Questions related to Cell Line Culture
Two or three days after removing the differentiation medium, the cells are detached from the culturing plates. I have tried several times, ending up with the same problem. Does anyone have the same experience or have any suggestions to fix this problem?
I cultured the Hela cell line for anticancer testing of a plant extract. On the 3rd day of subculture and changing the media, the cell culture showed that the DMEM medium was cloudy and looked like small particles when observed with an inverted microscope at 40X (as seen in the attached picture). What do you think contaminated the HeLa culture cells? Has anyone experienced the same as me?
I have recently embarked into mammalian cell culture using mutiwell culture plates (e.g. 6-well plates, 12-well plates, etc.) as opposed to previous studies where I have used individual flasks (e.g. T25, T75, etc.). When using flasks, I was accustomed to lysing cells in somewhat of a more "low-throughput" method in a lysis buffer containing 1% Triton X-100, applying mechanical shearing force by passing the cells through an 18G needle.
Moving to multiwell culture plates, the needle method is quite tedious. Having to pass the cells from each well through an 18G needle, one at a time, is very time consuming and counterbalances much of the time that is saved by using a multiwell plate in the first place.
Is there another method that is friendlier to "high-throughput" multiwell plates, that anyone might suggest which lyses cells without having to pass them through a needle one-by-one -- yet does not interfere with downstream assays for protein concentration?
Thanks in advance,
I am working with primary adipose-derived stem cells and have the problem that after plating the cells on 12-well-plates they all move to the center of the wells. I tried several things (8-shaped shaking, different volumes and seeding densitites, incubating the cells on a flat bench for 60 min before putting them into the incubator etc.) which all did not solve the problem. Plating other cells (for example NIH-3T3 cells does not lead to the problem - the distribution is very even so that I assume that it's a cell type-specific problem). Now my hypothesis is that the vibrations of the used incubator is the main issue. Are there any possibilities to somehow catch the vibrations inside the incubator? I found some "anti vibrating pads" in the internet for example. Maybe someone has another idea.
Do I need to add osteogenic induction media to hFOB1.19 (CRL-11372) cell line to hFOB1.19 (CRL-11372) cell line if I want the cells to differentiate to mature osteoblast? or these cells are osteoblast already? This is my first time hOB cell line culture. Some studies incubate at temperature 39.5°C for differentiating temperature. Is it necessary? or is it OK with Growth Conditions at temp of 34°C? ( recommended by manufacturer )
- If osteogenic media need to be added the media would contains Ham's F12 Medium Dulbecco's Modified Eagle's Medium, with 2.5 mM L-glutamine (without phenol red), 0.3 mg/ml G418, 10% fetal bovine serum supplemented with 0.1 µM dexamethasone, 50 μM ascorbic acid and 10 mM β-glycerol phosphate
I have been trying to passage ma-mel-66a cell line, however, I am having zero luck. I have used RPMI + 10%FBS and RPMI + 5%FBS, I even plated them (straight from LN2) in a small p10 to help them out. Nope, nodda, nothing. Can anyone recommend me a useful protocol or media formulation for this cell line?
I am starting working with E0771 cell line since I have to establish an orthotopic breast tumor model in c57 mice but I have no experience with this cell line so I would really appreciate any advice that you can give me.
In particular, I saw the ATCC website and they say that it is better to culture these cells in a t-75 corning flask, maintaining cultures at a cell concentration between 6 x 10^4 and 8 x 10^4 cells/cm2, is this true also for your experience? How many cells do you plate in a 75 cm2 flask?
Do they grow fast? How many times per week do you subculture them?
Sorry for all of these questions but I am new with these cells and so I would really be very grateful for all your advice,
Thanks a lot,
I have experienced this problem previously with a commercial cell line, however in this case, a well established immortalized cell line when cultured from frozen stock has cell populations that have a bright border around them. This has never been the issue with this particular immortalized cell line. I am wondering if there could be possible mycoplasma contamination. Any comments on this are highly appreciated. Thanks!
I need to prove that my protein is localized on the cell-surface but not inside the cell in soluble form.
How can I make it?
There is an opinion to wash cells, to label all cell-surface proteins with NHS-biotin, and to collect labelled fraction of cell-surface proteins with streptavidin magnetic beads. After that I can make western blot with specific Ab.
Maybe there is another way?
Protein and cell lines are human.
Wondering if TrypLE Express cell dissociation agent is significantly better than other options for culturing mammalian cell lines. Currently mainly using Accutase (for fibroblasts and epithelial cells, NOT stem cells) and sometimes also trypsin for specific cell lines. Any positive/negative experience with TrypLE Express enzyme? Main attraction is that it does not require neutralization as trypsin does, but has longer incubation times. Would appreciate any comments and suggestions.
Does anyone have experience with CAEV in immortalized cell lines? Some publications show use of goat synovial membrane cells or goat milk epithelial cells, but these cell lines are not available through commercial companies.
The Boyden chamber protocol requires a high budget for us. We want to try modifying this test instead. Can we combine it with a protocol like the filter diffusion protocol?
I am currently culturing NK92 cell line in complete RPMI supplemented with 500u/ml IL-2. Observing steady-state cells by fluorescence microscopy, I observed that the majority of them have a rounded but sometimes irregular shape, with a single nucleous showing lobes. However, about 10% of them showed a more regular rounded shape and a perfectly spherical nucleus.
Since this morphology correlated with the expression of a protein of interest, I was wandering if someone could tell me if the "more regular" shape could correlate to any functional state for these cells.
According to the analysis of cell-surface markers using flow cytometry, is it possible that trypsinization would block or even digest the markers?
If so, based on your experience, what is the best way to dissociate the adherent cells from the flask? Thanks
I am culturing the primary cells (Monocytes) from the patient blood samples. During the culture (2-3 weeks), I have seen the long tape shape black color filament (1 or 2 in number). Is it a type of contamination? How to overcome this?
Thank you in advanced.
I am working with M93-047 cells and I have cell lines cultured in T75 Flasks, I want to seed cell for Pull down IP experiments. I want to seed cells either in 3 or 4 10cm plates, so that I can have 3 or 4 samples of the cell line to work on.
I am confused that what dilution of RPMI media I should use to seed M93 cells, so if someone can help, and then secondly after counting the viable cells using hemocytometer, what factors we should keep in mind for seeding.
If someone can share good protocol for seeding cells, I'll appreciate.
I have seen some very strange debris in my U87MG (glioblastoma cell line) culture. It seems that there are shards/fragments of glass that are not contaminating the cell culture and the cells are potentially trying to adhere to them. I attached some pictures of the debris, taken shortly after passaging, so they are not fully adhered.
When I first saw this, I bleached the cells, threw out the package of flasks I was using, made new media, etc. I figured that this originated from a glass pasteur pipette, so I thawed another vial of cells that was frozen down at a different time than the previous ones I used. However, I'm still seeing the same debris in the freshly woken up cells.
I'm confused because I didn't see the debris in the first set of cells until weeks into using them, and I didn't see them in the second set of cells until about two weeks since I thawed them. I may have just not seen it until two weeks in, but it just seems unlikely due to how much and how obvious it is.
This is the only cell line I use EMEM (w/ penstrep & 10% FBS) for, and I have never seen this in any of my other cell types that use other media (completed with the same batch of penstrep and FBS).
Please let me know if you have ever seen anything like this before! Any comments are appreciated - thank you all in advance.
I am trying to induce inflammation in the cell lines (Beas 2B & NHBE) by using the dust but haven't worked with cell lines previously. Kindly help with how to identify the cell lines whether it is inflammated or not?
What are all the parameters I have to consider?
How does the passage number of a cell line affect experiment results including toxicity assays? Which characteristics of cells are changing as the passage number increases?
What is the most efficient or optimum passage number of cells (for example, for cancer cell lines such as HepG2, A549 etc. or for healthy cell lines like HEK293) for setting an experiment?
I've been trying to differentiating SH-SY5Y cells with brain-derived neurotrophic factor (BDNF) in 8-well chamber slides but I seem to be having problems with cells detaching over the course of the 7 days in serum-free media with BDNF. The problem seems to be more apparent after changing the media but I am unsure as to why this would occur and would value any insight from people with similar problems with SH-SY5Y or other cell types. My first thought was that I was removing too much media from the well, so I began to leave around 1/4 of the old media and add 3/4 new media and pipette the media off very slowly and add the new media very slowly (to the side of the well, not directly on to the top of the cells) but I still seem to be having the same issue. I thought there might be a problem with the incubator I was using so I changed incubators, and yet the problem persists.
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?
I recently sequenced (Sanger) an exon from single alleles of a CRISPR-Cas9 mutated cell line (diploid) to find whether the mutation is homozygous or heterozygous and what the mutation is. What I got is 3 different results, a WT, and two diff mutations, a 1 base del and a 7 base del. My lab has come to the conclusion that it must somehow have 3 alleles instead of 2, and suggested looking into this. I can't really find anything relevant to this find, so I'm wondering if anyone might know of such research articles or point me in the right direction, and if this is even possible?
My original thought was that perhaps the 7 base deletion is somehow an error due to the crispr process? But more experienced people than me have suggested it means there are 3 alleles, although like I said I'm struggling to find relevant literature to support this.
Any help would be much appreciated, thanks.
I have difficulty every time whenever I subculture C6/36 Cell line and vero cell line.
Can anyone suggest how to spread cells evenly in growth media.
In my experiments, I found that different monoclonal cell lines from the same host cell produce similar concentrations of ammonium ion when using the same medium; but when the same monoclonal cell line cultured with different types of fermentation medium, the concentrations of ammonium ion vary greatly. Is it because which of the ingredients in the medium is different that makes this difference?
I am looking for suggestions for HL60 cell line culture. Our lab is new to suspension cell culture. we bought the cell line from National Center for Cell Sciences (NCCS) Pune. Upon arrival, the cell was absolutely fine, having a doubling time of around 48 hours and we were able to freeze 10 vials with 10^6 cells/ml (90% FBS + 10% DMSO). But after 6 months in liquid Nitrogen when we started culturing from those vials it seems that the cells are not proliferating. We are seeing cell clumps forming and after 3-4 days all of these cells are dying (when checked using trypan blue). Can you help me with this problem? We are unable to point out the exact cause of the situation.
I want to know the expression change of my target protein by western blot from the control and drug-treated cells. 48hrs. after the treatment, I have pelleted the cells and extracted the proteins using RIPA buffer (50mM Tris-HCl pH 7.6, 150mM NaCl, 2% NP-40, 1% SDC, 0.1% SDS, 2mM EDTA). After denatured the proteins using SDS-PAGE loading buffer, I have loaded the samples in SDS-PAGE wells. But, some of the samples leaked from the resolving gel and it did not stack properly. The samples spread laterally as soon as they entered resolving gel. I have prepared fresh RIPA also, but the result did not improve. As a control, I have run my old extract in the same gel. but it did not leak. My labmates use the same reagent, but they do not face any problems. Previously I have thought that it may be due to sample overloading so I have reduced the volume. But it did not seem the actual cause. It would be very helpful if anyone inform me of the root cause of this type of sample spreading in the gel. Please find the images, attached below.
Hi everyone, I am doing 2D monolayer cell culture on HeLa cells and A549 cells purchased from ATCC. I have had some problems after a few passages that cells occasionally all become apoptotic-ish looking. The cell confluence is normally high in that case, and instead of a more elongated morphology like what they normally look like, the cells can become round and flat (I cannot distinguish the contrast between nucleus and cytoplasm using bright field anymore) within one or two days. I have attached a picture of the HeLa cells that went wrong/abnormal looking below.
The media I am currently using is DMEM Phenol red free.
Can someone please provide some suggestion on this wether it could be caused by infection, medium related issue, or related to sub-culture procedure.
Thank you very much!
I am using G418 to select for stably transduced 22Rv1-luciferase expressing cells. Would anyone know of the minimal effective concentration of G418 required to kill 22Rv1 parental cells? I am currently using 1 mg/ml in my media, but I'm not seeing much cell death.
I'm working with BJAB lymphocytic cell line, culturing them with RPMI media (with L-glutamine) + 10%FBS + 1% antibiotics. After two days of changing media by centrifugation, they started to adhere to the T flask and a monolayer has formed. Does someone know what could be happening? Is the media lacking something or should it be a step in the handling that I'm missing?
I'm focusing on L929 cells to collect the supernatant using for Bone-marrow-derived macrophage (BMDM) cell culture.
Our L929-Conditioned Medium was out of stock and I started thawing an old L929 stock (in -80°C fridge) (it is written 2014 but maybe even before that time).
I followed the protocol of our lab to grow the cells and collect the supernatant.
Our lab used that protocol for very long time and it still worked well.
However, now the BMDM cells which I cultured by using that supernatant grew not as well as normal, the morphology changed.
Then, one of my senior who has been working long with L929 cells said that my L929 cells after 100% confluent continue growing too strongly, which forms several layers of cells.
She said it was abnormal because based on her experience, after 100% confluent, L929 cells will be inhibit somehow to hardly multiple as many as that.
(The L929 supernatant stock I used before was from her and It did work well, now I started thawing new one. )
That might be the reason why my L929-conditioned medium was so poor in nutrition, and might be also the reason why my BMDM growth medium didn't work well as normal.
We think it is because of the degeneration of L929 cells.
Does anyone have any experiences with this? Please let me know. Thank you so much.
I am trying to culture dental pulp stem cell line from our storage. The cell line was preserved on 2019. However, the cells aren't adhere to the flask. I used DMEM with 20% FBS, 1% penicillin and streptomycin, 10mM L.Ascorbic, 1mM L.Glutamin. I tried different cryovial from our storage but none of them tend to adhere to the flask.
We are preparing a project to access a grant, but we are missing a lab that can help in the first phase of Proof of Concept and Safety.
The lab should be European (But not spanish, and sorry, but not UK) and be seasoned in brain samples handling, and also if possible having experience in cell morphology study (Neuron, Microglia, Astrocytes) or metabolic study (on said samples)
The project includes the development of a protocol in humans but we need to try it before on brain samples, and we are in a hurry to find a lab that can provide experience in this field.
Since we are in a hurry, you might contact me directly or respond in the thread.
Sincerely, J Vigil
I want to do some cell migration assays and I would like to know if it's possible to grow these cells in suspension
Suppose i got a cell line say for example (CHO) and i want to maintain it for my experiments, I'ld like to know what all the properties/characteristics I've to know about the cell line, I'm thinking few points like :
Doubling time, It's morphology, To know whether it is adherent/suspension cell line, It's Suitable media.
I'ld like to know what all other things I should know ?
I am doing a cell culturing experiment in a fluid flow chamber made of PDMS material. I was wondering how to sterilize the pipes and the flow chamber before/after the experiment, in case I want to use the same chamber for the second and third biological repeats?
Hi, I had a lot of problems with this cell line. When I count the number of cells, I can't do it, because the cells form a lot of cell aggregations and i can't separate them. The cells was incubated with trypsin for 2 minutes, and the cells were detached of plate, but the cells form aggregations. So, i have a lot of problems with the MTT assay due to the number of cells is not very real. If somebody can help me... I dont' know if the problem could be the medium, because i was used DMEM instead of McCoy...and i don't know if could be this the problem.
Using the recommended media and temperature to grow the cells but they die following the first sub-culture. When revived from frozen they look very healthy but as soon as they are split most of the cells die. Have tried different batches of FBS to no avail. Does anyone have any ideas?
I have been culturing MC38 cells for a few months and I am experiencing some problems. The cells aggregate a lot and generate clumps on the flask surface. I am using Trypsin to split them but it is really hard to de-attach them once they have created these clumps. I started splitting them 1/10 every three days and then switch to 1/20 to see if it improved. The viability tends to be low as well, around 50-70%. I am using DMEM+10%FBS as culture medium. If anyone has any suggestions and recommendations on how to culture these cells it would be really helpful. Thank you
I am currently culturing some HEK cells with absolutely no issue, they are being cultured with the standard DMEM + 10% FBS media.
For a particular experiment, I need to introduce exogenous insulin for a variety of different time points and I am noticing that my cells are dying within 5-10 min of insulin addition.
I am adding insulin in at a concentration of 10ug/ml. The cells are grown in a 6 well and all those that dont have insulin added to them are surviving but the addition of insulin is resulting in all the cells lifting within 5 minutes. If someone could provide some guidance.
I tried staining MCF7 and MB231 for Vimentin and Cytokeratin antibodies, and surprisingly, I saw both cell lines showed both markers. According to the literature, MB231 is a mesenchymal type cell and MCF7 is luminal, so shouldn't it be that MB231 shows more cells positive for Vimentin as compared to MCF7?
Recently I thawed couple of cell lines, RPMI8226 and JJN3 and they both didn't do well during my first passage.
I thawed the vials in 37 deg water bath for couple of minutes. Transferred the cells with media into a conical tube. Centrifuged the tube at the lowest speed for 5mins. Aspyrated the supernatant and resuspended the cells in 10mL media in T-75 flask for 3 days.
I use the following media:
Gibco RPMI 1640 with L-glutamine + Penstrep + Glutamine + 10% FBS
My percent live cells was very low when I checked them 3 days later. I am thinking of using 20% FBS next time I thaw cells. Is there anything else I can do differently to keep the cells alive?
I am culturing NK3.3 cell line. I do not have any experience with them. Unfortuately, they do not grow and I do not know why.
I am using AdvancedMEM media, 10%FBS, 1%P/S, 1% glutamine and I add IL-2 separately.(I aliquoted IL-2, so I heat my media in water bath, také out how much I need and add IL-2 directly there to avoid degradation of IL-2 while it is in the water bath. I dont know if this is the best solution?). I use100U/mL of IL-2.
When I count my cells, it has never been more than 250 000 NKs there (its been three weeks already, I should have more!!). When passaging, I leave 600mL of old media and add 2mL of new media with IL-2. I put the cells so that T-25 flask is "standing" with the cap up.
I passage them 2x per week. Whenever I count them there are quite a lot of dead cells visible.
Any idea on what am I doing wrong? Any suggestions/advice would be amazing.
Thank you so much!
I searched a few papers but nowhere could find the answer. Asking out of curiosity. I went back to the first few papers that talked about RAW 264, RAW 309 cells etc. also I checked on the official webpage of Abelson's Lab from NCI, NIH. I couldn't find the answer.
I am working with raw cell line from last one year and they were fine before .But since last month they keep on self activating.. ( without treating any activating agent like LPS) even after thawing during 1st passage their morphology changes completely from round to star shape.
I have tried using different stocks...
All other cell lines in our incubator are fine so I think its not incubator problem, even then i tried to clean it all over again but all in vain..
what should I try .. ??
I had a contaminated cell culture I suspected several things, including the media. I put the media (only) in a T-25 flask and used 2 agar plates to check bacterial contamination (one opened in the laminar air flow the other in the CO2 incubator) and left them is in the CO2 incubator at 37 oC (after I cleaned it),
two days later I didn't find any thing on the agar plates and no changes in color or clarity of the media in flask however I examined the flask under the microscope and found these things .. What could it be?
I have to treat cell line for 6 days with compound to see its deleterious effects. But the thing is within 48 hours cells got confluenced. So how to do this experiment? Should I just decrease serum concentration or any other way to complete this type of study?
I have been working with 3d spheroids for a few months. I started with HCT-116 colon cancer cell line and have not had any problems, this cell forms tightly aggregated spheroids. However, I have not been able to form spheroids from HT-29 cell line. I've tried different agarose concentrations (1-3%) and different cell concentrations (800-10.000 per well) and this cell line won't form the spheroids, just cell clumps. Does anyone have any idea what could be happening? I've seen in many papers spheroids from HT-29 and theorically they are formed easily. I use the following protocol for the HCT-116:
96 well plate flat bottom coated with 50uL of 1.5% agarose. I plate 2000 cells and centrifuge 1000rpm for 5 minutes. Then, I incubate for 4 days untill the spheroids are formed.
Thanks in advance.
Hello I am having a bit of an issue with my sf9 cells and trying to express the human PTH receptors. So:
I transform DH10 cells which give me blue/white colonies, which I also do a colony PCR to see if my insert is there and it is. I then proceed to do a bacmid purification (precipitate over night in isopropanol and 70% ethanol just before transfection) and transfect my cells on a 6 well plate. After 72 hours I harvest the media,replace it with 3ml new media and store the baculovirus in the fridge. 72 hours from that I do a western blot which shows my protein was there.
I infected 25ml of sf9 cells with 3ml of my virus on saturday at a density of 1x10(6) they had stopped dividing on monday (48 hours) and then today (72hours) I was going to harvest the virus. However my cells look incredibly stressed with lots of debris, but the media they are in looks perfectly normal, after centrifugation the pellet is normal coloured etc etc. I see no signs of contamination.
Even on the previous 6 well plate my cells looked incredibly stressed with lots of debris, but the media control was normal and insect control looked perfectly fine. I'm assuming I have a contamination that has led to this so I have filtered the virus and will run some more tests but any ideas on what is happening? I'm quite new to insect culture but assumed contaminations would be very apparent?
Thank you very much for reading!
I am currently using MRC-5 cell lines and they are at 17th passage. I tried to collect and count them via trypan blue cell counting but I could not see enough cells even though they looked fine in the microscope. I used almost 16 T-25 flasks but couldn't catch enough cells. What could be the problem?
I need expert's suggestions about my research in Oral Squamous Cell Carcinoma of cell line culture. I will measure the cytokines of P53, BCl2, FasL, and Casp 3 to know the mechanism of apoptosis in this cell line culture after treatment. Please, how to measure using ELISA kits for these cytokines. I am not sure to measure these cytokines from supernatant or cell lysate.
I am establishing a polarized gastric-epithelial monolayer culture on transwell system for bacterial infection studies. I use NCI-N87 cells and I culture them by replacing medium on every alternative day for 21 days. Later, I confirmed the expression of ZO-1 on 100% methanol (-20C) fixed cells. However, I face the following issues during this process.
1. How to avoid membrane curling while I cut off the membrane from the insert to mount on glass slide?
2. When I used 4% formaldehyde as a fixative to stain cell surface proteins, I found few cells or small cell clusters lying over the tight monolayer.
3. Is it necessary to use 21 days grown cells for bacterial infection studies? Because I see many highly cited papers also have used lesser days grown cells.
How to overcome these technicalities?
Any help is highly appreciated.
Thank you in advance.
I am having difficulties in reviving the chicken embryonic fibroblast cell line DF-1. I froze down the cells in complete DMEM medium plus 5% DMSO (half of a entire T75 flask) and thawed following the same procedures as other cell lines. However, even though the cells would attach on the next day, they started to die and float on day 2. I am thinking that the antibiotics (pen-strep) in the culture medium may affect DF-1 growth but shouldn't they only inhibit prokaryotic cell's protein synthesis (as for streptomycin)?? Has anyone come across the same problem?
Dear all, recently I used a TCR- murine T cell hybridoma 58-/- CD8+cell line to do the calcium flux assay. But Prof. Bernard MALISSEN, the creator of this cell line, told me the cell line can not flux much calcium. So does anyone know a nice TCR- murine T cell hybridoma TCR alpha-/beta- CD8alpha+/beta+ cell line which can get a robust calcium flux when I use anti-CD3(145-2C11) and anti-IgG to trigger them? I need to transfect BglII/SalI-linearized pBJNeo-TCR alpha and SalI-digested pSH-TCR beta into the cell line and get a stable T cell transfectant.
Thank you very much!
It seems that resazurin-based viability assays (such as Alamar Blue) are frequently used with hepatocytes or hepatic cell lines cultures, but the effect of resaruzin on the induction of cytochroms P450 on the samples is barely discussed. As these tests are usually non-destructive, it is appealing to use the same sample for the viability assay then CYP induction analysis, but I can't find an actual proof that resazurin won't affect the CYP activity. Does anyone have feedback or insight on this question? Thanks a lot!
I would like to add some guanine, cytosine and adenine, independently, to cell culture. I can't dissolve them in DMSO or in water, even turning up the temperature until 95°C. Only cytosine dissolve at 55°C but precipitate again when the temperature goes down. On the specification sheets of the products, it is written they are soluble in HCl.
- HCl 0.5M for cytosine up to 100mM
- HCl 0.5M for adenine up to 20mg/mL
- HCl 5M for guanine up to 25mg/mL
At the concentrations I would like to use them on the cells, even trying to prepare these bases at a 1000X concentration, two problems come to me:
- I'm not even sure they can dissolve in the appropriate volume for the 1000X concentration: the mass/volume being higher than the specifications
- I tried to dilute, even the lowest HCl concentration (0.5M), a thousand time in my cell culture medium and measured the pH with a pH paper --> pH came down to 6. This is not suitable for cell culture.
Do you have any idea how to dissolve these bases?
I would like to work on THP-1 as adherent cells without changing its natural. Is there any method or protocol for develop adherence cell from suspension cell lines?
Regarding price issue, I want to use NBCS instead of FBS for the culture of THP1 cell lines. If any body has experience (good/bad), please share it.
I need to culture mammalian cells on a 60mm culture dish in an airtight container with 5% CO2 gas (5mL growth medium).
Are there any rules / published data on the ratio of medium : atmosphere that is required for optimum cell growth in closed environments? E.g. 1:20 ratio maintains stable gas concentrations for 24 hours.
Also, are there any publications describing the effects of closed culture on mammalian cells?
So recently I started working with HepG2 cell line and I'm having the hardest time to evaluate their confluency due to their morphology and the formation of aggregates, so I was wondering if anyone can tell me what's the confluency of the hepG2 in the photo and if I should sub-cultivate them in this state.
I want to analyze apoptosis using AnnexinV/ PI. I am working with various neuroblastoma cell lines such as SKNAS, SHSY5Y SKNBE(2), Kelly and several other.
I am using the FITC-AnnexinV/PI kit from BD.
The assay worked always fine when I analyzed apoptosis in SKNAS.
However, when I used the same protocol for SKNBE(2), I always got approximately 80% Annexin potitive cells. And these cells were not treated- thus, these cells were healthy cells that should not have more than 5-10% apoptotic cells.
Today, I analyzed apoptosis of Kelly and SHSY5Y cells. Here, I also got 70-80% Annexin positive cells in untreated cells.
Might the cell membrane of these cell lines have phosphatidylserine in the outer leaflet of the plasma membrane even if cells are not apoptotic?
If so, the assay would not work for these cell lines...Did you have similar problems when using this assay or read about it?
This is my protocol:
- Transfer medium to 1.5 ml tubes
- Wash with 300 μl PBS and transfer PBS to 1.5 ml tubes
- Add 200 μl trypsin and incubate 3 minutes
- Use the medium to inactivate trypsin
- Transfer the cell suspension back to 1,5 ml tubes
- Centrifuge at 200 g 5 minutes
- Resuspend cells in 500 μl PBS (2 wells are merged)
- Centrifuge at 200 g 5 minutes
- Resuspend in 100 μl Annexin V binding buffer
- Add 5μl FITC-Annexin V and PI
- Gently vortex cells and incubate 15 minutes at RT in the dark
- Add 400 μl 1x binding buffer to each tube
- Analyze by flow cytometry
PI: Laser 561 nm; Filter 670/30
AnnexinV: Laser 488nm; Filter 530/30
- unstained cells (to set gates)
- PI only
- AnnexinV only
Dear colleagues, currently I am doing an inflammasome-related experiment using cells. I have a substance as a candidate for anti-inflammasome. However, I am a little bit confused about which step I should administrate my candidate regarding the inflammasome molecular pathway. Is it better after primary inducer (LPS) and before secondary inducer (ATP) or simultaneously administrate with the secondary inducer (ATP)? I beg your suggestion and further consideration. Thank you.