Culture conditions for new cell lines

Setting up Drosophila primary cultures from Ras^V12-expressing genotypes

This is a detailed protocol for establishing primary cultures that has more details than the original method in a paper describing derviation of continuous lines from Drosophila culures (https://elifesciences.org/articles/85814#s4).

Author for enquiries: Amanda Simcox, Department of Molecular Genetics, Ohio State University, Columbus OH 43210. simcox.1@osu.edu

Materials and reagents:

Egg laying medium: Separately autoclave solutions A and B, combine and add food color for contrast so that eggs and hatched larvae are more visible. Pour (sterilely) into 60mm petri dishes and store at 4°C. 

• Solution A: 83 g dried bakers yeast, 75 mL water, 175 mL 4% acetic acid
• Solution B: 17 g agar 485 mL water

TXN: This is used to wash eggs and keep them from adhering to each other. Make a large volume for convenience and store at room temperature.

• NaCl (0.7%), Triton X (0.02%) in water

Cell culture medium: Many companies make media suitable for Drosophila cell culture including Schneider′s Insect Medium. The Drosophila Genomics Resource Center Cell Line pages have information on media types (https://dgrc.bio.indiana.edu/Protocols). Medium is supplemented with 10% Fetal Bovine Serum (FBS). Many sources of FBS are available and the Ras-method produces robust cells that in our experience have grown well in all the different serum lots we have used over the years. Antibiotics although not strictly required with good sterile technique can also be added. We use antibiotics; however, the Drosophila Genomics Resource Center do not routinely use antibiotics in the cell cultures grown there. The following are the sources we typically use:

• Schneider′s Insect Medium Sigma Aldrich Cat. # S0146
• FBS, Gibco ThermoFisher Cat. # 26140-079
• Penicillin-Streptomycin (10,000 U/mL), Gibco ThermoFisher Cat. # 15140122
• 0.05% Trypsin–EDTA (1×) Gibco Thermo Fisher Cat. # 25300–120

Equipment:

Fly cage

• Flystuff® Drosophila Embryo Collection Cages CAT#: 59-100 Size Small Fits 60mm Petri Dish, 4 Cages/Unit

Homogenizer

• Corning (or similar) CAT#: 7725T-5 PYREX® 5 mL Potter-Elvehjem Tissue Grinder with PTFE Pestle

Transfer Pipettes: These are used for all liquid handling. They are very efficient to use and fully depressing the bulb will take up about 4 mls of liquid. When seeding cells, one drop from the pipette is about 50ml.

• Fisherbrand 13-711-9CM (500—25 bags of 20—sterile)

Procedure

Establishing Primary Cultures

1. Set up cages of flies of the desired cross. When using the Ras-method one stock should have the UAS-RasV12 transgene and the other the Gal-4 driver with expression in the target cell type. The stocks we generated that also include an RMCE site for inserting single transgenes are available from the Bloomington Stock Center (https://bdsc.indiana.edu/stocks/misc/celllines.html).The small cages can accommodate large numbers of flies, we typically use approximately 200 males and 200 females. Fluting a piece of Whatman 3MM paper and inserting it into the cage gives the flies a more extensive surface to stand on. 
2. Egg collections and harvesting: Eggs are collected overnight at 17-19°C or through the day at room temperature or at 25°C. Add approximately 3mls of TXN to the plate and use a microscope to view and remove any hatched larvae, which rise to the surface and can be seen moving. Dislodge the unhatched embryos using a large soft paint brush to gently release them from the surface. Embryos are tipped off with the liquid into a sieve. We make these by cutting the end off a 15 ml Falcon tube, cutting a hole in the lid and then securing a piece of mesh that retains eggs (Genessee 57-103). Additional rinsing and brushing ensures most embryos are dislodged and collected in the sieve. After thorough rinsing of the embryos with TXN from a squirt bottle, the sieve is upended over a 15ml Falcon tube and a stream of TXN is used to transfer the embryos into the tube. Once the embryos have settled, the TXN is removed and replaced with 7mls of 50% bleach (Clorox) in water. The tube is capped and inverted 3-5 times and subsequently the embryos are handled using sterile techniques. The embryos are allowed to settle at the bottom of the tube and the bleach removed after 3-5 minutes. The bleach dechorionates and surface sterilizes the embryos. The embryos can be checked with a microscope to ensure the egg shells are removed and they appear shiny because the vitelline membrane is exposed. 
3. Setting up primary cultures: The tube of embryos in bleach are taken to a sterile hood and treated sterilely from now on. The embryos are rinsed 2X with 4 ml of sterile TXN and transferred to a fresh tube of TXN to minimize bleach contamination. The embryos settle rapidly in TXN and can be transferred in a small volume of TXN by ‘vacuuming up’ from the bottom of the tube. After 2 additional TXN rinses the settled embryos are transferred to TXN in a 5ml glass homogenizer. Embryos are rinsed in 3ml of water followed by one rinse in 1 ml of Schneider’s S2 medium (supplemented with 10% heat inactivated Fetal Bovine Serum and 1X Pen-strep solution). Embryos tend to clump in the Schneider’s S2 medium and stick to the sides of the homogenizer and pipette, and care is needed to remove the medium without disturbing the embryos. 3ml of fresh Schneider’s S2 medium is added to the homogenizer and the embryos are disrupted by 3 gentle strokes with the pestle. Care should be taken to minimize bubbles by not withdrawing the pestle beyond the surface of the liquid. The homogenate is allowed to settle for 2 mins and the supernatant is transferred to a 15ml Falcon tube leaving the large cell clumps and any whole embryos in the bottom of the homogenizer. 3ml of fresh Schneider’s S2 medium is added to the homogenizer and 3 more strokes, with a twist at the bottom, are used to disrupt remaining tissue and embryos. The second homogenate is added to the Falcon tube. The tube is centrifuged in a benchtop centrifuge at 1,400 x g for 3 mins. The supernatant is discarded by pouring off, and the pellet was resuspended first in the small volume remaining in the tube and then in 4ml Schneider’s S2 medium. The centrifugation step and washing with Schneider’s S2 medium is repeated twice more. The final pellet size is estimated and plated in 1 or more 12.5cm² T-flasks with 3-4 ml Schneider’s S2 medium. The number of flasks needed for a given pellet size can also be estimated from the volume of packed embryos with approximately 30µl of packed embryos being sufficient for one flask. Judging the plating density is important and can be learned with experience.
4. Caring for primary cultures: A typical egg collection generates 2-3 flasks of primary cultures. If two collections are made each day, a week of work will generate sufficient primary cultures to determine if a given genotype is conducive to cell line production. In general, we found that egg collections made during the day were more successful likely because the embryos are younger; however, overnight collections were also successful so both can be used. The embryonic cells soon attach and many differentiated cell types like nerve and muscle can be seen—they are very interesting to observe. Infections are rare but will usually be obvious by approximately 3 days when the medium will appear cloudy. In a few days (7-10 days) patches of proliferating cells are visible and if these are marked to express GFP as well as Ras, the cells will be GFP positive. Depending on the genotype, the cultures will be about 70-90% confluent within a month. During this growth period, we change the medium about every 7-10 days. This can be done to replace half the medium; however, Ras^V12-expressing cultures are robust and can withstand a complete medium change.
5. Early passages: Determining when to make the first passage is important: The flask should have large colonies of proliferating cells that are starting to make a contiguous patch of multiple colonies contacting each other. For the first passage, remove the culture medium and put it into a 15 ml Falcon tube. Rinse the cells in the flask with 1 ml 0.05% Trypsin–EDTA, discard the rinse and add approximately 3 mls of 0.05% Trypsin–EDTA. Leave the cells in incubate for 3-5 mins. You can observe them with a microscope (inverted) and will see sheets of cells and individual cells freed from the flask surface. Squirt the cells gently by sucking up and releasing the trypsin solution. Once most cells have been dislodged add the cells in trypsin to the tube containing the ‘old’ cell culture medium. This dilutes and inactivates the trypsin. Add medium back to the primary culture—it will likely regrow because not all cells are removed and serves as a backup. Centrifuge the harvested cells in a benchtop centrifuge at 1,400 x g for 3 mins. Pour off the supernatant and resuspend the cells in the remaining liquid (usually about 200 ml or 4 drops). Seed two drops of the cell suspension in a new flask with approximately 4mls of fresh medium. This is passage 1 and if healthy will grow to confluence in 2-4 weeks. Check on it periodically and set up passage 2 when it is 70-90% confluent. As with the primary culture, we put new medium on passage 1 and keep it as a backup. We find that 10 primary cultures will yield 5-10 lines provided the genotype is conducive to cell line propagation. Given this success rate, it is typically unnecessary to keep multiple sublines from a given primary culture—this creates a lot of work and complicates the derivation of independent lines that can be distinguished by coming from a given primary culture. As passage number increases cells can be plated at larger dilutions (1 in 4).
6. Different methods: We generated lines from a number of cell types and intestinal cell lines have been generated (https://www.nature.com/articles/s41598-025-17336-z#Sec2). In this latter publication, embryos were homogenized in 0.2% Trypsin (Thermo Fisher 12563011), which may promote tissue dissociation. This method also uses sieving to remove large clumps.