Questions related to Cell Adhesion
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 culturing MSCs on 3D-printed hydroxyapatite scaffolds. We need to detach cells from the 3D to analyze/quantify overall DNA content using Quant-iT PicoGreen dsDNA Reagents and Kit. However, these protocols have not been tested or adapted for complex 3D cultures. We aren't sure what the best method would be to detach & cells from the 3D scaffold and lyse the cells. We also need a technique to verify that our adapted method is effective. I'm interested in hearing what techniques/protocols others are using or any recommendations. Thanks!
Our currently drafted protocol, which is subject to change, involves the following steps:
1. Get DNA standard lysates using PureLink Genomic DNA Kit of cells prior to seeding.
2. After culturing cells seeded on 3D scaffolds for _____ days, at different timepoints, transfer the scaffolds to new wells in 24-well plates so that cells adhered to the wells are excluded.
3. Add TrypLE to the scaffold wells and incubate them, on an oscillating shaker to promote detachment, for 20 minutes.
4. Collect the trypsinized cells and transfer to centrifuge tubes.
5. Add TrypLE to the scaffold wells again and incubate for 10 minutes. Then, repeat collection & transfer of trypsinized cells.
6. Do a 2x rinse using trypLE to try to "knock off" remaining cells and collect as many cells as as possible from the matrix. Repeat until the TrypLE collected is clear, not turbid, hinting that there are little cells remaining in trypsinized suspension.
7. Centrifuge the trypsinized cells to isolate the cell pellet.
8. Resuspend cells in PBS.
9. Follow the protocol in the PureLink Genomic DNA kit to prepare unknown content of DNA in the cell lysates.
10. Follow the Quant-iT PicoGreen Kit protocol to complete the reactions & quantify dsDNA in the samples.
I'm newly working with MDA-MB 231 cells. I have sub-cultured cells using 4. 0 mL L-15 media with 10% FBS in a T-25 culture flask. The cells have incubated them incubator at 37°C, with (0 % CO2, recommended company for L-15 media). After the 48 incubations, I checked the cells under the microscope, and the cells were dead. I checked the flask also, and some of the white precipitated parts were attached to the flask. For reference, I have attached the cells Images. This is the 4th time I face this issue and I cannot figure out why. I would appreciate any suggestions/tips on what I might be doing wrong. Thanks in advance!
Articles have only mentioned that cell-laden hydrogel scaffolds were lyophilized before SEM analyses for cell adhesion. However, no details were mentioned.
Hello. So I had seeded some 48-well plates with 70,000 cells/well (human osteoprogenitor) on a calcium phosphate substrate scaffold.
After 24 hours I wanted to study the overall attachment so basically I incubated I removed the scaffolds from the wells and placed in to a new well plate.
I washed with PBS and then incubated in a trypsin 1x solution for 30 minutes to allow for the cells to deattach from the scaffold.
I then neutralized with media and centrifuged the solution to form the smallest dot sized pellet.
I removed the solution and reconstituted in 1 mL of media and then using 10 uL of the solution, mixed it with 10uL of trypan blue.
Transferred 10 uL to side A and side B of a chambress counting slide and used the corresponding automated cell counter by life technologies to count the cells.
I somehow got a bigger number - for example 4.5x10^5 cells alive/ mL
The time point was only 24 hours so it’s not possible that the cells divided that quickly and multiplex in that number so fast.
Can someone please help me understand the principles behind automated cell counting because I believe the machine maybe possibly multiplying by a factor to estimate the number of cells? Please help because clearly the machine won’t count cells which aren’t there, I just don’t understand why it’s spitting out such larger numbers.
Please it’s my last experiment of my thesis and I just need a little help please.
I am culturing Jurkat T cells E6.1 obtained from ATCC. This cell line is known to be in suspension forms. The cells were expanded as as suspensions for the first few days after reviving but then became adherent. The culture media used for culturing is made by ATCC protocols (RPMI + 10% FBS). Please let me know if anyone also experienced this and if there are any suggested treatment. Thank you for the help!
I'm inducing M0 macrophages from THP-1 by using Phorbol 12-myristate 13-acetate (PMA)
But frome time to time, I encounter induction failure (twice now), which could be fixed by new PMA stock.
I would like to have some suggestions on storing PMA?
Currently I preserve 5000x and 1000x stock,
5000x stock dissolved in DMSO, and 1000x was diluted from 5000x with PBS.
All stock was store in -20°C.
Is there any suggestions about how many freeze-defreeze cycle PMA could encounter, or if PMA can't left on ice for too long?
I have been bringing up SH-SY5Y cells for a year now and have never had this cell death problem. The cells are cultured from frozen cells using full growth medium (100mlDullbeccomedium, 10ml HamsnutrientF12, 10ml FBS, 1ml sodium pyruvate, 1ml PenStrep, 1ml Glutamax). I believe it may be an issue with the media created, however I have made fresh media from fresh bottles and have the same thing happening. Has anybody else experienced cell death like this?
Cells begin to adhere and then by day 3 a criss-cross mesh seems to spread across the entire flask with cells dying (round cell morphology). See image below.
Any ideas would be great!
We have to use a non-enzymatic cell dissociation to detach cells and EDTA/versene is taking far too long.
What concentration of sodium citrate is usually recommended to detach cells?
I am a physical chemistry PhD student currently working with 3D cell culture on polycaprolactone UNTREATED scaffoldings, for NMR and MRI purposes only.
I am not particularly interested in cells physiology, althouh I understand this is important to optimise cell adhesion on the scaffolds I plan to do imaging on.
I am working with mesenchymal stem cells, which differentiate into osteoblasts after the seeding.
My scaffolds are disc shaped, 1.5 mm thick, kept in 96-well plate. I seed each scaffold with a 20 uL drop of cell suspension, topping up the wells with media after 3-5 h of incubation.
I do know I can make the scaffolds more appealing to the cells by coating them.
However I wondered:
- can I make the bottom of the plate LESS appealing for the cells, in order to convince them to attach to the scaffold from the get-go?
- would that damage the cells, as they sink to the bottom for gravity and they find an unconfortable environment to attach to? and
- could that be avoided by keeping the suspension gently agitated so that cells have more chances to come in contact with the scaffold?
I tried UNTREATED wells too, but cells seem to like those anyway, no difference with the treated wells.
Control wells in 96 well plate does not show any absorbance at 570nm
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.
I have isolated mitochondria from tissue. I would like to check them by fluorescence imaging. Can I use Mitored dye for these isolated mitochondria?
Hello to all,
Lately the HEK293T cell line I am using does not adhere very well to T75 and T175 flask. I wanted to analyse possible factors and conditions that could affect adhesion (trypsin concentration, medium...) and I wanted to ask you, if you know of any way to quantify adhesion. Maybe by measuring the cells in suspension, or give some kind of shock or vortex to the T flask and do a count to see how they have adhered and withstood the impact... I don't know.
I would appreciate your ideas.
Thank you very much,
I am currently trying to culture murine fibroblasts and HaCat cells from a cryo-frozen back-up. The murine fibroblasts are at passage 3 and the HaCat cells have not been passaged priorly.
After thawing, I have seeded them onto a T-25 vial with 10% FBS enriched DMEM with antibiotic , L-glut and NEAA at 37^C and 5.5 -5.7%CO2.
Media was changed 24h after culture initiation- only a handful of cells (of either variety) had adhered to the plate. Further, media has been changed every 48h/ 72h, depending on the state of the cells. They still look highly stressed and do not show cell-specific morphology (21 days, today)
I do not have any further back-up of the cells left, nor funds to procure more vials. So I desperately need these cells to survive.
Would be grateful if anyone could suggest ways to ensure survival (propagation) of this batch.
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.
We’ve started to work with this cell line, and it is driving us crazy. We are unable to making them attach to the plate and when we do, we see a huge amount of apoptosis and cell death.
The coating is performed with PDL (50 ug/mL) and laminin within a range of 6 to 12 ug/mL. Those parameters are what it’s written in most of the limited bibliography that exist about this cell line, so we are unable to find what’s the problem.
Has anyone experienced the same problem as us? Did you manage to solve it somehow?
The pics are showing how our cultures looks like in bright field microscope from 2 – 5 days after passage approx.
Thank you in advance!
I am recently started working with THP-1 cells and still in the phase of gaining experience in handling of THP-1.
Current obstacle I am facing is fixation. After differentiating THP-1 to M1 macrophages, I am trying to do fixation and ICC, but during 1st wash step during fixation most of macrophages are lifted and washed away. I have tried to fix the cells using several ways with no improvements.
Any suggestions on where I could look at the problem? Thank you in advance.
(I have added images of cells before and after wash step)
- The protocol I have used for M1 differentiation is: 48h incubation with 25ng/ml PMA, 48h resting incubation with serum free RMPI media and 48h incubation with 25ng/ml LPS and IFNy.
- I also checked M1 cell viability by briefly adding live/dead staining solution into culture before wash step and found out macrophages were still alive.
Dear readers, I am a master degree student currently working on phototoxicity assessment using Balb/c 3T3 cell line. However, I have encountered some technical issue during the running of this assay, for both UV- and UV+ exposed well plates. The cells (passage number 20, cultured in DMEM + 10%Calf serum +1% Pen-Strep 5% CO2, 37°C ) get rounded and detach after the first and in particular after the second washing step required by the test guideline at day 2 after the 1h and 50 min incubation with the test substance (1h in incubator and 50min in dark or under UV-light exposure). I would underline that, in my case, the test substance consist of only 1% DMSO in HBSS (Mg+Ca+) because the issue of detachment occurs even with this vehicle (we have tried also with just HBSS but we obtained the same issue).
Do you have any clue on how to troubleshoot this problem? Thank you in advance.
- 96-well plate NUNC 96-well plate (#167 008) (I have tried also with poly-lysinated or collagenated plates, both were unsuccessful)
- Buffer for washing: warmed HBSS (Mg+ Ca+)
- 8-pin manifold to remove buffer (we have tried also to invert the plate over a absorbent paper, but was unsuccessful)
- Serum used: Calf serum (we have tried also with FBS or with combination of Calf+FBS, both were unsuccessful).
Currently i work with metatranscriptomic data from environmental samples and wanted to have a look at the expression of genes involved in adhesion to surfaces. The genes are annotated with KEGG, eggnog and InterPro, but i struggle to find relevant genes which i should have a look at and the respecitve identifiers.
Does anyone has experience with that?
Are there any types of cylindrical cell culture inserts that are designed to confine cell seeding area? Say, if I'm using a 12 well plate (3.5 cm^2/well), but only want to seed cells in the middle 1.9 cm^2 area. If so, are there any reusable insert options or only disposable ones? Thanks in advance.
Hello, I am new to biofilm research and have a question about growing and measuring biofilms.
We want to research the effect of a type of enzymes in breaking down biofilms in showers, baths, hot tubs, and hot springs.
Therefore we must devise a way to grow and measure biofilm that is close to natural biofilms in showers etc.
Are there any existing protocols or any other researches that may be useful?
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.
Has anyone came across an extracellular-tagged cell adhesion molecule.
Ideally in the axon and ideally in the growth cone such as L1CAM or NCAM.
I would like to tag the extracellular domain and need some advice on signal peptides and correct protein translocation.
I am very appreciative for any info on this matter.
Thank you very much
I have HBMEC cells that are not sticking to the plate. I already increased the fetal bovine serum concentration, but nothing worked. I have already collected the cells that did not adhere and transferred them to a new plate with a higher concentration of fetal bovine serum, and even then it does not get successful. What could be happening?
Hi Guys I am facing problem as my MDCK cells are dying when I stain after the plaque assay is done. I do the infection in 1X MEM, 0.2%BSA, with TPCK 1µg/ml, sod bicarbonate, HEPES). i tried to change many things, including the cells, BSA and AGAROSE etc. Still my cells are dying. I make sure Agarose is not too hot. I don think this is the problem. The cells are mostly dead from the sides even in the highest diluted wells. After changing the agarose, the plaques were visible though, but the cells were anyways dying like the previous observations. I had no problem doing plaque assay in my previous lab.
Please suggestion suggestion would be helpful.
I am conducting a experiment where I am attaching fibroblast cells to a PEDOT:PSS film surface. Unfortunately the cells are more attracted to the surface of the 24 well plate in which the PEDOT film is located.
Is there a simple coating I can apply to the PEDOT film to increase cell adhesion or is there a coating I can apply to the 24 well plate to inhibit cell adhesion.
I have looked at using bovine serum albumin or Angiogenin, but using proteins are very expensive to coat surfaces in. So far poly-l-lysine seems like the front runner for increasing cell adhesion to the PEDOT film.
I would appreciate any other suggestions.
Can I use 0.2% Gelatine coated, glutaraldehyde cross-linked glass coverslips as an alternative to Poly-L- lysine coated coverslips?
I tried plating the same batch of cells on a 24-well plate and a cell-stack 2. Both surfaces are tissue culture treated and coated overnight with laminin. The cells plated very well on the 24-well plate, but were loosely adhered on the CS2 and are falling off as I move the CS. Any ideas on why this would be? I'm not sure if the sheer forces of media in the larger format could account for the poor adhesion. Thanks for any thoughts!
We have cultured MCF-7 cancer cells in microfluidics several times and incubated them for 24 hours, but as the nanomaterial passed through the cells, the cells moved and no cell remained in the canals. How can cell adhesion be increased?
Can cells be washed with distilled water instead of phosphate buffer solution (1x)?
The secretion system of Escherichia coli is important for the adherence and invasion of epithelial cells. I need to find out a way to measure the interaction of Escherichia coli with mammary epithelial cells in mammals. Could anyone indicate a simple, standard, and straightforward epithelial cell adhesion (and invasion) in vitro methodology? I was wondering what types of methods people usually use in this situation. Thank you very much.
I want to siliconize my cell culture bioreactor vessel, and buying sigmacoat is a little expensive by considering the other materials which I need and my budget. So I was thinking if it's ok to use dimethyldichlorosilane for siliconization of the vessel? also do you know any alternative material or method for this purpose?
I want to prevent cell adhesion from my product surface, just like the 384-well plate. The product was a 3D printing product with resin, which is hydrophobic. The challenge is the improvement of the surface to form cell spheroids under a constrained environment. PEG is a kind of ultra-low adhesion material, but I'm wondering how to apply it.
Hi I have searched a lot about finding a polymer or any material or peptide to have good cell adhesion properties and also have thermal resistance properties that could be sterilized in autoclave. I haven't got answer yet. Maybe I missed something in my search. I found in one article that RGD peptide could resist 120 C temperature without losing its cell adhesion properties (RGD Surface Functionalization of the Hydrophilic Acrylic Intraocular Lens Material to Control Posterior Capsular Opacification). I hope someone have experience in this issue
I'm working with large pore (800 um) titanium samples. After seeding, and waiting 30 min. for cell adhesion, most of cells attaches on polystyrene plate. Considering that, the evaluation of alkaline phosphatase is not valid in my opinion, since we aim to evaluate differentiation of cells adhered to the sample. If we seed cells and after seeding change the sample to another well I will not have the same amount of cells every time I perform a new experiment.
How could I solve this problem?
Actually I am confused about the activation of CCR2.
1. I would like to know that does CCR2 need to be activated for the binding of MCP-1; for example, integrins in leukocytes are needed to be activated to be bound with the adhesion molecules of the endothelium?
2. If not, then can MCP-1 stimulates monocytes to express CCR2 in higher extent?
What do you think regarding this?
Would you like to share your idea?
We knew that inflammasome event induced by LPS and extracellular ATP could lead to the cell pyroptosis and death, but what about the effect of only ATP administration to the cell culture? Is it also toxic and induce cell apoptosis? Currently, I am doing an experiment regarding the inflammasome mechanism in cell culture. I observed that the only ATP triggers loss of cell adhesion and morphological changes. I am just wondered whether it was normal or not.
the cells are recovered just a week ago, and the cells on the plastic dish are growing well. But once the cells transferred into the new bought glass dish, even without the further treatment of cell synchronization, the cells cannot get adherent to the bottom of the dish well, and can be easily washed away by fresh PBS.
Sphere formation is known among normal or cancer stem cells. But whats the underlying mechanism of cells to come together and make sphere?
How to estimate more precisely the real surface area relating to different degree of roughness?
It is known that coating pluronic F127 inhibit cell adhesion on PDMS or glass. I just curis about the mechanism of the suppression. Thanks in advance!
We are culturing BJ fibroblasts in MEM on TPP yellow culture plates (10 and 15 cm). Recently our cells started to proliferate at a much lower rate than usual, and we checked that the problem was not coming from the incubators, medium, serum... Even freshly thawed cells eventually stop proliferating after 2 passages. When routinely looking at the cells under the microscope, I noticed that the number of dividing cells in the plate was normal, the same as you would expect if the cells were proliferating at their normal rate. However, many mitotic cells were detached and floating in the medium (particularly in telophase). I suspect a problem with the coating of the plates (I mean the normal surface treatment performed by the manufacturer, we don't additionally coat the plates ourselves for culturing the cells), but culturing the cells on different TPP plate lots didn't solve the problem. However, the cells grow normally in 96- or 24-well plates... Have you dealt with similar issues with BJ cells or other cell lines? Or had a problem with culture plate adhesion? Any feedback or help would be greatly appreciated!
Hi, I have grown adherent cells on glass coverslip and would like to use them for Immuno Fluorescence experiment later. I would like to know if it is possible to dehydrate the cells gradually using different percentage of ethanol and store them for now, which I can fix later and use. I heard that fixing the cells and then storing them might lead to detachment of the cells if the fixed slides are more than a week old. Can someone help with the protocol, if any please?
Hi, folks! I am currently working on MB231 culturing in 96 well plate for fluorescence study. The thing is when I wash the cells with PBS, some cells start to shrink and become a chunk; meanwhile, some cells are lost (as from microscope clearly cells become less than the time before washing). I check the research gate, people only mentioned Caco cells having the similar problem, so I am very sincere and eager to find out what is going on with the cells. My senior told me when they reach a certain confluency they become easy to shrink and flip apart from the well-plate. But my judgment is there was only 60% confluency when I was doing the washing process. So Could there be some other mysteries which are beyond my knowledge?
So basically what I did was I just gently pipetted up the medium and added in the PBS. I even tilted the well-plate to a certain angle and make sure the pipette tips never touch the cells.
When I cultured cells on PEGDA/GelMA hydrogels (75%PEGDA/1%GelMA), cells can't adhere to the gel surface. Is it because there is too much PEGDA making the surface hydrophobic and GelMA isn't enough for cells to attach?
I have been using an EVOS fluorescent microscope to look at my dental pulp stem cells every day. To monitor cell attachment, I look at how many cells seem to be resting on the steel substrate that I plated them on. This is clearly a qualitative method, but is this reasonable for assessing cell attachment?
I want to use centrifuge assay to quantify cell adhesion strength to substrate. In one review, there is a equation to calculate the body force exerted on each cell
F = ( density of cell - density of medium) * Volume of cell * RCF
Does this mean when I centrifuge to detach the cells, the cells are still in medium?
In one reference I find, they remove the lid, cover the plate with sealing tape, then centrifuge. If they didn't leave medium in the plate, they don't need to seal the plate right?
Please share with me your experience. Thanks so much
We want to stimulate the lymphatic endothelial cells (LECs) with Ang2. We will seed the cells on glass cover slip and then stimulate with Ang2 for sometime. After stimulation we will fix the cells and check the cell-cell junction proteins such VE-cad, ZO-1, Claudin5 etc. How long we should stimulate the LECs?? Is there any article regarding this?
Hi everyone, I'm encountering a problem in fixing the cells after the Glyco and the Mito stress assays in Seahorse XFe96 analyser. I start with monolayer of 100% confluent before the assay and after the assay most of the cells are detaching from the wells. And that eventually this affects the measurements of OCR and ECAR. I'm working on adherent cell line. Did any of you noticed this before?
I use EGTA/EDTA solution (in HBSS-) for 3 min at room temperature for the detachment of monocytes from the HUVEC in a cell adhesion assay. After collecting the detached cell aliquot, I observed the well-plate under microscope. It was seen that so many monocyte cells were attached to the HUVEC, the detachment was not complete. What would be the possible reasons for the incomplete cell detachment? Actually, I just want the monocytes, no HUVEC contamination.
Please provide your important suggestion regarding this.
I want to prevent cell adhesion from my polymer surface. What is the best way to treat the surface. The challenge here is to keep the surface hydrophilic.
Note the surface is cellulose.
I'm using MG-63 cell line for cell adhesion study on bioactive glass scaffold and it is non-porous. PLA and PDLLA are hydrophobic in nature so it will not encourage cell attachment. But my sample bioactive glasses contain salts which are hydrophilic. So what could be the possibility?
Failure in cell detachment after trypsinization!
I use trypsin to harvest my cells (4T1), but they don't detach after trypsin incubation. I usually use 5ml of trypsin per 75cm2 T flask and incubate for 7 mins at 37C. Has anyone encountered the same problem? How did you solve it?
We are working on adipogenesis using various in vitro models. One of them is 3T3-F442A cells. In the literature, there are numerous protocols for differentiation of these cells. However, mostly they say that they start differentiation using differentiation media 2 days after confluency as in 3T3-L1 cells. We tried the same protocol but the cells detached from the dish surfaces at day 2 after confluency. What would be possible reasons for this undesired situation?
We purchased the cells from Sigma and cultured as it was recommended. We used newborn calf serum as a supplement during propagation and never allowed cells to reach over 80% confluency.
We tried corning cell culture treated 35mm, 60mm and 100mm dishes along with 6-well plates.
Thanks in advance.
For a microfluidic device of mine, cells are sticking to a PDMS surface (not oxidized). Are there any surface treatments that can be used to reduce this effect? It may also be that the surface of the PDMS is not perfectly smooth. In that case how can I ensure the PDMS being spin coated is smoother?
how to predict homophilic and heterophilic interactions for cell adhesion molecule. Is there a software that predict homophilic or heterophilic interaction from protein sequence ?
I have prepared Hydrogel desks (6 mm) and placed then in 24 well-plate, afterwards, I seeded MSCs cells on top.
there is no cells been attached to the hydrogel desk?????
- cell density was 50000/well.
- I could not observe any cells adhere on the hydrogel desks?
I would like to perform a battery of anxiety, depression and social interaction tests on the same animals that have a deletion in one synapse cell adhesion gene but not sure if I should do it in the same day or in a particular order (open field test, elevated plus maze, social interaction test, tail suspension test, light-dark box). Could anyone help me with that?
Thanks a lot in advance!
I'm trying to make an adult (!) primary astrocyte culture from mouse cortex. I am doing this with a very experienced person who has done this before in a different lab. I am using her reagents and protocol, and at the last stage before plating we are using a viability assay and automatic cell counter, we have a lot of alive cells (around 70%) and then we are plating them at the density suggested by her protocol. All of this seems to be going well.
But then, if I check 4h or 24h later, none of the cells attach. Sometime I see very small cells attaching which I guess are bacteria (looks like it). Here are the things I tried: Poly-L-Lysine coated cell culture flasks (the "normal" T25 flasks), Poly-L-Lysine coated dishes, Fibronectin coated flasks, uncoated flasks, Poly-L-Lysine coated glass coverslips, Poly-L-Lysine and Laminin coated glass coverslips.
Nothing attached to any of these surfaces! I also tried different media that were tried and tested and used in other labs (Astrocyte Basal Medium and DMEM-based recipes), but I think the problem is rather in the attaching.
Other primary cultures in my lab are doing fine on the PLL coated surfaces, so there doesn't seem to be a problem with the coating itself (I read PLL can be toxic to neurones if not done properly).
We are all out of ideas.
I know I haven't given details about my culturing protocol, I think that is fine, so did anyone ever have an overcome this problem with attaching? Or do you think it can be due to the medium after all?
I am doing my master of science project and facing a cell culture adhesion problem. In my study, hydrogel scaffolds are made of polyvinyl alcohol and prepared by freze-thaw method.
Is there any suggestion in this field???
I have found that my cells, NIH3T3 fibroblasts, do not do great on gelatin. The number of cells that adhere or spread after 24 hours and even further are far less on gelatin (5 and 10%) than on chitosan (2 and 4 %). The expected outcome is generally the opposite. All the gel samples were given same treatment.
Any suggestion is appreciated!
i use head and neck cancer cell line. using media DMEM-F12 with 2%FBS, 2%N2 and 1%B27. my seeding density is 20000/well in 1.5 ml media.
We use coating media consisting of fibronectin, collagen and BSA on dishes overnight before growing cells. For microscopic studies like IF these cells are not sticking well to the plastic slides, coverslips etc. After the treatment cells lose their morphology and become rounded. Does anyone come across this problem?
Even for DCF staining im having an issue with them. I used 10uM of DCFDA for 30min but it did not work. Can anyone suggest the concentration of DCFDA on Beas2B cells?
I'm doing apoptotic analysis (Annexin V - PI) by flow cytometry on serum-starved HeLa Kyoto cells treated with different combinations of growth factors to induce apoptosis/necroptosis. I use Tryple/EDTA solution to detach cells from the dish rather than using the conventional trypsin/EDTA, and I will call it 'trypsinization' in the remainder of my post.
The serum starvation procedure itself makes the cells attach to the dish very strongly and therefore the trypsinization procedure takes longer than usual. Regularly growing HeLa in serum-containing medium detach quickly in 5 min. If the total time the cells spend in serum-free media is 1 day, the cells detach in about 15 min. If two days, then it takes almost half an hour to detach. And I don't have a problem with that. My problem is with the cells treated for induction of apoptosis/necroptosis because they form unbreakable clumps during trypsinization despite the fact that I don't shake or hit the dishes during their detachment. On the other hand, only serum-starved cells detach without forming any clump. So I think this clumping issue has to do with the cells being apoptotic/necroptotic in my experimental setup.
What I do is I first remove the medium from the dishes and collect all the floating dead cells, then wash with PBS w/o calcium and magnesium, then I add the Tryple/EDTA solution. Once detachment is complete, I add serum containing medium and pipette up and down. But it seems like I won't be able to break up the clumps without killing the cells. Do you think some DNA from the floating dead cells are not removed completely and causes the cells stick to each other after detachment? Shall I wash the cells with 1% BSA in PBS before the regular PBS wash step to make sure that all crap is removed?
Clumping upon trypsinization occurs during regular culturing of HeLa growing in serum-containing media but one can very easily break up those clumps. My problem is exactly with apoptotic/necroptotic cells under serum-starved conditions. I fear that I'm selectively losing certain populations into those clumps.
Any help will be greatly appreciated!
I'm produced collagen scaffolds and am planning to do some AFM (in liquid) to understand their mechanical properties. However I am struggling to immobilise them on a surface. I have tried Cell Tak and super glue but both seem to be absorbed by the scaffold. Does anyone have any ideas?
Thanks for any help!
Carboxyl-anhydride and amine plasma coating of PCL nanofibers to improve their bioactivity
Free download via link below or please contact me.
The plasma modification of biodegradable nanofibers is of great interest for improvement of their biocompatibility. However, there are no systematic studies regarding the influence of plasma polymer deposition onto the surface of nanofibers to improve cell adhesion. In the present study, homogenous and reproducible modification of polycaprolactone (PCL) nanofibers by amine and carboxyl/anhydride groups was achieved. The concentration of amines NH2/C and C(O)O contribution were up to 2.9 and 14.1%, respectively. Regardless the plasma conditions, the deposition of amine and carboxyl-anhydride plasma coatings onto the PCL nanofibers sufficiently improved the cell adhesion and viability, as was evidenced by microscopy observations and ATP assay results. It should be emphasized that the deposition of negatively charged carboxyl-anhydride coatings resulted in slightly better cell adhesion compared to the positively charged amine plasma coatings, unlike the widespread opinion that COOH modification has less effect on myoblasts adhesion
We seem to be having trouble getting 30-70 kDa poly-L-lysine (Sigma) to remain well bound to a glass coverslip. Cells (protists in this case) won't stick if the coverslip is rinsed multiple times with PBS, or even twice with 1% heat-inactivated serum in DMEM. In fact, the serum protein makes the poly-L-lysine "ball up" into ~2 micron spheroids on the surface, suggesting that it's not well bound.
We've tried a few things to solve this:
- from 1-10 mg/ml concentration poly-lysine in water
- 1 mg/ml dissolved in 50 mM pH 8.5 borate buffer
- drying the poly-lysine onto the coverslip before rinsing
- Corning versus Fisher sourced coverglass
In none of these circumstances do the cells remain well adhered, and the spheroids always form if the rinse contains protein. The odd thing is that one year ago (and one lab move) it worked acceptably well. Glass formulation has been suggested as a possible cause, though the Corning product numbers are the same before and since.
We would appreciate any thoughts, coating recipes, or other suggestions.
If you know of another non-specific adhesive surface for protists on glass, that too we'd love to know!
I'm having trouble getting chemically (Lipofectamine 2000) transfected cells to grow on glass cover slips. I transfect on a 10cm plate and after 8 hours split and seed 1x10^5 cells on glass cover slides. Instead of seeing happy growing cells. I usually see unhappy balls, that are attached and growing slowly. I have been trying this with both HEK293 and HCT-116.
I'm guessing maybe seeding, then transfecting and then fixing and probing the cells might be my best bet....
Is polylysine coating needed?
Anyone have a great protocol/advice for growing, fixing and imaging transiently transfected mammalian cells?
I have no trouble with the transient transfection, I usually get beautiful protein expression on normal tissue culture plates, even after splitting them. I normally use HEK293 and 293T cells.