Dear Rohan

Firstly, why are you using a pre-embedding technique? Have you tried post-embedding IHC with acrylic resins such as LR White or Lowicryls in conjunction with light fixation and/or low temperature embedding such as PLT?

Regarding IHC with peroxidase/DAB, this can be rendered electron-opaque with gold chloride or osmium tetroxide, the former giving greater contrast of the polyDAB deposits. The peroxidase/DAB system is, however, incompatible with Lowicryl K4M. Alternatively, use colloidal gold.

If you are restricted to a pre-embedding protocol, for whatever reason, it might be worth trying light fixation to begin with, followed by IHC, and then further fixation and embedding.

I can provide some references for most of the above if you need them.

Cheers

Chris

Hi!

I would also suggest LR white. In case of LR white you do not even need to remove the resin to do your staining.

Would it be possible for you to try freeze substitution following high pressure freezing? THis would give you the perfect morphology.

We have good results with fixation of tissues/ cell pellets/bacteria using PHEM buffer containing either 1 % PFA for immunostaining or 2,5% GA for fine structure.

I would recommend using a gold-labeled sec. AB, either colloidal gold, or nano-gold. If you need to enhance the signal you can also do this with silver enhancement.

I could provide protocols for a lot of stuff in case you're interested.

Cheers

Elisabeth

I concur with Chris. Trying to do pre-embedding on tissue pieces is very limiting in itself due to the problem of reagents defusing into the tissue at a consistent rate. You may get labeling in the outermost areas but these are most likely to have the most mechanical damage from cutting. The best fixation would be perfusion fix and glutaraldehyde should be kept to a minimum (excluded initially) as it often interferes with labeling. High pressure freezing may be desirable but is difficult with nervous tissue and sample size must be very small...thus more chance of mechanical damage.

If perfusion is an option than you might try a perfusion fixation technique, used by the well-known Hungarian scientist, Dr. Zsolt Lipositz, that involves perfusing the animal with a low-pH fix followed by high pH that promotes cross-linking. I do not have the details (used many years ago) but you can find the references via the web. Look for ICC papers done in the 80's. I would then recommend embedding in HM-20. You can try this with infiltration and polymerization, using UV light, at -20oC. Then section (sections well) and react with your antibody using 10nm colloidal gold as the marker. The HM-20 at -20oC has worked well for me in the past for membrane protein localization when RT fix and embedding in LRW or other resins did not.

I agree with most comments and keeping GA as low as possible if you fix your sample at room temperature. Embedding in acrylic resins (LR White or Lowicryls) is also a good idea. Best possible structure preservation you will get with high pressure freezing and freeze substitution and believe me or not you also get labeling if you finally embed in Epon - see Ref:

High-Pressure Freezing and Freeze-Substitution of Native Rat Brain: Suitability for Preservation and Immunoelectron Microscopic Localization of Myelin Glycolipids by Eckhard Kirschning,* Gabriel Rutter, and Heinz Hohenberg - Journal of Neuroscience Research 53:465–474 (1998).

Good luck with best regards Roger

I won't be original and also agree onthe previous comments with the emphasis on the last one from Dr. Wepf about HPF. The only remark I can make - if you are trying to label some soluble cytoplasmic protein it can be very tough. We tried to do that for AP1, got some results, but even ourselves we did not consider those results convinsing and presentable... Membrane proteins are easier to label.

We believe the problem are the detergents, at any concentration. We perfuse-fix with 4%PF-0.2% glute. 50-100 micron vibratome sections are cut and processed free floating. No detergents, but with extended incubation times, 2-4 days for each primary, secondary, and ABC or immunoglobulin second antibody method. Flat embedded sandwiched between a liar film. If you take ultrathin sections on edge you will see that there is staining on each side extending about 5 microns deep , the center unstained. The surface 0.5 micron or so is frazzled. So, the goal is to take ultrathin sections at about 1-4 microns deep. The ultrastructure is pretty good.

EM Processing Protocol of Myelinated Tissue

Anesthesia: 0.5 cc of 2.5% Avertin injected IP

Perfusion: Perfuse for 30 seconds in rat ringers at 37˚C. The rat ringers also contains heparin(2.0 ml/L) and 2% lidocaine(3.0 ml/L) .

Perfuse for 7-10 minutes at 37˚C in 2%glutaraldehyde/1%paraformaldehyde in 0.15M sodium cacodylate buffer, pH 7.2.

1˚ Fixation: Immersion fix 2-4 hours at 4˚C with 2%glutaraldehyde/1%paraformaldehyde in 0.15M sodium cacodylate buffer pH 7.2.

Rinse: 6X20' each followed by an overnight rinse at 4˚C in 0.15M sodium cacodylate buffer pH 7.2 .

2˚ Fixation: Fix for 4 hours at 4˚C with 1% osmium-ferrocyanide in 0.15M sodium cacodylate buffer . Final concentration is 1% osmium tetroxide and 1.5% potassium ferrocyanide. Make up by adding 0.015grams of potassium ferrocyanide per ml of 1% osmium tetroxide/0.15M caco.

Rinse: 3X10' each in millipore filtered water at 4˚C.

En Bloc Stain: Stain for 1 hour at 4˚C with 2% uranyl acetate(aq.).

Rinse: 2X5' each in millipore filtered water.

Dehydrate: 1X8' each in 30, 50, 70, 95, 2X 100% ETOH(on ice at 4˚C).

2X8' each in propylene oxide at RT.

Infiltration: 1X60' in 25/75 ratio of propylene oxide and Durcupan(Epoxy Resin).

3X120' each in Durcupan.

Embed: Freshly made Durcupan is poured into the labeled embedding molds, then the sciatic nerves are put into the proper orientation for polymerization. The specimens are then cured for 24-36 hours at 65˚C.

1µm Thick Sections: The resin blocks are trimmed by hand using acetone cleaned teflon coated razor blades and then sectioned using glass knives. The 1µm sections are cut and mounted on glass slides then stained with Toluidine Blue.

Silver Sections: 700 Å thin sections are cut using a diamond knife. These sections are picked up and supported on 3mm diameter, circular, 200 mesh, copper grids.

Post-stain: The silver sections are stained in 2% uranyl acetate(aq.) for 30', rinsed in millipore filtered water, stained in Reynold's Lead Citrate for 8' in a carbon dioxide free environment with a final rinse in millipore filtered water. These sections are air dried and then viewed at 60kv on a JEOL 100 CX conventional transmission electron microscope.