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 border="1">
<tbody>
 <tr>
 <td>Name 			of reagent/equipment</td>
 <td>Company</td>
 <td>Catalogue 			number</td>
 <td></td>
 </tr>
 <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) 
Breeder 			Tank-2 (2L)</td>
 <td></td>
 </tr>
 <tr>
 <td>Red 			sea salt</td>
 <td>Red 			sea salt, USA</td>
 <td>Local 			pet store/ 
 <a href="http://www.redseafish.com" target="_blank">www.redseafish.com</a></td>
 <td></td>
 </tr>
 <tr>
 <td>Filters 			(Canister, filter pads, and UV) </td>
 <td>Aquatic 			Habitats, USA</td>
 <td><a href="http://www.aquatichabitats.com" target="_blank">aquatichabitats.com</a></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><a href="http://www.aquatichabitats.com" target="_blank">aquatichabitats.com</a></td>
 <td></td>
 </tr>
 <tr>
 <td></td>
 <td></td>
 <td></td>
 <td>Table 1. Table of specific reagents and equipment used in this protocol.</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="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="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="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="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 &deg;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 &mu;S</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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88.6K Views.  Edith Cowan University. 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.

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

Gross and Fine Dissection of Inner Ear Sensory Epithelia in Adult Zebrafish (Danio rerio)

Neurosensory epithelia in the inner ear are the crucial structures for hearing and balance functions. Therefore, it is important to understand the cellular and molecular features of the epithelia, which are mainly composed of two types of cells: hair cells (HCs) and supporting cells (SCs). Here we choose to study the inner ear sensory epithelia in adult zebrafish not only because the epithelial structures are highly conserved in all vertebrates studied, but also because the adult zebrafish is able to regenerate HCs, an ability that mammals lose shortly after birth. We use the inner ear of adult zebrafish as a model system to study the mechanisms of inner ear HC regeneration in adult vertebrates that could be helpful for clinical therapy of hearing/balance deficits in human as a result of HC loss.
Here we demonstrate how to do gross and fine dissections of inner ear sensory epithelia in adult zebrafish. The gross dissection removes the tissues surrounding the inner ear and is helpful for preparing tissue sections, which allows us to examine the detailed structure of the sensory epithelia. The fine dissection cleans up the non-sensory-epithelial tissues of each individual epithelium and enables us to examine the heterogeneity of the whole epithelium easily in whole-mount epithelial samples.

Gross and Fine Dissection of Inner Ear Sensory Epithelia in Adult Zebrafish Danio rerio

The inner ear sensory epithelium of adult zebrafish is a good model system for understanding the mechanisms of hair cell regeneration in adult vertebrates. This protocol demonstrates the fine dissection of the epithelia, through which we can get tissue samples for studying the regenerative events at cellular and subcellular levels.

Neurosensory epithelia in the inner ear are the crucial structures for hearing and balance functions. Therefore, it is important to understand the ...

Heart Dissection in Larval, Juvenile and Adult Zebrafish, Danio rerio

Zebrafish have become a beneficial and practical model organism for the study of embryonic heart development (see recent reviews1-6), however, work examining post-embryonic through adult cardiac development has been limited7-10. Examining the changing morphology of the maturing and aging heart are restricted by the lack of techniques available for staging and isolating juvenile and adult hearts. In order to analyze heart development over the fish's lifespan, we dissect zebrafish hearts at numerous stages and photograph them for further analysis11. The morphological features of the heart can easily be quantified and individual hearts can be further analyzed by a host of standard methods. Zebrafish grow at variable rates and maturation correlates better with fish size than age, thus, post-fixation, we photograph and measure fish length as a gauge of fish maturation. This protocol explains two distinct, size dependent dissection techniques for zebrafish, ranging from larvae 3.5mm standard length (SL) with hearts of 100&mu;m ventricle length (VL), to adults, with SL of 30mm and VL 1mm or larger. Larval and adult fish have quite distinct body and organ morphology. Larvae are not only significantly smaller, they have less pigment and each organ is visually very difficult to identify. For this reason, we use distinct dissection techniques.
We used pre-dissection fixation procedures, as we discovered that hearts dissected directly after euthanization have a more variable morphology, with very loose and balloon like atria compared with hearts removed following fixation. The fish fixed prior to dissection, retain in vivo morphology and chamber position (data not shown). In addition, for demonstration purposes, we take advantage of the heart (myocardial) specific GFP transgenic Tg(myl7:GFP)twu34 (12), which allows us to visualize the entire heart and is particularly useful at early stages in development when the cardiac morphology is less distinct from surrounding tissues. Dissection of the heart makes further analysis of the cell and molecular biology underlying heart development and maturation using in situ hybridization, immunohistochemistry, RNA extraction or other analytical methods easier in post-embryonic zebrafish. This protocol will provide a valuable technique for the study of cardiac development maturation and aging.

Heart Dissection in Larval, Juvenile and Adult Zebrafish, Danio rerio

A clear, standardized method for dissection and isolation of the zebrafish heart at multiple developmental stages are described. Annotation and quantification techniques are also discussed.

Zebrafish have become a beneficial and practical model organism for the study of embryonic heart development (see recent reviews1-6), however, work ...

In Vivo Modeling of the Morbid Human Genome using Danio rerio

Here, we present methods for the development of assays to query potentially clinically significant nonsynonymous changes using in vivo complementation in zebrafish. Zebrafish (Danio rerio) are a useful animal system due to their experimental tractability; embryos are transparent to enable facile viewing, undergo rapid development ex vivo, and can be genetically manipulated.1 These aspects have allowed for significant advances in the analysis of embryogenesis, molecular processes, and morphogenetic signaling. Taken together, the advantages of this vertebrate model make zebrafish highly amenable to modeling the developmental defects in pediatric disease, and in some cases, adult-onset disorders. Because the zebrafish genome is highly conserved with that of humans (~70% orthologous), it is possible to recapitulate human disease states in zebrafish. This is accomplished either through the injection of mutant human mRNA to induce dominant negative or gain of function alleles, or utilization of morpholino (MO) antisense oligonucleotides to suppress genes to mimic loss of function variants. Through complementation of MO-induced phenotypes with capped human mRNA, our approach enables the interpretation of the deleterious effect of mutations on human protein sequence based on the ability of mutant mRNA to rescue a measurable, physiologically relevant phenotype. Modeling of the human disease alleles occurs through microinjection of zebrafish embryos with MO and/or human mRNA at the 1-4 cell stage, and phenotyping up to seven days post fertilization (dpf). This general strategy can be extended to a wide range of disease phenotypes, as demonstrated in the following protocol. We present our established models for morphogenetic signaling, craniofacial, cardiac, vascular integrity, renal function, and skeletal muscle disorder phenotypes, as well as others.

In Vivo Modeling of the Morbid Human Genome using Danio rerio

Here, we present a systematic approach for developing physiologically relevant, sensitive and specific in vivo assays for interpreting variation in human pathology. Transient genetic manipulation via microinjection of WT and mutant human mRNA and morpholino (MO) antisense oligonucleotides harness the tractability of the developing zebrafish embryo to rapidly assay pathogenic mutations, especially, but not exclusively, in the context of human developmental disorders.

Here,  we present methods for the development of assays to query potentially clinically  significant nonsynonymous changes using in  vivo complementation ...

High Throughput Danio Rerio Energy Expenditure Assay

Zebrafish are an important model organism with inherent advantages that have the potential to make zebrafish a widely applied model for the study of energy homeostasis and obesity. The small size of zebrafish allows for assays on embryos to be conducted in a 96- or 384-well plate format, Morpholino and CRISPR based technologies promote ease of genetic manipulation, and drug treatment by bath application is viable. Moreover, zebrafish are ideal for forward genetic screens allowing for novel gene discovery. Given the relative novelty of zebrafish as a model for obesity, it is necessary to develop tools that fully exploit these benefits. Herein, we describe a method to measure energy expenditure in thousands of embryonic zebrafish simultaneously. We have developed a whole animal microplate platform in which we use 96-well plates to isolate individual fish and we assess cumulative NADH2 production using the commercially available cell culture viability reagent alamarBlue. In poikilotherms the relationship between NADH2 production and energy expenditure is tightly linked. This energy expenditure assay creates the potential to rapidly screen pharmacological or genetic manipulations that directly alter energy expenditure or alter the response to an applied drug (e.g. insulin sensitizers).

High Throughput Danio Rerio Energy Expenditure Assay

Zebrafish are an important model organism for the study of energy homeostasis. By utilizing a NADH2 sensitive redox indicator, alamar Blue, we have developed an assay that measures the metabolic rate of zebrafish larvae in a 96 well plate format and can be applied to drug or gene discovery.

Zebrafish are an important model organism with inherent advantages that have the potential to make zebrafish a widely applied model for the study of ...

Maintenance of a Drosophila melanogaster Population Cage

Large quantities of DNA, RNA, proteins and other cellular components are often required for biochemistry and molecular biology experiments. The short life cycle of Drosophila enables collection of large quantities of material from embryos, larvae, pupae and adult flies, in a synchronized way, at a low economic cost. A major strategy for propagating large numbers of flies is the use of a fly population cage. This useful and common tool in the Drososphila community is an efficient way to regularly produce milligrams to tens of grams of embryos, depending on uniformity of developmental stage desired. While a population cage can be time consuming to set up, maintaining a cage over months takes much less time and enables rapid collection of biological material in a short period. This paper describes a detailed and flexible protocol for the maintenance of a Drosophila melanogaster population cage, starting with 1.5 g of harvested material from the previous cycle.

Maintenance of a Drosophila melanogaster Population Cage

This manuscript reports a detailed protocol for culturing, on a regular basis, a population of Drosophila melanogaster using a fly population cage.

Large quantities of DNA, RNA, proteins and other cellular components are often required for biochemistry and molecular biology experiments. The short life ...

Nephrotoxin Microinjection in Zebrafish to Model Acute Kidney Injury

The kidneys are susceptible to harm from exposure to chemicals they filter from the bloodstream. This can lead to organ injury associated with a rapid decline in renal function and development of the clinical syndrome known as acute kidney injury (AKI). Pharmacological agents used to treat medical circumstances ranging from bacterial infection to cancer, when administered individually or in combination with other drugs, can initiate AKI. Zebrafish are a useful animal model to study the chemical effects on renal function in vivo, as they form an embryonic kidney comprised of nephron functional units that are conserved with higher vertebrates, including humans. Further, zebrafish can be utilized to perform genetic and chemical screens, which provide opportunities to elucidate the cellular and molecular facets of AKI and develop therapeutic strategies such as the identification of nephroprotective molecules. Here, we demonstrate how microinjection into the zebrafish embryo can be utilized as a paradigm for nephrotoxin studies.

Renal injuries incurred from nephrotoxins, which include drugs ranging from antibiotics to chemotherapeutics, can result in complex disorders whose pathogenesis remains incompletely understood. This protocol demonstrates how zebrafish can be used for disease modeling of these conditions, which can be applied to the identification of renoprotective measures.

The kidneys are susceptible to harm from exposure to chemicals they filter from the bloodstream. This can lead to organ injury associated with a rapid ...

Detecting Estrogenic Ligands in Personal Care Products using a Yeast Estrogen Screen Optimized for the Undergraduate Teaching Laboratory

The Yeast Estrogen Screen (YES) is used to detect estrogenic ligands in environmental samples and has been broadly applied in studies of endocrine disruption. Estrogenic ligands include both natural and manmade &#34;Environmental Estrogens&#34; (EEs) found in many consumer goods including Personal Care Products (PCPs), plastics, pesticides, and foods. EEs disrupt hormone signaling in humans and other animals, potentially reducing fertility and increasing disease risk. Despite the importance of EEs and other Endocrine Disrupting Chemicals (EDCs) to public health, endocrine disruption is not typically included in undergraduate curricula. This shortcoming is partly due to a lack of relevant laboratory activities that illustrate the principles involved while also being accessible to undergraduate students. This article presents an optimized YES for quantifying ligands in personal care products that bind estrogen receptors alpha (ER&#945;) and/or beta (ER&#946;). The method incorporates one of the two colorimetric substrates (ortho-nitrophenyl-&#946;-D-galactopyranoside (ONPG) or chlorophenol red-&#946;-D-galactopyranoside (CPRG)) that are cleaved by &#946;-galactosidase, a 6-day refrigerated incubation step to facilitate use in undergraduate laboratory courses, an automated application for LacZ calculations, and R code for the associated 4-parameter logistic regression analysis. The protocol has been designed to allow undergraduate students to develop and conduct experiments in which they screen products of their choosing for estrogen mimics. In the process, they learn about endocrine disruption, cell culture, receptor binding, enzyme activity, genetic engineering, statistics, and experimental design. Simultaneously, they also practice fundamental and broadly applicable laboratory skills, such as: calculating concentrations; making solutions; demonstrating sterile technique; serially diluting standards; constructing and interpolating standard curves; identifying variables and controls; collecting, organizing, and analyzing data; constructing and interpreting graphs; and using common laboratory equipment such as micropipettors and spectrophotometers. Thus, implementing this assay encourages students to engage in inquiry-based learning while exploring emerging issues in environmental science and health.

This article presents an optimized yeast estrogen screen for quantifying ligands in Personal Care Products (PCPs) that bind estrogen receptors alpha (ER&#945;) and/or beta (ER&#946;). The method incorporates two colorimetric substrate options, a six-day refrigerated incubation for use in undergraduate courses, and statistical tools for data analysis.

The Yeast Estrogen Screen (YES) is used to detect estrogenic ligands in environmental samples and has been broadly applied in studies of endocrine ...

Anaerobic Growth and Maintenance of Mammalian Cell Lines

Most mucosal surfaces along with the midpoints in tumors and stem cell niches are geographic areas of the body that are anoxic. Previous studies show that the incubation in normoxic (5% CO2 in air) or hypoxic (low oxygen levels) conditions followed by an anoxic incubation (an absence of free oxygen) results in limited viability (2&#8211;3 days). A novel methodology was developed that enables an anoxic cell cultivation (for at least 17 days; the maximum time tested). The most critical aspect of this methodology is to ensure that no oxygen is introduced into the system. This is obtained by the degassing of media, and by flushing tubes, dishes, flasks, and pipettes with an anaerobic gas mixture (H2, CO2, N2) followed by permitting the materials to equilibrate to the anoxic (non-oxygen) environment prior to usage.&#160;Additional care must be exercised when acquiring photomicrographs to ensure that the micrographs obtained do not contain artifacts. In the absence of oxygen, cell morphology is significantly altered. Two distinct morphotypes are noted for all anaerobically-grown cells. The ability to grow and maintain mammalian cells in the absence of oxygen can be applied to the analysis of cell physiology, polymicrobial interactions, and the characterization of biosynthetic pathways for novel cancer drug development.

Here, we describe a new method that enables the anaerobic long-term cultivation of established cell lines. The maximum survival time that was tested is 17 days. This method is suitable for the testing of cytotoxic agents and exploring the physiology of anoxically replicating cells.

Most mucosal surfaces along with the midpoints in tumors and stem cell niches are geographic areas of the body that are anoxic. Previous studies show that ...

Establishment and Maintenance of Gnotobiotic American Cockroaches (Periplaneta americana)

Gnotobiotic animals are a powerful tool for the study of controls on microbiome structure and function. Presented here is a protocol for the establishment and maintenance of gnotobiotic American cockroaches (Periplaneta americana). This approach includes built-in sterility checks for ongoing quality control. Gnotobiotic insects are defined here as cockroaches that still contain their vertically transmitted endosymbiont (Blattabacterium) but lack other microbes that normally reside on their surface and in their digestive tract. For this protocol, egg cases (oothecae) are removed from a (nonsterile) stock colony and surface sterilized. Once collected and sterilized, the oothecae are incubated at 30 &#176;C for approximately 4&#8722;6 weeks on brain-heart infusion (BHI) agar until they hatch or are removed due to contamination. Hatched nymphs are transferred to an Erlenmeyer flask containing a BHI floor, sterile water, and sterile rat food. To ensure that the nymphs are not housing microbes that are unable to grow on BHI in the given conditions, an additional quality control measure uses restriction fragment-length polymorphism (RFLP) to test for nonendosymbiotic microbes. Gnotobiotic nymphs generated using this approach can be inoculated with simple or complex microbial communities and used as a tool in gut microbiome studies.

Establishment and Maintenance of Gnotobiotic American Cockroaches Periplaneta americana

This protocol is used in establishing and maintaining gnotobiotic American cockroaches (Periplaneta americana) by surface sterilizing the egg cases (oothecae) prior to hatching. These gnotobiotic insects contain their vertically transmitted Blattabacterium endosymbionts but have axenic guts.

Gnotobiotic animals are a powerful tool for the study of controls on microbiome structure and function. Presented here is a protocol for the establishment ...

Fluorescence In Situ Hybridization on DNA Halo Preparations to Reveal Whole Chromosomes, Telomeres and Gene Loci

The genome is associated with several structures inside cell nuclei, in order to regulate its activity and anchor it in specific locations. These structures are collectively known as the nucleoskeleton and include the nuclear lamina, the nucleoli, and nuclear bodies. Although many variants of fluorescence in situ hybridization (FISH) exist to study the genome and its organization, these are often limited by resolution and provide insufficient information on the genome&#39;s association with nuclear structures. The DNA halo method uses high salt concentrations and nonionic detergents to generate DNA loops that remain anchored to structures within nuclei through attachment regions within the genome. Here, soluble nuclear proteins, such as histones, lipids, and DNA not tightly bound to the nuclear matrix, are extracted. This leads to the formation of a halo of unattached DNA surrounding a residual nucleus which itself contains DNA closely associated with internal nuclear structures and extraction-resistant proteins. These extended DNA strands enable increased resolution and can facilitate physical mapping. In combination with FISH, this method has the added advantage of studying genomic interactions with all the structures that the genome is anchored by. This technique, termed HALO-FISH, is highly versatile whereby DNA halos can be coupled with nucleic acid probes to reveal gene loci, whole chromosomes, alpha satellite, telomeres and even RNA. This technique provides an insight into nuclear organization and function in normal&#160;cells and in disease progression such as with cancer.

Combining DNA halo preparations with fluorescence in situ hybridization enables high-resolution analysis of genomic interactions with the nucleoskeleton. Attached genome leads to hybridized fluorescent signals located within the residual extracted nuclei, whereas non-attached genome is in the halo of DNA surrounding the residual nuclei.