Cell Imaging - Science method

If you have a cells monolayer, 30 min is a good time. If you have something like a tissue developing, with a lot of collagen, then you need 1 hr. HA is soluble in water (very slow, but still...). So if you culture started generate small centers of mineralization, you do not want to keep it too long (days, weeks) in water solutions. From the other side prolonged storage in desiccator can lead to fungus growth. Some desiccators are badly infested with fungus and need through cleaning and disinfection. From my opinion the best way to store specimens is when their preparation is complete, i.e. they are dehydrated and coated with conductive coating.

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

I also have a Zoe imager and I am also experiencing this issue. Renaming the directory or burying the default export folder doesn't address the issue. Bio-Rad technical support notes that other people have reported this issue, and I am working with them to try and troubleshoot the issue.

Dear Ayman

Check that the condenser iris aperture is set to the objective numeric aperture of 40x. In another case, I suggest checking that you are using the proper technique. Check if using phase contrast or DIC is no longer suitable for visualizing your cells.

Hi usually we do imaging a day after the transfection because sometimes after the transfection if you leave it for 3 days. The non transfected cells can over grown on top of the transfected cells which would then be on your way. So ideally we do imaging the day after transfection.

You can calculate the chromatin condensation parameter (CCP) simply by staining the cells with DAPI.

Dear i want to detect and segment each blood cell from the blood cell image using neculie segmentation but i want to use SBF and Laplacian Gauss Algorithm

Optimizing the loading of AM dyes on the microscope stage is an excellent suggestion, as cell types show tremendous variability. Loading times tend to be in the 10-30 minute range. At temperatures approaching 37C, AM dyes can selectively load into organelles like the ER and mitochondria causing damage and/or non-cytosolic Ca2+ signals. You can use the microscope to check the dye distribution, a dark nucleus would indicate some ER loading for example. After a wash, you can also check to see if your cell type actively pumps out the dye, if so you may need to include something like sulfinpyrazone.

Make sure the DMSO used is OK.

Keep the dye around 1uM - loading is dependent on esterase activity, so higher conc. doesn't help.

Add pluronic to ~10% volumes of dish media, then add Fluo-AM to that, then add all to dish

You don't need to worry about -AM dyes binding Ca2+. The acetoxymethyl (AM) groups prevent that. Even if the dyes did bind Ca2+, then the effect would only be in the low micromolar range while extracellular Ca2+ is ~1000fold above that at 1-2mM.

Hi Adem,

You can differentiate U-937 cell line using "a differentiatial mark" that we had utilized in our experiments with PMA-stimulated U937 macrophageal cell line. Namely, as U937 cells respond to the presence of LPS with a marked increase in the rate of cellular AA metabolism and cytokine release into the extracellular space, we measured activity of cyclooxygenase 2 (COX), which catalyze the key step in the conversion of cellular arachidonic acid into prostaglandins. The COX-2 expression is strictly coordinated with up-regulated in PMA-differentiated U937 cells stimulated with lipopolysaccharide (LPS). At the same time, COX-1 is not up-regulated at all. In turn, another "market sign" we'd used from time to time was enzymic activity of the Mg+2-dependent phosphatidic acid phosphohydrolase 1 (PAP-1), because up-regulation of COX-2 strictly depends on the PAP-1.

In regard to your request on seeding concentration of cell, PMA concentration for treatment, and on phenotypic and morphologic changes observed in U937 cells after treatment with PMA (100 nM) in the presence of Haemophilus influenzae, please read this paper of Dr. Jahn and colleagues 

Best wishes,

Ilya

Hi Mohammad, tank for appreciating our past discussion. As far as cel fixed are concerned, I've mostly used PFA4%. It is the one of the best chemicals, used for cell fixation, recommended for AFM. I suggest you to scanned the cell at least 24 hour later cell fixation, unless the protocol required differently. AFM preserves cells from any kinds of changes made by time. I scanned cell samples fixated 8 years beforehand. Concerning the essication, made by biological cabinet airflow (30 minutes, no more), it depends what you want to look for. In my latest AFM experiments I fixed cells by airflow, and the samples were perfect. I would like to suggest to consider also the airflow fixation, not for long time or you must seriously shrink samples. I hope my reply will be helpful. :-)

Cheers

You might want to take a look at the attached paper. The principle and implementation of VCA and other techniques were mentioned.

The cell line was prepared as described in Section 2.1.

Hi Agata,

I was wondering if you got/have a MitoSOX Red protocol for suspension cells.

Thank you.

KJ

Hi,

I am currently facing a similar issue. I believe Spinning disk confocal microscopy can help imaging (especially if you need a Z stack) at faster rates, but I have not tried that myself. Have you been able to find something that helps? I do not want to image my cells at 4 degrees as that affects the uptake of my compound by the endocytic pathway, so I am trying to find techniques with better resolution (unsuccessfully).

Well, if the microscope is not damaged then the most likely culprit is that it is not calibrated correctly. Have you gone through the entire process of aligning the optical path? Are your phase plate and annulus properly matched? Is your Kohler illumination aligned? Are your samples sitting flat on the stage? Is your camera in the correct alignment in the optical port?

The microscope itself is professional grade. You should be able to take clear images in any mode so long as it is in working order.

Glad it worked.

I would recommend taking an image of anything with each of the objectives and at each bin setting using your system. Then embed the Scale bar using the FLiod software as you did and measure the length in ImageJ. Then just make a table of the conversion factor between pixels and microns for each objective and binning for future reference.

Sounds like something that could easily be done with ImageJ. I like the FIJI distro.

If I understand correctly, you are trying to analyse co-localisation in an enzymatically stained sample? So the problem is that you can't distinguish the 2 separate stainings in separate channels? In that case I believe you can try to use an ImageJ colour deconvolution plugin, such as is available in FIJI. After that, if the deconvolution was successful (it might need optimisation), you might be able to do colocalisation analyses. I hope that helps.

I don t know for mitoTracker, but if the dye is not stable enough for a 48h video, you can try to transfect/infect your cells with mtDsRed plasmid/virus. the fluorescence might bleach at some point so you should try to acquire at a low frequency. For that, you need to be sure that this frequency is appropriate for analysing the biological process (velocity, morphology, fusion/fission) you want to study.

Yes, looking at the figure, it is clear for me that authors used purple instead of blue to show DAPI.

It is common now to see such color in publications. Blue may be difficult to see on dark background and purple is also used for the colorblind people.

Try to use the red channel/filter (eg. Texas Red) if it is available on the microscope. 

 Fluorescence Image Acquisition for BX53

1.Click the FL button on the remote controller box to use mercury arc lamp light for fluorescence.  The light can be on or off by pressing FL Shutter button on the remote box.   

2.Select an appropriate fluorescence mirror cube by pressing Mirror buttons in the controller box or clicking any one of the Observation Methods buttons   (except BF and DIC). The selected mirror will be in position 1.  Make sure that the light path bars are in correct position (the upper bar at the mid or extended position and the lower bar pushed in) and camera is selected.  

The cited statement is right only in pure theory.

In practice it means that beam intensity (called "spot size" by some manufacturers) should be changed with magnification - higher intensity for lower magnifications. In this way operator can achieve right balance between image sharpness and noise level.

Thankyou, but will glass pasteur pipette be visible under the objective lens? what is the minimum objective lens required for that? 40x or 400x ?

Dear Mou. The columns you use to set a calibration, for example: if what is black in the image represents 0 mg of calcium (just an e.g.), and the strongest red is 25 mg of calcium. In this case, you can make a calibration (in this case using only 2 points, the zero and 25... ). So, as black is represented in ImageJ (8 bits images) by zero, and white (red in the case you choose it in the lookup table) is 255.. so you can make

in the left column:

0

255

in the right column:

0

25

and choose the straight line for the function. 

After calibrated, you can select an ROI (region of interest) by choosing the forms (rectangle, circle, etc.) available, and then go to Analyze>measure (CTRL+M) and you will have a statistics of the region you selected.  You can also go to SET Measurements to select what do you want to measure...

Hope it is what you are looking for...

All the best

Hi Guillaume,

Unfortunately, it's practically unreliable assessing the ATP quantification in the cell lysates in which ROS accumulation was undertaken. Just compare the toughness of the later procedure, in order to imagine the inability of coexistence for delicate ATP reaction.

Intracellular ROS levels usually assess using the CM-H2DCFDA probe. Cells grown in 96-well flat clear bottom black polystyrene microplates are washed twice with 150 μL of HBSS to remove culture medium. Then, 50 μL of the probe (7.21 μM in HBSS) added to each well and the plates are incubated for 30 minutes at 37°C. Excess probe was discarded and 100 μL of the prooxidant solutions prepared in HBSS added. After the incubation, cells are lysed by adding 50 μL of 0.5% Triton X-100 in PBS. Cellular levels of ROS are visualized under the microscope using CM-H2DCFDA fluorescence live cell imaging; cell fluorescence images captured using an epifluorescence microscope.

At the same time, intracellular ATP determination performed by a bioluminescence assay based on the ATP-dependent luciferin-luciferase reaction. A new internal calibration standard of ATP usually prepared each day in a range from 1 to 100 μg ATP/mL prior to readings. To determine the cellular ATP content, cells grown on 96-well white clear-bottom plates were first incubated for 3 minutes with 25 μL somatic cell ATP releasing reagent and then for 3 minutes with 25 μL of sterile water. The plates placed in a Luminoskan RS luminometer and 50 μL of the luciferase-containing buffer (adenosine 5′-triphosphate assay mix) added to each well just before measurement of the light emitted, which is proportional to the ATP concentration. 

Best wishes,

Ilya

Thanks so much for all infos

Thank you Dr. Rad for sharing the article. Appreciate your help!

Hi Md. Amran,

To practically solve your issue (that requires an accurate cell segmentation by image processing), I suggest that you follow these links:

Here is an excerpt from (1) Carpenter et al, 2009: “…It is sometimes unnecessary to precisely identify the cell boundaries: for instance, to determine whether a protein is predominantly in the nucleus or the cytoplasm, it can be sufficient to measure the average intensity of a protein in the nucleus and in a ringshaped region around the nucleus, as a proxy for the cytoplasm…”

Note: this computational technique may be extended to calculate fluorescence intensities within 3D cell structures via ImageJ plug-in “WatershedCounting3D” ( https://www.ncbi.nlm.nih.gov/pmc/articles/PMC3607619/pdf/cl-2-176.pdf ).

Here are some complementary references of interest:

Finally, in addition to the here-above valuable reference suggested by Hermance Beaud ( https://blogs.qub.ac.uk/ccbg/files/2014/05/2014-05_Analyzing_fluorescence_microscopy_images.pdf ), some readings about the –comparative- use of ImageJ and Fiji: 

Hoping it will be helpfull

We use a super resolution microscopy (STORM) which is more sensitive to fluorescence that a normal fluorescence microscope. With the polymer, which is label with Rhodamine, I can see exactly the same profile. In fact at first I thought that I did a mistake with the negative control and it was not the real sample. 

Short answer, your experiment failed.  Have the reagents you used given positive results before?  Were your samples and reagents fresh?  Have you done a successful dot blot before or is this a new technique for you?  You are going to have to give more details to get specific help.

Your best bet is to just do total fluorescence intensity in the mitotracker channel. It's always difficult to count mitochondria by a total mitochondrial stain partially because they can span imaging planes fairly easily.

If you wanted discrete counts, best to use something like a mitochondrial DNA stain that should only correspond to one spot per mitochondrion.

Split and segregate the z-stacks for each time point. Then take each of the z-stacks and go to Iamge - Z porject - maximum intensity. You will get a maximum intensity projection for each of your time point from the z-stacks. in this way. Now go to Image - Stack - Images to Stack. You will get a stack with all the max. projections from each z-stack as a time series.Hope it works for u.

http://www.4ti.co.uk/seal/adhesive-seals/gas-permeable-imaging-seal ?

All I did was google your question.

Dear Attila, thank you, I will buy labelled one. Do you have a clear protocol for studying the uptake by labelled dextran?

Thanks

Alaa

There are many nuclear stains but the two best dye versions (and easiest, no transfections required) are Hoechst  and SIR-DNA (from Cytoskeleton).  Both are bright dyes that when used at the appropriate dose (I use 1ug/ml for Hoechst and 250nM for SIR-DNA) you can timelapse image for 24 hours or more without significant photobleaching.  Neither dye (at low doses) inhibits cell division or cell migration. Both are great for cell tracking. Hoechst excites in the 365nm  range while SIR-DNA excites with 642nm.

Dear John and Hesham,

thank you very much for your comments.

Try the rolling ball background subtraction in ImageJ.  It does wonders for unevenly illuminated fields.

The better solution is to use a Chroma slide to check that your beam is centered.  TIRF uses single mode fiber with a Gaussian beam profile, so it will always be brighter in the center and fall of at the edges.  Put the cells in the center of the field. 

In the following paper by ZaritskyEmail et al., "Benchmark for multi-cellular segmentation of bright field microscopy images" BMC Bioinformatics201314:319 DOI: 10.1186/1471-2105-14-319, they talk about a dataset of 171 manually segmented images of 5 different cell lines at diverse confluence levels, acquired in several laboratories under different imaging conditions in the first figure.  HTH.

10

Dear Celine Choo

You can join this ImageJ Forum, we discussed alot of issues about imageJ in relation to Image-Pro Plus and you will find your answers.

This ImageJ Forum for those interested in morphometry. the link is provided:

Are there any plugins for Imagej or a method for Image-Pro Plus to measure the mean/average diameter of polygons?. Available from: https://www.researchgate.net/post/Are_there_any_plugins_for_Imagej_or_a_method_for_Image-Pro_Plus_to_measure_the_mean_average_diameter_of_polygons [accessed Apr 15, 2017].

Thanks, Michaela and Usama, It helps a lot, I'll try to use the vaseline frist, if it doesn't work well, I'll buy the Fluro-dish. Thank very much.

Thank you so much. Indeed, knowledgeable information. I am quite familiar with Brd-U labeling of different cells including cardiomyocytes of my interest. I am seeking BrU labeling to analysee RNA stability via pulse-chase.. which can be alternative of usage of intercalator like actinomycin D.... https://www.ncbi.nlm.nih.gov/pmc/articles/PMC3583853/pdf/rna-9-1233.pdf

Thanks again.

Hi Justing:

No sure if you are aware or not, but they are commercially available (albeit overpriced!) https://www.mattek.com/store-category/cultureware/glass-bottom-multi-well-plates/.

A cheap alternative we have done in the past, is to simply heat up a cork borer (i.e. the metal tool for cutting a hole in a cork or rubber stopper) of the desired diameter and simply punch/melt a hole through the bottom of plate/well. Don't get the metal too hot or you will end up melting away a larger hole; with the right temperature you will get enough heat into the borer to help it move through the plastic smoothly.

Then just glue a round coverslip to the bottom (outside face) of the plate using sylgard 184 or your favorite adhesive. Works great for plating and imaging or as a recording chamber for patch clamp experiments. 

Hope this helps. Good luck

I recommend scanning slides with a ScanScope XT and analyse images with an eSlide Imager (Aperio, Vista, US). Hope this helps, Tom

Is what you tried to use HPLC reversed with solvent mixture in gradient mode

Hi,

Please note that:

We suggest using two cytotoxicity assays instead of just one. This is because certain chemicals have been reported to give divergent results in different toxicity tests including the NRU and MTT assays (Olivier et al 1995; Chiba et al 1998). Besides, Evans et al (2001) have recently found that in some cases one of the NRU or MTT assays can be more sensitive in detecting the toxicity of non-viral transfection reagents.

Evans, A.R., Alexander, D., Burke, P. and Reed, C.J. (2001) Toxicity and transfection efficiency: comparison between four commercially available non-viral transfection reagents in a human bronchial epithelial cell line. British Pharmaceutical Conference Science Proceedings 2001,  106.

Pls also see attached research article.

Have you access to a fluorescence microscope? I would suggest that you look at single bacteria first.

 Dear, Babu Mia, my task is to quantify the content of phosphatidylserine in the outer layer of the membrane. I can not find a technique how to do it = (. I do not have the opportunity to do this with flow cytometry.

Find the "brightest" FPs and choose those - http://www.fpvis.org/FP.html

mNeonGreen looks good.

Modern CFPs are mTurquoise2 and/or mCerulean3, CyPet or mVenus for yellow. mCardinal or mSmurFP maybe for red

You could try fluorescence imaging. Check for fluorescence emission from the drugs loaded in nanoparticles if the nanoparticles are non-fluorescent. In the absence of fluorescence from nanoparticles, the fluorescence has to come from the drug present in the nanoparticles.

But there are other methods such as infrared spectroscopy. You could identify unique characteristic IR bands of the drug molecule and check their presence in the sample confirming the presence of drug inside the nanoparticle.

Thanks

We are planning to do high Resolution Imaging (STORM) and confocal Imaging. So normal analyzers like JuLI are no option. That's why I need a thin microscope slide that suits very sensitive cells and high Resolution imaging.

Hi Magdolna, I had the same before, used alpha-acetylated tubulin antibody on human airway epithelial cells grown in liquid culture and I only saw cytoskeletal structure (with the confocal). Cilia would normally appear in air/liquid interphase cultured, therefore structually differenciated cells. So if you dont see cilia on the wide-field I think the chances are good that there are no cilia. Best, Anita

@Srinivasa Rao Palla:  even if the specialist knows that you wanted to name the dye....

would you - please - mind to correct (using the edit-function top right edge/margin to your question-form....)   the wrong / misspelled "according" in the title of your question to "acridine orange" (this honestly asked for better searchability in RG-archives....)  Thank you! 

Hi,

have you tried confocal microscopy?

Thanks! Good luck!

Hi,

You can do it easily with the ImajeJ program. You can set an threshold for the roundness of the adipocyte size and tell the program to count every cell that have x and z size and the ones in between will also be included. The program is automated but you can add and exclude cell once you are done. It will give you the celk area, cell number and perimeter. You do need to apply and color deconvolution on your image and use the  color that will suit better for your analysis. 

Erika Barboza

Hello

The book and discussion on RG link will help you

Good Luck 

Dear Fatima,

thank you really for detailing your trials. I imagine the problems you are facing (I have to admit:  imagine "theoretically" for AFM-examination) but as you certainly found out I was working in the "branch" of diagnostic TEM formerly and also have a wide background to SEM, AFM and other microscopical techniques as read by literature rather than personal experience).  So your approach, to include your Arabidopsis thaliana cells into liquid low-melt agarose at first glance seems interesting and promising....   the problem will be the intermediate drying and then re-wetting of the cells (in appropriate buffer..., guessing you can not fix your plant cells prior to examination because you want to find out not only morphology but also  the Young modulus in living cells). Other AFM articles / work report adhering cell material (e. g. homogenized and therefore 'hackled' prior to imaging) to mica-sheets.......    since you intend to use a cell suspension under liquid conditions for measurement I guess that literally physical "adhering" will be impossible...  the only way of keeping your cells alive for measurement in the low melting agarose film (low melting gelatine too ??) IMHO would be seeding your cell material to the moist agarose layer - stirring and cooling down until the agarose is solid...attempts to measure then would need a cooled chamber/ setting....this might be difficult and delicate circumstances....Hope there are out other ( the real )experts, wish you all the best, good luck, WM

I would say, it depends on the way, how cells have been fixed. Some fixatives destroy or alter lipid structures, which would result in disturbed lipid staining by Nile Red. If possible, I would suggest labeling with Nile Red before fixation and use a gentle fixation method, such as paraformaldehyde, 2% 10 min on ice.

Thank you Jan,

      Ya i am getting the emission and excitation for quantum dots at a time. I dont know how to rectify this issues. I tried green LED for excitation and used the standard filter#4. Also How to get the 640BP35 filter active. There is no display of this filter. It seems, in manual they mentioned "AN OPTION 640BP35 FILTER IS AVAILABLE". please guide me, how to proceed with this instrument for selective filter emission for particular wavelength

Hi Joe, in my opinion this is the way it should be.

You can also do statistics within your groups, but this will only show your inter-experimental variation (technical n).

If you want to measure puncta specifically, it's not as simple as the above response. In that case, can use automatic threshholding (Image > thresholding on Image J toolbar) to essentially eliminate the background fluorescence, such that your image will only display the puncta. From there you can quantify the puncta. You will have to play with the thresholding a bit to get your desired result. Another possibility method is to use the "substract background function" (process > subtract background) to remove the diffuse fluorescence such that only the puncta are visible. To quantify the puncta, you can "analyze particles" (analyze > analyze particles). Otherwise, if you want to simply calculate the overall fluorescence you will first need to "set measurements" to determine what info you'd like (I suggest checking the following boxes: area, min/max gray value and mean). Then you can measure (analyze > measure). You will need to copy and paste the raw data into an Excel spreadsheet from there.

Hi Vipin,

the accurcy of your localization is strongly depending on signal to noise, pixel size and the size of the point spread function. You might want to take a look into this publication doi:10.1038/nmeth.1447

The experimental localization accuracy is often measured as the FWHM of the distribution of a big number of localizations from one emitter. For example: localize one bead a thousand times and analyze the distribution of the thousand localizations.

Best,

Thorge

It is just idea below not an experience :)

Do incubation chamber as usual and use not microscope but cover slide for later steps.

A couple tape on microscope glass slide + drop of binding media + inverted cover slide with array.

Have a nice day!

I have never observed such kind of inclusion body in A431 cells. In my opinion, the contamination is the most likely. How about the doubling time of the cells?

Given the centres of the cells you could look at the cumulative distribution of the distances between all pairs of cells.

According to Inouye et al, 2015 (http://dx.doi.org/10.1016/j.pep.2015.02.002), the coelenerazine-h is also a very good substrate for NanoLuc/NanoKAZ.

Although, I didn't see a direct comparison. Furimazine, probably, has lower background, anyway.

PS. According to Promega, Furimazine gives slightly higher initial bioluminescence and the signal then decays much slower compared to CTZ-h.

Dear Dan: If you are working on mature B cells; CD19;  B220 are good markers (both for human and mice). Percp is very easy to be bleached under focus, never try this fluorochrome; FITC, PE and APC are good for FACS, but not ideal for Confocal; tandom Abs (say, PE-Cy5) is a bit more stable; but my favourite is alexa fluro confugated Abs which are very tolerant.

Good luck

Hello Aditya,

it's quite simple, once you open your sequence as a stack, you can select the rectangular region you want to "zoom in",and then select Image->crop. It will crop the area you selected, so then you can just save it as AVI and have your video.

OK no, you don't need to calibrate. Your image is calibrated since you have µm mentioned in your image header.

Dear Sumit,

I have few other suggestions:

1) I think that one key important thing is that the acquisition parameters are the same across the images. I know that is an obvious point, but I saw so many people measuring and comparing fluorescence from image acquired with completely different parameters.

2) Be sure you are not saturating the pixels on your CCD, otherwise you will lose power in your quantification. It does not seems the case in the sample image you posted, but I thought it was worth to mention.

finally for the quantification... I think that the answer on your question depends on what you would like to quantify.

1) The answer above are all valid, however, personally I would stay away from normalisation, Adam is correct in saying that there is variation between coverslips and samples, but I usually deal with this variability making (when possible) more sample for each condition and averaging them across.

2) another point to consider is if you are interested in the total amount of fluorescence, I mean a bigger mitocondrium with the same amount of marker will have a lower mean intensity... You can use the total (or integrated) fluorescence intensity but in this case you have to be careful and remove the background, (a bigger object have a bigger integrated background).

Hope this help,

good luck

Max

I always randomly select regions on my coverslip for imaging - to avoid biasing, you can refrain from looking down onto the coverslip while you move in the XY plane. You can do the corners and centre but as your cells clump in middle it's not a good representation.

Just as a matter of interest, when you seed your coverslips with cells, do you swirl the cell mix or do you rock the cell mix once it's in the tissue culture dish/plate? Normally one should not swirl the mix as it causes a vortex in the medium thereby pulling all the cells to the centre of the plate - resulting in uneven distribution and clumping. If you gently rock from left to right, up and down it should give a better distribution. Also preparing the cell mix prior to addition to coverslips is better than having coverslips in fresh medium and adding a small volume of concentrated cells directly to it.

Thanks 

MATLAB

Try to use phase contrast to see and move a tip of your capillary to the cells. Switch to fluorescence regime just for a short time to catch it. Algorithm: 1) find some cell in fluorescence and keep in mind this area in the view, 2) switch to phase contrast, 3) move a capillary tip close to this cell, 4) switch to fluorescence again (to be sure if this "right" cell) and 5) collect this cell. Such approach may work for small amount of cells...

Probably, a better option is collecting the cells under combined low-intensity phase contrast and fluorescence microscopy.

Dear Neha,

You are right. However, that factor of predictability on when such morphological changes will begin to appear is quite difficult to have. Even minor things trigger changes. Let's hope you can resolve these issues with developing fresh cultures from new cell lines.

Regards, Sourav