Dear Saeed,

sorry for delay with my promised answer. Concerning your statement:

1) Can I use formaldehyde (FA) instead of Glutaraldehyde (GA) ?

2a) Or can I substitute osmium tetroxide with another material? 2b)osmium tetroxide is necessary?

3) What Hexamethyldisilazane do?

4) (a)Fixation with formaldehyde and (b) dehydration with ethanol may work?

my reply is:

ad 1) for sure, you / one can, as using any other fixative you intend to use… BUT: have in mind that the classical preservation of ultrastructural details (EM, SEM, TEM) always is done by fixation with at least glutaraldehyde (means: always appropriately and well adjusted buffered solutions of…!) These mixtures may vary a) with respect to pH and b) to osmolality) with the type of tissue / cells to be fixed.

If you read specific literature on fixation you’ll find out that only (buffered) formaldehyde (FA) as a non coagulant fixative will not be the right one for your task.

(e.g. cf. also ResearchGate postings https://www.researchgate.net/post/Principle_behind_cell_fixation?; https://www.researchgate.net/post/Cell_monolayer_Electron_Microscope_protocol?; https://www.researchgate.net/topic/Histology/post/4_paraformaldehyde_for_fixing_cells-4_degrees_or_room_temp?; http://www.researchgate.net/topic/Histology/post/Alternatives_to_formalin_fixation?

Typically for cultured cells (cf. also the method/schedule presented by Bhuminder Singh above) those preparations don’t need long fixation (e.g. flushing the cells if you want with iso-somotic or SLIGHTLY hyperosmotic buffered fixative, then substituting with fresh e.g. 1.5 – 3% buffered GA for 10-15 min @RT or at 4 degr C over night will be sufficient, after that carefully washing and proceeding to dehydration (see below (4) )… the duration of initial and primary/secondary fixation ought to be changed (prolonged) if the supporting layer is Chitosan and you need to demonstrate/image cells also with the attached Chitosan scaffold.

ad 2a) It might be possible to substitute osmium with another material or to process cultured cells just by omitting OsO4 completely - BUT IMHO (in my honest opinion) I doubt you’ll get optimal fixation especially of lipid(ic) material / membrane preservation (in the classical sense) of your cells and imaging by SEM might be low in contrast (description by far not exhaustive).

ad 2b) IMHO: YES. There have been publsihed a lot of articles using so called (= ) like G-OTO or G-OTOTOTO (G= GA, O= Osmium, T= Tannic acid)-sequences (for rendering samples/images with more contrast in SEM). Using such techniques might depend on what your expectations on SEM imaging were.

ad 3) Buminder Singh already has answered that. In addition I enclose the link for the suggested paper(PMID: 14599106):

http://www.ncbi.nlm.nih.gov/pubmed/?term=14599106

J Electron Microsc (Tokyo). 2003;52(4):429-33.

.

Araujo JC, Téran FC, Oliveira RA, Nour EA, Montenegro MA, Campos JR, Vazoller RF.

Departamento de Hidráulica e Saneamento, Universidade de São Paulo, Escola de Engenharia de São Carlos, Av. Trabalhador São Carlense 400, São Carlos-SP, 13566-590, Brazil. [email protected]

[Abstract

We present a fast procedure for scanning electron microscopy (SEM) analysis in which hexamethyldisilazane (HMDS) solvent, instead of the critical point drying, is used to remove liquids from a microbiological specimen. The results indicate that the HMDS solvent is suitable for drying samples of anaerobic cells for examination by SEM and does not cause cell structure disruption.

(if you are in need of the article, write to me - I could provide a pdf).

ad 4)

a) Yes and No (see above) - b) YES

for me: ethanol always is the preferred dehydration agent, but other chemicals have been used too (acetone, methanol, isopropyl-alcohol, 2,2-DMP acidified, etc.)

% Ethanol ascending series:

Buminder in his previous post has shown how this is to be done (a possibility out of several)..

• 35% Ethanol (due to a possible of tissue which is said to occur when starting with 30% EtOH I personally would start with 50% EtOH. Starting dehydration with 70% could favour precipitation of PO4 = phosphate from buffer on and in the tissue)…

• 50% Ethanol

• 70% Ethanol

• 95% Ethanol

• 100% Ethanol

• 100% Ethanol (store tissue here if critical point or chemical drying will not be performed immediately)

7. Replace the final 100% Ethanol wash with hexamethyldisilazane (HDMS), let HDMS evaporate in the hood. …..

It could be that you are in doubt about using hazardous substances like GA or Osmium, or – haven’t worked with those chemicals either, perhaps you’ll be afraid of handling and using these chemicals in your lab. Or is there any special cause you want to omit GA and/or OSO4? Any restrictions for the lab you are working? Maybe you also don’t have access to Critical Point Drying in your lab at all (examination of specimens in an outside facility?). It would be fortunate and of benefit to get some (your) answers therefore. Unfortunately we (at least I) do not know your institutional affiliation and country so we have to guess a little bit about the capacities you might have to solve your task.

Best wishes and regards, Wolfgang

Dear Saeed,

I think "aldehyde(s)" are the right way for fixation.Whether "plain" formaldehyde (FA) solution will work, can be doubted about. For SEM AND Imaging your specs it is advisable to think at least about "Glutaraldehyde"(GA) (perhaps initially a mixture of FA&GA), and - not to be forgotten: all fixatives in the right buffer (tonicity, pH, etc...)

Perhaps as only one example to start with reading about your sepcific matter / request I would recommend (knowing out there will be found several specific articles dealing with chitosan and "cells") the following article (open access):

http://www.groupes.polymtl.ca/tissue/Publications/PDFs/MB91.pdf,

M. Iliescu, C. D. Hoemann, M. S. Shive, A. Chenite, M. D. Buschmann:

Ultrastructure of Hybrid Chitosan-Glycerol Phosphate Blood Clots by Environmental Scanning Electron Microscopy (2006) (first place in Google search for: < Chitosan specimens SEM fixation > followed by similar articles) Also of interest (concerning properties of Chitosan would be perhaps http://trace.tennessee.edu/cgi/viewcontent.cgi?article=1629&context=utk_gradthes&sei-redir=1&referer=http%3A%2F%2Fwww.google.at%2Furl%3Fsa%3Dt%26rct%3Dj%26q%3Dchitosan%2520specimen%2520preparation%2520sem%2520fixation%26source%3Dweb%26cd%3D2%26ved%3D0CD0QFjAB%26url%3Dhttp%253A%252F%252Ftrace.tennessee.edu%252Fcgi%252Fviewcontent.cgi%253Farticle%253D1629%2526context%253Dutk_gradthes%26ei%3DB3nyUfW7I4LdOrelgLAN%26usg%3DAFQjCNE8RBUrB1v4Sew1-qxLzxlBxwzr5Q#search=%22chitosan%20specimen%20preparation%20sem%20fixation%22,

= Cao, Zheng, "Developing Chitosan-based Biomaterials for Brain Repair and Neuroprosthetics. " Master's Thesis, University of

Tennessee, 2010.

http://trace.tennessee.edu/utk_gradthes/609

and

http://library.iyte.edu.tr/tezler/master/malzemebilimivemuh/t000452.pdf,

Oktay YILDIRIM: Preparation and Characterization of Chitosan /Calcium Phosphate Based Composite Biomaterials.

Regards, Wolfgang

Pasting my protocol below. Definitely recomment using glutaraldehyde.

Preparing tissue/cells for SEM

SEM buffer: 0.1 M Na-phosphate buffer, pH 7.4 containing 0.1M Sucrose

Protocol:

1. Rince tissue with warm HBSS /and slice in small chunks

2. Fix with warm 3% glutaraldehyde in SEM buffer. Shift to cold for overnight (4C)

3. Washout fix with SEM buffer (2 x 5 min each)

4. Fix with 2% osmium tetroxide in SEM buffer on ice for 1 hr

5. Washout osmium with SEM buffer (2 x 5 min each)

6. Dehydrate tissue with the following series (15 min in each wash):

• 35% Ethanol

• 50% Ethanol

• 70% Ethanol

• 95% Ethanol

• 100% Ethanol

• 100% Ethanol (store tissue here if critical point or chemical drying will not be performed immediately)

7. Replace the final 100% Ethanol wash with hexamethyldisilazane (HDMS), let HDMS evaporate in the hood.

8. Mount on stub and coat with gold or platinum in a sputter coater.

Dear Bhuminder,

thank you for posting your protocol.

Allow 2 questions just for clarity:

1) SEM-Buffer:

1a) 0.1M Na-phosphate buffer, guessing self made Na-Na-buffer or purchased '0.1M Na-PO4 buffer soluton' ?

1b) any thoughts on use of Na-Cacodylate or a zwitterionic (e.g. HEPES-)buffer ?

1c) any suggestion/thought about why using 0.1M Sucrose (= 3.423%) instead of using 0.13M (isotonic) Na-PO4-buffer?

1d) any contradiction to use the 0.1 M SEM-buffer to wash the specimens after culturing instead of 'warm'(=RT?) HBSS?

2) 'warm 3% GA in SEM buffer': with warm you mean 'room temperature' or another elveated temperature (if the latter, what temperature you recommend?)

Thanks, best regards, Wolfgang

Dear Wolfgang Muss,

1a. Prepare buffer yourself (using mono and dibasic salt forms)

1b. Phosphate buffer works just fine, cacodylate is toxic also.

1c. Either sucrose concentration 0.1M and 0.13M should work.

1d. HBSS keeps cells happier before fixing. Recommeded if you have many samples, otherwise phosphate buffer should be fine also.

2. Warm means RT.

Good luck

Bhumi

Dear Buminder (hope you'll accept this entitling),

thank you for the straight advices and explanation....

Hope that Saeed will follow these posts....

Best regards, Wolfgang

Dear Wolfgang,

Hope it helps Saeed or other interested readers.

Regards

Bhumi

Dear Bhuminder and Dear Wolfgang

Thank you very much for your exellent responses and comments

Actually I was some kind of busy and I will do this protocol in next week. My cells will be ready in next week. I am gathering protocol for that.

Thank you very much for protocols

Now I have some questions

Can I use formaldehyde instead of Glutaraldehyde?

Or can I substitute osmium tetroxide with another material? osmium tetroxide is necessary?

What Hexamethyldisilazane do?

Fixation with formaldehyde and dehydration with ethanol may work?

Thank you again

Hi Saeed,

HDMS is an alternative to critical point drying (PMID: 14599106)

I recommend glutaraldehyde and osmium tetroxide

Dear Saeed, will comment on tomorrow morning, best, Wolfgang.

Unfortunately I deleted some second ago the whole text of my reply post here, so I have to do it again....please allow for another hour...

best, Wolfgang

Dear Saeed,

sorry for delay with my promised answer. Concerning your statement:

1) Can I use formaldehyde (FA) instead of Glutaraldehyde (GA) ?

2a) Or can I substitute osmium tetroxide with another material? 2b)osmium tetroxide is necessary?

3) What Hexamethyldisilazane do?

4) (a)Fixation with formaldehyde and (b) dehydration with ethanol may work?

my reply is:

ad 1) for sure, you / one can, as using any other fixative you intend to use… BUT: have in mind that the classical preservation of ultrastructural details (EM, SEM, TEM) always is done by fixation with at least glutaraldehyde (means: always appropriately and well adjusted buffered solutions of…!) These mixtures may vary a) with respect to pH and b) to osmolality) with the type of tissue / cells to be fixed.

If you read specific literature on fixation you’ll find out that only (buffered) formaldehyde (FA) as a non coagulant fixative will not be the right one for your task.

(e.g. cf. also ResearchGate postings https://www.researchgate.net/post/Principle_behind_cell_fixation?; https://www.researchgate.net/post/Cell_monolayer_Electron_Microscope_protocol?; https://www.researchgate.net/topic/Histology/post/4_paraformaldehyde_for_fixing_cells-4_degrees_or_room_temp?; http://www.researchgate.net/topic/Histology/post/Alternatives_to_formalin_fixation?

Typically for cultured cells (cf. also the method/schedule presented by Bhuminder Singh above) those preparations don’t need long fixation (e.g. flushing the cells if you want with iso-somotic or SLIGHTLY hyperosmotic buffered fixative, then substituting with fresh e.g. 1.5 – 3% buffered GA for 10-15 min @RT or at 4 degr C over night will be sufficient, after that carefully washing and proceeding to dehydration (see below (4) )… the duration of initial and primary/secondary fixation ought to be changed (prolonged) if the supporting layer is Chitosan and you need to demonstrate/image cells also with the attached Chitosan scaffold.

ad 2a) It might be possible to substitute osmium with another material or to process cultured cells just by omitting OsO4 completely - BUT IMHO (in my honest opinion) I doubt you’ll get optimal fixation especially of lipid(ic) material / membrane preservation (in the classical sense) of your cells and imaging by SEM might be low in contrast (description by far not exhaustive).

ad 2b) IMHO: YES. There have been publsihed a lot of articles using so called (= ) like G-OTO or G-OTOTOTO (G= GA, O= Osmium, T= Tannic acid)-sequences (for rendering samples/images with more contrast in SEM). Using such techniques might depend on what your expectations on SEM imaging were.

ad 3) Buminder Singh already has answered that. In addition I enclose the link for the suggested paper(PMID: 14599106):

http://www.ncbi.nlm.nih.gov/pubmed/?term=14599106

J Electron Microsc (Tokyo). 2003;52(4):429-33.

.

Araujo JC, Téran FC, Oliveira RA, Nour EA, Montenegro MA, Campos JR, Vazoller RF.

Departamento de Hidráulica e Saneamento, Universidade de São Paulo, Escola de Engenharia de São Carlos, Av. Trabalhador São Carlense 400, São Carlos-SP, 13566-590, Brazil. [email protected]

[Abstract

We present a fast procedure for scanning electron microscopy (SEM) analysis in which hexamethyldisilazane (HMDS) solvent, instead of the critical point drying, is used to remove liquids from a microbiological specimen. The results indicate that the HMDS solvent is suitable for drying samples of anaerobic cells for examination by SEM and does not cause cell structure disruption.

(if you are in need of the article, write to me - I could provide a pdf).

ad 4)

a) Yes and No (see above) - b) YES

for me: ethanol always is the preferred dehydration agent, but other chemicals have been used too (acetone, methanol, isopropyl-alcohol, 2,2-DMP acidified, etc.)

% Ethanol ascending series:

Buminder in his previous post has shown how this is to be done (a possibility out of several)..

• 35% Ethanol (due to a possible of tissue which is said to occur when starting with 30% EtOH I personally would start with 50% EtOH. Starting dehydration with 70% could favour precipitation of PO4 = phosphate from buffer on and in the tissue)…

• 50% Ethanol

• 70% Ethanol

• 95% Ethanol

• 100% Ethanol

• 100% Ethanol (store tissue here if critical point or chemical drying will not be performed immediately)

7. Replace the final 100% Ethanol wash with hexamethyldisilazane (HDMS), let HDMS evaporate in the hood. …..

It could be that you are in doubt about using hazardous substances like GA or Osmium, or – haven’t worked with those chemicals either, perhaps you’ll be afraid of handling and using these chemicals in your lab. Or is there any special cause you want to omit GA and/or OSO4? Any restrictions for the lab you are working? Maybe you also don’t have access to Critical Point Drying in your lab at all (examination of specimens in an outside facility?). It would be fortunate and of benefit to get some (your) answers therefore. Unfortunately we (at least I) do not know your institutional affiliation and country so we have to guess a little bit about the capacities you might have to solve your task.

Best wishes and regards, Wolfgang

Dear Saeed,

Following excellent advice from Wolfgang and Bhuminder you should get really good results.

Just a few comments.

1. Fixation at room temperature for 1-1.5 hrs with glut is sufficient, but you can leave specimens in fixative in a fridge if needed.

2. In most cases you can skip OsO4 completely. Sometimes it can help, but too often it is useless for SEM. For TEM Os is important: it helps to preserve inner morphology of cells, but in SEM we see only shape of cells. From the other hand Os can make specimen more rigid, so it may help to preserve shape of delicate features. So, if you have access to OsO4 you may want to use it, otherwise do not bother with it. As for Os-induced conductivity and contrast, it is negligible in comparison with coating (Au-Pd, for example).

3. If your culture is mineralizing culture and you are looking for mineral distribution, you may want to simplify protocol and work as fast as possible: all water-based solutions slowly dissolve mineral. You can start with 20 min fixation (increasing time if needed). Do not use Os at all, it will alter chemical composition and will be visible on BSE micrographs along with mineral. Decrease time in water-alcohol solutions to 5 min each(for example 33%, 66%, 85%, 95%); 100% alcohol will not dissolve mineral, so use 3 changes for 20 min each. HMDS also changes composition, but much less than OsO4. So, CPD is better, but if you do not have it, HMDS will work also (or you can skip this step completely if you can tolerate some additional cracks in cells)

Good luck.

Try methodology described in my ResearchGate page as brief book on "Electron Microscopy".

Thank you all

I have done the protocol with regard to these comments and I have got so nice and suitable photos.

Best Regards

Dear Dr. Vladimir

Can I only use the cyanobacterial lyophilized form for SEM or I must use the fixing step

thank you

Dear Lamia,

I do not work with cyanobacteria, so I do not know particularities of your specimens.

Just in general:

If you want to observe bacteria while attached to substrate, efforts should be made not to disturb (deattach) specimen, so you may want to avoid standard methods of biological specimen preparation (too many steps involving liquid replacement).

If you specimens are already frozen, then only freeze drying is advisable (thawing can damage them).

In general standard bilogical method (fixation, alcohol dehidration and CPD/HMDS) provide better preservation of cells, but some bacteria are tough enough to be just air dried. It's better to try several methods to choose the best for your case.

Hi everyone,here are some excellent preparation techniques as mentioned above.

I have a question to ask,mainly to Vladimir if possible.

I am fixing clots to analyse under SEM and I am not using Osmium at all.The end result is really good.As I am mainly interested in fibrin density and not directly at the cells I have good results.Is there a paper available to highlight that we can skip the fixation with Osmium especially when gold palladium is used as it will add nothing to fixation?

Your help is highly appreciated. 

Hi Nicolaos,

I did not work with fibrin, but try to google "fibrin sem" and you can find a lot of papers. Fibrin is a tough thing, it definitely can be easily preserved without OsO4. Actually, most SEM protocols do not include Os. It's either for those who do not know what are they doing, or for rare specimens with "special needs".

Thank you ever so much Vladimir.I understand this but in the medical field once someone has used one protocol (this is the end )and it seems everyone uses Osmium which I really dont understand as you ve previously highlighted as it doesnt offer anything for SEM.Thank you again,at least I can find some people to communicate as others, including my supervisor, do not seem to get it. 

Hi everyone

I had some bacterial cultures grew on sand grains. Since following the common protocols like what mentioned here would disturb my sample , I froze them for 2 hours in liquid nitrogen and then freeze-dried it for 48 hours. This method does not provide photos as good as using solvents but I couldn't find any better fixing protocol for small sand grains.

Now I am wondering that how long I can keep fixed samples. If I want to repeat the SEM analysis what the maximum time between fixation and SEM analysis can be.

Dear Nafise, it might depend on the way you store the specimens (with "keep fixed grains" you mean frozen & finally freeze-dried samples). I am sure, Vlad will chime in and provide real knowledge about storing and maximum storage time (;:-))  Best wishes and regards, WM

Thank you Dr. Muss

Yes by "fixed grains" I mean after freeze-drying. I have stored the freeze-dried samples at room temperature in capped vials and I am wondering what the maximum storage time(for SEM analysis) is.

Dear Wolfgang H. Muss,

After the dehydration in 100% Ethanol, I need to wash with any solution or directly freeze samples to then put in Freeze Dryer Machine? 

(Apologize for lengthiness): in Re to Nafise JamiAlahmadi  and Karen Yarasca

@Nafise JamiAlahmadi: I apologize.... didn't want to let you think that I am/was unfriendly...  but I just did not receive a notification that you answered the thread again long ago and requested a clue for the "maximum storage time [of FD-samples prior to SEM-imaging].

I confess: I don't have a solution or can sell you "the truth"...  but I guess   [thinking only practically, since I have not searched for specific literature or technical information on such matter]   that under certain conditions (i. e. proper decoration / sputtering and respective handling of your samples, further cleanliness, dryness of sample as well as the air volume  [eventually an overlay of inert, dry gas]   in your tightly locked capped vials)    the storage time might be a week, or even longer. As I once had colleagues working with such methods I got aware that they used to use either sophisticated or even "commercially available" dessicators, which contained a kind of dessication material (e.g. silica gel, generally hygroscopic material)* and were evacuated, at least up to approx. 2.5-4 x 10^-2 mbar, using either a dry or a rotation pump (NOT a water-jet-vac-pump!). I have seen those colleagues using such samples for multiple examination and analysis in SEM....and some of them stored samples longer than a year and more....

* cf. GOOGLE search for:

| desiccators and vacuum storage |        or also                                                         | desiccators and vacuum storage to preserve samples/chemicals|                           (only as ONE possibility):  https://www.tedpella.com/desiccat_html/desicatr.htm

also, please (if you haven't yet) try to read: Fischer ER et al, 2012: Scanning Electron Microscopy. Curr Protoc Microbiol. Author manuscript; available in PMC 2013 May 1.   https://www.ncbi.nlm.nih.gov/pmc/articles/PMC3352184/ ;  pdf: https://www.ncbi.nlm.nih.gov/pmc/articles/PMC3352184/pdf/nihms-375012.pdf

Published in final edited form as:

Curr Protoc Microbiol. 2012 May; CHAPTER: Unit2B.2.

doi: 10.1002/9780471729259.mc02b02s25

-------------------------------------------------------------------------------------------------

@Karen Yarasca, fortunately I found your reply some minute(s) ago, which, indeed, also turns out to be a question....     which unfortunately I cannot understand quite well. 

I wonder about your method [you are talking about CPD=Critical Point drying or FD/Lyophilization?] since you report on a method which I am not familiar with:  

you have [I am guessing, after proper fixation] "dehydrated a sample via ....alcohols / ethanols, ending in 100% ethanol..."  I am a bit confused...

With what - with which solution - or why -  should you need to wash the "totally dehydrated" sample? or, even in the case you NEED NOT to wash after full dehydration in 100% Ethanol, how would you [be able to]  freeze your sample effectively to put it then in a Freeze Dryer machine...     Unfortunately I cannot follow this your regimen, at least unless you describe your (needed or anticipated ) method more detailed ....

Please, consult / read:

e. g.:  http://www.spscientific.com/freeze-drying-lyophilization-basics/; 

http://www.eurotherm.com/freeze-drying

https://archive-resources.coleparmer.com/MoreInfo/Labconco_guide_freeze_dry_in_lab.pdf; 

http://freezedrying.com/freeze-dryers/general-principles-of-freeze-drying/;

http://journal.pda.org/content/39/3/115.short;

https://en.wikipedia.org/wiki/Freeze-drying:   'Freeze Drying' certainly is a kind of "dehdyration" process, but not by exchanging (cellular or tissue) water content with increasing alcohol concentrations (as used in histology or classical ultrastructural studies)  but [cit.] "is a dehydration process typically used to preserve a perishable material or make the material more convenient for transport. Freeze-drying works by freezing the material and then reducing the surrounding pressure [=under vacuum conditions] to allow the frozen water in the material to sublimate directly from the solid phase to the gas phase.

Regarding a sample in 100% ethanol:  cf. [-citation-]

Top 5 mistakes made in the lyophilization process

By Jenny Sprung, Senior Application Specialist

On Thursday, February 23, 2017

It’s been said that freeze drying is an art, not a science, but there are ways to help improve your artistic capabilities. 

Some of the top mistakes of the freeze dry process are…

1)     Not knowing your sample’s melting point

Without knowing what temperature your sample melts at, you can’t choose the correct lyophilizer for your needs, and your samples may melt during the process. A freeze dryer requires a temperature differential between the sample’s eutectic temperature and the freeze dryer collector. The collector must be 20 degrees colder than the eutectic temperature to allow for proper sublimation during lyophilization.

Example: Ethanol has a freezing point of -114C. If used in a freeze dryer, the collector temperature would need to be -134˚C. Unless you’re using a liquid nitrogen freeze dryer, freeze drying a sample in pure ethanol would be impossible. In fact, just freezing pure ethanol is difficult.  If you dilute ethanol with water, you can raise the sample’s eutectic temperature to a point that it could be freeze dried using a -105C freeze dryer.   

2)....

3).....    [end of citation].

[found @: http://www.labconco.com/news/top-5-lyophilization-mistakes

Personal remark: bold and italic fonts set by myself...

I admit that I have no practical experience with these techniques but somehow was instructed about these techniques and their physical background.  Hope this helps at least anyway, best regards and good luck, Wolfgang