Our protocols

1. Introduction.

Here we describe protocols to maintain and transfect cell lines derived from zebrafish embryos (36 hours post fertilization). We have the most practical experience with a line called PAC-2 that was originally obtained from Nancy Hopkins’ lab.

2. Materials

2.1 Routine passaging of zebrafish cell cultures

• Sterile, disposable tissue culture flasks (Greiner, cell culture grade, no specialized coating, caps NOT vented) and Petri dishes (Falcon, Cell culture grade), conical tubes and pipettes.

• Laminar flow hood for sterile tissue culture work.

• L15 complete culture medium:

L15 (Leibovitz) culture liquid medium (Gibco),

15% fetal bovine serum (Biochrom KG),

100 u/ml Penicillin / 100mg/ml Streptomycin (Gibco),

50 mg/ml Gentamicin (Gibco).

• 1X PBS (phosphate buffered saline) without calcium and magnesium (Gibco).

• 1x Trypsin-EDTA (diluted in 1x PBS) (10x stock from Gibco).

2.2 Electroporation of zebrafish cell lines.

• Gene pulser apparatus (Bio Rad).

• Haemocytometer.

• 4mm sterile electroporation cuvettes (Peqlab).

2.3 Stable transfection of PAC-2 zebrafish cells:

• G-418 resistance encoding plasmid (based upon pcDNA3,1, Invitrogen).

• Geneticin G-418 sulphate (Gibco).

2.4 Long term storage of cells:

• Freezing medium

L15 culture medium,

30% fetal bovine serum,

10% DMSO (Sigma),

100 u/ml Penicillin / 100mg/ml Streptomycin,

50 mg/ml Gentamicin.

3. Methods

3.1 Routine passaging of zebrafish cell cultures:

All cell culture work should be performed in a laminar flow tissue culture hood using sterile solutions and plastic ware.

1. Aspirate the medium from a confluent cell monolayer and then gently wash the cells twice with 1x PBS.

2. Detach the cells from the culture flask substrate by a 5 minutes treatment with 1x trypsin-EDTA at room temperature. Use a volume of trypsin just sufficient to cover the cell layer. Dilute the suspension of detached cells with L15 complete culture medium and pipette the cell suspension vigorously up and down through a 5 ml or 10 ml pipette to break up cell clumps.

3. Transfer 1/5th of the suspension to a new flask and dilute with more L15 complete culture medium to ensure that the medium completely covers the new growth surface.

4. Seal the flasks and incubate at room temperature or in a 25°C incubator.

5. Passage the cells again when they reach confluence. Alternatively, change the culture medium each 7 days until ready to repassage the cells. The confluent cell monolayers remain viable for up to 1 month.

3.2 Protocol for transient transfection of DNA into zebrafish cells:

Contrary to what we describe here, we now use Fugene HD as our standard transient and stable transfection reagent – as efficient and a lot easier than electroporation. We follow the protocol described by the company. I can send you the precise details whenever you might need them.

Here, we describe a simple and reliable procedure for the introduction of DNA into zebrafish primary cultures by electroporation. Electroporation involves the exposure of cells to a pulsed electric field to create transient pores in the plasma membrane and thereby facilitate access of DNA. We have studied the PAC-2 cell line extensively and so have determined a set of electroporation conditions that provide optimal transfection efficiency.

1. Plate cells at 40-50% confluence in L15 complete culture medium 2 days before electroporation. This ensures that the cells are in exponential growth phase the day of the transfection.

2. Aspirate the medium from the cell monolayer and then detach the cells from the flask by trypsin treatment to produce a suspension of single cells (see section 3.1).

3. Determine the cell density by counting cells in a haemocytometer. Enough cells for several electroporation reactions (1×107 cells are required for one transfection reaction) are centrifuged at 200g for 5 minutes at 4°C and washed twice with 1x PBS. Each cell pellet is re-suspended by pipetting in serum-free medium (0.5 ml of medium for each transfection reaction) and stored on ice for a maximum of 30 minutes before electroporation.

4. Dissolve 35 g of plasmid DNA containing 25 g of the experimental plasmid and 10 g of the DNA carrier in 50 l of water and mix with a 500 l aliquot of resuspended cells.

5. Transfer each aliquot of cells mixed with DNA to a cuvette for electroporation and perform the electroporation at 0.29 KV, with capacitance set to 960 F and resistance set to 0, at room temperature using a Bio-Rad gene pulser apparatus. Following electroporation dilute each cell aliquot to 10 ml with L15 complete culture medium and then transfer to a 10 cm tissue culture Petri dish to allow attachment of viable cells.

6. The day after transfection, remove the culture medium, wash the cell monolayer twice with 1x PBS in order to eliminate cell debris and then add fresh complete medium.

7. Maximum levels of transient expression are detected between 48 and 72 hours following transfection. Therefore, cells should be harvested at this stage.

3.3 Stable transfections:

1. Harvest the cells and prepare them for electroporation exactly as described in section 3.2.

2. To each 500 l of cells, add 35g of plasmid DNA, containing 10 g of carrier DNA and 25 g of experimental plasmid (when the neo cassette is integrated into the same plasmid) or alternatively 22.5 g of experimental plasmid and 2.5 g of plasmid containing a neo resistance cassette. Also, include a negative control cell aliquot lacking plasmid DNA.

3. Perform the electroporation at 0.29 kV, 960 F, R = 0. Following electroporation dilute each cell aliquot to 10 ml with L15 complete culture medium and then transfer to a 10 cm tissue culture Petri dish to allow attachment of viable cells.

4. The day after transfection, remove the culture medium, wash the cell monolayer twice with 1x PBS in order to eliminate cell debris and then add fresh medium.

5. 72 hours following electroporation start the selection for antibiotic resistance by supplementing the medium with the highest antibiotic concentration, 800 g/ml of G-418.

6. Each five days wash the cells with 1x PBS and change the selection medium.

7. After a period of 10 to 15 days, all non-resistant cells on the negative control plate (non-transfected cells) should have detached. At this stage, reduce the concentration of G-418 to 400 g/ml.

8. Following 1 month of selection reduce the G-418 concentration to 250 g/ml (the maintenance concentration – as used for DAP49). At this stage, 100 to 200 colonies of resistant cells should be clearly visible for each electroporation reaction. Colonies typically consist of a monolayer of densely packed cells unlike other mammalian transformed cell lines where colonies are typically composed of multiple layers of cells.

9. When positive clones contain several hundred cells, they are large enough to be individually trypsinized and transferred to single wells of a 96-multiwell plate. Upon reaching confluence, transfer the cells to larger multiwell plates and then ultimately maintain them in 25 cm2 flasks. Alternatively, the clones can be trypsinized, mixed together and subsequently propagated as a pool.

10. The clones should be propagated always at the maintenance concentration of G-418 and the medium changed each 5 days.

3.4 Protocol for long term storage of cells:

1 Wash a subconfluent cell monolayer twice with 1x PBS, trypsinize and then resuspend the cells in 10 ml of L15 complete culture medium.

2 Determine the density of the cell suspension by counting an aliquot with a haemocytometer and then harvest the cells by centrifugation at 200g for 5 minutes at 4°C and wash twice with serum-free medium.

3 Finally, resuspend the cells at a density of 2×106 cells/ml in freezing medium (lacking G-418). Transfer 1 ml aliquots to cryopreservation vials and place them in a -80°C freezer for 2-3 days.

4 Transfer frozen aliquots to liquid nitrogen for storage.

3.5 Thawing cryopreserved cells:

1 Remove frozen aliquots from liquid nitrogen and place them immediately on ice for 10-15 minutes.

2 Remove the thawed cell suspension from the vial and immediately dilute it in 10 ml of complete culture medium at room temperature.

3 Centrifuge the cells at 200g for 5 minutes at 4°C and discard the supernatant.

3 Resuspend the cells in L15 complete medium (lacking G-418) and transfer them to a tissue culture flask. Typically, more than 50% of the cell should attach.

4 After 24 hours change the medium for fresh L15 complete culture medium (include 250 g/ml G-418 for resistant clones) to remove dead, floating cells.

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