NOAA Technical Memorandum NMFS-NWFSC-100

Zebrafish (Danio rerio)

Husbandry and Colony Maintenance

at the

Northwest Fisheries Science Center

May 2009

U.S. DEPARTMENT OF COMMERCE

National Oceanic and Atmospheric Administration

National Marine Fisheries Service
NOAA Technical Memorandum

NMFS-NWFSC Series

The Northwest Fisheries Science Center of the National 

Marine Fisheries Service, NOAA, uses the NOAA Technical Memorandum NMFS-NWFSC series to issue scientific 

and technical publications. Manuscripts have been peer 

reviewed and edited. Documents published in this series 

may be cited in the scientific and technical literature.

The NMFS-NWFSC Technical Memorandum series of the 

Northwest Fisheries Science Center continues the NMFSF/NWC series established in 1970 by the Northwest & 

Alaska Fisheries Science Center, which has since been 

split into the Northwest Fisheries Science Center and the 

Alaska Fisheries Science Center. The NMFS-AFSC Technical Memorandum series is now being used by the Alaska 

Fisheries Science Center.

Reference throughout this document to trade names does 

not imply endorsement by the National Marine Fisheries 

Service, NOAA.

This document should be referenced as follows:

Linbo, T.L. 2009. Zebrafish (Danio rerio) husbandry and 

colony maintenance at the Northwest Fisheries Science 

Center. U.S. Dept. Commer., NOAA Tech. Memo. NMFSNWFSC-100, 62 p. 
NOAA Technical Memorandum NMFS-NWFSC-100

Zebrafish (Danio rerio)

Husbandry and Colony Maintenance

at the

Northwest Fisheries Science Center

Tiffany L. Linbo

Northwest Fisheries Science Center

Environmental Conservation Division

2725 Montlake Boulevard East

Seattle, Washington 98112

May 2009

U.S. DEPARTMENT OF COMMERCE

National Oceanic and Atmospheric Administration

National Marine Fisheries Service
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NMFS-NWFSC are available online at the 

Northwest Fisheries Science Center

web site (http://www.nwfsc.noaa.gov)

Copies are also available from:

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Table of Contents 

List of Figures...............................................................................................................................................v 

Executive Summary ....................................................................................................................................vii 

Acknowledgments........................................................................................................................................ix 

Introduction...................................................................................................................................................1 

General Maintenance Overview....................................................................................................................2 

Water and Water Quality ..............................................................................................................................4 

Water System............................................................................................................................................4 

Zebrafish Module Water Quality..............................................................................................................6 

Zebrafish Module System Maintenance .....................................................................................................11 

Filter Maintenance..................................................................................................................................11 

UV Light Maintenance ...........................................................................................................................14 

Cleaning..................................................................................................................................................14 

Culturing Zebrafish.....................................................................................................................................18 

Food for Adult Zebrafish........................................................................................................................18 

Spawning Fish and Collecting Embryos.................................................................................................21 

Raising Larvae and Beyond....................................................................................................................23 

Paramecium Culture ...............................................................................................................................27 

Maintaining a Breeding Zebrafish Colony .............................................................................................30 

Biosecurity ..................................................................................................................................................34 

Record Keeping ..........................................................................................................................................35 

Zebrafish Room Power Failure Plan...........................................................................................................36 

When the Power Goes Out .....................................................................................................................36 

After the Power is Restored....................................................................................................................37 

Expanding–New Fish and Systems.............................................................................................................38 

Bringing in New Fish from a Different System......................................................................................38 

Setting Up and Maintaining a Quarantine Tank .....................................................................................38 

Bringing in New Embryos from an Outside Source...............................................................................40

Preventing the Spread of Pathogens .......................................................................................................40 

Conclusion ..................................................................................................................................................42 

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References...................................................................................................................................................43 

Appendix A: Ordering Information ............................................................................................................45 

Fish Food................................................................................................................................................45 

General Fish and Aquarium Supplies.....................................................................................................45 

Zebrafish Hardware ................................................................................................................................45 

Appendix B: General Fish Aquaculture Information..................................................................................47 

Appendix C: Examples of Zebrafish Records.............................................................................................49 

Appendix D: Zebrafish Diseases.................................................................................................................55 

Common Diseases of Zebrafish..............................................................................................................55 

Routes of Disease Transmission.............................................................................................................57 

Diagnostic Techniques ...........................................................................................................................57 

Treatments and Controls.........................................................................................................................58 

Prevention...............................................................................................................................................59 

Zoonosis .................................................................................................................................................59 

Appendix E: Training Checklist for New Zebrafish Users.........................................................................61 

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List of Figures 

Figure 1. Schematic diagram of the reverse osmotic and deionization system............................................4 

Figure 2. Schematic diagram of the self-contained, modified Z-Mod system.............................................9 

Figure 3. The nitrogen cycle in aquaculture systems and aquarium tanks.................................................13 

Figure 4. Siphon for cleaning out Z-Mod tanks.........................................................................................17 

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Executive Summary 

This technical memorandum describes the protocols and procedures for maintaining a 

small-scale zebrafish (Danio rerio) operation at the Northwest Fisheries Science Center, 

Environmental Conservation Division. Detailed descriptions are presented for Zebrafish Module 

maintenance, spawning, larvae rearing, adult feeding, quarantine procedures, and general fish 

health. These protocols may not be appropriate for all zebrafish system sizes and configurations; 

however, small zebrafish laboratories may find that this technical memorandum provides useful 

guidance for managing a healthy, breeding zebrafish colony. 

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viii
Acknowledgments 

This guide is a gathering of invaluable information and experience gained over many 

years of working, consulting, and troubleshooting with Laura Swaim and David White of the 

University of Washington Zebrafish Facility, Tor Linbo of the University of Washington 

Department of Biological Structure, and Carla Stehr, Megan Bushnell, and other colleagues of 

the Northwest Fisheries Science Center, Environmental Conservation Division. Laura Swaim 

and Carla Stehr developed and set up our system in 2001. Nat Scholz obtained funding through 

a Northwest Fisheries Science Center internal grant to establish the system. The information on 

general fish physiology and fish diseases was obtained from the Health Management of 

Laboratory Fish course at the Mount Desert Island Biological Lab in Maine. 

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Introduction 

The Environmental Conservation Division of the Northwest Fisheries Science Center 

(NWFSC) has developed protocols to manage and maintain a small-scale zebrafish (Danio rerio) 

colony. Small-scale refers to less than 3,000 zebrafish adults. These protocols were specifically 

developed for the aquaculture system used by the NWFSC, a modified Zebrafish Module (ZMod) manufactured by Marine Biotech (Appendix A: Ordering Information). 

Zebrafish are a useful research model because of their high year-round fecundity, wide 

array of molecular and genetic markers, rapid development, and relative ease of maintaining a 

breeding colony. Also as a teleost, the zebrafish may be used as an experimental model for other 

fish, such as salmonids. At the NWFSC, zebrafish are used to study the developmental, 

behavioral, and physiological effects of anthropogenic contaminants (e.g., polycyclic aromatic 

hydrocarbons, metals, pesticides, and estrogenic compounds), natural toxins (e.g., biotoxins from 

harmful algal blooms), and toxic bacteria (e.g., Vibrio). Since embryos, larvae, and adult fish are 

used in our studies, it is important to maintain healthy zebrafish throughout their lifecycle 

without introducing pollutants, chemicals, or antibiotics. 

One of the most important aspects of keeping a healthy colony is providing consistent 

care to achieve a stable environment for the fish. Some of the protocols described here involve 

additional experience in recognizing the needs of the fish. For example, how much food to feed 

and prepare for fish requires firsthand experience to know when fish are satiated. Tank cleaning 

requires the ability to identify when bacterial growth could become a health hazard. A dedicated 

zebrafish caretaker with the ability to recognize and address the many zebrafish needs is vital to 

the overall health and productivity of the colony. 

The products and supplies mentioned in this guide are routinely used for NWFSC’s 

zebrafish husbandry. However, they may not be suitable for all zebrafish laboratories; therefore, 

information on product suppliers is not limited to what can be found in Appendix A: Ordering 

Information. 
General Maintenance Overview 

Daily (listed in the order the duties should be performed): 

Morning: 

• Clean larval tanks (see Cleaning) and feed larvae (see Raising Larvae and Beyond). 

• Feed adult fish using salmon starter and Artemia sp. (brine shrimp) nauplii. Feed the dry 

food first (see Food for Adult Zebrafish). 

• Take readings for ammonia, conductivity, pH, and temperature. Make adjustments if 

necessary (see Zebrafish Module Water Quality). 

• Check sump water level. Add reverse osmotic and deionized (RO/DI) water if needed 

(see Water System). 

• Record feeding and water quality measurements on daily logs. 

Noon: 

• Feed larvae (see Raising Larvae and Beyond). 

Afternoon: 

• Feed larvae (see Raising Larvae and Beyond). 

• Feed adult fish salmon starter and Artemia (see Food for Adult Zebrafish). 

• Take readings for pH and temperature. Make adjustments if necessary (see Zebrafish 

Module Water Quality). 

• Prepare batch of Artemia for next day’s feeding and start a new batch of Artemia (see 

Food for Adult Zebrafish). 

• Record feeding and water quality measurements on daily logs (see Record Keeping). 

• Check sump water level. Add more RO/DI water if water level has decreased due to 

evaporation (usually requires 3–4 gallons [gal] of RO/DI water per day) (see Water 

System). 

• Refill the RO/DI water bucket. It will take about 90 minutes (min) to fill a five-gal 

bucket (see Water System). 

Once a week:

• Measure nitrite levels in Z-Mod system (see Zebrafish Module Water Quality). 

• Clean the tops of the tanks (remove old food debris) and make sure all water outlets are 

clear and not clogged with food debris (see Cleaning). 

• Check to see if the shelves and back of the Z-Mod need to be cleaned (see Cleaning). 

• Record all cleanings on log sheets (see Record Keeping). 

• Flush out RO membrane filter (see Water System). 

• Change Net Soak solution (see Cleaning). 

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Once every 2 weeks: 

• Siphon debris out of tanks (see Cleaning). 

• Clean sump tank (see Cleaning). 

• Inoculate paramecium (Paramecium multimicronucleatum) bottle culture (see 

Paramecium Culture). 

• Change and clean sump sock if necessary (see Cleaning). 

Once a month: 

• Check for chlorine and total dissolved solids (TDS) (see Zebrafish Module Water 

Quality). 

• Calibrate pH meter and conductivity meters (refer to instructions in instruction manual 

folder). 

• Change paper filter (#3) of Z-Mod. Replace carbon and crushed coral (see Filter 

Maintenance). 

• Start new paramecium Tupperware culture (see Paramecium Culture). 

• Change water in bleach bath (see Cleaning). 

Once every 3 months: 

• Clean biofilters (sponge filters #1 and #2) on Z-Mod (see Filter Maintenance). 

• Set up spawns to maintain zebrafish colony (see Maintaining a Breeding Zebrafish 

Colony). 

• Replace RO/DI system’s sediment and carbon cartridges (see Water System). 

Once a year: 

• Replace UV bulb (see Filter Maintenance). 

• Replace RO/DI deionizing filters (see Water System). 

• Replace Ammonia Alert (see Zebrafish Module Water Quality). 

• Order new adult and larval fish food (see Food for Adult Zebrafish). 

Once every 2 years: 

• Replace RO membrane (see Water System). 

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Water and Water Quality 

Water System 

Zebrafish systems require a controlled, clean source of water for basic facilities 

operations. Municipal water that has passed through a reverse osmotic and deionization (RO/DI) 

system is used to make system water, Artemia water, and for general maintenance in the 

zebrafish facility. 

Reverse Osmotic and Deionized Water and the RO/DI System 

RO/DI water is dispensed from the AquaFX’s Mako 5-filter RO/DI system (Figure 1) 

mounted on the wall. The RO/DI water is stored in two 5-gal buckets labeled “RO/DI water 

only” (nothing else is put in these buckets). 

Figure 1. Schematic diagram of the RO/DI system. From the right, city water is filtered through a 25-

micron sediment filter before entering the RO/DI system. A water timer controls the length of 

time the system runs. Water enters the system and travels through a 1-micron sediment and a 

carbon filter. It then travels through the RO membrane. Waste from the RO membrane is routed 

through the waste water line (yellow line) that should be directed into the sink. Water processed 

by the RO membrane continues through two DI filters and exits through the RO/DI water blue 

line. Two cutoff valves are located at the start and end of the system to allow for system 

maintenance. A flush valve is located on the waste water line to back flush the RO membrane for 

cleaning. 

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Before entering the RO/DI filter system, the city water is run through a 25-micron 

sediment filter, which is provided by NWFSC maintenance, to remove large particulates. 

Typically the filter is changed every 2–3 months. If there are problems with this filter (e.g., 

leaking), contact building maintenance. 

After passing through the 25-micron filter, the water goes through a water timer, which 

controls the length of time that water passes through the RO/DI system. Water is first filtered 

through the sediment filters: a 1-micron filter and a carbon filter. It then goes through the RO 

membrane on top of the unit. Wastewater from the RO process goes out the yellow drain hose 

(that drains into the sink), while the RO water continues through the two deionizing filters. 

Finally, the RO/DI water comes out the blue hose and collects in 5-gal RO/DI buckets. 

To turn on the RO/DI water system, turn the cutoff valve that comes out of the 

deionization filters so it parallels the blue tube, then open the main cutoff valve located before 

the 25-micron filter (Figure 1). Now turn on the water timer (located between the 25-micron 

filter and the RO/DI system). Normally, all valves are open and only the timer needs to be 

turned on. The timer can be set for up to 120 min (or it can be turned to manual, which will run 

indefinitely until turned to the off position. If it is turned to manual, remember to turn it off so 

the room does not flood. The RO/DI system is rated to produce RO/DI water at 100 gal per day. 

Actual use suggests that it produces just under 3 gal an hour (h) or 5 gal in 90 min. 

Replace the two sediment filters every 2–3 months (depending on the season’s influence 

on city water). Replace the RO membrane every 2 years. Replace one of the deionization filters 

once a year. The deionization filters are rotated through the two canisters, spending 1 year in 

each canister, so filters are used a total of 2 years. Specifically, remove and discard the first 

deionization filter from the first slot when is has changed color completely (from blue to orange 

over about 2 years) and replace it with the deionization filter that was in the second slot (after 1 

year). Put a new deionization filter into the second slot (refer to AquaFX directions online at 

http://www.aquariumwaterfilters.com/library/manuals/5%20Stage%20Mako%20Manual.pdf). 

A flush valve is located on the top of the RO/DI water filter system (Figure 1). The 

membrane should be flushed once a week or more, depending on RO/DI water usage. To flush 

the membrane, allow water to run through the RO/DI system and turn the blue valve on the flush 

valve kit so it parallels the white tubing. Water flow out of the blue hose (where the RO/DI 

water comes out) should stop, and water flow out of the yellow hose (the non-RO water) should 

increase. Let the water flush out the system for about 30 seconds (refer to AquaFX directions 

online at the URL above). To resume RO water filtering, turn the blue valve perpendicular to the 

white hose. 

System Water 

System water is RO/DI water amended with artificial sea salt (Instant Ocean, Appendix 

A) to a conductivity of 1,500–1,600 μS/cm and a pH of 7.0–8.0. It is stored in the 50-gal tower. 

System water is used in the Z-mod and for water changes, paramecium cultures, spawning, and 

other needs. 

5
To prepare system water, first add RO/DI water to the system tower while keeping track 

of how much water has been added. Dissolve salt in a small amount of RO/DI water and add it 

to the tank at a rate of about 3 tbsp Instant Ocean salt per 20 gal RO/DI water, or 1 tbsp for every 

2 h the RO/DI system is running. The tower is marked with a guide for the amount of sea salt to 

add when filling. 

A submersible pump on the bottom of the tank is on 24 h a day to circulate the water. 

Check the pump periodically to make sure it is working. After adding salt, let the water stir for a 

while, then test the conductivity. It should be around 1,500 μS/cm. If below 1,400 μS/cm, add 

more salt. If above 1,600 μS/cm, add more RO/DI water. 

Check the pH of the system water as well. RO/DI water has a pH of approximately 5.5, 

however system water should have a pH of 7.4–7.8. To increase the pH of freshly made system 

water, add sodium bicarbonate to the system water tower (approximately 100 ml of 1 molar 

sodium bicarbonate solution to 50 gal). This will also increase conductivity, so recheck the 

conductivity of the system water. Dilute with more RO/DI water if conductivity is too high. 

Filling the system water tower can take an entire day, so prepare accordingly. A 5-gal 

bucket (labeled System Water) should be filled as a reserve before adding RO/DI water to the 

system water tower. The reserve system water in the bucket can be used while the tower is filled 

and the system water is adjusted for conductivity and pH. 

Zebrafish Module Water Quality 

Maintaining optimal water quality is vital for fish health. These water quality parameters 

are interconnected and a sudden change in one could signify a larger problem (Appendix B: 

General Fish Aquaculture Information). The following water quality parameters need to be 

monitored and kept stable. 

Conductivity 

Check daily. Conductivity should be 1,500–1,600 μS/cm. 

If conductivity is too high, first check to see whether the water level in the sump is low. 

Water evaporation will lead to an increase in salt concentration. Add RO/DI water if needed and 

recheck the conductivity after several hours. If the conductivity is still too high, empty some of 

the water from the sump and replace it with RO/DI water. Give the system several hours to 

circulate and check the conductivity again. 

If the conductivity is too low (this is usually not a problem), check the water level and 

add system water if needed. If the conductivity is still too low, dissolve a little salt (1 tbsp or 

less) in RO/DI water and add to it the sump. Give the system several hours to circulate and 

check the conductivity again. 

pH 

Check twice daily. The pH range of the Z-Mod should be between 7.0 and 8.0, but the 

optimum range is 7.0–7.4. 

6
If it gets more acidic than 7.0 (the pH of the Z-Mod will naturally go down as ammoniaand nitrite-metabolizing bacteria lower the alkalinity of the water, Appendix B: General Fish 

Aquaculture Information), try gentle changes, such as adding crushed coral to the mesh bag in 

the filtration system (see Filter Maintenance) or replacing some system water to increase the pH. 

Sodium bicarbonate can also be added using a 1 molar stock solution of sodium bicarbonate 

dissolved in RO/DI water. Depending on how low the pH level is, start by adding 50 ml of the 

stock solution to the sump. However, do not add large amounts of sodium bicarbonate because 

sudden changes in pH could be stressful to the fish and overload the capabilities of the biofilter 

bacteria to convert ammonia (NH3). Under acidic conditions, the biofilter cannot convert the 

nontoxic ammonium (NH4+); instead, it can only convert NH3 (Appendix B: General Fish 

Aquaculture Information), causing an accumulation of this ion. A sudden rise in pH, however, 

will cause the accumulated NH4+ to shift to excessive amounts of toxic NH3, which the biofilter 

may not be able to handle. Therefore, increases in pH should be kept to a rate at which the 

biofilter can convert the resulting NH3. 

If the pH is too basic (more than 8.0, which is unlikely to occur), try changing the system 

water or removing the crushed coral. 

Temperature 

Check twice daily. The temperature should be around 26oC. Although zebrafish are 

hardy and can withstand typical fluctuations in room temperatures, two submersible heaters are 

kept in the sump to provide a consistent water temperature. It is ideal to keep the environment as 

stable as possible for the fish, biofilters, and water parameters (Appendix B: General Fish 

Aquaculture Information). 

If the water temperature falls below 22oC, check the heaters in the sump. Replace any 

that are not working or add a higher-powered heater to the sump. If the temperature rises above 

30oC, check the heaters in the sump. Make sure the heater temperature is not set too high. If the 

room temperature is also higher than normal (≈23–24oC), contact Maintenance to check the air 

conditioning units and the ventilation ducts. In the meantime, fill the sump with RO/DI water, as 

the evaporation rate increases during higher temperatures. 

Ammonia (NH3) 

Check daily. Check the Ammonia Alert disc (located in a Z-Mod tank without fish) to 

monitor ammonia levels. The disc should be yellow (safe). The disc can detect levels as low as 

0.05 ppm. Ammonia levels 0.8 ppm or higher are toxic to the fish. The Ammonia Alert sensor 

is replaced once a year. 

If the center disc changes color from yellow to light green (safe to alert), this indicates 

that ammonia is detected in the water and the population of ammonia- and nitrite-metabolizing 

bacteria in the biofilter may have been harmed (likely due to sudden changes in water quality). 

The fish can survive with this amount of ammonia, but system water changes are needed 

immediately to dilute the ammonia. Allow the bacteria on the biofilters to repopulate while 

constantly monitoring the ammonia and nitrite levels (check three times a day) and performing 

7
system water changes (once a day change out 5–10% of the system water) until the biofilters are 

working again. 

If ammonia levels increase to blue (alarm), then more corrective actions are need. Use 

products such as Ammo-Lock (Appendix A) to temporarily reduce ammonia levels. However, 

this is not a long-term solution. The biofilters need to be repopulated with ammoniametabolizing bacteria. Reduce the amount of ammonia introduced to the system by feeding the 

fish only once per day and, if possible, remove some of the fish from the system. Continue 

monitoring the ammonia and nitrite levels and performing system water changes. 

Nitrite (NO2) 

Check weekly. Follow the directions of the test kit to check the nitrite levels. Nitrite 

levels should be zero. 

If nitrite is detected, a water change is necessary. Recheck the levels after water changes. 

If nitrite is still detected in the water, then the biofilters are not working properly. Treat the 

system the same as if the ammonia levels were too high (see above). Monitor the ammonia and 

nitrite levels and perform water changes until the biofilters are working again. 

Chlorine 

Check monthly. To check the level of chlorine in Z-Mod water, follow the directions in 

the chlorine test kit. The RO/DI system should be removing the chlorine, so no chlorine should 

be in the Z-Mod water. However if chlorine is present, then the RO/DI system is likely not 

functioning properly. 

Chlorine is harmful to fish; corrective actions are required if chlorine is detected. If 

chlorine is in the water, add Genesis Chlorine Neutralizer at 2 drops/gal to neutralize the 

chlorine. For the Z-Mod system, use 40 drops for 20 gal. Check the RO/DI water for chlorine. 

If chlorine is detected, change the carbon filter in the RO/DI system. 

Total Dissolved Solids 

Check monthly. To check the level of TDS in the RO/DI water, follow the directions in 

the TDS test kit. The TDS of the RO/DI water should be 000–001, indicating that the RO/DI 

system is working properly. 

If dissolved solids are detected in the RO/DI water, check the RO/DI water filters. Check 

the gauge to make sure the water is being filtered within the operating range. If necessary, 

change RO/DI water cartridges (usually changing the sediment filters will correct the problem). 

Sump Water Level 

Check daily. Water level in the sump should be near the level marked by the tape that 

reads “maintain water level here,” located on the outside of the sump. The water level needs to 

be above the outlet to the water pump and 2–3 inches below the lip of the filter bag or sump sock 

(Figure 2). 

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Figure 2. Schematic diagram of the self-contained, modified Z-Mod system. Arrows indicate the 

directional flow of the water. Water leaving the Z-Mod tanks (effluent) enters the sump, where it 

is filtered through the filter bag or sump sock to remove larger debris. The water pump powers 

the water flow through a modified three-step filtration system. Water passes through two sponge 

biofilters then through the paper filter, which also contains activated carbon (for chemical 

removal) and coral (for pH balance). Finally the water goes through a UV sterilizer and enters 

the Z-Mod tanks again. Individual row valves control water flow into the rows. The line on the 

sump indicates where the water level should be maintained. The bottom line on the sump 

indicates how low the water level is drained when doing maintenance (the water level should be 

kept above the drain to the water pump). 

9
If water is low due to evaporation, add RO/DI water. When water evaporates, the salt 

stays in the system, so do not add system water or the conductivity will be too high. The normal 

evaporation rate is 3–4 gal a day. 

Flow Meter and Bypass Valve 

Check once per week and before filter maintenance. The flow meter should read 

approximately 5–7 gal per min. 

The water flow of the Z-Mod system is a dynamic balance between the individual row 

valves and the bypass valve. Water flowing to each row can be adjusted by turning the red 

valves at the end of each row. To adjust the entire flow pressure, adjustments can be made to the 

bypass valve (white ball valve) that is closest to the sump on the PVC piping (Figure 2). The 

bypass valve alleviates backpressure on the sponge biofilters (see Filter Maintenance) and affects 

the pressure of the water entering the tanks. As more tanks are added to the Z-Mod, the pressure 

may need to be increased (i.e., the bypass value almost closed) to compensate for the additional 

rows being used. During sponge biofilter maintenance, however, it is important to open the 

bypass valve to relieve the initial water pressure when water reenters the filter canisters (see 

Filter Maintenance). 

10
Zebrafish Module System Maintenance 

Filter Maintenance 

Since the Z-Mod is a closed system (water recirculates within the system), the water 

entering the tanks must first pass through a filtration system to remove particulates, ammonia, 

and chemicals. The modified filter system (Figure 2), which is not the Z-Mod filtration system 

offered by Marine Biotech, consists of a sump (or a reservoir of water), a water pump that 

circulates water through two sponge biofilters, a paper filter with carbon and coral, and the UV 

sterilizer. To remove large debris (e.g., uneaten food, bacterial growth) and escaped fish, Z-Mod 

effluent is first filtered through a filter bag or sump sock as it enters the sump. Then the water is 

pumped through two sponge biofilters (#1 and #2), which remove ammonia and nitrite. Next the 

water goes through the paper filter (#3) for fine particle removal, pH balance (coral), and 

chemical removal (activated carbon). For the final recirculation step, the water passes through a 

UV light sterilizer, then the filtered water reenters the Z-Mod tanks. 

Maintenance on the Z-Mod is preferably scheduled earlier in the day. If canisters are not 

tightened properly, a leak can be seen and corrected later in the day. It is better to catch the 

problem earlier (on the same day) than later (the next morning after the Z-Mod has been running 

and leaking overnight). 

Filter Bag or Sump Sock 

Clean every 2–3 weeks or as needed (when the water level in the filter bag is higher than 

the water level of the sump or when the bag is overflowing). Remove the felt filter bag or sump 

sock and quickly exchange it for a clean one. Use the tap water hose to blast debris off the used 

sump sock. Soak the sump sock in a 5% bleach solution (100 ml/5 gal or ≈250 ppm active 

chlorine) overnight. It may need two nights of 5% bleaching to get rid of the grime. The next 

morning, rinse the sump sock with tap water for a few minutes. Soak the sock in clean water for 

several hours. Rinse again in running water and hang dry. 

Paper Filter 

Clean once a month or when pressure is less than 5 pounds per square inch or flow rate 

has dropped. Turn off the pump. Attach the hose to the sump and drain the sump water down to 

the green tape mark. Open the air bleed valve on the filter casing (white valve on top of the lid). 

Attach the hose to the drain valve on the Ocean Clear (Appendix A) filter casing #3, and open 

the valve to drain water from the filter (the other filters will drain part way down as well). 

Unscrew the lid (use the rubber mallet to help loosen the lid). 

Remove the carbon in the center and throw it away, keeping the mesh bag. Remove the 

coral bag, but do not throw it away. Put the coral in a “used coral” container (to be reautoclaved) 

11
and rinse out the bag. Reused crushed coral is good for up to 6 months. Remove the used 

circular paper filter. 

Rinse the filter casing (using RO/DI or system water) and use paper towels to remove 

bacterial growth inside the case. Let the dirty water drain from the valve. When the water has 

completely drained, remove the hose and close the valve on the casing. 

Place a clean paper filter in the casing. Fill the carbon bag with about 3–4 cups of new 

Kent Marine Reef Carbon. Rinse the carbon with RO/DI water. Add 1⁄2 to 1 cup of autoclaved 

coral to the coral bag. Put the carbon and coral bags inside the circular filter. 

Wipe down the O-ring and lid of the Ocean Clear case with a paper towel. Put a small 

amount of silicon grease (safe for fish) on the O-ring and screw the lid on the filter. Use the 

rubber mallet to help tighten the lid (do not over tighten). 

Fill the sump with system water. Have extra water in a 5-gal bucket to top off the sump 

as water circulates back into the filters. Turn on the pump. When water reaches the top of the 

paper filter #3, it will come out the air bleed valve. Close the air bleed valve. Air will slowly 

continue to bubble and build up at the top of the canister case. Open the air bleed valve to 

remove this air. 

Clean the dirty paper filter by rinsing it with warm tap water. Put the soiled filter in a 5-

gal bucket. Let it sit in a 5% bleach solution (100 ml bleach/5 gal tap water) overnight. Next 

day refresh the 5% bleach bath for a second night of soaking. On the third day, rinse the filter 

with tap water. Add new tap water and 30 g of sodium thiosulfate to chelate the bleach left in 

the filter. Soak overnight. Rinse completely with tap water and air dry. 

Make sure to log all maintenance information in the logbook (Appendix C: Examples of 

Zebrafish Records). 

Sponge Biofilters 

Sponge biofilters serve an important role in maintaining the nitrogen cycle (Figure 3 and 

Appendix B). The biofilter provides a large surface area for the ammonia- and nitritemetabolizing bacteria (our biofilters are polystrand dual filter pads). Sudden changes in the 

water quality or chemistry (e.g., pH or the presence of chlorine) can kill the bacteria and stop the 

nitrogen cycle, which will lead to the build up of harmful ammonia and nitrites. Therefore it is 

important to keep water conditions stable for the bacteria and ultimately the fish. 

Clean biofilters every 3 months. Turn off the water pump. Attach the hose to the sump 

valve and drain the water in sump to the green tape (above the drain to the pump). Open the air 

bleed valve (white valve on top of the lid) on Ocean Clear filter casing #1. Attach the hose to the 

drain valve on Ocean Clear filter casing #1, open valve and drain water from the filter (the other 

filters will also drain part way down). Unscrew the filter lid (if necessary, use the rubber mallet 

to help loosen it). 

Transfer the sponge pads from the filter case and place them in a bucket of clean system 

water so they do not dry out. Using RO/DI or system water, rinse down and use paper towels to 

12
Figure 3. The nitrogen cycle in aquaculture systems and aquarium tanks. Ammonia is built up from fish 

excretion and uneaten food. Nitrosomonas, a bacterium, converts the ammonia into nitrite, which 

is converted into nitrate by nitrobacter, another bacterium. During both chemical conversions by 

the bacteria, H+

 is released causing a reduction in pH. A reduced pH will change ammonia into 

ammonium, which cannot be converted by the bacteria. Ammonia and nitrite are toxic to fish; 

nitrate is also toxic to fish, but at a much higher level. To reduce nitrate levels, routine water 

changes are required. 

remove bacterial growth inside the filter case. Let the dirty water drain out of the valve. 

Remove the hose and close the valve once the water has drained. Wring excess bacteria from the 

filter pads in the bucket of system water. Removing the excess bacterial growth will allow water 

to flow through the filters efficiently, while the remaining population of ammonia-metabolizing 

and nitrite-metabolizing bacteria will continue the nitrogen cycle (Figure 3). 

Wipe down the O-ring on top of the canister and the canister lid with a paper towel. 

Apply silicon lubricant in a light layer on the filter casing’s O-ring. Screw on the lid. Use the 

rubber mallet to help tighten the lid. 

Repeat the process with sponge filter #2. 

Fill the sump with system water so the pump does not go dry when it is turned back on. 

Keep buckets of water nearby to keep the sump full as the water refills the filters. Check to 

make sure the bypass valve is open. 

Turn on the water pump and close the air bleed valves on filter casings #1 and #2 when 

the water reaches the top of the filter and starts to come out of the air bleed valves. 

Readjust water flow into the tanks via individual row valves and the bypass valve if 

necessary (see Flow Meter and Bypass Valve). 

13
Log all maintenance information in the logbook (Appendix C: Examples of Zebrafish 

Records). 

UV Light Maintenance 

A UV sterilizer kills pathogens before the water enters the fish tanks and is essential in 

preventing the spread of disease within the zebrafish colony. The efficacy of the sterilization 

process depends on keeping the unit well maintained. The UV sterilizer (Emperor Aquatics 

Smart High Output UV Sterilizer) should be on at all times (except when replacing the bulb). 

The bulb should be replaced every 12 months to ensure the effectiveness of the sterilization 

process. 

Consult the UV sterilizer manual (online at http://www.emperoraquatics.com/ 

SMARTUV_Instructions_07.pdf) for full instructions and diagrams. Four feet of clearance is 

needed to remove the bulb. Unplug the light before replacing the bulb. Drain the water from the 

sump, keeping the water level above the outlet to the water pump. To drain water from UV unit, 

open the valve to the UV sterilizer drain (it is the ball valve above the pump, towards the back of 

the Z-Mod, on the tubing). Tilt the UV unit and drain the water out. A little bit of water will 

remain and drip out when the unit is opened, so be prepared for some water to leak. Unscrew the 

end (black retaining nut) of the UV light. Unscrew the internal white quartz sleeve retaining nut 

(it holds the quartz sleeve to the ballast). Unplug the ballast from the UV light. Carefully 

remove the bulb from the quartz sleeve. 

Check to see whether the quartz sleeve is clear. If there is any opacity, clean it off with a 

scrub brush and a little water; opacities on the sleeve decrease the efficacy of the bulb. Slide a 

new UV bulb into the quartz sleeve. Plug the ballast back into the UV light. Hand tighten the 

white internal and black external retaining nuts into place, making sure they are well sealed. 

Turn the UV light back on. Close the ball valve to the UV sterilizer drain. 

Fill the sump with RO/DI or system water. Turn on the pump to start water circulating 

into the UV light. Add more RO/DI or system water to the sump as water circulates through the 

UV sterilizer. 

Leave the flow control valve by the UV light open (i.e., do not adjust it). It affects how 

fast the water goes through the UV system, which in turn affects the length of time the water is in 

contact with the UV light. In some circumstances (if a disease outbreak occurs) it may be 

desirable to partially close the valve to increase the time the water is exposed to the UV light. 

However, partially closing the valve also increases water pressure against the water pump. This 

will in turn add unwanted pressure throughout the filtration system. 

Cleaning 

Zebrafish equipment and the Z-Mod need constant cleaning to reduce chances of 

spreading disease among the fish and to reduce harmful levels of nitrates (Appendix D: Zebrafish 

Diseases). 

14
It is important to note that the only disinfectants used are alcohols, bleach, benzalkonium 

chloride, and methylene blue. Items cleaned with alcohols and bleach should be completely dry 

before use. Benzalkonium chloride and methylene blue are the active ingredients in Net Soak 

(see Nets) and are not harmful to fish at the concentrations used. Detergents such as hand or dish 

soap should never be used on any zebrafish equipment. However, hand soap may be used to 

prevent the spread of diseases. Hand washing is encouraged before and after feeding or handling 

the fish. 

Countertops 

Countertops should be sterilized with 70% ethanol or isopropanol after fish tanks have 

been on the counter and before making paramecium cultures. This will reduce the spread of 

diseases among fish. 

Also, no chemicals such as toxins, contaminants, or fixatives are allowed on the 

countertops in the zebrafish room, especially near the Z-Mod. Laboratory experiments utilizing 

zebrafish embryos, larva, or adults must take place away from the Z-Mod with no chance of 

introducing a chemical to the system. Exposed fish are not allowed to return to the zebrafish 

colony unless all traces of the exposure chemical are removed (and cannot be excreted by fish or 

transmitted with the fish water). 

Nets 

Use a clean net for each Z-Mod tank (i.e., do not double dip). Clean, dry nets are located 

on the drying rack. 

Sterilize nets between each use. After using a net, rinse it with tap water. Place the net 

into the designated beaker of Net Soak (Appendix A) solution (≈1.5 tsps/5 L). Soak the net 

overnight. The following morning, remove the net from the Net Soak solution, shake off excess 

solution into the sink, and hang up the clean net on the drying rack. The Net Soak solution can 

also be used to sterilize sponges and bottlebrushes. Change the solution once per week or more 

often when use is heavy. 

Bleach Bath 

Use the bleach bath to sterilize spawning cages and other equipment that will come in 

contact with the fish (e.g., turkey basters, larval tanks, Z-Mod tanks, tank lids) or any cultured 

organism (e.g., paramecium Tupperware, paramecium beakers). Mostly glass and plastic items 

will be placed in the bleach bath. Use tap water to rinse and scrub off feces, old food, etc., from 

tanks and lids before fully submersing items into the bleach bath. Do not put turkey baster bulbs 

in the bath as the bleach will break down the rubber. 

Change bleach bath water every 2–3 months or sooner if use is heavy. To make the 

bleach bath, add 750 ml of bleach to 10 gal of tap water (20% bleach solution or ≈1,000 ppm 

active chlorine). Leave items in the bleach bath for at least 1 h or overnight. Triple rinse the 

items with plenty of warm tap water in the sink. Air dry the items separately on the drying rack. 

Bleach can kill fish and embryos, so it is very important to rinse well and completely air dry. 

15
Tank Lids and Outlets 

Clean lids once a week or sooner if dirty. Using a paper towel or sterilized bottlebrush, 

wipe away food left on tank lid. If the tank lid is particularly dirty, switch out the dirty lid with a 

clean one. Scrub the dirty lid and put it in the bleach bath. 

Check the outlet where the water flows out of the tank daily. It should be clear of debris. 

If bacteria and gunk are growing on the outlet mesh, use a finger or a sterilized bottlebrush to 

remove the material from the mesh (and out of the tank) or replace the dirty outlet with a clean 

one. Scrub the dirty outlet and put it in the bleach bath. 

Nursery Tanks 

Check nursery tanks (bottom row of the Z-Mod) daily and clean as needed. It is easy to 

overfeed fish in the nursery tanks, so check daily to make sure fungus is not growing on the 

bottom of the tanks. 

Every other day the dirty tanks should be removed from the Z-Mod and fungus on the 

bottom of the tank taken out. Use a turkey baster to gently remove the fungus, being careful not 

aspirate small fish. Do not use a siphon to clean the nursery tanks, as the water flow would be 

too great for the small fish and would likely aspirate them. 

Siphoning Adult Tanks 

Check adult tanks for build up of debris on the bottom and clean with a siphon when 

dirty. Fuzzy buildup on the tank bottom is fungus growing on old food; the darker materials are 

feces. These can be harmful to zebrafish and should be siphoned out of the tank. To clean the 

tanks, remove the tanks to be siphoned from the Z-Mod and place them on the counter. It is 

easier to set several tanks on the counter at the same time. 

The siphon tool (Figure 4) is a vacuum-shaped nozzle (a Nalgene T-type connector with 

the length of the top of the “T” cut off to form a small slot opening and aquarium safe silicon 

sealing the ends of the top of the “T”) attached to a rigid tube. The other end of the rigid tube is 

connected to a flexible length of tubing with an inner diameter of 3/8 inch. A tubing pinch 

clamp is located on the flexible part of the tubing close to the rigid tube. 

To siphon, fill the tubing with water (use system water in case there is back flow) by 

submerging the siphon tool in system water or pouring it through the tube. Once there is water 

in the siphon (with no large air bubbles), use the clamp on the tubing to pinch off the flow so the 

water cannot escape. Then place the nozzle in the tank and put the tube end in an empty bucket 

on the floor. Release the clamp and the water should start flowing into the bucket. 

The water will flow from the high spot (the tank on the counter) to the lower spot (the 

bucket on the floor). Siphon out debris by moving the nozzle over the bottom of the tank, 

scraping, and loosening the debris to be siphoned off by the tube. To move from tank to tank 

without starting the siphoning process again, simply clamp the tubing and move the nozzle. 

16
Figure 4. Siphon for cleaning out Z-Mod tanks. This easily assembled siphon makes removing detritus 

and feces from Z-Mod tanks more efficient. 

Be careful not to siphon out all the water (there will still be fish in the tank during this 

process) or fish (smaller fish might get drawn into the siphon). Always check the bucket for fish 

before siphoning a different tank and before emptying the bucket. 

Back of the Z-Mod 

As part of its design, effluent tank water runs down the back, inside wall of the Z-Mod. 

Bacteria build up on the inside wall, making it brown and slimy. Once a year, use RO/DI or 

system water (not tap water) to clean a portion of the Z-Mod. Do not clean the entire Z-Mod at 

the same time or the sump sock will quickly become clogged with bacterial mats. Use a sponge 

that has not been treated with a fungicide (if a new sponge feels damp coming out of the packet, 

it has probably been treated with a fungicide), a sponge that has been well rinsed of the fungicide 

(i.e., the sponge is held under running tap water, while being repeatedly wrung out for 5–10 

min), or paper towels. After cleaning the back of the Z-Mod, change the sump sock (see Filter 

Maintenance). 

17
Culturing Zebrafish 

Food for Adult Zebrafish 

Adult zebrafish are fed dry salmon starter meal (Bio-Oregon Bio Vita starter, Appendix 

A) and Artemia sp. twice a day. Feed the dry salmon meal first, as it is more nutritious than the 

Artemia. 

Dry Salmon Food 

One pinch (0.05–0.07 g) of dry salmon meal is fed to each tank (of ≈15 fish). The 

salmon starter should be kept dry and well preserved; it is stored in a –20°C freezer. In order to 

ensure the zebrafish are given the right nutrients and vitamins (which degrade with time), new 

food should be ordered every year. 

Since our zebrafish are used for toxicology research, each new bag of salmon starter 

should be tested in-house for contaminants (e.g., polycyclic aromatic hydrocarbons, persistent 

organic pollutants, and metals) to determine background levels and whether the food is suitable 

for use. These precautions are taken because fish processed by commercial fish food makers to 

make salmon meal could contain these pollutants. 

Artemia Culture 

Adult zebrafish are fed Artemia that are approximately 40–48-h old. Therefore, Artemia

cultures will be fed to fish 2 days after they have been started (e.g., a culture started Monday 

afternoon will be fed Wednesday morning and afternoon). 

Artemia culture setup 

To hatch a small amount (at most 2 tsps) of Artemia, two 2-L Artemia cones are needed. 

One cone is designated the “0–24 hour” Artemia; the other is “24–48 hour” Artemia. An air 

pump with one air hose, which consists of aquarium tubing and a 1-ml glass serological pipette, 

is used for each cone to keep the Artemia eggs (cysts) well aerated and suspended in the water. 

Both cones of Artemia must be aerated at all times except when harvesting or preparing the 

Artemia. 

Artemia hatch at around 20 h at 25oC. However, at room temperature (23oC) the hatching 

rate will be slightly delayed. Therefore, we culture our Artemia for more than 24 h. The 

additional time will allow any delayed Artemia cysts to hatch. Additionally, the two-cone 

system allows for more of the eggshells to be removed from the culture. While using a heater to 

increase the temperature of the Artemia culture will cause them to hatch sooner (see Starting 

Larger Amounts of Artemia Cysts), our two-cone method of hatching Artemia at room 

temperature meets our needs for a small zebrafish colony. 

18
Feeding Artemia to the fish 

To harvest the Artemia, remove the air hose from the 24–48-h cone. This cone contains 

Artemia that have been growing for nearly 2 days. After approximately 5 min, the Artemia

culture will settle into three layers. Most of the unhatched cysts will settle to the bottom, the 

hatched Artemia will settle just above the bottom layer of unhatched eggs, and the eggshells will 

float to the top. To dispose of the unhatched eggs, open the bottom valve and release a small 

amount of brown liquid (containing the unhatched cysts) down the sink drain. Close the valve as 

soon as the water turns orange (containing Artemia). 

To collect Artemia for feeding, first dampen the designated Artemia net with system 

water to allow the Artemia water to pass through more easily. For the first feeding of the day (in 

the morning), open the valve to drain one-half of the Artemia into the Artemia net. Return the 

remaining one-half of the Artemia in the 24–48-h funnel to the holder and put the air hose back 

in the funnel. 

Rinse off excess salt water from Artemia in the net with a squirt bottle of system water. 

Turn the net inside out and rinse the Artemia off the net into a 100-ml glass evaporation bowl or 

similar size beaker using the squirt bottle of system water. Dilute the Artemia out to 

approximately 100 ml (if 2 tsp of Artemia cysts were hatched). Allow the Artemia to settle for 

1–2 min. Any unhatched Artemia that have made it through the cone will settle on the bottom 

and the eggshells will form a brown layer floating on top of the bowl. When pipetting out the 

Artemia, avoid the eggs and shells as much as possible. Feeding the indigestible unhatched cysts 

and eggshells to fish, young ones in particular, can lead to gut obstruction and ultimately death. 

Use a pipette (plastic or glass, depending on preference) to remove Artemia from the finger 

bowl, and distribute them among the fish tanks. Feed according to the number of fish per tank 

(i.e., feed more if there are more fish in a tank, less if there are fewer). Feed approximately 1 ml 

of Artemia to a tank of 15 fish. 

For the second, afternoon feeding, drain the remaining Artemia through the dampened 

Artemia net and close the valve before draining the eggshells on top. Collect the Artemia into 

the evaporation bowl and feed them to the zebrafish as described above. After the second 

feeding of the day, empty and scrub the cone using tap water and a sponge (see sponge 

comments under Cleaning the Back of the Z-Mod). The clean cone will be used to hold Artemia

for the next day’s feeding.

Preparing 0–24-h Artemia for next day’s feeding 

Every afternoon, after the second feeding or single feedings on the weekends, Artemia

are prepared for the next day’s feeding. 

Remove the air hose from the 0–24-h Artemia cone, which contains Artemia that have 

been culturing for 24 h. The unhatched eggs and the hatched Artemia will settle to the bottom 

and the eggshells will float to the top. It is important not to transfer the majority of the eggshells 

at this stage. Open the bottom valve and drain the unhatched eggs and Artemia from the 0–24-h 

cone into the cleaned 24–48-h cone, but do not drain the eggshells floating on top into the 24–48-

h cone. Close the bottom valve to stop the flow before reaching the eggshells on the top. 

19
Artemia are able to start feeding at 32 h. Since the Artemia are fed to the zebrafish at 40–

48 h, the Artemia are fed an enrichment to load their guts (bioencapsulation) to maintain their 

nutritional value. Add a couple dashes of Artemia food (a dry powder food mixture containing 

carotenoids and highly unsaturated fatty acids or a commercially available Artemia enrichment) 

to the 24–48-h Artemia cone. Replace the air hose in the 24–48-h cone to aerate the Artemia for 

next day’s feeding. 

Discard the eggshells from the 0–24-h cone. Rinse and scrub the cone under tap water 

with a sponge (see sponge comments under Cleaning the Back of the Z-Mod) to start a new 

Artemia culture. Bleach and dry the cones every 2 weeks to reduce bacterial growth, which will 

cause the eggs to stick to the sides of the cone (the eggs will not hatch). 

Starting Artemia from cysts 

A new batch of Artemia is made every afternoon. 

Artemia require a salinity of about 1 tbsp of sea salt (Instant Ocean) per 1 L of RO/DI 

water (about 4 tbsp/gal). In the 2-L plastic container labeled “Artemia Water,” add about 2 tbsp 

of sea salt to 1.75 L of RO/DI water. The solution should be saturated, so there will be some 

nondissolved salt on the bottom of the container. Decant the saltwater into the clean 0–24-h 

cone (try not to add the nondissolved salt to the Artemia water). Do not overfill the funnel above 

the fill line. If the water level is too high, Artemia cysts will stick to the lid of the funnel and will 

not hatch. 

For our system containing approximately 2,000 fish, add about 2 tsp of Artemia cysts to 

the 0–24-h funnel filled with the Artemia water. The amount of eggs used may change 

depending on the number of fish in the breeding colony, but in general use about 1 tsp of eggs 

for 1,000 fish. Replace the air hose in the cone, and make sure the airflow is sufficient for the 

cysts to stay suspended in the water (the best position for the hose is on the bottom side of the 

cone). The Artemia will go anoxic and not hatch if aeration is not sufficient. 

Artemia cysts are stored in the refrigerator (4oC). 

Starting larger amounts of Artemia cysts 

If more than 2 tsp of Artemia are required, then additional hatching cones or a larger 

Artemia hatchery are needed. The volume of water should be scaled up at the same salinity, and 

the egg density (1 tsp cysts/1 L Artemia water) should be approximately the same or lower. If 

the density of the cysts is too high, the hatching rate will decrease. 

To hatch a larger amount of cysts, eggs should first be decapsulated (see following 

subsection). Add an aquarium heater set to 26–28°C to the water so the Artemia will hatch at a 

faster rate. This way the Artemia will be ready to feed to the zebrafish within 24 h (ideally 

Artemia should be fed to the adult or larval zebrafish immediately after hatching). Harvest the 

Artemia within 24 h using the same protocols as above. The Artemia can be diluted into a small 

squirt bottle with system water for quicker feeding. 

20
Starting Artemia using decapsulated cysts 

When a larger amount of Artemia is required to feed a bigger zebrafish colony, Artemia

cysts need to be decapsulated. The advantages of decapsulated eggs are a higher hatching rate 

and the softened eggshells are easier on the digestive systems of the both adult and larval 

zebrafish. However, the process of decapsulating cysts requires greater effort. When hatching 

smaller batches of cysts (see Starting Artemia from Cysts), the procedure described above is 

sufficient for removing indigestible eggs without taking additional time. 

To decapsulate the eggs, add several teaspoons of cysts to a dish with system water. Let 

the eggs soak for at least 30 min. Pour the eggs into the Artemia net for cysts. Immerse the net 

with the eggs into a bowl of full strength bleach, while keeping the eggs contained in the net. 

Using a disposable pipette, stir the eggs until they start to change color from brown to a pale 

orange color (≈1.5 min). Rinse the eggs (still in the net) thoroughly with RO/DI water (pour at 

least 1 gal over the eggs) to remove all the chlorine (trace amounts of chlorine will kill the 

hatched Artemia). Add eggs to the hatchery cone filled with Artemia water, then add the air hose 

and ensure there is enough air movement to keep the eggs suspended. 

Weekend Feedings 

On weekends, when there are no larval fish in the husbandry facility, adult fish are fed 

once a day. Under these circumstances, weekend feedings are like afternoon feedings. The fish 

are fed dry salmon food and all of the Artemia in the 24–48-h cone are drained and fed. The 0–

24-h Artemia are drained into an emptied and cleaned 24–48-h cone and a new Artemia culture is 

started in a cleaned 0–24-h cone. If larval fish are being raised over the weekend, then two 

feedings per day are required (see Raising Larvae and Beyond). 

Spawning Fish and Collecting Embryos 

The day before zebrafish embryos are needed, adult males and females are combined in 

spawning tanks in the afternoon. They are left overnight in the tanks together. Dawn (i.e., when 

the lights turn on in the morning) cues the fish to start spawning, which generally occurs during 

the first 4 h of the day. After spawning, the fish are returned to their respective Z-Mod tanks, 

and the embryos are collected, cleaned, and stored in the incubator. 

Spawning fish 

Set up fish to be spawned in the afternoon, before the last feeding of the day. This 

reduces the amount of feces on the bottom of the cage with the eggs (thus making the embryos 

easier to clean). 

Before setting up the spawning tanks, sterilize the counter with 70% alcohol prior to fish 

transfer (if a fish falls onto the countertop, it does not get contaminated). To set up a spawn, fill 

the spawning tank (which consists of a plastic mesh bottom insert that fits into an outer plastic 

tank) with system water from the system water tower. Once the spawning tanks are filled with 

water, place them on the counter where the spawning will take place. 

21
Select the tanks containing fish to be spawned from the Z-Mod. Fish can be spawned at 

most once per week, otherwise protein and lipid reserves in the fish become exhausted, resulting 

in a poor egg yield. Check the logbook to choose fish that have not been spawned within the last 

week. 

Use nets that have been sterilized with purple Net Soak solution (see Nets) to transfer fish 

from their original tanks into the spawning tanks, using a different net for each tank. Depending 

on the number of adult fish available and the number of eggs needed, set up the spawns using the 

following ratios. To yield the greatest amount of eggs, the best spawning ratio is 2:3, males to 

females. Ratios of 1:1 or 1:2 also work (and are also less stressful for all fish involved). 

In general, wild-type zebrafish can be sexed by color and body shape. Females have 

large bellies that descend from the body in the anterior region. Males have a red-yellowish hue 

in the anal and caudal fins and tail, and are more streamlined in shape (i.e., they lack a belly). 

Sexing fish takes practice and is greatly aided by direct guidance from someone who is 

experienced. 

After placing males and females in the spawning tanks, clean the nets by rinsing out 

debris with tap water, then place them into Net Soak. Put covers on the spawning cages and 

stack the cages on the sterilized counter away from general use. 

Label the spawning tanks with the fish tank names (see Maintaining a Breeding Zebrafish 

Colony). This ensures the fish are returned to their correct tank and avoids mix-ups. 

The lights in the room are programmed to turn on at 800 hours and go off at 2200 (14-h 

light cycle). The next morning, zebrafish spawning behavior will be triggered by the lights 

coming on (simulates dawn). The fish will generally spawn during the first 4 h after the lights 

come on (between 800 and 1200). Occasionally, the fish will spawn the same evening the tanks 

are set up. This can be avoided by setting up the fish later in the day, after noon at the earliest. 

To reduce the amount of feces mixed with the eggs and to stimulate spawning, move the 

inner tank with the fish to a fresh tank of system water early in the morning before spawning 

occurs. 

Once the fish have spawned, they cease spawning behavior (males pursuing females). 

Record the spawning activity (which fish tanks were spawned, egg quality and quantity, time 

spawned) on the log sheets (Appendix C: Examples of Zebrafish Records). Put the fish back into 

their correct respective tanks using a clean net. Place them back on the Z-Mod to allow them to 

recover. 

Embryo Collection 

To harvest the embryos, remove the insert from the spawning tank. Once the embryos 

and feces have settled to the bottom of the outer tank, slowly decant the water from the spawning 

tank into the sink. When only the embryos and debris remain in a small amount of water, pour 

the remaining water through the embryo strainer (a plastic tri-pour beaker with the bottom 

removed and a piece of 500-μm mesh netting attached to the top with silicone aquarium sealer). 

Wet down the mesh with system water to allow the water to pass through more easily. Embryos 

22
will catch on the mesh and smaller debris will pass through. Use a squirt bottle of system water 

to wash any remaining embryos from the bottom of the spawning tank into the strainer. Gently 

rinse the embryos on the strainer using the squirt bottle. This will push the softer remaining 

feces through the strainer, leaving cleaner embryos. 

Turn the strainer over and gently wash the embryos into a clean plastic petri dish with the 

squirt bottle of system water. Fill the petri dish half way with system water to keep the embryos 

in water. 

After spawns are taken down and embryos are harvested, wipe down the countertops and 

sterilize the surface with 70% alcohol. 

Clean the embryos under a stereomicroscope by removing remaining feces, unfertilized 

eggs, and scales with a wide mouth glass pipette and a pipette pump. Unfertilized eggs can be 

distinguished by comparing the embryos to the stages as documented by Kimmel et al (1995). 

Remove embryos that are not developing normally (e.g., asymmetrical cleavage) or that do not 

appear to be developing (i.e., if they look like they are in the one-cell stage when all other 

embryos are in the 48-cell stage, it is highly likely they are unfertilized). Changing the system 

water after cleaning the embryos is also recommended. 

Daily water renewals and removal of dead embryos are imperative for keeping embryos 

and larvae healthy and less susceptible to Coleps sp. infestations (a single cell organism that can 

feed on zebrafish embryos). Also limit the number of embryos to a maximum of 50 for each 100 

mm petri dish; 30 embryos per dish are optimal. 

Label, date, and initial the petri dishes and place the embryos in the incubator for 

consistent development. The incubator is set at 28.5oC, which is used as the standard for 

zebrafish development (Kimmel et al. 1995). Later in the afternoon, check the embryos again to 

remove any missed unfertilized eggs. 

If the fish are not spawning or are not producing enough embryos, refer to Maintaining a 

Breeding Zebrafish Colony. 

Raising Larvae and Beyond 

Raising zebrafish larvae is an intensive process requiring preparation and planning. At 4 

days postfertilization (dpf), larvae are moved from the incubator into a lighted water bath, where 

they will be fed a special diet, raised in a larger volume of water, and given daily water changes. 

When they are large enough to regularly eat Artemia, they are moved to the Z-Mod. 

Setup 

Zebrafish larvae absorb most of their yolk by 6 dpf. However, food should be introduced 

to the larvae at 4 dpf to encourage a better growth rate and survival. It is best to start feeding 

larvae at the beginning of the week (Monday/Tuesday), when they will get better care during that 

critical period of development. Therefore, it is best to spawn adult fish on Wednesday to 

produce larvae that will be ready to be fed on Monday. 

23
To raise larvae, prepare a water bath to maintain the temperature of the larval rearing 

tanks. Use a large shallow plastic container (such as an inexpensive storage tub) partially filled 

with system water. Use system water when filling the water bath to prevent accidental water 

contamination. For example, when removing the larval containers from the water bath, water 

could drip off the outer surface of the larval tank into another tank. Use one or two submersible 

heaters to maintain the water bath temperature between 26–28oC. Keep a thermometer in the 

water bath to check the temperature. The heated water will increase the growth rate and provide 

a stable environment for the larvae. Place the water bath in a lighted area (ideally with the same 

14-h light/10-h dark cycle as the adults). The light will allow the larvae to more easily detect 

and capture their prey. 

Next, fill the larval rearing tanks, which are clear, 1-L square salad bar containers, to 

approximately a 2-inch depth with system water (≈500 ml). Put the larval tanks in the water 

bath. Adjust the water bath level so the tanks are not freely floating but the heaters are still 

submerged. The day before the larvae are transferred to the larval tanks, turn on the heaters so 

the water bath will come to the desired temperature. 

When the larvae are 4 dpf, transfer them from the petri dishes in the incubator to the 

larval tanks. Place 20 to 25 larvae in each 1-L larval tank. Higher densities will diminish 

growth. To transfer the larvae from the petri dish to the tank, immerse the petri dish containing 

the 4 dpf larvae into the larval tank water. Remove the dish and check for any larvae still on the 

dish. 

Label each larval container with the type of stock or transgenic line and the date of 

fertilization. Complete a new larvae fish feeding log sheet (Appendix C: Examples of Zebrafish 

Records) for each stock or transgenic line; include the parental lineage used to generate the new 

larvae. 

Every 2–3 days refill the water bath with RO/DI water, as the water will evaporate. 

Feed Schedule 

Feed the dry larval food first, then the paramecia (see Paramecium Feeding), and finally 

Artemia, if needed. 

Age (days) Morning Noon Afternoon

1–6 Dry larval food Dry larval food Dry larval food 

3,000 paramecia 3,000 paramecia 

 7–12+ Dry larval food Dry larval food Dry larval food 

3,000 paramecia 3,000 paramecia 

3 drops Artemia 3 drops Artemia 3 drops Artemia

Dry Larval Food 

The dry food is a 1:1:1:1 mixture of Spirulina, Artificial Plankton, Micro-Food, and 

Cyclop-eeze. Alternatively, use Zeigler Larval AP100 (Appendix A), which is a complete 

mixture of dry food formulated for fish larvae. A stock of the dry food is stored in the 

refrigerator. As with adult fish food, larval fish food should be ordered every year to ensure 

24
proper nutrition. For feeding, larval fish food is stored in a 50-ml conical test/centrifuge tube 

with an approximately 2-mm hole in the lid. Give one shake of the tube over a larval tank to 

dispense the dry food. 

Paramecia 

See Paramecium Feeding. 

Artemia 

At 24-h old, Artemia sp. are smaller and still have yolk, which is a better size for the 

small mouths of larval zebrafish and is nutritionally desirable. Unlike adult zebrafish, which are 

fed 40–48-h old Artemia, Artemia fed to larvae should be 24-h old. Therefore, an Artemia cone 

needs to be set up specifically for the larvae. Harvest and set up the larval Artemia cone each 

morning before the larvae are fed. Harvest all the 24-h old Artemia in the morning using the 

same protocol described for harvesting Artemia for adult zebrafish. Once diluted with system 

water in an evaporation bowl, the Artemia can be kept in the bowl for the rest of the day. There 

will still be eggshells and unhatched eggs in the bowl. Avoid feeding these to the larvae. 

Prepare the cone for the next day’s Artemia culture using similar protocols for setting up the 

Artemia for adult zebrafish. Use only 1 L Artemia water (1 tbsp Instant Ocean/1 L RO/DI water) 

and only 1⁄4 to 1⁄2 tsp of Artemia cysts (depending on number of larval tanks, 1⁄4 tsp is sufficient to 

feed eight tanks). 

Cleaning Larval Tanks 

Start cleaning the tanks when the tank bottoms collect noticeable amounts of leftover 

food or on the third day of feeding (when larvae are 7 dpf). Clean the containers once a day in 

the morning before feeding. 

Use a clean (bleach-sterilized) turkey baster to remove debris and most of the water, 

leaving enough water so the larvae can swim freely. Use a beaker to contain the dirty water. Be 

careful not to aspirate the larvae. Another method of cleaning larval tanks is to slowly pour off 

the water, being careful not to lose any fish. Use a clean pipette to “chase” fish away from 

where water is being poured off. Swish fresh system water around on the bottom of the larval 

tank to loosen moldy food, then pour off the fouled water. 

For either method, fill the larval tank with clean system water back to a 2-inch depth or 

about 500 ml. Repeat cleaning if the water is still cloudy. Between each larval tank cleaning, 

check the dirty water in the beaker for larvae before dumping it down the sink. Empty the 

beaker between each tank cleaning so the origin of stray larvae is not in question. Use a clean 

pipette to retrieve fish. 

If dead larvae are found, remove these and record the number of mortalities on the larval 

fish feeding log sheet (Appendix C: Examples of Zebrafish Records). 

25
Controlling Coleps

The protozoan Coleps is ubiquitous in zebrafish colonies and typically eats decaying 

matter. However, large outbreaks of Coleps kill and eat live larvae. There are two main sources 

of Coleps: the water with the embryos and the paramecium cultures. Ideally zebrafish should not 

be raised in Coleps infested waters. To prevent Coleps from growing in petri dishes with the 

embryos, water changes should be done daily and the density of embryos should be kept low. To 

prevent Coleps from being introduced through contaminated paramecium cultures, check the 

cultures before feeding (see Paramecium Culture). If there is a high population of Coleps, it is 

best to raise embryos only when a clean culture can be produced. 

With great care, however, it is possible to raise embryos in high Coleps conditions. First 

make a high salinity (conductivity is ≈ 2,500 μS/cm) Instant Ocean sea salt solution in a 5-gal 

bucket. Use this water to raise the embryos. Complete water changes are essential and should 

be done every day (do not wait until 7 dpf). When the larvae look like they will survive (i.e., are 

eating Artemia), slowly lower the conductivity each day by increments of 200–250 μS/cm until it 

is the same as the Z-Mod (1,500 μS/cm). Then the larvae can be moved to the Z-Mod. 

Moving Fry to the Z-Mod 

Fry are moved to the Z-Mod around day 13. They must pass an “orange belly test” 

wherein the larval tank is held to the light and the bellies of the fry are examined. If they are 

bloated and orange, this indicates they are eating Artemia and can be moved to the Z-Mod 

system. Fry should be moved to the bottom shelf of the Z-Mod, the “zebrafish nursery,” where 

the water flow is adjusted to a slower flow rate (≈1 ml/sec). 

When they are young and small (<1 month old), fry can be kept in groups of up to 20 fish 

per tank. As they get older and bigger, they will be separated into smaller groups of up to 15 fish 

per tank (see Maintaining a Breeding Zebrafish Colony). 

Once the fish are introduced to the Z-Mod, they should be fed twice daily. Typically, 

during the first week that they are in the Z-Mod, start feeding them a tiny pinch (about the size of 

a grain of rice; smaller than the amount fed to adult tanks) of small, fish crumble (Bio-Oregon 

Bio Vita dry salmon starter that is processed in a coffee grinder so that it is small enough for the 

fry to eat) and 5–7 drops of Artemia. Increase the amount of dry salmon food and Artemia as 

they get older. The amount of food is a judgment call that relies on experience. Overfeeding can 

be detrimental to the fry, as leftover food can quickly be overgrown by fungus. The tanks should 

be checked every other day and cleaned when necessary (see Cleaning). It is best to avoid 

overfeeding all together by adjusting the amount of food so that the young fish are satiated, 

without leftover food. 

Tank Outlet Size 

Use the appropriate outlet with each tank. There are three types of outlets for the 

different stages of zebrafish life. They are color coded orange, white, and blue or red. For small 

fry in the nursery, use the orange outlets (mesh size 500 μm). This mesh is fine, so fry cannot 

pass through the holes. The mesh also catches most of the food and quickly becomes clogged; 

make sure the mesh is clear of debris so the tank does not overflow. When fish are slightly 

26
bigger (≈1 month), the white outlet can be used (mesh size 1 mm). When the fish are almost 

adult size (≈2 months) use the blue/red outlets with the largest mesh size (4 mm). The 

appropriate size mesh allows for the safety of the fish and proper water flow. 

Out of the Nursery 

When the fish are large enough to handle a blue/red outlet (≈2 months), they are also able 

to deal with a stronger water flow (≈2 ml/sec). Then they can be moved from the nursery to a 

different shelf in the Z-mod where the flow is faster. 

Paramecium Culture 

Culturing paramecia (Paramecium multimicronucleatum) involves supporting a food 

chain: boiled wheat pellets are broken down by bacteria, which are eaten by flagellates, which 

are eaten by the paramecia. Sustaining this food chain can be very sensitive; cultures can 

become contaminated, paramecium populations can grow too dense and crash, or paramecium 

cultures can go anaerobic (and smell like rotten eggs). It is important to plan accordingly by 

using aseptic, sterile techniques when preparing the cultures and solutions and by checking the 

cultures before using them. 

The general process of growing paramecia to feed larvae involves keeping stock cultures 

in bottles. Bottle cultures are created by inoculating a bottle of new media with an existing bottle 

culture of paramecia. Bottle culture propagation should continue at all times, even when no 

larvae are being fed. To create larger working cultures, the paramecia from the bottle cultures 

are used to inoculate Tupperware containers filled with culturing media. These Tupperware 

cultures are then fed to the larvae when their density reaches 150–200 paramecia/ml. 

Prepare cultures at least 1 week (2 weeks ideally) before zebrafish larvae are fed at 4 dpf. 

When making cultures, stagger them by 1 week so densities do not drop while larvae are being 

raised. Make cultures according to how many larvae are being raised or expected. For example, 

when raising 16 containers of 20–25 larvae, make two Tupperware cultures (see below) 1 week 

before the larvae start feeding. Then 1 week later (when the larvae are feeding), make at least 

one more Tupperware culture. 

Needed for 16 Larval Tanks 

Autoclaved system water (10 L in a 25-L carboy) 

Pellet stock solution (five 1-L bottles) 

Paramecium bottle culture (four 1-L bottles) 

Paramecium Tupperware culture 

Autoclave 

Autoclaving liquid takes approximately 1.5 h, plus 10 min for cooling. A few rules about 

the autoclave: 1) make sure all plastic and glassware are autoclavable; 2) when autoclaving 

27
liquids, there must be an equal volume of air space as there is volume of liquid; and 3) do not 

tighten caps on bottles or the carboy before autoclaving. 

Pellet Stock Solution 

For making approximately 2 L: 

• Obtain a clean 2-L glass beaker. On a hot plate, boil 1 L of RO/DI water with 20 

protozoan pellets (Carolina Biological) and 2 grams of NaCl. 

• Boil for 10 min and allow sediment to settle for 10 min. 

• Decant supernatant into five 1-L glass bottles (≈160–200 ml per bottle). 

• Top off bottles to 400 ml with RO/DI water. 

• Add about 1 tsp of the remaining pellet sediment (the residue leftover after the 

supernatant was poured off) to each bottle. Discard remaining sediment. 

• Label and date bottles. 

• Autoclave pellet stock solution for 20 min. 

Assessing the Purity and Density of Paramecium Cultures 

For assessing the cultures: 

• The purity and density of the bottle and Tupperware cultures should be assessed before 

using them. 

• The density of the paramecia will change daily. The paramecium population will grow 

steadily, plateau at the highest density, then drop off. Cultures should be used at 

maximum density (150–200 per 1 ml). 

• Use a clean, unused pipette to stir the culture. 

• Remove 1 ml of the culture and observe it under the microscope. 

• Slow down paramecium movement with MS-222 (tricaine). 

• Under a stereomicroscope, count the number of paramecia to get an estimate of the 

density. 

• Observe the paramecium culture under high magnification to see whether there are alien 

or unwanted protozoa, such as Coleps. If there are Coleps in the bottle culture, discard 

the culture and order more paramecia or create a new culture (see following subsection). 

Do not use Tupperware cultures that are infested with Coleps. If there is no choice and 

the larvae must be raised with Coleps, refer to Controlling Coleps. 

Paramecium Bottle Culture 

For making approximately 2 L: 

• Shake an autoclaved pellet stock solution to suspend sediment. 

• Aliquot 40 ml of pellet stock solution into each of four or five 1-L glass bottles. 

28
• Add 320 ml of RO/DI water to each bottle. Label each bottle with the date the culture 

medium was made. 

• Autoclave bottles for 20 min. 

• Assess the purity and density of the existing paramecium bottle culture. The culture 

should be between 7–10 days old or when the density of the paramecia is the highest. 

• Inoculate at least two bottles to start future paramecium Tupperware cultures. For each 

new bottle culture, thoroughly mix the existing bottle culture, measure out 40 ml, and add 

it to the new bottle culture. 

• Label the bottle culture with date inoculated. Store bottle cultures at room temperature 

with a loose lid (paramecium cultures are aerobic). 

• Ready for use in 7–10 days. 

Paramecium Tupperware Culture 

For making approximately 4 L: 

• Make cultures at least 1–2 weeks ahead of time before raising larvae. 

• Disinfect the countertop and two Tupperware containers (containers are the size of a 

shoebox with fitting lids, but are not air tight) by spraying them with 70% ethanol and 

allowing to air dry or wiping down with a clean paper towel. 

• Boil 15 wheat berries per Tupperware container (30 total for 2 cultures) in RO/DI water 

for 7 min. 

• Add 200 ml of pellet stock solution, 200 ml of inoculated paramecium bottle culture 

(assess the health first), and 15 drained, boiled wheat berries to 2 L of autoclaved system 

water for each container. 

• Keep the lids on the Tupperware. 

• Label the Tupperware with the date. One week later, assess the purity and density of 

paramecia. Note density and observations on the label. 

• Tupperware cultures can be kept for up to 1 month. Check purity and density of the 

paramecia before feeding. 

Paramecium Feeding 

For feeding proceed as follows: 

• Use the oldest and densest paramecium culture for feeding. 

• Remove lid for only short periods, because the culture can be contaminated by airborne 

particles. 

• Use a stainless steel spatula sterilized with alcohol or a clean pipette to stir the culture to 

evenly distribute the paramecia. 

29
• Feed approximately 3,000 paramecia to each tank of larvae. Calculate the volume to feed 

a single tank by dividing 3,000 by the density of the Tupperware culture. Multiply the 

calculated volume by the number of larval tanks to get the total volume to feed the larvae. 

Pour the calculated total volume through the designated tight mesh paramecium net into a 

beaker. Measure out the volume to feed a single tank into a smaller (100 ml) beaker for 

each larvae tank. For example, if the paramecium density of the Tupperware culture is 

100 per 1 ml, 30 ml of the culture is fed to each larval tank. If there are 8 tanks to feed, 

then 8 tanks × 30 ml = 240 ml. Therefore for each feeding, 240 ml of the Tupperware 

culture should be strained through the net and 30 ml measured out to each of the 8 tanks. 

• Clean the Tupperware container with a sponge when the culture is exhausted and place it 

in the bleach bath. 

Creating a Clean Culture 

If the bottle cultures become contaminated, order new stock cultures. However, ordering 

new, clean cultures is not always reliable. Check incoming cultures for alien protozoa before 

culturing them. 

Creating a pure stock of paramecia takes considerable time, but the end result is worth it. 

In a sterile, 6–well plate filled with sterile/autoclaved system water, transfer a single paramecium 

through each well using a micropipette and a clean pipette tip between each well. At the end of 

the wash, put the paramecium into a bottle culture with boiled wheat berries. Repeat the process 

with another paramecium. Add at least two cleaned paramecia to start the parent cultures. Start 

several parent cultures at the same time as backup. Once the parent cultures are set up, wait for 

them to multiply to a higher density. 

Rotifers 

Currently rotifers are not cultured or fed to the larvae at our facility, but they may be 

easier to culture than paramecia. The upsides of using rotifers are that they are direct filter 

feeders (and do not require a food chain as do paramecia), they can grow to a greater density 

than paramecia (without the population crashing), they can be collected in a 57-μm screen (and 

rinsed, decreasing the chance of introducing other unwanted protozoans or bacteria to the fish 

larvae), they can be easily fortified with Roti-Rich (Appendix A), and the protocols for rotifers 

are very similar to those for paramecia. 

Rotifers are something to consider if or when the facility has a greater demand for larval 

food sources. 

Maintaining a Breeding Zebrafish Colony 

Maintaining the colony is mainly performed by the primary zebrafish caretaker. It 

involves determining when to raise a new generation for breeding, ensuring that the next 

generation of fish produces viable eggs for studies and future generations, and keeping a healthy, 

genetically diverse colony. 

30
When and How to Breed 

Zebrafish are able to breed when they are 3 months old (initial spawns can be as early as 

2.5 months, with the zebrafish producing small eggs). As a general guideline, they will be able 

to breed until they are 1.5 years old. (Zebrafish usually live up to 2.5 years, but their gametes 

start deteriorating after 1.5 years.) A good breeding population should be maintained so that 

there are always zebrafish within breeding age. 

Start a new generation of breeders every 3 to 4 months. Roughly 100 new fish should be 

raised each generation to maintain the breeding colony. This number can decrease or increase 

according to project needs. The breeding fish should always have a good age range to choose 

from. 

To prepare for a new generation, make room on the Z-Mod, prepare paramecium 

Tupperware cultures, and check to see whether it is a good time for labor intensive larval rearing 

(e.g., larval rearing over holidays is not good timing). When starting a new generation, choose 

parents randomly to avoid inbreeding. Genetic diversity is important in the zebrafish colony. 

Set up several spawns from different tanks. When gathering each clutch or the eggs produced by 

a spawning group, keep the eggs separate. Raise eggs from clutches that are from strong 

spawners (i.e., fish that provide eggs of good quantity and quality) to be the next generation. 

If the quality of zebrafish breeders starts to deteriorate, try pair-wise spawning (i.e., one 

male and one female). Set up many pairs and collect the clutches of eggs while keeping them 

separated from each tank. Assess the number of unfertilized eggs and developmental 

abnormalities for each clutch. Raise only the clutches that have eggs of high quality and 

quantity. 

Labeling Generations 

Labeling tanks is a somewhat arbitrary process. A good way to keep track of the 

generations is to color code tanks, so that a single generation will have the same colored label. 

Different simple names are assigned to each tank, thus a record can be kept as to which tanks 

have been spawned. More importantly, keep the date of fertilization on the label, so the age of 

the fish is readily apparent. 

Grouping Fish into Z-Mod Tanks 

At 3 months of age, the fish can be distinguished as male or female. Separate males and 

females into groups of up to 15 per 2-L tank. Keeping the sexes separate makes setting up 

spawns easier. 

To keep reproductive hormones elevated, tanks with females require the constant 

presence of at least one male in the tank. When setting up female tanks, add one or two males 

(preferably of the same clutch) to the tank. Male tanks do not require the presence of females. 

Males and females may also be grouped together. Keep in mind that small groups of six 

or less will lead to constant chasing, a high stress situation, and perhaps death (most likely for 

31
the females). If there is no choice but to have a small group in a tank, place a piece of mesh in 

the tank to act as a hiding place for the fish to find refuge. 

Factors that Affect Quality and Quantity of Eggs 

Many factors affect how well zebrafish will spawn. 

1. Age: Generally 6-month-old to 1-year-old zebrafish spawn the best. Animals older than 

1.5 years will not spawn as many eggs or they may be of poor quality (e.g., eggs may not 

be viable or the number of abnormal embryos may be high). 

2. How often the zebrafish are spawned: 

• Overspawning the fish decreases the quantity of the eggs in the clutch. Fish should 

be given at least 1 week of rest between spawning events. 

• If fish are not spawned often, eggs are reabsorbed or deteriorate in the female fish. 

Females should be spawned at least once a month to maintain egg quality. 

3. Diet: If zebrafish do not receive the right nutrients or are not fed enough, the quality and 

quantity of the eggs will be affected. 

4. Light cycle: A change in light cycle affects whether zebrafish spawn. It may take up to 1 

week for them to acclimate to a new light cycle (e.g., daylight savings time). 

5. Genetics: 

• A bottleneck in the gene pool can severely affect spawning capabilities. Outcrossing 

(crossing fish to a different genetic background) transgenic or mutant fish with the 

wild-type background (e.g., AB strain) is important to keep these lines viable. 

Bringing in a new source of wild-type zebrafish is a way to keep the gene pool larger 

(see Expanding–New Fish and Systems). 

• “Bad” genes in the gene pool can affect egg viability and egg production. This can be 

overcome by using the pair-wise spawning technique described above. 

6. Disease: Some diseases can affect spawning. Unhealthy fish are less likely to produce 

viable eggs. Fish with obvious signs of disease must be removed from the system. 

If there are periods of time where zebrafish are spawned infrequently, set up routine 

spawns to keep the female zebrafish “in shape.” Keep good records of when and which females 

and males have been spawned. Refer to the records when setting up routine spawns, making sure 

to spawn females every 4–5 weeks. 

Keeping a Healthy Colony 

Maintaining the health of the colony is the responsibility of all zebrafish users. 

Unhealthy fish should be immediately removed and euthanized to reduce the chance of spreading 

disease to healthy fish. 

Preventative measures should also be taken to keep the colony healthy. When fish are 

past 1.5 years in age, diseases and degeneration can become problematic. As fish age, their 

immune systems lose strength and diseases can appear (e.g., curled back, tumors, thin, off color, 

32
retinal degeneration, Appendix B: General Fish Aquaculture Information). To reduce the chance 

of spreading diseases from older fish, fish that are more than 1.5 years old should be removed 

from the Z-Mod. 

Euthanizing Adult Fish 

A standard method of euthanizing zebrafish as described by the Institutional Animal Care 

and Use Committee is to use an overdose of 200–300 mg/L tricaine methanesulfonate (MS-222). 

To euthanize adult fish, move them from the tank or tanks into a beaker or container that 

contains system water with the overdose concentration of MS-222. After a few minutes, the fish 

will stop respirating (i.e., opercular movement will cease). Leave the fish in the solution for 10 

min, then drain the water and store the fish in the freezer (in a bag or petri dish). They can be 

sealed in another plastic bag and placed in the refuse later. 

Record of the number of fish euthanized, the tank number, and the reason for euthanasia 

on the mortality log sheet (Appendix C: Examples of Zebrafish Records). 

33
Biosecurity 

It is important to keep the zebrafish colony safe from pathogens. There are several 

control points to consider to prevent pathogens from being introduced to the colony. 

1. People: Limit access to authorized personnel. This ensures no equipment is borrowed. 

Zebrafish users must be properly trained on the procedures (Appendix E: Training 

Checklist for New Zebrafish Users). 

2. Fish: Know the health status of the fish. Quarantine fish obtained from outside sources 

(see Expanding–New Fish and Systems), do not house other fish species or aquatic 

organisms in the same room as the zebrafish, and bring only disinfected eggs into the 

facility (even these can carry disease). 

3. Water: Know the source of water or make sure the filtration system is working properly, 

or both. 

4. Equipment: Clean, maintain, and store equipment properly. Use one net per tank, and 

ensure that nothing is shared or borrowed. 

34
Record Keeping 

Keep records of all maintenance and feedings in the zebrafish room. If it is not written 

down, assume it has not been done. Filling out records is a good way to follow a checklist of 

what needs to be done, and it allows monitoring for sudden changes (such as water quality or a 

spike in mortality). Records can help prevent and solve problems (Appendix B: General Fish 

Aquaculture Information). 

Fill out log sheets (Appendix C: Examples of Zebrafish Records) for: 

• Z-Mod zebrafish feeding 

• Z-Mod water quality 

• Spawning record 

• Larval fish feeding 

• Filter change 

• Mortality record 

• Calibrations of pH and conductivity meters 

• Chlorine and TDS test 

• System water change 

35
Zebrafish Room Power Failure Plan 

An emergency plan for power failures is an important part of zebrafish colony 

maintenance. Every zebrafish user in our facility should be aware of this plan. 

When the Power Goes Out 

There is a small flashlight located inside the zebrafish room by the door. 

Using the two extension cords located in the box outside the fish room, plug both 

extension cords into the outlets outside the south door of the freezer room. The outlets are 

orange, indicating they convey power from the generators. 

Plug the following into the extension cords: 

1. The Z-Mod is running on a power strip, located in the corner of the room to the right of 

the Z-Mod, southwest corner of the room. 

2. Additional tanks on the shelves (if applicable) run on two power strips, which are 

plugged into the outlet on the west side of the room. 

3. The larval tank heaters (if applicable) are on a power strip. 

4. If there is still room on the extension cord, plug in the refrigerator and the Artemia air 

pump. 

Perform a quick check of the system to make sure everything is running, nothing is 

leaking, etc. 

Find the ceiling light timer, which is installed on the wall next to the light switch. Peel 

off the tape to open the cover. Next, remove the “on” and “off” clamps in the metal timer box, 

that control the ceiling lights. The metal clamps can be unscrewed and taken off the metal timer 

disk (it will be obvious once the box is opened). Leave the clamps inside the box. The clamps 

must be removed so that when the electricity does come on, the lights will not turn on when they 

are not supposed to. It is also important to switch the light toggle (located near the lower right 

corner of the timer disk) to the “off” position. It is best to have the ceiling lights off when the 

electricity comes back on, that way the lights will not turn on at the wrong time if electricity 

returns during the night. 

Do not feed the fish. Get the water flowing again and cleaned up. Feeding after the 

system has been off for a period of time can add too much ammonia to the system. 

Keep the door closed as much as possible, since the extension cords will be in the way, 

and to the extent possible keep the power cord connections off the floor or at least dry. 

36
After the Power is Restored 

Perform the following steps: 

1. Plug all cords back into the wall sockets. 

2. Correct the time in the ceiling light timer. Replace the “on” and “off” clamps to the time 

disk, so that the timer turns the lights on at 800 hours and off at 2200. Toggle the on/off 

switch located to the lower right of the time disk to “on” if the current time is between 

800 and 2200 or “off” if it is between 2201 and 759 hours. 

3. Double-check that everything is working properly. 

4. Bundle up the extension cords and place them in the storage box outside the zebrafish 

room. 

37
Expanding–New Fish and Systems 

When new fish or embryos are required for an expanding zebrafish system, precautions 

must be taken to protect the health of the existing colony. New fish must be quarantined and 

only bleached embryos are allowed in the zebrafish facility. 

Bringing in New Fish from a Different System 

All nonembryo fish brought from another facility must be quarantined. No matter how 

reputable, every major zebrafish lab has or had cases of disease. It may take months for a 

disease to surface; so to prevent new pathogens from being introduced to the system, new fish 

must be quarantined. The quarantine area should be an entirely separate room away from the 

main zebrafish room (i.e., no adjoining entrances). 

To ensure the health of the zebrafish, several rules must be followed: 

1. Nothing is shared between the zebrafish room and the quarantine area. It is important 

that nothing goes back and forth between the two. The quarantine room is equipped and 

entirely independent of the zebrafish room (i.e., separate meters, nets, tanks, spawning 

tanks, etc.). Take only disposable ware (e.g., pipettes, tri-pour beakers) into the 

quarantine room. 

2. Feed the quarantined fish after the zebrafish room. It is best not to reenter the zebrafish 

room immediately after working with the quarantined fish. 

3. Wash your hands, especially after feeding or dealing with quarantined fish. 

4. Be mindful. Think about what you are doing while in the quarantine area and consider 

yourself contaminated after you have been in the quarantine area. 

When quarantined fish are spawned, the spawns must be set up in the quarantine area and 

the embryos must be bleached (as described in Westerfield 2007) before leaving the area. 

If the original purpose of obtaining adult zebrafish is to expand the colony’s gene pool or 

add special strains, then only the offspring of the quarantined adults can be introduced to the 

zebrafish room. To add progeny of quarantined adults, eggs must be bleached before being 

introduced to the zebrafish room (see Bringing in New Embryos from an Outside Source). 

Quarantined adults are never brought into the zebrafish room and must remain in quarantine for 

the remainder of their lives. 

Setting Up and Maintaining a Quarantine Tank 

Currently, two quarantine tanks are set up in the quarantine area. They are 15- and 20-gal 

tanks. Each has a three-filter system, a heater, and an air stone, which is connected to an air 

38
pump with capacity for more air stones. This is not a typical household aquarium tank; the water 

and the environment are closely monitored and controlled. 

If setting up a tank from scratch, set up the tank and allow water to go through the 

nitrogen cycle before adding the quarantine fish. For the quarantine tank, make system water 

using Instant Ocean salt and distilled water. Distilled water is used instead of the RO/DI water 

from the zebrafish room. This reduces chances of cross-contamination by eliminating a shared 

water source. 

To encourage and speed up establishment of the nitrogen cycle (Figure 3 and Appendix 

B), seed the tank’s sponge filter by adding water and bacteria from the Z-Mod’s sponge filter. 

Next, add seed fish (fish that are no longer needed; e.g., too old to spawn) from the Z-Mod 

system (they will be euthanized once the quarantine fish arrive). Initially after the tank is set up 

and the seed fish are added, ammonia levels will be high. Do not be alarmed; it takes time for 

the bacteria to grow and start the nitrogen cycle. Slowly, ammonia levels should go down once 

the bacteria are growing. Ideally, set up the tank 4–6 weeks before the quarantine fish arrive. 

This should be enough time for the sponge filter to build a good population of bacteria, which 

will control ammonia and nitrite levels. 

When the tank is initially being set up, check the pH, temperature, conductivity, 

ammonia, and nitrite levels daily. Add water amendments (e.g., sodium bicarbonate) and allow 

them to mix for at least 24 h before adding more. The goal is to get the bacteria growing and the 

water quality stabilized before adding the quarantine fish. 

If a tank must be set up quickly, use an ammonia removing filter medium (e.g., zeolite). 

The drawbacks are that the water will have to be changed frequently in large amounts and the 

biofilter will not have a chance to grow bacteria. Maintenance tasks include: 

1. Feeding (daily): First, feed the Z-mod, reserving some Artemia in a disposable pipette for 

the quarantine fish, then feed the quarantine tanks. A small container of dry salmon 

crumble is kept on the countertop next to the quarantine tanks. Dispose of the pipette that 

was used to feed Artemia in the quarantine room. 

2. Water measurements (daily): As with the Z-Mod, water parameters of all tanks must be 

checked and recorded. Check the conductivity and Ammonia Alert disc once a day. 

Check the pH and room and water temperatures twice a day while feeding. 

3. Nitrite check (weekly): Use the kit to check the nitrite levels of all tanks. 

4. Water changes (when needed or weekly): To remove water from the quarantine tank, 

siphon off debris and water, taking care not to siphon up fish. Remove 5–10% of the 

tank water. To reduce cross-contamination, dissolve Instant Ocean salt in distilled water 

(instead of using the RO/DI water system in the zebrafish room) to the appropriate 

conductivity in the quarantine system water bucket. Add water amendments if needed. 

Add the water to the tank. 

5. Water amendments (when needed): If the pH is low (<7.0), add about 5 ml of 1 molar 

sodium bicarbonate solution to a 25-gal tank. Add these amendments slowly to the filter 

so it can mix thoroughly. Ideally, the tank will be stabilized before quarantined fish are 

39
added. Amendments can also be mixed in the quarantine system water bucket when 

doing water changes—this is a better and subtler change for the fish. 

6. Filter changes (monthly or more depending on the number of fish): The sediment/carbon 

filter should be changed monthly. Do not rinse the sponge filter at the same time the 

sediment/carbon filter is changed. The sponge filter may be lightly rinsed if bacterial 

buildup decreases water flow. 

Bringing in New Embryos from an Outside Source 

To bring in new embryos, the eggs must be bleached before taken into the zebrafish 

room. Bleaching protocol is found in Westerfield (2007). 

Embryos should be at least 6 h postfertilization (hpf) and no more than 24 hpf (hatched 

embryos cannot be bleached) for bleaching. Bleaching embryos is a delicate operation and 

should be done very carefully. Overbleaching can kill them. 

Make a bleach solution in a 250-ml beaker by adding 100 μl of 5.25% sodium 

hypochlorite (bleach) to 170 ml of system water. Use a pipette to transfer up to 30 embryos to 

the bleach solution. Keep the embryos in the solution for 2 min while occasionally swirling the 

beaker to allow the entire surface area of the chorions to come in contact with the bleach 

solution. With a clean pipette, transfer the embryos from the bleach solution to a 250-ml beaker 

of clean system water. Swirl the embryos in the system water. Repeat the wash in system water 

one more time, then transfer the embryos back into petri dishes with fresh system water. 

When the bleached embryos are 24–48 hpf, they should be manually dechorionated as 

bleaching will cause the chorions to become tougher. Keep in mind that even bleaching does not 

guarantee that no diseases will be transferred (Appendix D: Zebrafish Diseases). 

Preventing the Spread of Pathogens 

Adding a new aquaculture system is an opportunity to make a fresh start at establishing a 

colony with minimal amounts of pathogens. Since a new system will share the same space as the 

old one, it is highly unlikely that the new system will be kept entirely pathogen free. However, it 

is ideal to reduce the transfer of pathogens between systems. In the unlikely event of a disease 

outbreak, at least both systems will not succumb to a massive die-off or other deleterious effect. 

Several things should be considered to minimize the spread of pathogens to the new 

system and to monitor the health of a larger colony. Typically, a new aquaculture system is 

seeded by transferring several tanks of healthy young fish from the old system to the new 

system. The introduction of fish to the new system will produce ammonia, which will start the 

nitrogen cycle (Figure 3) and grow the ammonia- and nitrite-metabolizing bacteria in the sponge 

biofilters. However, adding fish to the new system will carry over any pathogens from the old 

system. To eliminate transfer of pathogens to the new system, seed a system without using fish 

by adding large amounts of fish food to the new system daily. This should build up ammonia 

and start the nitrogen cycle. However, it will also take longer to achieve a good population of 

ammonia- and nitrite-metabolizing bacteria in the sponge biofilters without fish. 

40
Once the biofilters are established, pathogen-free fish should be introduced to the new 

system. They can be grown from pathogen-free eggs. Such eggs are difficult to find, especially 

since not all pathogens are killed by bleach (Appendix D: Zebrafish Diseases). The Zebrafish 

International Research Center (ZIRC) in Oregon is working to achieve this. If pathogen-free 

eggs are not available, the next best step is starting with bleached eggs (see previous subsection, 

Bringing in New Embryos from an Outside Source). 

To keep the new system free of pathogens, all zebrafish users should be familiar with 

ways to minimize potential pathways for pathogens to travel between the old and new systems. 

Sharing nonsterilized equipment and moving fish between the two systems is not allowed; these 

practices would guarantee the spread of pathogens. Only clean, sterilized equipment (e.g., nets, 

spawning tanks) can be used for both systems. Separate equipment that cannot be easily 

sterilized, such as meters and siphons. 

Unlike a smaller colony, where more time can be devoted to monitoring the health of 

each tank, tracking the health of individual tanks in a larger zebrafish colony is impractical. To 

monitor the general health of zebrafish in a large colony, sentinel fish should be used in each 

system. Sentinels are fish whose sole purpose is to be sacrificed at a later date and analyzed for 

pathogens. There are two groups of sentinels: one group should be placed at the influent of clean 

water (after filtration and UV sterilization) and the other should be placed at the effluent of the 

waste water (before filtration and UV sterilization). The groups should each consist of 

approximately 10 young, pathogen–free fish (grown from the pathogen-free eggs). Every 3 

months, these fish should be sent away for out-of-house pathology tests. 

41
Conclusion 

The zebrafish husbandry methods presented here are continuously evolving to meet new 

aquaculture technology, zebrafish capacity, spawning and breeding needs, maximum colony 

health, and cost-effectiveness. The success of our healthy colony relies on consistent care and a 

solid knowledge of fish health and aquaculture. At the NWFSC, our research using zebrafish 

embryos, larvae, and adults requires that no factors such as disease or pollutants confound our 

results. Thus we use the techniques presented here to minimize these concerns. This technical 

memorandum provides detailed documentation of our protocols for small-scale zebrafish 

husbandry. It may also be a useful reference for other facilities that have their own small-scale 

systems. 

42
References 

Kent, M. L., J. M. Spitsbergen, J. M. Matthews, J. W. Fournie, and M. Westerfield. 2007. Diseases of 

zebrafish in research facilities. Zebrafish International Resource Center (ZIRC) Health Services 

Zebrafish Disease Manual. Online at http://zebrafish.org/zirc/health/diseaseManual.php 

[accessed 10 April 2009]. 

Kimmel, C. B., W. W. Ballard, S. R. Kimmel, B. Ullmann, and T. F. Schilling. 1995. Stages of 

embryonic development of the zebrafish. Dev. Dyn. 203:253–310. 

Westerfield, M. 2007. The zebrafish book: A guide for the laboratory use of zebrafish (Danio rerio). 5th 

edition. University of Oregon Press, Eugene. 

43
44
Appendix A: Ordering Information 

Fish Food 

Brine Shrimp Direct, Ogden, Utah: Artemia nauplii premium grade, Spirulina. Online at 

www.brineshrimpdirect.com [accessed 17 April 2009]; phone (800) 303-7914. 

Bio-Oregon, Longview, Washington: salmon starter–Bio Vita #2. Online at 

http://www.bio-oregon.com/ [accessed 17 April 2009]; phone (800) 962-2001. 

Carolina Biological Supply Company, Burlington, North Carolina: paramecia 

(Paramecium multimicronucleatum), protozoan pellets. Online at: www.carolina.com [accessed 

17 April 2009]; phone (800) 334-5551. 

General Fish and Aquarium Supplies 

Drs Foster & Smith, Rhinelander, Wisconsin: conductivity meter, solutions, Instant 

Ocean, general aquarium and fish supplies. Online at www.DrsFosterSmith.com [accessed 17 

April 2009]; phone (800) 381-7179. 

That Pet Place, Lancaster, Pennsylvania: general fish supplies, test kits, Ocean Clear 

filters, Micro-Food, Cyclop-eeze. Online at www.thatpetplace.com [accessed 17 April 2009]; 

phone (888) 842-8738. 

Aquatic Eco-Systems Inc., Apopka, Florida: general fish supplies, Artemia cones, 

Artificial Plankton, Larval Diet AP100, Reef Carbon, Roti-Rich. Online at www.aquaticeco.com

[accessed 17 April 2009]; phone (877) 347-4788. 

PetStore.com, Garden Grove, California: Ammo-Lock, Net Soak. Online at 

http://www.petstore.com/ps_ViewItem-idproduct-AP8173.html [accessed 18 May 2009]; phone 

(888) 993-2378. 

Zebrafish (Danio rerio) Hardware 

AquaFX, Winter Park, Florida: RO/DI system and filters. Online at 

www.aquariumwaterfilters.com [accessed 17 April 2009]; phone (877) 256-3467. 

Emperor Aquatics Inc., Pottstown, Pennsylvania: UV light bulb and UV sterilizer parts. 

Online at http://www.emperoraquatics.com [accessed 17 April 2009]; phone (610) 970-0440. 

45
AZPonds and Supplies Inc., Birdsboro, Pennsylvania: alternative source for UV light 

bulb and parts. Online at http://www.azponds.com/uv_parts.htm [accessed 17 April 2009]; 

phone (800) 722-8877. 

Marine Biotech, Beverly, Massachusetts (recently acquired by Aquatic Habitats): Z-Mod 

system and tanks. Online at http://www.marinebiotech.com/ [accessed 17 April 2009]; phone 

(888) 624-8265. 

Thoren Aquatics Systems Inc., Hazelton, Pennsylvania: spawning cages and lids. Online 

at http://thorenaquatics.com/index.htm [accessed 17 April 2009]; phone (570) 454-3517. 

Cambro Manufacturing Company, Huntington Beach, California: larval tanks. Online at 

www.cambro.com/ [accessed 17 April 2009]; phone (800) 833-3003. 

46
Appendix B: General Fish Aquaculture 

Information 

Below is background material on fish aquaculture to help understand why the Northwest 

Fisheries Science Center zebrafish (Danio rerio) facility uses the protocols described in this tech 

memo. 

The basics of fish physiology: 

1. Cold-blooded: Temperature affects the fish’s immune function and metabolic rates (e.g., 

high temperature increases the demand for oxygen, food, and growth). 

• High temperatures may cause stress and death. 

• Low temperatures may cause slower growth and sometimes death. 

2. Osmotic regulation: With freshwater fish, water moves into the fish. The kidneys excrete 

high volumes of dilute urine and the skin is a barrier to the uptake of water. With 

pathogens or rough handling, the kidneys must work harder (sometimes leading to kidney 

failure). 

3. Gills: The essential functions of the gills are respiration, excretion of nitrogenous waste 

(ammonia), and ion exchange. They are especially vulnerable to various forms of 

damage such as from bacterial or fungal infections. 

Important water considerations: 

1. Dissolved oxygen (DO): As water temperature increases, DO decreases. 

Temperature (°C) O2 at 100% saturation (mg/L) 

5 12.76 

10 11.28 

15 10.07 

20 9.08 

25 8.24 

30 7.54 

As temperature increases, the oxygen requirement (metabolism) of the fish increases. 

The movement of water (e.g., via air stones) will keep oxygen in the water. The factors 

that affect DO are the air-water interface, utilizers of oxygen (fish), and producers of 

oxygen (mechanical or photosynthetic). 

2. Temperature: The optimum temperature for zebrafish is approximately 26°C. As 

mentioned previously, temperature can affect DO, and sudden changes in temperature 

and DO can be a stressor. 

47
3. Source of the water: It is necessary to filter incoming water for particles and 

contaminants (such as chlorine). 

4. Hardness and alkalinity: Hardness is the amount of cations in the water and can have 

interactions with other water parameters or disease treatments. Alkalinity is the buffering 

capacity of the water (resists changes in pH) and is related to the amount of bicarbonate 

in the water. 

5. The nitrogen cycle (Figure 3): Ammonia and nitrite are toxic to the fish; nitrate is toxic at 

very high concentrations (which is why water changes are important). Nitrosomonas and 

nitrobacter—bacteria that convert ammonia and nitrite—remove the buffering capacity 

(alkalinity) so the pH will naturally go down. Under low pH conditions, ammonium 

(NH4+) will accumulate, as the nitrosomanas cannot convert this ion. An increase in pH 

will cause nontoxic ammonium to convert to toxic ammonia. The nitrogen cycle 

(nitrosomonas and nitrobacter) takes time to establish (4–6 weeks). The reaction is 

aerobic, so the biological filter must not become anaerobic. Also the bigger the surface 

area of the filter, the higher the fish capacity the system can support. 

6. Filtration: This is required for closed systems. There are three parts to filtration. 

• Biological filters grow bacteria for the nitrogen cycle. Sudden changes in water 

quality, therapeutics, and antibiotics can destroy the biofilter. Examples of 

biofilters are foam, floss, and gravel (they provide surface area for bacteria). 

• Mechanical filtration removes particulate matter. Examples are foam, filter floss, 

and gravel (1/4–1/8 inch diameter). 

• Chemical filters remove small molecular weight compounds. Examples are 

activated carbon, dolomite, zeolite, ion exchangers, and peat moss (but it also 

lowers pH). 

7. Water sterilization: It is important to remove pathogens from the water source in flow 

through systems or in closed recirculating systems. Water should be UV or ozone 

sterilized. 

8. Water changes: They are helpful in diluting waste products (e.g., nitrate) and correcting 

the pH level. 

9. Carrying capacity of the system: The number of fish sustainable in a closed system is 

determined by the amount and capabilities of filtration. 

10. Chlorine (and chloramines): It is toxic to fish. Chlorine can be removed by sodium 

thiosulfate, allowed to dissipate with time, aeration or agitation (to speed release), or with 

an activated carbon filter. 

11. Other toxicants: Toxicants such as pesticides, fertilizers, metals, gases, and smoke affect 

the health of the fish. 

12. Change is not good: Sudden changes in water quality stress the fish and can have 

detrimental effects on biological filters. 

48
49

Appendix C: Examples of Zebrafish Records 

What follows are examples of forms used in record keeping for the Northwest Fisheries 

Science Center Zebrafish (Danio rerio) colony. They are: 

• Z-Mod Zebrafish Feeding 

• Z-Mod Water Quality 

• Spawning Record 

• Larval Fish Feeding 

• Filter Change 

• Mortality Record 

• Calibration of pH and Conductivity Meters 

• Chlorine and total dissolved solids (TDS) Tests 

• System Water Change 
Z-Mod Zebrafish Feeding

Month/year: ________________________ 

AM PM

Date SS A Time Initials SS

A Time Initials

Notes:

1 2 3 4 5 6 7 8 9 

10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 

50
Z-Mod Water Quality

Month/year: _____________________ 

AM PM 

Date Room temp H2O temp pH Room temp H2O temp pH Conduct. NH3 NO2

RO/DI H2O 

added Comments: 

1 2 3 4 5 6 7 8 9 

10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 

51
52

Spawning Record

Date Males Females 

spawned # Tank name # Tank name 

Spawning 

time 

(approx.) Study 

No. of eggs 

(approx.) Initials Notes: 

(Form extends to fill one page.) 

Larval Fish Feeding Tank # :_____________________________ 

Fill out one sheet per zfish strain/stock Parent#: _____________________________ 

Date Fertilized: _______________________ 

AM Noon PM 

Date Dry Para. A Time Initial Dry A Time Initial Dry Para. A Time Initial Water change Notes: 

(Form extends to fill one page.) 
Filter Change

Once a month or depending on density of fish in system and psi

Follow instructions in zebrafish maintenance guide. 

Date Initials Pressure Filter # Notes: 

(Form extends to fill one page.) 

Mortality Record

Date Tank # Dead Notes: 

(Form extends to fill one page.) 

Calibration of pH and Conductivity Meters

Once a month

Follow instruction manuals for each meter. 

Date Initials pH Conductivity Notes: 

(Form extends to fill one page.) 

53
Chlorine and TDS Tests

Once a month

Follow instruction manuals for each test kit. Test chlorine levels in Z-Mod. Test TDS of the RO/DI water. 

Date Initials Chlorine TDS Notes: 

(Form extends to fill one page.) 

System Water Change

To be changed every 1–2 weeks

Turn off pump. Empty out sump to above the drain. Fill sump with system water from the 50-gallon 

tank. Check ammonia levels and conductivity after 3 hours. 

Date Initials Notes: 

(Form extends to fill one page.) 

54
Appendix D: Zebrafish Diseases 

Zebrafish (Danio rerio) are susceptible to many diseases. Transmission among fish can 

happen quickly, especially when they are in an unnatural, stressful, and crowded environment. 

Understanding types of diseases, how they are transmitted, and steps to prevent outbreaks can 

keep the fish healthy. This is a brief introduction to zebrafish disease, diagnosis, and prevention. 

For a more extensive review, see the Zebrafish International Research Center (ZIRC) Health 

Services Zebrafish Disease Manual (Kent et al. 2007). 

Common Diseases of Zebrafish 

The majority of bacteria, fungi, and parasites that cause disease in aquaculture and in 

nature are opportunistic pathogens that are ubiquitous in aquatic environments. Stress from 

transport, endocrine changes (e.g., spawning, aging), handling, crowding, and dramatic 

fluctuations in water quality may trigger disease outbreaks. 

Bacterial Disease, Mycobacteriosis 

Forty percent of zebrafish facilities are infected (ZIRC). 

It causes serious diseases, tuberculosis and leprosy. 

It is widespread and ubiquitous in aquatic ecosystems. The bacteria colonize 

asymptomatically, with chronic and acute infections. 

Conditions that encourage mycobacteria growth are low DO, low pH, warm water, high 

organic load, and clean water (less competition). 

It survives in surface biofilms (tank surface), which harbor and protect mycobacteria. 

Signs of mycobacteriosis: 

• External signs: swollen belly, skin ulcers, exopthalmia (pop eye), and general malaise. 

• Internal signs (target tissues): granulomas throughout viscera, swollen spleen (sometimes 

gone), kidney, and liver. 

Other target tissues affected less often are gills, skin (unless visible lesions), muscle, 

nervous tissues, and the gut (contains other bacteria). 

Infections are notoriously difficult to treat because mycobacteria are naturally resistant to 

antibiotics. 

55
Protozoan Parasite, Microsporidia (Pseudoloma neurophilia) 

Fifty-nine percent of facilities are infected (ZIRC). 

Signs of microsporidia: 

• In adults and juveniles, chronic infections cause emaciation and skeletal deformities 

(skinny fish disease), reduced growth, and fecundity problems. 

• In larval fish, acute infections cause mortality. 

Primary sites of infection are the spinal cord and the hindbrain. 

Its life cycle takes place intercellularly and produces infectious spores. 

Infectious spores within the eggs may not be killed by chlorine treatments. 

It can be treated with fumagillin, an antibiotic taken orally. 

Noninfectious Diseases 

Nutritional imbalances: 

• Vitamin E/selenium deficiency causes degenerative muscle disease. 

• Vitamin C deficiency (also called broken back disease) causes scoliosis (misshapen 

vertebrae at major pressure points on spinal column). Vitamin C participates in 

conversion of cartilage to bone. 

Methemoglobinemia (nitrite toxicity) or “brown blood disease” is caused by excess 

nitrite binding to hemoglobin (inhibiting the ability of blood to carry oxygen). This is a reason 

why water quality must be monitored. To treat the toxicity, add NaCl (the chloride will reverse 

the reaction), stop feeding, and determine why nitrite levels are high. 

Ammonia toxicity or “callus and blister” causes a proliferation of cells and an 

accumulation of fluid in the gill. At low concentrations, it causes a diffuse gill epithelial tissue 

proliferation. At high concentrations, a typical response to the toxin in the water is lifting of the 

gill epithelium from the supporting structure. 

Nephrocalcinosis is the formation of calcium in kidneys, which indicates too much 

calcium in the water. Mineralization on kidney tubules is associated with high concentrations of 

dissolved carbon dioxide in water or excessive calcium or magnesium in the diet. It is important 

to use sodium bicarbonate—not calcium carbonate—when buffering fish water. 

Gill hyperplasia is associated with poor water quality such as active chlorine, excess 

metals such as copper and zinc, or excess ammonia particulates. 

Gas bubble disease is caused by supersaturated concentrations of gases, photosynthesis, 

imperfections in plumbing fixtures, or rapid heating of water. 

56
Egg associated inflammation and fibroplasia (EAIF) occurs when females retain eggs, 

which degenerate and eventually push out the abdominal wall. EAIF is very common and often 

associated with secondary bacterial infections, including mycobacteriosis. The exact cause is 

unknown. 

Routes of Disease Transmission 

Direct transmission: Pathogens can pass between live fish. This is why fish coming from 

other facilities must be quarantined and why equipment must not be shared between fish 

facilities or individual tanks. Direct transmission also occurs when fish eat the infected tissues of 

other fish; therefore, it is important to remove sick and dead fish. 

Vertical transmission: Pathogens can be passed from the mother into the egg. Since the 

pathogen is in the eggs, bleaching cannot kill all pathogens. 

Biofilm transmission: Biofilm is the layer of material that builds up on the inside of 

tanks. Pathogens can grow and multiply in biofilm. 

Aerosol transmission: Some pathogens are small enough to drift in the air, which is why 

it is important to put lids on the tanks. 

Diagnostic Techniques 

It is imperative to diagnose disease outbreaks as soon as possible, so a proper treatment 

can be quickly implemented. Diagnoses can be done in-house or be sent to ZIRC 

(http://zebrafish.org). Samples can be shipped to ZIRC and technicians can perform the 

techniques described below. They are experienced and reliable at identifying pathogens. 

Wet mounts are easy and can be done in-house. The best results are found on freshly 

dead fish. To take wet mounts, place the tissue sample in a drop of water on a microscope slide 

and cover slip. Examine at all powers of magnification, looking for external parasites (e.g., 

protozoans and monogeneans), bacteria (e.g., Flavobacterium, a long flexing rod-shaped 

bacterium), and fungi (e.g., Saprolegnia). Take wet mounts of the external and internal parts of 

the fish. 

• External exam: Look for behavioral signs, body condition, swollen abdomen, and 

abnormal growths (e.g., tumors). Photograph any gross abnormalities. Tissues to sample 

and wet mount: 

o Gills, examine the individual filaments. 

o Fins, examine the end of the pectoral fin. 

o Skin scraping, use a cover slip to scrape behind the pectoral fin and any external 

lesions (on the skin, fin, or gills). 

• Internal exam: Disinfect the external surface (with ethanol). To avoid contamination, 

aseptically open the abdomen with sterile instruments and avoid cutting the digestive 

tract (do not release the gut flora). Do a gross examination of organs. Tissues to sample 

and examine microscopically: 

57
o Lower intestine, chop up and wet mount on to a slide. 

o Brain, smear and wet mount on to a slide. 

o Contents of any cysts (helminthes and sporozoa). 

Bacterial culture should be done with the bacteria from the posterior kidney (bacteria are 

more likely to be here) and any visible lesions on the organs. Using aseptic techniques, collect 

kidney tissue from a small fish by entering the fish from the dorsal side downward to the kidney. 

Culture the bacteria on the appropriate agar. 

Viral culture can be done by collecting selected internal organs (usually kidney, spleen, 

and liver) and inoculating the appropriate fish cell culture. Use aseptic techniques. 

Histopathology can be performed by removing pieces of organs (tissue size no greater 

than 1 cm for rapid fixation) and placing them in a fixative such as Dietrich’s fixative (use a 

fixative volume of 10:1 fixative:tissue). Dietrich’s fixative is used because it softens scales, 

decalcifies bone, and provides good nuclear staining. Storage in plastic jars is also suggested, 

especially if the samples will be shipped. 

Treatments and Controls 

Below is a brief overview of the treatments commonly used to combat diseases in 

aquaculture. However, make sure to obtain an accurate diagnosis before proceeding with a 

treatment, because an incorrect treatment could harm the fish. 

External treatments are generally needed for external pathogens. The treatment 

compound is added to the water. 

• Types of treatment and duration of dose: 

o Dip is for a short time at a relatively high concentration. 

o Bath is a longer time at a lower concentration. 

o Prolonged treatment is over a long duration (the compound decomposes with time). 

• Formalin treatments must have adequate aeration and proper storage (above 40°F or else 

it turns into paraformaldehyde). Formalin is caustic. 

• Copper sulfate is an algaecide that is effective against some external parasites. Toxicity 

increases with low alkalinity (toxic at alkalinity < 50 mg/L). 

• Potassium permanganate is a strong oxidizing agent that is effective against some fungal 

and bacterial diseases. The dose is related to the amount of organic matter in the water. 

• Sodium chloride causes osmotic shock on external parasites. The salt is also a general 

stress alleviator for freshwater fish (the elevation of salinity reduces osmotic stress on 

fish). 

Internal treatments are generally needed for internal pathogens. The treatment compound 

(typically antibiotics) must get inside the fish (e.g., via the feed). To treat with antibiotics, there 

58
must be a diagnosis that includes the resistance pattern of the bacterial pathogen. Since the most 

common dosing method is via the feed, the treatment must start in timely manner (e.g., fish must 

be feeding). Feed for the entire duration of the treatment. 

Eradication is the depopulation of the facility and chlorination of the system (a fresh 

start). Exhaust all other possibilities before eradicating the system. This treatment is the last 

resort when a disease outbreak cannot be controlled. Eradicating the system means destroying 

all fish and sterilizing the system to wipe out the disease. This will also destroy the biofilters, 

meaning the entire system will have to be reseeded with new fish or fish grown from embryos to 

repopulate the system. 

Prevention 

Remember that pathogens are ubiquitous in aquaculture and fish can live with a level of 

pathogens. Keeping fish healthy and eliminating the introduction of new pathogens are the best 

ways to prevent disease outbreaks. Use the following steps to prevent an outbreak: 

• Quarantine incoming fish. 

• Maintain equipment (e.g., UV system) and make sure all equipment is working properly. 

• Remove sick and dead fish immediately. 

• Keep good records of mortality and water quality (to look for patterns or sudden 

changes). 

• Properly clean, disinfect, and store nets and equipment (to prevent tank-to-tank spread). 

• Do not share equipment between aquaculture systems or within the aquaculture system 

(e.g., do not use the same net between tanks without sterilizing it). 

Zoonosis 

Human infections are rare (usually only in people who are immune compromised). Some 

strains of mycobacteria can cause “fish handler’s disease.” Hand washing and good hygiene are 

the best prevention techniques. 

59
60
Appendix E: Training Checklist for New 

Zebrafish Users 

Use this checklist as a guide when introducing new zebrafish (Danio rerio) users to the zebrafish 

room. 

F General room set up 

–RO/DI versus system water 

–How the Z-Mod works: plumbing, filters, sump, etc. 

–Location of nets, tanks, logbook, food, etc. 

–Location of the procedure book 

–Location of emergency plan 

F Biosecurity/health management 

–Hand washing 

–Quarantine rules 

–No sharing equipment 

–No outside equipment 

–Bleach/sterilize everything that comes in contact with fish between uses 

–No chemicals/toxins in the fish room 

F Making system water 

F Spawning procedures 

–Records: location of spawn logbook and log sheet (on wall), refer to it before setting up a 

spawn (do not use fish that have been spawned within 1 week), good for future reference 

–Spawning cycle: set up spawns in evening, spawn occurs about 3 h after lights come on in 

the morning 

–Light cycle is important: on from 800 hours to 2200, changes in cycle will affect spawning 

–Tanks and lids 

–Organization of zebrafish: male and female tanks, female tanks have one male to keep 

females fertile, mutants and transgenic strains are mixed sexes 

–How to sex a fish 

–Nets: one net per tank, rinse net after use, Net Soak solution 

–Use black mesh in tanks with less than six fish; this prevents fighting among fish 

F Collecting eggs 

–Put fish back into correct tanks, then back on the Z-Mod 

–Use strainer, rinse eggs, wash into a petri dish 

–Sort eggs 

61
62

F How to recognize fertilized versus unfertilized eggs 

F Feeding the fish 

–Typical weekday feeding 

–two times/day (morning and afternoon) 

–salmon starter and Artemia

–Typical weekend feeding: 

 –feeding 

 –water quality 

 –start new Artemia

 –fill sump with RO/DI water 

 –larvae feeding if applicable 

–Harvesting Artemia: two-cone system (0–24 h and 24–48 h), allow shrimp to completely 

settle (≈5 min), use spigot to drain 24–48-h Artemia into net, rinse off salty water, dilute 

Artemia into bowl, drain 0–24-h Artemia into 2-day-old cone via spigot 

–Setting up new Artemia: scrub cones with a sponge between uses, Artemia water, placement 

of aeration hose 

F Larvae care and feeding 

–Larvae set up–water bath, temp at ≈28°C 

–Water changes every morning before feeding 

–Feeding schedule: dry food, paramecia, 24-h Artemia (at 7 dpf+) 

–Larvae Artemia cone 

–Larval feeding log sheet 
Recent NOAA Technical Memorandums

published by the

Northwest Fisheries Science Center

NOAA Technical Memorandum NMFS-NWFSC99 Rawson, K., N.J. Sands, K.P. Currens, W.H. Graeber, M.H. Ruckelshaus, R.R. Fuerstenberg, and J.B. 

Scott. 2009. Viability criteria for the Lake Ozette sockeye salmon evolutionarily significant unit. U.S. Dept. 

Commer., NOAA Tech. Memo. NMFS-NWFSC-99, 38 p. NTIS number pending. 

98 Johnson, L.L., G.M. Ylitalo, M.S. Myers, B.F. Anulacion, J. Buzitis, W.L. Reichert, and T.K. Collier. 

2009. Polycyclic aromatic hydrocarbons and fish health indicators in the marine ecosystem in Kitimat, British 

Columbia. U.S. Dept. Commer., NOAA Tech. Memo. NMFS-NWFSC-98, 123 p. NTIS number pending. 

97 Roegner, G.C., H.L. Diefenderfer, A.B. Borde, R.M. Thom, E.M. Dawley, A.H. Whiting, S.A. 

 Zimmerman, and G.E. Johnson. 2009. Protocols for monitoring habitat restoration projects in the lower

 Columbia River and estuary. U.S. Dept. Commer., NOAA Tech. Memo. NMFS-NWFSC-97, 63 p. NTIS

 number pending. 

96 Currens, K.P., R.R. Fuerstenberg, W.H. Graeber, K. Rawson, M.H. Ruckelshaus, N.J. Sands, and J.B. 

Scott. 2009. Identification of an independent population of sockeye salmon in Lake Ozette, Washington. 

U.S. Dept. Commer., NOAA Tech. Memo. NMFS-NWFSC-96, 18 p. NTIS number PB2009-108060. 

95 Harms, J.H., J.A. Benante, and R.M. Barnhart. 2008. The 2004–2007 hook and line survey of shelf 

rockfish in the Southern California Bight: Estimates of distribution, abundance, and length composition. 

U.S. Dept. Commer., NOAA Tech. Memo. NMFS-NWFSC-95, 110 p. NTIS number PB2009-106740.

94 Fleischer,G.W., K.D. Cooke, P.H. Ressler, R.E. Thomas, S.K. de Blois, and L.C. Hufnagle. 2008. The 

2005 integrated acoustic and trawl survey of Pacific hake, Merluccius productus, in U.S. and Canadian 

waters off the Pacific coast. U.S. Dept. Commer., NOAA Tech. Memo. NMFS-NWFSC-94, 41 p. NTIS 

number PB2009-104774.

93 Keller, A.A., B.H. Horness, E.L. Fruh, V.H. Simon, V.J. Tuttle, K.L. Bosley, J.C. Buchanan, D.J.

 Kamikawa, and J.R. Wallace. 2008. The 2005 U.S. West Coast bottom trawl survey of groundfish resources off Washington, Oregon, and California: Estimates of distribution, abundance, and length composition. U.S. Dept. Commer., NOAA Tech. Memo. NMFS-NWFSC-93, 136 p. NTIS number PB2008-113766.

92 Levin, P.S., M.J. Fogarty, G.C. Matlock, and M. Ernst. 2008. Integrated ecosystem assessments. U.S. 

Dept. Commer., NOAA Tech. Memo. NMFS-NWFSC-92, 20 p. NTIS number PB2008-113765.

Most NOAA Technical Memorandums NMFS-NWFSC are available online at the

Northwest Fisheries Science Center web site (http://www.nwfsc.noaa.gov).