Science topic

Cell Adhesion - Science topic

Adherence of cells to surfaces or to other cells.
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Hello everyone,
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
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The quality of analysis in terms of detection of surface markers using flow cytometry will be determined by the cell detachment method.
The commonly used methods of adherent cell detachment include enzymatic and mechanical treatments. Enzymatic detachment, however, can lead to significant changes in cell membrane protein structure and composition leading potentially to significant experimental bias.
Accutase which is an enzyme mixture with proteolytic and collagenolytic activities is usually considered a less damaging agent than trypsin and is recommended for the treatment of sensitive cells as well as for analysis of surface markers using flow cytometry.
On the other hand, detachment by the mechanical method is achieved using the so-called ‘rubber policeman’ which is a rubber or plastic scraper attached to a glass rod. But again, mechanical methods can lead to cell membrane damage and to substantial changes in its structure.
Thus, choosing an inappropriate detachment method may cause unintended effects biasing experiment results. So, I would suggest that you conduct a pilot experiment to assess the optimal cell harvesting method (either accutase treatment or mechanical detachment using a cell scrapper) for your cells to avoid experimental bias and obtain reliable results.
Best.
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Hi Everyone,
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.
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Doesn't seem like a contamination, but more of a poly(propylene)fiber filament, a result from plastic injection of pipette tip during production. We usually ignore it, through centrifugation it will disappear, eventually. Else keep an eye on the tip, if one showed with filament seems going to detach, please discard it and choose a new one.
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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.
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Benjamin Fournier I was considering doing so but I wasn’t sure if cells should/could be lysed straight off the scaffold without detaching first. I am concerned that I won’t extract all of the DNA if cells are trapped on the internal area of the scaffold. I will have to make sure that each internal pore/cavity is exposed to the lysis buffer but I believe it should work. Have you ever tried to lyse cells directly from a scaffold? Thanks!
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Hello,
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!
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hi,
the white layer on the flask is debris of dying cells. Are you sure, that the flasks are Tissue culture grade/ coated for adhesion cell culture? For me it looks like an attachment issue and all non-suspension cells die fast in the wrong type of bolltes
Sven
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Articles have only mentioned that cell-laden hydrogel scaffolds were lyophilized before SEM analyses for cell adhesion. However, no details were mentioned.
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Samples were fixed by adding 5% glutaraldehyde solution overnight which was replaced with a fresh sterilised solution of PBS and changed three times before soaking in fresh sterilised deionised water for one hour twice. Then, the CCC samples were frozen at −80 °C for three hours before placing them into a Christ ALPHA 2–4 freeze-dryer for 24 hours.
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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.
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Dear researcher
See the following papers method
Cell viability analysis using trypan blue: manual and automated methods
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Hi everyone,
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!
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One way to avoid adhesion of Jurkat cells is to culture them in flasks in upright position, which I always did. Nevertheless as they divide, they adhere to each other and form floating round aggregates.
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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?
Thanks!
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Here are some suggestions.
You make a stock of 5000x in DMSO and then make aliquots of this stock and store in -20 degree C under dark conditions. This stock will last for least 6 months. Make aliquots of this stock in such a way that it could be used only for single use so that repeated freeze-thaw cycles could be avoided.
Do not preserve 1000x diluted stock made in PBS. You will have to make fresh diluted stock in aqueous medium each time you perform your experiment. Such diluted stocks are not stable when prepared and stored.
The diluted stock and the working solution of PMA should be prepared and used on the day of the experiment and could be reused on the same day if you are planning for another experiment on that day. Do not store the diluted stock and working solutions of PMA.
Best Wishes.
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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!
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There may be two prime reasons, if you say you tried with fresh (contamination free) media -
1) The cell stock has some contamination, which is though not clear in attached photograph, as debris or Yeast?? Probably you should discard this stock and start afresh with new stored vial, of some other lot. If cells are rare at your end (widely available if fresh stock to be procured), just try trypsin wash; dont wait for cells to detach; and keep changing fresh media daily for 2-3 days. If morphology is same, then you will have to discard. Alternatively, seed in some smaller flask, as very less no. of cells are viable, and concentration of cells is less. they need to grow densely to look good.
2) The cells might have undergone many many passages and may undergoing senescence. In that case you will have to discard the stock and start afresh with a old stored vial.
I hope this works. Do let me know the progress and get back to me if you have some further questions!!
Regards
Dr. Satyam
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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? 
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Hi all,
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.
Many thanks
Giulia
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Rough surfaces encourage the entrapment of fibrin protein, adhesion of osteogenic cells and mechanical stability of implants in host bone [23–29]. Results from in vitro studies suggested a positive correlation between surface roughness and cellular attachment and cell proliferation.
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Control wells in 96 well plate does not show any absorbance at 570nm
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Dear Mude Hemanjali,
unfortunately you added no additional information concerning your assay..!
One explanation for your observation could be that control cells simply have overgrown and started to detach (did the medium already turn yellow)? If treatment is growth limiting for example, cells in those treated wells would still give a signal with the MTT assay.
Maybe try seeding less cells or shorten the incubation time?
Hope that helps,
best regards,
Christian
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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.
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It seems that there is some issue either with the cell line or the culture flask itself. Did you check the viability of non-adharent cells in the flask after 24 hr of seeding? Did you find any cell adhared to the flask? If no, kindly get one vial from other sources to rule out any issue related to the cell line storage/damage. You can also check the final pH of the culture medium. With this, hope you may get the issue resolved. Good luck
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I have isolated mitochondria from tissue. I would like to check them by fluorescence imaging. Can I use Mitored dye for these isolated mitochondria?
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Debaprasad Koner if you are seeing fluorescence with mitoRed then you are on the right track. Mitotracker red is membrane potential specific so what you see are mitos with reserved MP, and the stain is retained post fixation. However if you want to do a mito count then mitotracker green is the best since it is not mp dependent. Some mitos will lose MP based on treatment, mito fitness, lysis etc. MitoGreen also retains well post fixation.
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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,
Ismael
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Hello,
The HEK293T cells attach poorly to regular uncoated flask. Coated flask/dish will help in culturing HEK293T cells.
Attachment of cells to the substratum requires interaction of healthy cells with a range of adherent protein molecules present in the serum and supplements added to the culture medium.
Cells fail to attach or show poor attachment to the substratum under sub-optimal growth conditions. Failure to attach to the substratum could also be due to environmental stress caused because of contamination, improper freezing or thawing of cells, using cells with higher passage number, temperature fluctuation in the incubator.
Also, when cells are over trypsinized and when they are subcultured at over confluency, they attach poorly to the surface of the flask.
You just need to know these points whenever you carry out cell culture activities.
I hope this will be helpful.
Good Luck.
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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.
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1st, you need to remove antibiotics during the subculture of the cells.
2nd, minimize CO2 level up to 4.5-5.0%.
3rd, The next day of thawing, you need to change the DMEM media completely and put new DMEM-10% FBS/LG (Low Glucose)/ No antibiotics. This kind of cell usually prefers Low glucose DMEM media. I think it will work for sure.
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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.
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Dear Smarth!
Plese You look a t the article
Perhaps insulin through its receptors on HEK293 cells enhances the accumulation of APP, which is toxic to these cells
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Hello everyone!
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!
Kind regards
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Dear Alejandro!
Please You look at the following protocol:
NSC-34 cells were cultured in differentiation medium consisting of minimum essential medium Eagle/alpha modification (Millipore-Sigma, Burlington, MA, USA) supplemented with 1% fetal bovine serum (Thermo-Fisher Scientific, Cambridge, MA, USA), 1% 100× MEM non-essential amino acid solution (Millipore-Sigma, Burlington, MA, USA), 1% pen strep (Thermo-Fisher Scientific, Cambridge, MA, USA). D
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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.
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A bit mysterious. It really looks like you can see the cellular contents bursting out of the cells at around 10sec. Presumably you are using TC treated material? THP-1 not adhesive unless differentiated, maybe you are washing the cells into the PBS? You could try fixing the cells by adding a high concentration of you fixative, for example leave 500ul of media in well (without disturbing cells) and gently add 500ul of 2x fixative.
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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.
Material:
  • 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).
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Hello Luca,
You can check on this paper for help.
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Hey.
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?
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I would compare them to/map them against a reference database such as in VirulenceFinder. The Center for Genomic Epidemiology website provides comprehensive databases.
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I have a problem with clear renal cell carcinoma cell lines Caki-I and Caki-II. I can not proliferate them. Are there anybody who studied these cell lines before? Can somebody give some advice. Because it is very critic for me.
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Dear Emine!
Please You look at these articles:
2.2. Cell Lines and Culture Conditions
The human ccRCC cell lines, Caki-1 (ATCC® HTB-46™) and Caki-2 (ATCC® HTB-47™) were obtained from American Type Culture Collection (ATCC, Manassas, VA, USA). Cells were maintained in McCoy’s 5A (modified) medium supplemented with 10% FBS, 1% l-glutamine and 1% penicillin/streptomycin. Cells were cultured in a 37 °C humidified atmosphere containing 5% CO2 and 95% air. All methods were conducted in accordance with the relevant guidelines and regulations of the institutional biosafety committee.
The RCC cell lines 786-0, Caki-1 and Caki-2 were obtained from American Type Culture Collection (Rockville, MD). The RCC cell lines were cultured in DMEM/F12 (Gibco, Invitrogen, CA, USA) supplemented with foetal bovine serum (10%), penicillin (50 U/ml), streptomycin (50 μg/ml) and amphotericin B (0.125 μg/ml) in a humidified atmosphere of 5% CO2 in air at 37°C. All cell lines were recurrently tested and determined to be mycoplasma-free.
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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.
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You can try adding a drop of medium in the centre of the well (Do not pre-wet the well before adding your drop!). Try different volumes so that you can obtain your desired sureface area.
After knowing the volume, prepare your cell suspension at a proper cencentration and add drops onto your plate. Avoid tilting or disturbing the plate too much so that the droplet can maintain the hemisphere shape. Wait until the cells adhere to the surface and you can replenish enough growth medium to the wells.
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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?
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Dear Nanami, the following papers maybe helpfull to find answer to your question.
You can isolate bacteria from surfaces and than you can characterize their biofilm formation capabilities on different materials surfaces like stainless steel or polymer surfaces. There are both quantitative and qualitative methods to evaluate biofilm formation.
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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.
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Dear Gabriel,
my team and I have recently published a paper on tumor spheroids (https://www.frontiersin.org/articles/10.3389/fimmu.2020.564887/full). We found that HT-29 cells form spheroids with lower weight, diameter and size compared to other CRC cell lines. In the case that you are interested in increasing the sphericity and compactness of HT-29 spheroids I'd suggest using micro patterned ULA plates (Elplasia, AggreWell, SP5D) or adding 1:1 fibroblasts to your 3D cell culture.
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Dear all,
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
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HI Gunnar,
I sent you the paper. Check figure 6.
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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?
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Dear Géssica!
Please You read this article:
Please You read this article.
This isn't about the FBS.
Try using plates or Petri dishes with increased adhesion (for example, I used Petri dishes).
Also, if cells do not adhere well, plates or Petri dishes covered with collagen or fibronectin can be used.
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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.
THANKS
ST
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Hey Smala,
Did you ever figure this out?
I may have a similar problem with my MDCK in plaque assays.
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Hi,
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.
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Thanks for all the answers guys. I've gone with the coating in collagen method as it seems to be the most established and I will probably use the drop culture method aswell.
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Hi All,
Can I use 0.2% Gelatine coated, glutaraldehyde cross-linked glass coverslips as an alternative to Poly-L- lysine coated coverslips?
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Dear Abir,
In my experience, glass coverslips could be plasma oxidized and then could be coated with 0.2% gelatin. There is no need of glutaraldehyde crosslinking. It would allow proper cell adhesion onto the glass.
However, while cellular intrinsic response on any 2 materials is unlikely to be comparable. So, if you are targeting molecular studies, the cells on gelatin could show slightly different phenotype than on PLL. This is particularly due to the differences in cell adhesion motifs in both the materials.
Thank you,
Best,
Tarun
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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!
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Hi Daisy
What is your cell type and both (24 well plate and cell stack) the culture dishes are the same brand? We have seen a huge difference in stem cell attachment between the brands despite all coated with laminin.
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Hello everyone.
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)?
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For cell adhesion on a microfluidic device, you need to be very careful with the protocol itself.
1. You need to ensure the cells adhered to the surface before conducting any experiment
2. To increase adhesion, you can use ECM proteins such as vitronectin, fibronectin, collagen, gelatin and others to coat the surface prior the cell culture.
3. Let the cells settled for at least 5-8 hours, then observe. You can use cell tracker on the cells and monitor the adhesion using a fluorescent microscope.
4. Check and optimize the flowrate for the media and when nanomaterials were injected into the canal - this could be washing the cells away.
All the best!
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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.
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E. coli can be added to a monolayer of epithelial cells (1 􏰃x10^5 cells/well) in a 24-well culture plate at a defined multiplicity of infection (can be variable) and incubate for 60 min (time can be variable) at 37°C in the presence of 5% CO2. Nonadherent bacteria can be removed by washing the cells after incubation. Disrupted the cells by addition of 100u􏰂l of distilled water and incubating at 37°C for 10 min. Serial dilutions of the disrupted mixture can plated on blood agar plates. E. coli colony count will help to determine the ratio of adherent cells to invading E. coli.
Alternatively, E. coli can be incubated with 0.1 mCi of [methyl-3H]thymidine for 24 h and labeled E.coli can be added to a monolayer of epithelial cells in the same fashion as mentioned above. The numbers of adhering and invading organisms can be determined using a liquid scintillation counter. The amounts of 3H detected from infected cells can be expressed as percentage of the total number of invading E. coli.
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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?
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Hi, it's ok, I recommend that you dissolve the antifoam with water to make it last longer and avoid clogged hoses. A silicone-based defoamer is sufficient, normally 0.2 mL is needed for each liter of fermentation (it depends on the amount of air you supply). Regards.
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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.
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Dear Jungen Chen,
You could use the polyvinyl alcohol (PVA) with a concentration of 10% (w/v) for 30-120 minutes to make it the culture plate with a hydrophobic coating surface. Best of luck!
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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
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Dear Chris,
Sorry that I do not have the answer but I am now struggling with the same question ...
But I found another article saying that RGD grafted PDMS can resist harsh cleaning and sterilization like UV or autoclave (RGD PEPTIDE CONJUGATION PROMOTES CELL ADHESION ON PDMS)
Have you find anything elseon the subject untill now ?
Thanks !
Phuong-Anh Dang
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Hi there,
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?
Best regards
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Caro Andre,
I hope you are well?
You will never have a defined number of cells on a scaffold. Even if all "remain", an undefined number will die, and proliferation will be different depending on the individual sample. Therefore, you have to normalize the ALP results to protein content (commonly using Bradford-Assay) and/or DNA - content. It is absolutely vital to switch plates before assaying, including cells adherent to the TC plate does not produce valid results!
In order to improve seeding efficiency, you can
- incubate the samples with proliferation medium or straight FCS for up to 24 h before seeding
- seed cells in a very small volume, e.g. 1 mio cells in 50-200 µl, only just covering the sample; after 20-60 min you can carefully add some cell culture medium.
But bear in mind: if the aim is testing implant suitability, seeding efficiency is one of the biological properties that you are wanting to measure.
Melhores cumprimentos,
Daniel
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Hi colleagues
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?
Thanks you.
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Dear Prof. Gheita sir,
Thank you for your very informative response. Hope you would like to response to other related problems in future.
Thank you!
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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.
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Make sure you buffer your ATP well - it's very acidic when dissolved just in the water. Otherwise, low pH might have unintended consequences... ATP alone shouldn't cause much normally in my experience. Of course, there is always a chance that P2YRs stimulation might induce something, as Hani pointed out
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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.
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I understand the headache here. Usually hela cells don't grow well in glass compared to plastic flask. Usually, the one may have to coat the glass overnight at 37oc.
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Sphere formation is known among normal or cancer stem cells. But whats the underlying mechanism of cells to come together and make sphere?
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A sphere is the lowest energy configuration for bound system (like a bubble). Other cell shapes require focal adhesion on the outside of the cell or asymmetrically organized structures like actin filaments on the inside of the cell. Cells often temporarily "abandon" the production of some adhesive and structural proteins when changing states to redirect energy to the new process and allow for some reorganization.
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How to estimate more precisely the real surface area relating to different degree of roughness?
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Please, excuse me for the late reply!
Many thanks for the useful information!
Maybe I have misled you with the poorly formulated question. I was looking for a simple practical solution to the problem by means of widely available equipment...
Thank you once again, wishing you success in your research efforts!
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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!
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Non-fouling created by peg moieties. Check this:
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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!
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Dear Valentin
you can check if your cells are cobtaminated by micoplasma.
a simple pcr protocool is available on my blog: ProteoCool ( https://proteocool.blogspot.com/?m=1)
ProteoCool n°18 Simple PCR test to detect Mycoplasma contaminations at Page 5
good luck
Manuele
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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?
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hi, during your protocol, did you carry out dehydration with increasing concentration of cold ethanol following FA treatment?
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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.  
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Hi,
I've had the same problem of cell detachment with the MDA-MB-231after gentle PBS washing. Alba Blánquez can you tell me if your Ca2+ or Mg2+ PBS formulation overcame this problem? Thanks a lot.
Daniel
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Which genes are suitable for evaluating cell adhesion on the scaffold?
for example
vcam 1
selectin
TLR receptor
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I follow Marcel Rodrigues Ferreira answer.
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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?
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Your question is difficult to answer. First of all it is important which cells you want to grow. This might make a difference. Then at least to my knowledge vertebrate cell adhesion to surfaces isn't fully understood yet.
Moreover, the percentages given in your question add up only to 76% . What is the rest of the network?
Since PEGDA is also hydrophilic I am not sure that this is the reason for the adhesion problem, but you could try to change composition and study this systematically if this wasn't done yet.
Our experience with hydrogel based surface coatings does show that the surface mechanics is probably more important compared to hydrophobicity. We used PNIPAM based microgel coatings and in the shrunken state (more hydrophobic but rather hard) cells grow nicely on it (e.g. mouse fibroblasts, HeLa cells, and some more). However, lowering the temperature makes these coatings swell in water (more hydrophilic and mechanically soft) and cells are detached.
Hence, I guess surface mechanics and steric repulsion is probably more important compared to hydrophobicity.
If you are interested in our works, you should have a look at:
  • DOI: 10.3390/polym10060656
  • DOI: 10.1021/acs.biomac.5b01728
  • DOI: 10.1002/adfm.201090084
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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?
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Hello Schneider
Through the following link you can find the answer to your question
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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
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It is of great significance in determining the adhesion behavior of cells toward treatments or different physiological conditions, understanding the mechanism of cell adhesion, analyzing the biocompatibility of bio-materials for tissue engineering, cancer metastasis studies, and also the potential of drug treatments.
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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?
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Hello Riaj,
Although the time and concentration of Ang2 varies between different endothelial cell types, but the attached manuscript will be helpful in determining the starting point for you. The authors have used 200ng/ml of Ang2 for 15 minutes and then stained the HUVECs for VE-cad and other proteins.
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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?
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I had the same issue with HEK293T cells but after coating the plate (poly-D-Lysine or Cell-Tak) prior to seeding and reducing the mixing to minimum during the seahorse assay I retain nearly all cells - and then I normalize to cell number post assay.
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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.
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You might have different populations of monocytes. I would do a pilot run where I incubate it at different time points at RT or 37C so zero in on the correct temperature/time for you to get all the monocytes but still avoiding the HUVEC.
Alternatively you can get everything and then sort the cells to separate the ones you want.
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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.
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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?
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PGA is more hydrophilic, while PLA is more hydrophobic
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Acceptance, Rejection, biocompatibility, cell adhesion, cell proliferation, etc.
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Recently,I need to do cancer cell adhesion assay. Please share the experiment steps for me.Thank you!
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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?
Kind Regards 
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I know it has been a while since this question was made but I would like to update here by saying that trypsin has never worked for me and it usually made my 4T1 culture die, so I have started using a scrapper instead and it has functioned perfectly for me.
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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. 
Any suggestions?
Thanks in advance.
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Hi, we regularly work with 3T3F442A cells, this cells don't have any trouble if the culture dish is 60mm or above, for smaller culture dishes or even slides, you need to coat the surface with poly-lysine, to help the cells to hold to the surface.
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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?
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Haven't tried myself, but you could try to plasma-coat the PDMS with a teflon-like polymer. This is a second half of the Bosch etching process for passivation: C4F8 gas, flow 50-100 sccm, RF 0-10W, ICP RF ~2000W, pressure some tens of mtorrs, or any conditions that give low bias voltage. A couple of seconds should grow a teflon-like substance on the surface to which nothing really sticks well.
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how to predict homophilic and heterophilic interactions for cell adhesion molecule. Is there a software that predict homophilic or heterophilic interaction from protein sequence ?
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Bioedit or Dnamana may can.
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Hi everyone,
I have adhesion problems to flask when I thaw the Beas-2b cells, someone has any advice for me? someone would be willing to give me a aliquot?
Thank you
Tommaso
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Hello Aubrey,
Thanks for your suggestions.
tommaso
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hey,
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?
Thanks
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Cells will not adhere to alginate. It does not present any ligands to which mammalian cells can bind. It also does not bind proteins to which cells can binds (e.g. fibronectin, collagen, laminin, etc.). It is possible to graft cell adhesion peptides to alginate to enable attachment. The chemistry is fairly straightforward. I am attaching a couple of papers that document how to make the modifications and the effect is has on cell adhesion.
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Hi,
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!
Eva
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Look at our mice tests:
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Hi,
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?
Cheers!
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i usually do a lot of astrocyte culture... i do not face that problem . i use PDL (poly- D- lysine) coating. i would like to share my protocol for your convenience
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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???
thank you
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Hello Mahtab,
Despite the cell type not clear in the question there are number of parameters involved, including but not limited to : The degree of polymerization of PVA, freezing temp, the freezing-thawing time, the water content of hydrogel, and type of cell lines.
In particular freezing-thawing process can be critical, the faster the freezing (-80 compared to -20 C) might result in smaller pore size and less water holding capacity or small pore size might prevent cell penetration and proliferation. Similarly, The Mw of PVA is also important.
I hope it will help
Amin
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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!
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It can't be unless you autoclaved gelatin sol, or you coated gelatin sol properly on cleaned glass coverslips by standing gelatin drop on the coverslip for 30 min, dried for 1h, washed with ddH2O, dried again. I found my students get this problems  with an easy prep. by soaking coverslips for a brief time or if gelatin is dissolved in salt sol.
1. Check your 3T3 cell are clean by staining with DAPI to check any bac or mycoplasma.
2. Check your medium whether it is sterile.
3. Prepare dust-free, clean coverslip, dried
4. Make sure your gelatin sol is new and clean, if you have any doubt or left from someone else , spin the sol in 1ml eppen tube at 12,000rpm for 10 min to remove any debris or contaminated microbes. Use the supernatant.
5. Put 50ul of gelatin sol on the middle of a small round coverslips, and leave them to dry completely under dust-free lamina flow within the 100ml culture dish. (give 1h)
6. Wash the coverslips in ddH2O (the stacked gelatin molecules loose should be washed away leaving a nice coating layer of gellatin molecules) and dry them and store in glass petri dish with lid.
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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. 
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Good news! finally you used corning ultra low cell attachment plate to solve your problem.
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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?
Thanks
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Hi Barbara, we used to freeze them in LHC-9 media +7.5% DMSO +1% PVP and it works fine for us. I think PVP is necessary. We also grow them in this media and for trypsinizing too we use PVP in our trypsin/EDTA as suggested by ATCC
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Hi,
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!
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Try adding some DNAze solution or passage cells with only EDTA solution or anesthetics solution like: procaine/lidocaine (It is slightly longer but the cells are not damaged like trypsin). You can also add some TNS after trypsynization, (trypsin:TNS - 1:1) (trypsin neutralizing solution; from Lonza) - I have observed that the cells are in much better condition than the inactivation of trypsin by 10% FBS-supplemented medium.
Good luck! ;) 
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Hi all,
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!
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Another option is to use a substrate that gets charged in solution. This could be accomplished with substrate and collagen having different isoelectric point values and choosing a pH value in between. Mica for example, is quite popular for this reason in the AFM investigation of biomolecules and polymers such as DNA. You also would like to keep in mind the roughness of the substrate which should be much smaller than the collagen fibers you aim at investigating. You can also consider using graphite (HOPG) which although it is chemically inert, the pi-pi interaction with organic molecules is often enough to ensure the mechanical stability needed for AFM imaging in liquids. Both of these substrates, HOPG and mica, can be atomically by simple mechanical cleavage with adhesive tape.
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Carboxyl-anhydride and amine plasma coating of PCL nanofibers to improve their bioactivity
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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
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Thank you Anton for sharing your nice paper
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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!
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Hi William,
Not sure it is of any help (we use different cells), but from our own experience, striping/cleaning the glass coverslips with high concentration HCl and then washing thoroughly with sterile distilled water before to apply the coating helps.
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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?
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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.
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