Danio rerio, commonly known as zebrafish, is an incredibly popular model organism in life science and pharmaceutical research due to their fast life cycle, high reproduction rates and transparent embryos that develop outside the mother. Additionally, zebrafish and humans share about 70% of their genes, including about 84% of genes associated with human diseases, and researchers have access to a wealth of published genomic information to reference for this species.
Many zebrafish studies begin with the spawning of a new generation of embryos that can then be manipulated through microinjection, or exposure to drugs and other chemicals, and imaged as they develop. Spawning starts with the pairing of a male and female zebrafish, which can be distinguished by their body shape and coloration: males are more slender, may have darker stripes and have a yellowish tinge to their anal fin, while females with eggs will have larger bellies and a duller silvery or blueish color. Once you can identify and pair male and female individuals, you’re off to a good start; here are some additional tips to keep in mind while conducting your zebrafish embryo studies.
1. Create the Best Conditions for Zebrafish Breeding Success
Light Cycle
Just putting a male and female zebrafish together in the same tank does not guarantee a fresh batch of fertilized eggs will soon appear. Laboratory zebrafish have a lot of instincts in common with counterparts in the wild and will prefer or require certain conditions be met before they are ready to spawn. In nature, zebrafish breed at dawn, so breeding should be set up to occur at the beginning of their light period. Moving your males and females into the divided sections of the breeding tank about an hour or two before the end of the previous light period and simply removing the divider when the lights come back on the next morning is a good way to time this.
Tilting Method
One popular method for stimulating breeding in zebrafish is to tilt the breeding tank in order to create a “shallow end” and “deep end,” with the raised (shallower) end simulating the shore, where wild fish like to spawn. Studies have shown that when this type of depth gradient is available, fish will deposit more embryos in the shallow area and fewer or none in the deeper area.1 Without this gradient, the fish will lay more fertilized eggs around each end of the tank and less in the center, but produce much fewer embryos overall.
Protect Eggs from Hungry Parents
It’s important to use a breeding tank with a mesh or slits at the bottom of the inner section for fertilized eggs to fall through. If eggs are allowed to settle at the bottom of the tank without any separation from their parents, the adult fish will happily eat all of the protein-rich embryos. If you are not using a tank with this built-in separation, a layer of marbles at the bottom of the tank can protect the embryos. The tiny fertilized eggs will fall between the marbles, making it harder for the adults to get to them.
Still Not Breeding?
Even if you have happy and healthy male and female fish together in a properly configured breeding tank, sometimes the fish will still not spawn. This can be due to a variety of causes, but there are a few other disturbances and sources of stress you can rule out. Zebrafish are very sensitive to vibrations, so excessive vibrations could be making your fish too stressed out to breed.2 Water quality in the breeding tank can also be a significant factor in pairs’ ability to breed; too much waste build-up in the tank overnight can cause fish not to spawn in the morning.
If there are no apparent disturbances that would be troubling your breeding pair, there are a few more things you can do to give them some extra encouragement. Fake vegetation is another thing you could add to the breeding tank to simulate their natural environment, and marbles can mimic the substrate females like to lay their eggs over in addition to protecting the embryos from cannibalism.
2. Get Your Embryos Off to a Good Start
Once your zebrafish pairs have mated and deposited fertilized eggs to the bottom of the breeding tank, it’s time to move the adults back to their usual tanks and collect the fast-developing embryos. You will want to thoroughly rinse your embryos with egg water or embryo medium to remove any debris. Keeping young embryos in egg water or embryo medium (E2 or E3) maintains a suitable pH level and other chemical conditions for this early stage of life. Recipes for these solutions are available from many sources, such as the Zebrafish Information Network.3 The recommended temperature for incubating embryos is about 28.5°C.
Prevent Mold and Infection
Methylene blue is a useful addition to embryo media as it helps prevent bacterial and fungal growth. It is also important to sort your embryos shortly after collection in order to remove unfertilized eggs and unhealthy or dead embryos. Healthy fertilized eggs will appear transparent and shiny - eggs that are cloudy, opaque or dark in color should be removed with a pipette. To further prevent mold or bacterial infection, embryos between 10 and 28 hours post fertilization (hpf) can be further treated with a bleach solution.4
3. Microinjection and Dechorionation
Microinjection of substances such as RNA, plasmids and morpholinos into embryos is a vital skill for any zebrafish researcher and can take some practice to master. Here are a few tips to make your injection process go smoothly:
Get as Much Done Ahead of Time as You Can
Zebrafish embryos develop extremely quickly, and you can expect a single-cell zygote to divide into two cells just 45 minutes post fertilization. If you’re hoping to have your embryos injected before that happens, completing as much preparation as possible ahead of time is crucial. There are many steps that can be completed the day before your zebrafish breed, such as needle pulling, dilution of reagents and preparation of injection plates. It’s a good idea to pull more needles than you will need to minimize the time spent replacing any needles that break.
Avoid These Common Pitfalls
There are a number of things that could go wrong during injection that could lead to the destruction of embryos upon injection or failure of the injection. Fortunately, many of the common problems are easily avoidable and correctable after the first bad injection. Firstly, make sure all excess egg water or embryo medium is removed from the injection plate, which will provide enough surface tension to prevent the embryo from sticking to the needle.
Secondly, make sure you break just a small piece at the tip of the needle at a diagonal angle so the tip is sharp enough to pierce the embryo but large enough to consistently release the desired volume of substance. Breaking off too far down the length of the tip can cause the diameter of the tip to be too large, which can damage the embryo or fail to pierce the chorion. Additionally, too big of an opening will cause too much of the substance to be released, which should become apparent during calibration.
Thirdly, avoid clogs by spinning the injection material before loading the needle to separate large particles.
Try Using Agarose Injection Plates
One simple method for positioning embryos for injection is to line the embryos up against the edge of a glass microscope slide, but other methods are available that can be more efficient and less error-prone. If you are not using them already, agarose molds that create wedge-shaped troughs for embryos to sit in during injection can increase the number injections you can perform per plate and allow you to deposit embryos in columns more quickly.5 These troughs can also be more effective at keeping embryos in place, and can be prepared easily the night before injection. In addition, the agarose substrate reduces the chances of needle breakage from hitting the plate surface.
Zebrafish embryos will hatch naturally from their chorions after about 48-72 hpf, but dechorionation can help with mounting and imaging embryos that are earlier in the development; removing the chorions could also improve chemical uptake in some experiments.6 Chorions can be removed manually by carefully tearing and pulling them apart using forceps under a microscope, or enzymatically by incubating and swirling several embryos in a pronase solution until the chorions soften and begin to fall off. Gently triturating embryos treated with pronase solution in a pipette tip should cause remaining chorions to fall away.7 Dechorionated embryos will be very fragile and should be handled with extreme care, especially embryos younger than 15 hpf.
4. Mounting and Imaging
When the time comes to get an up-close look at the results of your successful zebrafish breeding, embryo rearing and injections, there are a few considerations to make about mounting and other imaging preparations. Firstly, it’s important to minimize the embryos’ exposure to light prior to imaging, as this can cause darkening of the yolk sac and impact imaging quality.8 Next, embryos need to be immobilized with an anesthetic - tricaine keep the embryos immobilized for hours and is useful for long-term imaging. A couple of drops of 0.4% stock tricaine in a dish or a prepared working solution of 0.016% tricaine should be sufficient to anesthetize the embryos, but be sure not to use too much tricaine as this will kill the embryos.
Vessel Considerations
For low-throughput imaging in a standard glass microscope slide, a helpful method for preventing the embryo from being flattened is to place a barrier between the slide and coverslip. One simple technique is to cut a square-shaped hole into the middle of a strip of layered electrical tape. The square hole serves as a reservoir for the embryo to be mounted in the layered tape and creates enough space between the slide and coverslip to avoid flattening the specimen. Petroleum jelly, vacuum grease and modeling clay at the corners of the coverslip can also be used to create the necessary space.
For high-throughput imaging in 96-well or more microplates, well shape should be considered. Round bottom plates will allow embryos to naturally settle in the center of the well, but have the disadvantage of making it more difficult to have the embryos lie neatly on their side for lateral imaging. Conversely, flat-bottomed plates make it more likely embryos will settle flat on their side, but can cause embryos to collect around the edges of the well.
Choosing a Mounting Medium
Your choice of mounting medium will depend on factors such as how long you will spend imaging, whether you need to manipulate the embryos during imaging, and whether you are imaging live embryos or fixed embryos.
For long term imaging, low melt agarose is ideal, as it can keep live embryos in place for an extended period of time. When using agarose as a mounting medium, ensure you allow it to cool to an appropriate temperature before mounting, as excessive heat will kill the embryos.
For a less rigid medium that allows for embryos to be manipulated, 2-3% methylcellulose is a good choice. This medium will still keep the embryos in place long enough to be imaged, but is flexible enough to allow embryos to be moved and repositioned if necessary.
For imaging of fixed embryos, glycerol can be used. Like methylcellulose, glycerol allows for manipulation and repositioning while also holding the embryos still during imaging. An 80% glycerol solution provides a good balance of viscosity and fluidity.
Overcome Focusing Challenges With Image Stacks
Whole zebrafish embryos can be challenging to bring into proper focus due to being a relatively thick specimen. Because different parts of the embryo will be on different focal planes, it would be difficult to have every part show up clearly in a single image. This problem can be overcome by taking several images, creating an image stack that can then be processed to produce a final image with everything in focus. For fluorescent imaging, maximum projection stacks can give you the clearest view of your labeled structures.
References
1. Sessa AK, White R, Houvras Y, et al. The effect of a depth gradient on the mating behavior, oviposition site preference, and embryo production in the zebrafish, Danio rerio. Zebrafish. 2008;5(4):335-339. doi:10.1089/zeb.2008.0535
2. Lawrence C. The Art of Breeding Laboratory Zebrafish. Webinar hosted by Labroots. https://youtu.be/HrQHv5RLl0k
3. Westerfield, M. General Methods for Zebrafish Care. In: The zebrafish book. A guide for the laboratory use of zebrafish (Danio rerio). 4th ed., Univ. of Oregon Press, Eugene, 2000. https://zfin.org/zf_info/zfbook/chapt1/1.3.html
4. "Bleaching Zebrafish Embryos," Chen Lab, Stanford University. http://web.stanford.edu/group/chenlab/cgi-bin/wordpress2/wp-content/themes/chen/assets/docs/EmbryoBleaching.pdf
5. Westerfield, M. Cellular Methods. In: The zebrafish book. A guide for the laboratory use of zebrafish (Danio rerio). 4th ed., Univ. of Oregon Press, Eugene, 2000. https://zfin.org/zf_info/zfbook/chapt5/5.1.html
6. Wilson LB, Truong L, Simonich MT, Tanguay RL. Systematic Assessment of Exposure Variations on Observed Bioactivity in Zebrafish Chemical Screening. Toxics. 2020; 8(4):87. https://doi.org/10.3390/toxics8040087
7. "Dechorionating Zebrafish Embryos," Chen Lab, Stanford University. http://web.stanford.edu/group/chenlab/cgi-bin/wordpress2/wp-content/themes/chen/assets/docs/EmbryoDechorionation.pdf
8. "Considerations for Zebrafish Imaging," Biocompare. https://www.biocompare.com/Bench-Tips/342773-Considerations-for-Zebrafish-Imaging/