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Hello,

I want to express stable cells lines of S2 cells expressing my transgene.

My cells were seemed to grow in 6-well plate but after 4 weeks when I tried to expand them into 5-ml flasks they seem sick and small. I used Hygromycin B as a selection agent starting from the dilution of 1:250 and decreasing to 1:1000.

Do you have any suggestions why the cells are not healthy?

Thanks in advance

I was cultivating HEK-293s which were modified with CRISPR under the following conditions:

temp: 37°C

CO2: 5%

humidity(?): 98%

But out incubator failed during the night (completely failed, including the alarm that should have gone off once it lost 5% CO2) and the next morning (which is today) we came back to yellowed medium in our flasks and really bad-looking cells. I just put them back into a different (working) incubator with 37°C/5% CO2 again to try to rescue them but I'm not sure if there is a better way? Should I change the medium first, then put them back? Should I rather harvest them now and distribute the whole batch into 96-well plates to have them more compact together? Is there any other option to rescue them? A single surviving cell would be enough (I hope) to regrow the batch?

Hello all,

I've recently been trying to differentiate some 3T3 cells (below passage 20) and I have been having some issues. I seed 0.5x10^5 cells/well in 24 well plates and by 2 days the cells are confluent. Following this, I treat the cells with 1mL each media containing 10mg/mL insulin, 1uM dexamethasone and 0.5mM IBMX (MDI media) for 48hrs and then another 48hrs with media only containing 10mg/mL insulin (insulin media). Whenever I treat with the MDI media, the cells continue to grow over the next two days however during the culturing protocol, the cells start detaching and they look granulated and dead. I was wondering if anyone has experienced this and what I could do to prevent this from happening.

I have been doing long-term cultures of primary cortical neurons for several years, from 2018-2021, with no problem. I use Neurobasal medium with 2% B27, 1% glutamax, 1% penstrep. I would grow my neurons up to 31 days in culture at 30,000 neurons per well in a 12-well plate on a PDL-coated glass coverslip. I would change the entire volume of media on the 4th day in vitro (DIV), and then 50% of the volume every 3 days until the 16th day, after which I would change 50% of the media every 6 days.

Suddenly, at the end of 2021, my neurons died after DIV16. The neurons were beautiful at DIV16, and suddenly died the day after. I thought it was the B27 and glutamax and penstrep that were close to expiring, so I ordered new B27 and glutamax and penstrep and cultured again, but then the cultures started to die after DIV6.

In one of my recent cultures, one coverslip of neurons looked beautiful and just like how my neurons used to look in the past. However, the rest of the neurons in the other wells were dead. I used the exact same dissociation media, same neurobasal + supplements media, and same PDL coating on the same acid-washed coverslips from the same jar on every single well. So, I thought perhaps there was variation in the coverslips, so I washed them more intensively for my most recent culture, but the cells still died on DIV7.

What could be happening? I would appreciate any help.

I thawed a tube of Calu-3 cells from ATCC, following product instructions. The flask I used was not coated with any substrate. I used EMEM with 10% FBS and placed it in the usual 37degC incubator with 5% CO2 and water pan. The next day I discarded floating cells. Since then, these cells have always appeared round and bright (see attached photomicrograph). They appear to be able to change positions, moving from middle of vessel to the sides. At first the numbers grew slowly but later it started dying. I have changed the media to DMEM with sodium pyruvate and 10% FBS. I had transferred floating cells to another vessel coated with collagen IV. I had also trypsinized one flask and transferred all cells to one collagen IV-coated well of a 12-well plate, and refreshed media every 3 days. Each day I waited for it to adhere firmly and resemble the clumps of cells I saw in the ATCC photomicrographs. But after a month, they are still round and bright. Now, they are starting to die off. Can anyone advice or suggest what I can do to rescue these cells?

We are conducting a large-scale inhibitor-based screen in transfected BaF3 cells, where we seed the cells in 384-well plates in the presence of different kinase inhibitors. The issue we are having is that all the cells often die after the seeding into 384 plates in a new media (also control cells). It happens often, but not every time and we have ruled out the problems related to media and serum change, seeding density (we seed 2500 cells in 50ul), mycoplasma contamination.

Has anyone experienced something similar?

Hi all - first time poster!  Hope I can get some good feedback.  I am Ph.D. student and am working with an HN2-5 cell line (murine hippocampal neuron).  I've successfully done cell culture before and with the same experiment, but for some reason I'm having trouble keeping my cells viable.   We do EtOH research and so I have 4 groups - Control, DHA, EtOH, and DHA+EtOH.  I am using 12 well plates coated with Poly-L-lysine and seeding at 55,000 cells per well.  After reaching 70% confluence I treat with 5uM retinoic acid to differentiate.   After 24 hour treatment I begin my EtOH treatment which lasts 4 days.  Each day media is replaced with fresh media.  I grow cells in 10%, use 1% FBS while differentiating with RA, and 0.5% for treatment.  This is per a protocol a lab member of mine used in the past that was successful with these cells.  

I have done this experiment on 100mm petri dishes successfully. Maybe a month ago, but now the cells are clumping up on the last two days of treatment (in any group including control), they are shriveling up and there looks to be a good amount of cellular debris from dead cells.

Any idea as to what could be going on? 

Hi colleagues, I'm having problems sorting neural crest cells. I isolated NCC from the head of E13.5 embryos marked with tdTomato. The procedure for the isolation of the cells includes some steps using TrypLE for embryo digestion, followed by FBS supplementation, cell scraper to reduce clogs during sorting and I keep them on serum-free DMEM for sorting. Sorting takes ~20min and the cells are collected in DMEM supplemented with 10% FBS. I usually get 6 million td+ cells for 4 embryos.

I incubated them in 6-well plates coated with 2% gelatin and changed the media the next day, but almost 60% of them were dead. Does anyone have any tips on how to improve the cell's viability or if there is anything I should do to improve the sorting?

**This is the 3rd time I'm planning to do and previous lab members couldn't figure it out.

Hello everyone,

I'm a phD student doing primary cortical neurons culture for 3 years. I'm doing this protocol from embryonic mice (E16) and following these steps

-Dissection in cold HBSS 1X +mg +Ca

-Cortices are kept in DMEM high glucose + glutamax supplemented with 10% FBS and 1% P/S

- After DMEM complemented is removed and trypsin 0.05% is add and cortices are incubated for 15 min at 37°C.

- Trypsin is removed and cortices are washed with DMEM complemented to stop the reaction, then dissociated in 1 mL with a P1000 (10-15 up and down) and filtered with a 40µM cell filter.

- Then we add 9mL of DMEM complemented to dilute cells and we count.

- Cells are plated in precoated PLL plates (100mg/L, washed 3 times with H2O) with DMEM complemented, 37°C 5% CO2.

- After 4 hours, medium is fully changed with Neurobasal + B27+ gentamycin + Glutamax

This protocol was working really well in our hands but since this summer is not working anymore.

We have a lot cell mortality even before plating the cells and the ones that are surviving after plating are presenting vacuoles and are stressed. We continue to see debris and mortality.

We changed with new solutions all our media, we autoclaved our instruments, we changed many "lot" of FBS. We tested another incubator, changed our coating. We did also a mycoplasma test which is negative after 2 weeks of culture.

During summer we had some changes in our cell culture facility (temperature changes, pressure, ventilation filter changes) and we are 4 users with the same problem (knowing that before summer for all of us, all was working well).

We notice that our media change color really fast (after less than one week after opening)

We changed I guess all and we don't have ideas anymore of what could be the problem.

Any ideas ?

Thx a lot

I have a lentiviral transfection protocol on 293T cells that has been working for years, but for the past few months, my 293T cells starts de-attaching from the plate. I finally figured out it was the issue with my BGS serum so we switched to DMEM+10% FBS and they starts growing normally again and I noticed cells tend to adhere better in FBS media. Well, this time, they starts dying after transfection with lentival constructs at 5ug of plasmid. I changed media overnight after transfection and was bring very gentle. I have been using the same protocol for a few years now, and I can't seem to figure out what is the issue of them not growing well this time. Any advice would be appreciated!