We present an efficient and easy-to-use protocol for preparing primary cell cultures of zebrafish embryos for transfection and live cell imaging as well as a protocol to prepare primary cells from adult zebrafish brain.
This method provides the zebrafish community a tool for the the analysis of intercellular dynamics with a resolution unattainable in vivo. And also it allows to compensate for the lack of zebrafish cell lines. The main advantage of this technique is that you isolated and tranfect those primary cells directly from the original tissue so you do not need to keep cell lines.
To begin the protocol prepare the zebrafish embryos two days post fertilization, or dpf. On day one set up several crossings of the zebrafish strain of choice. On day two mate to the fish and collect the eggs directly after spawning in a plastic 10 centimeter Petri dish.
Remove the dead and contaminated eggs with a plastic Pasteur pipette. Wash the eggs once with Danieau 30%with 0.0001%weight per volume methylene blue. Then replace the medium with fresh Danieau 30%without methylene blue and incubate the eggs overnight at 28 degrees Celsius.
On day three remove the dead and contaminated embryos and exchange the medium to Danieau 30%Then incubate the embryos overnight at 28 degrees Celsius. On day four remove the chorions by removing the medium until approximately 10 milliliters are left in the dish. Add one milliliter of one milligram per milliliter of pronase and incubate the embryos on a shaker for approximately 30 minutes at room temperature until all the chorions are detached.
Next spread silicone grease around the hole on the underside of the dish bottoms and attach a cover slip using the grease as glue. Make sure that the grease seals the gap between the dish bottom and cover slip. Wash the glass bottomed dishes thoroughly but carefully with cold tap water and soap.
Rinse the glass bottom dishes three times with deionized water to remove the soap. Then air dry the dish lids and dish bottoms and store them in a clean box until further use. On the day of culture preparation moisten the inside of both dish lids and dish bottoms with 70%ethanol.
Place the dish lids and the dish bottoms facing the inner side up in a sterile workbench with laminar flow and UV light. Air dry the dishes until the ethanol has evaporated then apply UV light for 20 minutes. After this treatment dishes are assembled and sterile.
To coat the dishes pipette 200 microliters of 0.1 milligram per milliliter poly-L-lysine in the middle of each glass bottom dish and spread the liquid on the cover slip by breaking surface tension with a pipette tip. Then incubate dishes for 60 minutes under the bench at room temperature. After the incubation wash them once with sterile 1x phosphate-buffered saline or PBS and remove the liquid.
Keep dishes under the bench until further use. On day four the day of culture preparation transfer the embryos to a sterile cell culture dish with a fresh plastic Pasteur pipette. Remove the liquid step-wise until all the embryos are gathered in a big drop with a diameter of about two centimeters or less.
Place the dish with the embryos in a sterile workbench and add carbon dioxide independent medium until the dish is half filled. To remove the yolk pipette the embryos up and down using a 200 microliter pipette tip. Successful deyolking can be recognized by the cloudiness of the medium.
Next fill a cell culture dish with 70%ethanol and another cell culture dish with fresh cell culture medium. Use a cut off 1000 microliter pipette tip to transfer embryos into a sterile 40 micron cell strainer with a handle. Take the strainer by the handle and dip it into the dish with ethanol so that all embryos are submerged for five seconds.
After the ethanol treatment immediately submerge the strainer with embryos in the dish with fresh cell culture medium. Then transfer the embryos into sterile 1.5 milliliter reaction tubes. Add collagenase type 2 to the embryos.
Incubate the tubes on a vertical tube rotator. Dissociate the remaining cell clumps by pipetting the mixture up and down with a 1000 microliter pipette tip. Then filter the cell suspension through a sterile cell strainer with a venting slot into a 50 milliliter conical tube.
Rinse the strainer with 10 milliliters of fresh cell culture medium. Centrifuge the cells for three minutes at 180 times g. After centrifugation carefully check for pellet formation.
After that remove the supernatant and resuspend the cells in 200 microliters of fresh cell culture medium per 30 originally used embryos. Then pipette 200 microliters of the cell suspension directly on the glass area of a prepared poly-L-lysine coated glass bottom dish and let the cells attach for 60 minutes. Add six milliliters of fresh cell culture medium and incubate primary cells at 28 degrees Celsius.
Then perform the desired imaging application using an inverted microscope at 28 degrees Celsius. Determine the cell number in a counting chamber. Then in a 1.5 milliliter reaction tube mix 10 micrograms of ultra-pure plasmid DNA with 0.5 million cells and adjust the total volume to 100 microliters with sterile PBS if needed.
Thoroughly resuspend the solution for 20 seconds prior to electroporation. Next transfer the cell DNA mix immediately to an electroporation cuvette place the cuvette in an electroporation device and electroporate the sample. Directly after the electroporation transfer the cell DNA mix into a 1.5 milliliter reaction tube with 300 microliters of fresh cell culture medium.
Finally plate 200 microliters of the cell suspension as done in previous sections. Using the method in this video transfection of the pCS2 positive-based plasmids encoding fluorescent organelle markers such as endoplasmic reticulum-targeted red fluorescent protein or mitochondria-targeted yellow fluorescent protein subcellular structures were visualized in great detail. Simultaneous analysis was performed via co-transfection of the fusion proteins MitoTag-YFP with the microtubule marker human doublecortin fused to the red fluorescent protein tdTomato and the nuclear marker Histone H2B fused to a cyan fluorescent protein.
Selected images from a time lapse of the movement of vesicles positive for the membrane marker vesicle associated membrane protein one fused to the fluorescent protein mCitrine provides high temporal and spatial resolution. Morphological changes over time can be easily observed by labeling the whole cell by the expression of a bright fluorescent protein. They electroporation protocol is also suitable for combinatorial genetics.
The dissociation technique and the basic culture conditions were also applied to adult brain tissue from wild type zebrafish and shows the progressive development of an initially debris-coated culture to a complex neuronal-like network. Thanks to the availability of many zebrafish models of human diseases and tissue-specific reporter lines our method can be used to describe the intracellular mechanisms that underline biogenesis with focus at specific cell types and developmental stages.
