allow for replication of behavioral syndrome experiments.

This manuscript describes the setup, implementation, and analysis of boldness, aggression, and shoaling in zebrafish and testing for the presence of a behavioral syndrome. A standardized approach for behavioral quantification will allow for easier comparison across studies. Modifications to this protocol are possible as each assay can be run individually.

A behavioral syndrome exists when specific behaviors interact under different contexts. Zebrafish have been test subjects in recent studies and it is ...

Medicine

Zebrafish In Situ Spinal Cord Preparation for Electrophysiological Recordings from Spinal Sensory and Motor Neurons

Zebrafish, first introduced as a developmental model, have gained popularity in many other fields. The ease of rearing large numbers of rapidly developing organisms, combined with the embryonic optical clarity, served as initial compelling attributes of this model. Over the past two decades, the success of this model has been further propelled by its amenability to large-scale mutagenesis screens and by the ease of transgenesis. More recently, gene-editing approaches have extended the power of the model.
For neurodevelopmental studies, the zebrafish embryo and larva provide a model to which multiple methods can be applied. Here, we focus on methods that allow the study of an essential property of neurons, electrical excitability. Our preparation for the electrophysiological study of zebrafish spinal neurons involves the use of veterinarian suture glue to secure the preparation to a recording chamber. Alternative methods for recording from zebrafish embryos and larvae involve the attachment of the preparation to the chamber using a fine tungsten pin1,2,3,4,5. A tungsten pin is most often used to mount the preparation in a lateral orientation, although it has been used to mount larvae dorsal-side up4. The suture glue has been used to mount embryos and larvae in both orientations. Using the glue, a minimal dissection can be performed, allowing access to spinal neurons without the use of an enzymatic treatment, thereby avoiding any resultant damage. However, for larvae, it is necessary to apply a brief enzyme treatment to remove the muscle tissue surrounding the spinal cord. The methods described here have been used to study the intrinsic electrical properties of motor neurons, interneurons, and sensory neurons at several developmental stages6,7,8,9.

Zebrafish In Situ Spinal Cord Preparation for Electrophysiological Recordings from Spinal Sensory and Motor Neurons

This manuscript describes methods for electrophysiological recordings from spinal neurons of zebrafish embryos and larvae. The preparation maintains neurons in situ and often involves minimum dissection. These methods allow for the electrophysiological study of a variety of spinal neurons, from the initial electrical excitability acquisition through the early larval stages.

Zebrafish, first introduced as a developmental model, have gained popularity in many other fields. The ease of rearing large numbers of rapidly developing ...

Neuroscience

Removing Zebrafish Scales for Culturing and Bone Biology Studies

In Vitro Culturing Technique for Studying Cellular Dynamics in Zebrafish Scales

Zebrafish scales offer a variety of advantages for use in standard laboratories for teaching and research purposes. Scales are easily collected without the need for euthanasia, regenerate within a couple of weeks, and are translucent and small, allowing them to be viewed using a standard microscope. Zebrafish scales are especially useful in educational environments, as they provide a unique opportunity for students to engage in hands-on learning experiences, particularly in understanding cellular dynamics and in vitro culturing methods. The main objective of this protocol is to describe a method for collecting and maintaining zebrafish scales in culture for use in a variety of biological studies using basic laboratory equipment. Additionally, the protocol details their use in understanding bone homeostasis by examining the activity of bone cells involved in bone resorption and deposition. It also includes additional protocols for general techniques, such as the visualization of nuclei and apoptotic cells. The in vitro culturing protocol produces reliable results with minimal reagents and equipment. This article discusses the benefits of using in vitro cultures of zebrafish scales to foster scientific inquiry and outlines the resources needed to support their integration into educational settings.

Author Spotlight: Advancing Bone Cell Activity Research with Zebrafish Scales 

This protocol offers a method to study cellular dynamics using a simple in vitro culture technique. It provides an opportunity for zebrafish researchers and educators to study cellular processes, such as those related to bone homeostasis and basic cell biology, by visualizing fluorescent nuclei and apoptotic cells within the scales.

Zebrafish scales offer a variety of advantages for use in standard laboratories for teaching and research purposes. Scales are easily collected without ...

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 ...

Assessment of Swim Endurance and Swim Behavior in Adult Zebrafish

Due to their renowned regenerative capacity, adult zebrafish are a premier vertebrate model to interrogate mechanisms of innate spinal cord regeneration. Following complete transection of their spinal cord, zebrafish extend glial and axonal bridges across severed tissue, regenerate neurons proximal to the lesion, and regain their swim capacities within 8 weeks of injury. Recovery of swim function is thus a central readout for functional spinal cord repair. Here, we describe a set of behavioral assays to quantify zebrafish motor capacity inside an enclosed swim tunnel. The goal of these methods is to provide quantifiable measurements of swim endurance and swim behavior in adult zebrafish. For swim endurance, zebrafish are subjected to a constantly increasing water current velocity until exhaustion, and time at exhaustion is reported. For swim behavior assessment, zebrafish are subjected to low current velocities and swim videos are captured with a dorsal view of the fish. Percent activity, burst frequency, and time spent against the water current provide quantifiable readouts of swim behavior. We quantified swim endurance and swim behavior in wild-type zebrafish before injury and after spinal cord transection. We found that zebrafish lose swim function after spinal cord transection and gradually regain that capacity between 2 and 6 weeks post-injury. The methods described in this study could be applied to neurobehavioral, musculoskeletal, skeletal muscle regeneration, and neural regeneration studies in adult zebrafish.

Capable of functional recovery after spinal cord injury, adult zebrafish is a premier model system to elucidate innate mechanisms of neural regeneration. Here, we describe swim endurance and swim behavior assays as functional readouts of spinal cord regeneration.

Due to their renowned regenerative capacity, adult zebrafish are a premier vertebrate model to interrogate mechanisms of innate spinal cord regeneration. ...

An Introduction to the Zebrafish: Danio rerio

Zebrafish (Danio rerio) are small freshwater fish that are used as model organisms for biomedical research. The many strengths of these fish include their high degree of genetic conservation with humans and their simple, inexpensive maintenance. Additionally, gene expression can be easily manipulated in zebrafish embryos, and their transparency allows for observation of developmental processes.
This overview video first introduces basic zebrafish biology, including their phylogeny, life cycle, and natural environment, before presenting the features that make them so useful in the lab. A brief history of zebrafish research is also provided through a review of major discoveries made in fish, ranging from the early establishment of methods for efficient genetic screening to the discovery of novel therapeutics for human diseases such as cancer. Finally, some of the many avenues of experimentation performed in zebrafish are discussed, including immunological and developmental studies.

Danio rerio Zebrafish as a Model Organism

Zebrafish (Danio rerio) are small freshwater fish that are used as model organisms for biomedical research. The many strengths of these fish include their ...

Education

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Basic Biology

Biology II: Mouse, Zebrafish, and Chick

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 ...

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

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