what kind of coating did you use? because depending on the technology used, here can be the problem. Also, how many cells did you have? usually is better to use high confluency, because a certain cell loss is expected...another thing..glutaraldehyde is fresh? and how long did you fix the cells? usually is around 15'-30', if you fixed for less time you risk the loss of cells....

What types of biomaterial are you using to fabricate the scaffold? Previously I was trying to use the ethanol gradient method to dehydrate alginate fibers. But actually, I can see the alginate hydrogel is somehow react with ethanol. I am not sure whether it is chemical react or something else. But apparently, the biomaterial is affect by ethanol. So, are you sure ethanol is not react with the biomaterial you are using, and thus you loss cells? The ethanol gradient method successfully dehydrated our alginate fibers. And we obtained SEM figures by using this method. But I can see there are some reactions going on during the dehydration process and changed the fiber. So, I think this might cause your cells losing problem.

Maybe you should try other dehydration methods. Later, we used freeze drying method for dehydrating those fibers with cells encapsulation. And we viewed the seeded cells on the fiber surface sucessfully by SEM.

Hello Yahui,

As per my observation so far, my scaffold do not react with ethanol. Though, I have not checked for the changes in micro-structure. But according to your observation, this may be a reason for cell loss, so I will surely keep in mind and separately check these effects.

I might bother you again for details of process parameters used while freeze drying, if needed. Thank you so much for current suggestions.

hello Mr. Peter,

Thank you for your suggestion. I found that my samples were not completely dry. but then while imaging, we were able to see the scaffold surface clearly, only problem was that we dint spot cells on it. so I am suspecting that there is some missing link during sample processing.

Thank you Fulvio Sir, for pointing out the steps which are usually overlooked but play very important role in sample processing. well, in my case, I would like to answer your questions-

we used gold coating.

while seeding cells on the scaffold, they were well enough in number. We had performed florescence study for that sample, and spotted out many live cells which were later processed for SEM.

Glutaraldehyde was freshly prepared.

And now here I got some more doubts, it will be very kind of you, if you would solve my queries. I would like to know, that was it a reason for this problem, that we used same samples for SEM on which we had performed florescence study?

Secondly, could you tel me what is the maximum time period upto which we could keep cells fixed in glutaraldehyde before SEM imaging? I mean, does storing fixed cells for long time under refrigerated condition, also affect washing and dehydration steps later?

Hi. For well adherent cells, it usually isn't a problem! What kind of cells did you have on the scaffold? If it's a repetitive problem, you could try using an ESEM, and avoid the ethanol dehydration step completely by imaging under low vacuum conditions. Usually it's possible to image upto 25000x and get very good crisp images. I've looked at bone marrow derived MSCs and some bacterial cultures on various smooth and rough surfaces. At times, the washing/rinsing steps can be a bit harsh and wide areas of adherent cells can go missing, so I wonder if that is what has happened to your samples. Have you considered cryo-preservation of your samples? Usually that words quite nicely too.

You may want to consider performing critical point drying (CPD) after the ethanol dehydration. This will preserve the cell morphology and fine features that may be damaged by air drying. For CPD, you need a suitable setup (pressure vessel with vent) and you would take your samples in 100% ethanol, then exchange the ethanol for CO2, pass around the critical point of CO2, and vent. This produces dry samples and preserves the cell morphology. I have used this technique successfully with 3D porous scaffolds, 2D substrates, and other cultures.

You also asked about storage before imaging after fixing. Fix for your standard amount of time, then remove fixative and store the samples in sterile PBS in refrigerator until you are ready to process samples for imaging.

Florczyk Sir, I am extremely thankful for your valuable suggestions.

Hello Furqan Shah ji,

first of all thank you for suggestions. I will surely follow them.

Dear Paranita,

we incubate the samples additionally with hexamethyldisilazane overnight and obtain good results. Maybe it will help also in your case. I copy paste a part of a materials and methods section from one of our manuscripts for further details. Please note, that hexamethyldisilazane is toxic.

Good luck

Dirk

Surface topography, roughness and morphology of the biomaterials, as well as adherent BMC were assessed by scanning electron microscopy (SEM). 2 days after BMC seeding, untreated and treated scaffolds were fixed with glutardialdehyde (2%) for 30 min and subsequently dehydrated through ascending grades of alcohol (25%, 50%, 75%, 96 %, 100% ethanol) for 15 minutes each step. Scaffolds were then incubated overnight in 1,1,1,3,3,3-hexamethyldisilazane (Merck-Schuchardt, Hohenbrunn, Germany) and drained. Afterwards the samples were sputtered with gold (3 x 60 s, Agar Sputter Coater, Agar Scientific Ltd., UK) using a Hitachi FE-SEM S4500 (Hitachi, Dusseldorf, Germany) with a voltage of 5 kV. The images were digitally recorded using the Digital Image Processing System 2.6 (Point Electronic, Halle, Germany).

If the cells have not produced a suitable ECM and/or did not succeed to achieve stable adhesion to the available scaffold/substrate they could easily detach after deep dehydration following ethanol treatment. Particularly in case they form a monolayer and of a large differential hydration properties between cells and substrate. What kind of cell and substrate did you use?

@ Mr. Michel Spina .. In the current scenario of loss of cells occur after washing with ethanol and air drying (but major cell loss occur during SEM analysis only) , this might be due to low adherence property of cells to the scaffold material as it was suggested. We are working on Mesenchyma stem cells of human placental blood origin, which in general shows very good adherence (in culture flasks) and it is seeded to scaffold of a chitosan derivative with other biodegradable polymer. Scaffold has known bio-compatibility , still we are unable to get an image under SEM (Scanning Electron microscope).

Now the problem appears to be in focus. In order to get rid of ethanol you must perform CPD as suggested by Stephen Florczyk. Otherwise air drying will never remove completely ethanol. Moreover some water vapor present in air could be attracted in the sample by the avidity of residual ethanol. Therefore microresidues of ethanol/water in the samples could vaporize at the moment of air evacuation from the observation chamber and make the cells burst so that you do not see them any more. Good luck.

Dear Parnita,

For cell seeded hydrogels ESEM analysis under low vacuum will help.

Dear Parinita,

I will suggest you to freeze dry your samples instead of using ethanol.

Freezing-drying might be helpful however at the end: a) you'll have to heat the sample at least at 37-60 °C in high vacuum in presence of P2O5 in order to get rid of residual water usually present in freeze-dried materials; b) freezing itself is likely to induce some distortion within the sample as related to the degree of structural stabilization provided by the fixation process and the way the freezing process has been carried out.