Daphnia Embryonic Primary Cell Culture Protocol
Isern Lab (Last updated: 5-27-09)
Journal of Experimental Zoology 305A:62-67 (2006)
The main focus of the protocol is the sterile removal of the embryos from the females. Utmost care must be taken to keep all solutions sterile and clean all tools and working surfaces with 70% Ethanol including bench area and microscope stage. I recommend wearing gloves, a disposable gown and face mask.
75-100 Daphnia with embryos are filtered out of their media and placed in a 1.5ml Eppendorf tube, in the smallest volume possible. The surface of the tube is washed with 70% Ethanol before bringing it to the sterile working area. The Daphnia are surface sterilized by washing with a series of solutions (shown below). 500?l of each wash are placed in the Eppendorf containing the Daphnia, the tube is shaken for about 10 seconds and the liquid is carefully removed using a pipetman.
- 1 X 0.05% NaHClO4
- 2 X dH2O
- 1 X 70% Ethanol in dH2O
- 2 X dH2O
- 2 X Daphnia Cell Media
The Daphnia, for the most part dead at this point, are resuspended in 1-1.5ml of Daphnia Cell Media and decanted into a sterile 60mm plate. This may take a few tries as some of the Daphnia will stick to the walls of the tube.
The Daphnia are dissected (under a dissecting microscope) with a pair of pre-sterilized micro-forceps (Robotz). The most important aspect of the dissection will be to keep the female's gut intact while releasing the embryos into the solution. Once the embryos have been released the Daphnia's body should be taken out of the solution to be discarded.
Once all the embryos have been taken out of the females, they are transferred to a new Eppendorf tube using a pipetman. The Embryos are then surface sterilized by following the series of washes described above. This time, however, only 200?l of each solution are used for each wash. You will notice that the embryos are denser than the solutions and will therefore precipitate to the bottom of the Eppendorf, making it relatively easy to aspirate the supernatant with a pipetman after each of the washes. Some of the embryos will not precipitate as well after the Ethanol washes and will be aspirated along with the supernatant. This is alright since these are usually the more developed embryos which would otherwise contribute significantly to the mass of exoskeleton that will carry over into the primary tissue culture.
The remaining steps of the protocol are performed in a sterile tissue culture hood. The surface sterilized embryos are resuspended in 200?l of Daphnia Cell Media. Using a sterile tissue homogenizer pestle (micropestle) in an Eppendorf tube, the embryos are popped open to release the cells in them. This must be done will extreme care; too much force will lead to the shearing of cells and too little to whole embryos. Usually I will slowly and gently move the pestle up and down very slowly and gently about 30 times. I will take the pestle out and observe to see how much precipitate remains and repeat the process until most of the embryos have been popped and most of the tissue takes longer to precipitate. The sample is centrifuged at 1000rpm for 5 minutes and the supernatant is removed and the cells are resuspended in 1ml of Daphnia Cell Media. The cell suspension is then placed in a 60mm plate containing 1-2ml of Daphnia cell media. The plate is sealed with parafilm and placed away from the light at room temperature.
Frequently asked questions:
1) How important is it to keep embryos intact through the dissection
and washes? Should I remove embryos that have inadvertently been killed
(and are starting to turn white)?
The 'shell' on the embryos is pretty sturdy. In general, manipulation during the washes doesn't rupture them. The micropestle is required for the actual 'popping' of the embryos from their shell. If there happens to be a few ruptured embryos before then, I wouldn't worry about it.
2) How is the cell culture treated after the suspension is put on the dish? Were they grow in an incubator? What was the CO2 concentration?
We kept the cultures in a drawer or in an incubator at ambient temperature – around 22 to 24 C. No CO2 was used. It's important to keep the cultures from drying out. The plates edges were sealed with stretched out parafilm. Culture plates were placed in flat-bottomed plastic 'Tupperware' containers (food storage containers) lined with a thin layer of pre-moistened (with water) paper towels. The containers were closed tightly with their lids.
3) When should the medium be changed (if ever)?
We usually maintained our cultures for about 2 weeks (though initially we went out to about 1 month to see how long the cells would persist – they seemed to persist, but not divide). For our viral infection studies, we prepared the primary cultures and infected them immediately – Day 0. Since viral gene expression was evident in 24 to 48 hours, we did not change the media on the cells. If you plan to let the cultures go longer, then I suggest that you change the culture media once per week.
4) Will the cells stick to the plate?
The cells will be in suspension. The viable cells will be in large grapelike clusters. Single cells are less likely to be viable. It's important when using the micropestle to break the embryo shells, but not to macerate the embryos much more beyond that point.
5) Will the cultures be homogeneous?
When we initially started working with Daphnia primary cultures, we first attempted to get a homogenous looking population of cells – meaning that we tried to disperse them into individual cells. The viability of the cells was much compromised. We found that minimal disruption of the embryos gave the best results. The attached pdf file (daphnia primary cultures) illustrates the relative heterogenous look of the primary cultures. We did not filter the cultures nor avoid big pieces of embryos – we kept everything including empty 'shells'. Since they are sterile (after all the washes, plus you've added antibiotics/antimycotics to the media) they should not compromise the cultures.
6) Will the cells divide; how many times?
Difficult to tell. The cells were maintained and viable, but I'm not convinced that they divided to a great extent.
7) How do you know if the cultures are viable?
The best way I've found to check the viability of your cultures is to expose a small aliquot of your cultures to viability stains. I use SYBR14 and propidium iodide (see attached pdf file from Molecular Probes). SYBR14 is a membrane permeant nucleic acid stain that will stain all nuclei green whereas propidium iodide is a membrane impermeant nucleic acid stain (red) that will only stain the nuclei of cells that have compromised membranes. When cells are exposed to both stains simultaneously, the nuclei of 'dead' cells will stain red whereas the nuclei of live cells will remain green. Do this only to a small aliquot of your culture, as the dyes themselves are toxic to cells after a few minutes.
8) How large is a Daphnia cell?
To get an idea of the size of a Daphnia cell, we compared Daphnia primary cultures to human red blood cells (several drops of blood from a finger prick) under the microscope at the same magnification. We estimated human red blood cells to be about 10 microns in diameter. In comparison, individual Daphnia cells were slightly (but not too much smaller). You may want to try this to convince yourself that you are looking at the cells and not debris. Then compare the cell size to the size of the nuclei stained with SYBR14.