you can use different combinations of resin and do infiltration in vaccum. If your sample floats on the surface infiltration is bad. Try to settel your sample

Hi Ele, you don't mention the plant species you are processing, which may have a bearing on the properties of infiltration. I had experience at Scottish Crop Research Inst. with several plant and insect species and some including narcissus were particularily problematic during infiltration.

My suggestion would be to use smaller excised areas of material, and post propylene oxide dehydration, infiltrate in 1:1 PO:Araldite for >4 hours, 1:3 PO:Araldite - overnight and then Araldite for 16 - 96 hours with 2 changes minimum. Araldite Resin - Araldite CY212 (200 ml), DDSA (200 ml), DBP (20 ml) and BDMA (10 ml).

Regards, Euan

Hi Ele..I think you should fix the sample with a mixture of gluteraldehyde(2.5%) and paraformaldehyde (2%). After treatment with osmium tetroxide, start dehydrating your samples in gradation of alcohol in vacuum. Each 30 mins duration. Then infiltration with toluene, 2 changes 2-3 hrs each. Then toluene : resin (3:1), then toluene: resin (2:2), then in pure resin, 2 changes. the resin is Araldite CY212, (from TAAB Laboratories).

I think this will work with your samples.

Hi Ele,

You first need to get your glut into the samples. If you can do that then the resin infiltration should be straightforward (if you get the samples fixed OK I personally do not think that the resin type will make much difference - if you are after immunolabelling then use LR White or equivalent, but for good ultrastrucfture I would simply proceed as normal with OsO4, Epon/Araldite etc - whatever your preference).

The main barrier to infiltrating leaf tissue with aqueous solns (eg. glut in buffer of any type) is hydrophobicity of the leaf surface (largely due to waxy cuticles, but trichomes and glands will make this worse, as yoiu have found) and large volumes of air in the intercellular spaces, so I have always used vacuum infiltration to fix leaf material: - cut the leaves into small pieces (eg. 1 x 2 mm) and place them in a vial containing the glut. soln, and then apply a weak vacuum to the tube for a few seconds (place a stopper on the vial, but ensure that it has holes!). Release the vacuum and apply it again - repeat this on and off until the samples sink to the bottom of the vial containing the glut soln. Leave in the fridge for a few hours and check that none of the samples have floated back to the top. If all remain sunk then you should then be able to proceed with dehydration, resin etc.

One final point - if vacuum alone does solve the problem then add a very small amount of a detergent to the glut soln., such as 0.1% Triton-X100. This always does the job, BUT there is a slight deterioration in ultrastructure, so please be aware of that.

Good luck!

O.K., since I was asked (automatically) by the ResearchGate provider to react on the posting () I do so honestly and would like to chime in here fearlessly.

Dear Ele Mira, supporting you with any information how to get rid of or come over the problems you face with your leaf-specimen preparations it is of utmost interest to the colleagues (which are wanting to help you)

that you first of all give informations on:

i) the kind of plant tissue you are dealing with,

ii) previously used specimen preparation regime (starting with the size of your leaf specs – fixation – dehydration – intermedium – resin embedding – polymerization … i.e. the whole procedure in brief, naming also the substances / solutions you use (including company name(s) for the resin).

There are a lot of ways to ‚get to Rome’ or ‘Athens’, if you like, but you must tell us “which direction” you’ll start for… so there is no real starting point for a thorough discussion on the subject if most of necessary informations are lacking in your request. One item which certainly should be considered seriously is perhaps the humid climate in your area / environments (eventually also in your facility/lab): if it is too humid you can be sure that using not fresh solutions you’ll end up with bad preparation results due to deterioration of e. g. alcohols (or acetone, we don’t know what you are using), your intermedium (assuming epoxypropylene = propyleneoxide = PO, Acetone, or Acetonitrile= AN) and the correct designation of the resin you use (low viscosity resin from AGAR assumed to be SPURR’s resin, the new ‘formulation’ of which perhaps adds some other problems to the outcome of your embedding results). TOLUENE as intermedium, as Anirban Roy tells you/us, might not be a real choice, since it is a cancerogen, highly toxic and a hazardous chemical which should be abandoned for use. I agree mostly to the opinions of Euan Milne and Euan James but should be really careful when using vacuum during your DEHYDRATION steps (solutions will start to ‘cook’ over). If you do have access to the Internet you should google your matter, e.g. with a search phrase like: < Plant leaf AND Electron microscopy AND preparation > (yields over 1,2 Mio results), < "Plant leaf" AND Electron microscopy AND preparation > (yields 9.400 results), < "leaf" AND Electron microscopy AND preparation > (yields 468,000 results) or < "leaf" OR "leaves" AND specimen OR sample AND preparation AND "Electron microscopy" > yields some 428,000 results. Refinement, e.g. by choosing other index terms or adding the ‘plant’s name’ you use for your study perhaps might lower the number of results being then more specific to your problem. It might also be advisable to read specific literature, especially with emphasis on modern techniques other than “classical chemical fixation”, like physical fixation, i.e. Freezing/cryosectioning, Freeze substitution, HPF (High Pressure freezing), or a fixation with or without ‘Microwave assisted’ specific protocols, respectively. Looking forward to read your kind and expected answer…we all are interested in helping you seriously with your matter,

best regards and wishes, Wolfgang MUSS, PhD,

Electron Microscopist, EM-Lab, Univ.Inst.Pathology SALZBURG-Austria/Europe

Thanks for this Wolfgang. With regard to vacuum infiltration I actually only advocate it's use during the fixation step (and not the subsequent dehydration steps - for the very "overcooking" reasons that you mentioned). It has always seemed to me that if you still feel the need to use vacuum after fixation then you still have the same problem (floating samples that ultimately do not get infiltrated with resin and have poor ultrastructure). So, if after attempting to remove the air from the leaves in order to infiltrate them with fixative (via vacuum), you still have floating samples then there is no point in proceeding to dehydration and resin. Discard and start again (perhaps with the T-X100 I mentioned).

Cheers

Euan J

Hi

First, thanks for your reply

So, Mr Wolfgang Muss, i can tell you i made some research on google and specific litterature before sending this post, and try a lot of "recipies"but none of them gave perferct results.

In my lab, microscopists are only physicians or animalist so they can't help me with vegetal stuffs and the specialized labs provides a lot of expensive formation but keep their technical advices...

I agree that my post lack of precision an will fastly correct that.

First, I work on lamina of the leaves of angiosperm woody species,

i consider 20 different species of tropical dry, wet and coastal forest.In their leaves, i cut samples of 1mmx3mm, avoiding midrib

The best result obtained was following this protocol.

I do fixation in a glutaraldehyde 4% solution during 2h (at ambiant T° because cells of leaves of tropical species are known to collapse at 6°)

and change with a NaH2PO4 buffer at 25mM (3 baths of 20 min) as my field stations are far from my lab, i directlly do the fixation on the field, tho avoid any degradation in my tissus

i do deshydratation with successive bath of absolute ethanol (gradation of 10°, 30° 50° 70° 90° 100 100 100), but not under vacuum

i make a 1/2h bath of pure propylenoxyde and then, i begin the infiltration using PO and a fresh mix of resin (as I said, Agar resin kit, low viscosity)

1bath 24h 2PO:1resin; under agitation

24h 1PO:1resin under agitation

24h 1PO:2resin under agitation

24h x3 pure resin under vacuum in every bath, I put 10x the volume of my sample

Polymerisation is made at 70°during 12h

ialready try classical spurr resin , the impregnation was very bad so i thought it was because of the high viscosity of this product

I also already try short times of impregnation but the problem of penetration is worst

I really care about the flotabillity of my samples during all the operations, they never float when i change baths.

As i told you, that's particular structure wich are always bad embedded: cells with high cellular walls, glandular cells.

Maybe the using of T-X100 as suggested by Euan James could help to improve the permeability of theses structure.

It's hard to do the fixation under vacuum because some of my plants grow far away from my lab and the transport dry them (and i guess that proteic digestion begin few minutes after the cut)

so in order to preservate the integrity of my tissues, i make the fixation directlly in the field.

I'm the first to integrate microscopic approch in my lab so my financial means are very limited (i'm a little PhD student so my boss let me play with little cheap resins but that's all),

of course Freezing/cryosectioning or Freeze substitution sounds good but we doesn't have the equipment to consider them seriouslly and i'm in a little island far from everything, the transport of fresh leaves to an equiped lab is impossible so i make what i can with what i have.

So thank you very much for all your messages, that was my first post on this site and i'm surprised to receive so much answers.

Dear Ele Mira,

from your posting I see the tremendous troubles you could go in considering your work in "the fields". I have been going shortly through your spec.-prep-schedule-but not to depth (today is Sunday here and I am just for a very limited time in the Lab to sort some thing here). So, please, allow for one or two days to carefully evaluate your informations and perhaps - if my brain is working right - we'll find one or two points to be improved in your protocols.

From others I know too that (T-)EM of leaves, especially with "hard" leaves is very difficult. Since you have told us how you do it for now, we can imagine a little better what your work is....and - I assure you - it is very hard! Compliments that you haven't given up here, for now: best regards and have a good weekend too,

sincerely yours

Wolfgang (Salzburg, Austria)

Dear Dr. Wolfgang, I know that toluene is a carcinogen and not good for skin and eyes etc. Can you please suggest any alternative for this chemical? Mostly I do samples are of oral cancer tissues of humans.

Dear Anirban,

(as well as dear Ele Mira), first of all: thank you very much for your kind reply, also entitling me as "Mister" and "Dr. Wolfgang" MUSS (latter is my family name). I am a very informal person and therefore you do it quite right if you just address me as a colleague "Wolfgang", be it in success as well as in (scientific) doubt- as you all I understand too this Research Gate Forum to be an interesting possibility and opportunity for rapid informations on specific or global themes which might arise during your studies, research projects etc, and I think also this to be a very informal matter. So hopefully is not angry about being addressed/entitled only with his/her first name...that includes me too.

As to Anirban's question me suggesting an alternative for Toluene (AS INTERMEDIUM for resin embedding):

IMHO (in my honest opinion) there are several possibilities:

depending on the resin you use (yours is Araldite CY212 ):

a) without an intermedium from 100% EtOH (or acetone) into resin mixture (e.g. 1:1) : EtOH (needless to say that that approach needs good knowledge about the physics of dehydration-embedding in resins and chemicals/resins used, and at least the lab climate should be relatively arid)

b) from 100% EtOH (or Acetone) into propylene oxide (PO) OR acetonitrile (AN= methyl cyanide). To stay in Acetone for afterwards mixing it with Araldite (but perhaps as well as for other resins like Durcupan, Epon, Epon substitutes, etc.) is not really recommended as the literature read so far reports of brittleness of blocks produced (problems in polymerization).

PO is the chemical usually used as an intermedium in TEM preparations but meanwhile - at least in Western Countries - has been marked (not forbidden) too as being carcinogenic like Toluene, so you should take care of with safety actions when working with it.

Acetonitrile is a not commonly used candidate since it is miscible with EtOH, Acetone and other organic solutions, as well as the resins. It is not classified a carcinogen (despite being a cell tox, = methyl cyanide), has a remarkably lower vapor pressure than PO, so you need not stringently to apply harsh safety issues when working with it. On the other hand (especially important for tropic regions or areas with high relative humidity) the 100% miscibility with water is a weak spot since you won’t like to adsorb water again in your embedding procedure/resin and you should take action to get rid of the solvent out of the resin properly (i.e. by means of a light bulb over the open specimen vials in a specimen rotator). But for Labs with moderate room climate this should be no issue. If you would like to know more about this method I certainly can give you proper informations how this is done routinely in my/our Lab since 1985 instead of using PO any more.

c) as another method of processing AND embedding I could offer to you a “rapid processing/dehydration and embedding method” which is done by dehydration in/with 2,2-DMP (dimethoxypropane) acidified (method on request) which is as old as my “student and professional career”(if I had any): originally reported from the 1970ies, and used not much around the EM-Labs in the world, I guess, but always good to know such a method exists: the molecular stoichiometric dehydration 1:1 by acidified 2,2 DMP means that this organic solution brought into specimen vessels containing tissue specs and water/buffer, or also say 50% EtOH, will react 1:1 / molecule H2O in the tissue, generating Acetone and Methanol as reaction product. Acidified 2,2-DMP can also be used to be mixed with the pure resin (and therefore you don’t need toluene or PO for intermedium). Alas two points: if you use acidified 2,2-DMP as a dehydration agent you should be aware of the fact that your specs should be free of PO4-ions (which is of no means when using sodium-cacodylate buffer) since such would be precipitated by the generated Methanol (so after osmication you should do 1-3x a wash with A.(bi)dest.).

I understand that not only the non-rehydrating/immiscibility properties of Toluene might be an issue for using it in your area and on the other side also perhaps the cheapness of getting the substance. And I know by myself that it might be necessary to use that stuff anyway if there is no possibility of using a substitute for that chemical. Additional questions (e.g. concerning the techniques or how to do it I am willing to answer to the best of my knowledge. Best regards and wishes, Wolfgang