Technical Tips and Recommmended Article Index

1 Stability of Special Standing Solutions:

General Guidelines Concerning Solutions

Check stock solutions regularly for signs of deterioration.
All solutions should be dated and initialed by the person who prepared the solution.
Use caution to prevent cross solution contamination.
Use distilled or double distilled water to prepare solutions.
Use reagent grade chemicals.
Use dyes which are certified .
Keep solutions in well stoppered bottles at all times.
Refrigerated solutions should be poured into the staining jar, and the bottled solution returned to the refrigerator before it warms to room temperature.
Return all staining solutions to shelves soon after use to prevent mix-up.
Always use clean glassware.
Make sure that balance, pH meter, etc., are in proper working order.

THE FOLLOWING IS REPRINTED FROM MIKRO-GRAF, The Michigan Society of Histotechnologists, Volume XX, Number 8, June 1992.

2. Tips for BUTVAR B-98 Support Films:

At the University of California, San Francisco a simplified and less hazardous method from the original Handley method( Handley, D.A. et al Butvar B-98 as a Thin Support Film. Ultramicroscopy 4, 479 (1979)) has been employed for the fabrication of Butvar support films. Antonia M. Milroy, Anatomy Department, University of California, San Francisco, CA. 94143-0452

FOR REPRINTS OF THE PROCEDURE PLEASE CONTACT US.

3. Lead Staining:

Lead stains are very sensitive and will precipitate quickly upon contact with CO₂. To prevent this from happening either during preparation, storage, or staining the following steps should be followed:

Prepare CO₂ free water: boil double distilled water and store it at once(while still hot) in tightly capped bottles or glass stoppered bottles. When these bottles are opened you should hear a hissing sound which indicates the bottles were sealed properly. The water bottle should be opened only prior to use. The above procedure needs to be repeated if ample amount of water remains in the bottle and the bottle was left open for an extended period of time (more then 10 minutes).
Construct a staining apparatus: A sheet of dental wax is placed on top of wetted filter paper(1N NaOH) and it is surrounded with several NaOH pellets inside of a petri dish which serves as the staining chamber. The NaOH will quickly absorb CO2 in the chamber, giving a CO₂ free area for lead staining.

FOR A DETAILED DESCRIPTION AND EXPLANATION OF LEAD STAINING PLEASE REFER TO THE FOLLOWING BOOK:

ELECTRON MICROSCOPY PRINCIPLES AND TECHNIQUES FOR BIOLOGISTS
BY JOHN J. BOZZOLA AND LONNIE D. RUSSELL. 1992 PP. 115-118

4. The Preparation of Adhesives Coated Grids for Picking Up Carbon Film to Make Carbon Coated Grids:

The following steps should be followed in the preparation of adhesive coated grids:

Submerge about 2" of Scott clear tape (3M) into 10ml of Dichloroethane (Ethylene Dichloride); shake and discard the tape.
The solution now becomes "grid-glue"
Place the grids(dull side up) on a piece of filter paper (dust-free room).
Take a pipette and place a drop of "grid-glue" on top of each grid.
Let the grids dry.
The grids are now ready to pick up the carbon foil and make the carbon coated grids.

FOR A DETAILED DESCRIPTION AND EXPLANATION OF SUPPORT FILMS FOR GRIDS PLEASE REFER TO THE FOLLOWING BOOK:

ELECTRON MICROSCOPY PRINCIPLES AND TECHNIQUES FOR BIOLOGISTS
BY JOHN J. BOZZOLA AND LONNIE D. RUSSELL. 1992 PP. 83-86

5. Neutralization and Disposal of Osmium Tetroxide:

A 2% solution of Osmium Tetroxide can be fully neutralized by twice the volume of oil (corn oil is preferred because of its high percentage of unsaturated bonds).1 That is to say, for every 10ml of 2% Osmium solution, 20ml of corn oil is required.

Procedure:

Do all work under a fume hood.
Pour twice the volume of corn oil into used Osmium Tetroxide solution.
Wait for the oil to completely turn black.
Test to check that complete neutralization has taken place.
(To confirm neutralization; take either a glass coverslip coated in corn oil or a piece of filter paper soaked in corn oil and suspend it over the solution. Blackening indicates OsO4 is still present.)2
Dispose in accordance to local regulations. Contact your Disposal Agency or Environmental Health & Safety Office at your Institute to obtain local regulations.

References:

Cooper, K. Neutralization of Osmium Tetroxide in case of accidental spillage and for disposal. Bulletin of The Microscopical Society of Canada. 1988. 8:24-28
Lunn, G.; Sansone, E.B. Osmium Tetroxide. Destruction of Hazardous Chemicals in the Laboratory; Program Resources, Inc. Frederick, MD; pg. 211-213

6. Regarding Embedding Media and Methods:

Of late there has been much conversation and discussion regarding different embedding media and alternatives to many of the most commonly used components. We would like to recommend the following publications that may be of interest to you regarding this subject.

Glauert, A.M. Epoxy Resins: An Update On Their Selection And Use. Microscopy and Analysis, September 1991, pp. 15-20.
Mascorro, J.A. and Kirby, G.S. Alternatives To Traditional Epoxy Embedding "Kits": Novel Choices. Proc. Ann. Meet. EMSA, 50, 1992.

Preparation of Embedding Mixtures

It is essential to mix the components of the embedding medium very thoroughly to obtain uniform polymerization. Inadequate mixing is one of the main causes of sectioning problems with epoxy resins. The simplest way to obtain good mixing is to warm the epoxy resins, the hardener, and the containers to be used for measuring and mixing as well as the glass rod to be used for stirring to 60°C. A uniform mixture is obtained in a few minutes by this method and no special stirring apparatus is required. The warm components also are easily poured from its containers for measurement. It is best to prepare the complete mixture just before use but if necessary it can be stored at 4°C in a bottle with well fitting cap for several weeks, or for many months at -20°C. During infiltration of the specimens the embedding medium is kept at room temperature and warmed to 60°C and stirred well before change of solution. CAUTION: ALWAYS WEAR GLOVES AND WORK UNDER A FUME HOOD. Glauert, A. M.(Ed.). (1975). Practical Methods in Electron Microscopy. Vol.3, Part I. Elsevier North-Holland Inc. NY.

7. The Premixing of Resin for Later Use:

In the June 1992 edition of MICROSCOPY TODAY Issue #5 appeared an article regarding the preparation of large batches of resin (minus catalyst) which is stored in the freezer (-70°C) and when needed it is thawed, the catalyst added, and this is it. This method saves time and reduces the amount of contact time with the chemicals. John J. Bozzola, Center for Electron Microscopy, Southern Illinois University, Carbondale, IL 62901

The Storing of Embedding Resin Mixtures:

Our experience over the years shows that the best way to store remaining embedding resin mixtures is in disposable plastic syringes.

Procedure:

Take a fresh "all plastic" syringe (EMS Cat. #72520-72529-chose the appropriate size syringe depending on the volume mixture to be stored), remove the tip cover, slowly draw-in the mixture. Remove any air space by holding the syringe with the tip in the upward position pulling the plunger slowly to clear the resin from the tip as well as letting the air move toward the tip; slowly push the plunger upward until the resin appears at the end of the tip; put on the tip cover. Wrap the syringe with aluminum foil, label it (name of mixture and date) and store it in the refrigerator. This method protects the mixture from contamination by condensation, and is also more convenient for future use.

8. Epoxy Resins:

Common problems, causes, and solutions:

Problems associated with poor embedding in epoxy resins:

Blocks are difficult to section
Sections may disintegrate upon contact with water or the electron beam
Holes in the sections
Unevenly cured blocks
Blocks are either too soft or too hard

Factors that cause poor embedding in epoxy resins:

The presence of water in absolute dehydration solvents and in the ingredients of the embedding mixture.
High environmental relative humidity.
Inadequate dehydration of the specimen.
Inadequate mixing.
Inadequate addition of the accelerator to the mixture.

Solutions to the causes:

Use either fresh or a well capped bottle of dehydration solvents.
Embedding should be carried out at relative humidities below 50%. Dellman, H.D. and Pearson, C.B. (1977). "Better epoxy resin embedding for electron microscopy at a low relative humidity". Stain Technol., 52:5
Since epoxy resin is hygroscopic, extreme care should be taken to prevent contamination by water.
Accurate measurement of accelerators.
Complete mixing without beating air into the mixture (Use a TFE or glass rod for at least 20 minutes).

9. Accelerators for Epoxy Resins:

Embedding media for Electron Microscopy based on the epoxy resins Araldite and Epon (replacements) all require the addition on an amine accelerator to hasten polymerization (Glauert, 1974).

-DMP-30

-BDMA

Both of the above accelerators are equally efficient during polymerization, but BDMA has the great advantage of being much less viscous than DMP-30. This is important since accurate measurement of small amounts of the accelerator and its complete mixing with the other components of the embedding media are essential for the production of satisfactory blocks. See Audrey Glauert (Proc. Roy. Micros., Vol. 22/5 Sept. 1987)
NOTE: In order to achieve optimum results when using BDMA in place of DMP-30 you must double the quantity of BDMA.

10. The Search for the Epon-812 Replacement:

Since the discontinuation by Shell of Epon 812 many replacements have been introduced. Many of these replacements have shown much closer characteristics to the original than others. For an in depth study of the work that has been done regarding many of the replacements for the original Epon 812 we highly recommend the following publications:

Mascorro, J.A. and Kirby, G.S. Physical Characteristics of "Old" Epon 812 and Various Epon Like Replacements". Proc. 44th Ann. Meet. EMSA., pp. 222-223 (1986)
Mascorro, J.A. and Kirby G.S. Novel/ Epoxy/Anhydride Alternatives for Biological Electron Microscopy: Physical and Performance Characteristics of Embed 812 and LX112 in Combination with NSA/NMA/DMAE. Proc. 47th Ann. Meet. EMSA., pp. 1000-1001 (1989)
Mascorro, J.A. and Kirby G.S. Viscosity Characteristics and Hardening Rates For Embed 812 and LX112 With Alternative Anhydride and Catalyst Choices. Proc. 49th Ann. Meet. EMSA., (1991)

11. The Perfect Loop for Section Pick Up

The Perfect Loop (EMS cat # 70944) allows you to pick up sections consistently without causing any damage to the sections. It is the only loop that is currently available where the outside diameter of the loop is the same as the grid and the inside diameter is slightly larger than the observation area of the electron microscope. The thickness is about 40 microns. Due to the fact that the loop and the grid are of the same diameter they are attracted to one another when in water and attach together through the surface tension of the water.
Even if the section touches the inside of the grid during blotting the touching area is minor and, therefore, the section is not damaged.
When the grid is removed from the loop the section remains in place without fail. The area equals the observation field (about 2mm diameter) of the electron microscope; thus pieces can be fully observed.

12. Safety in the Electron Microscopy Laboratory:

All of us at Electron Microscopy Sciences are concerned for the safety and well being of the microscopy community. For this reason we highly recommend the following publications and tips regarding safety in the lab:

ELECTRON MICROSCOPY SAFETY HANDBOOK; Vernon C. Barber and Deborah L. Clayton, Editors. San Francisco Press Inc., 1985 ISBN 0-911302-56-5
ELECTRON MICROSCOPY PRINCIPLES AND TECHNIQUES FOR BIOLOGISTS; John J. Bozzala and Lonnie D. Russell. Jones and Bartlett Publishers., 1992 pp. 498-519.ISBN 0-86720-126-6
SAFETY IN THE SCANNING ELECTRON MICROSCOPY LAB; J. Bastacky and T.L. Hayes, Scanning 7:255-72., 1985

Safety Tips:

Latex Gloves: Allergies and Alternatives

In the past several years there has been documentation in the medical literature of numerous cases of latex allergy in medical and other laboratory personnel. Many people are experiencing itchy hands, shortness of breath or other signs of allergic reactions when they use latex gloves in their laboratory work. Some people are changing to powder free gloves with some relief; however, other people get relief only after changing to another type of glove.

E. Ann Ellis, Ellis EM Consulting Service, 1316 SW 61st Terrace, Gainesville, FL. 32607

Surgical Gloves: Safety

The standard latex surgical gloves do not offer adequate skin protection when handling Glutaraldehyde. Nitrile or Neoprene gloves provide the protection that is needed. However, dexterity is inhibited with these gloves. Therefore, double-gloving with latex gloves is recommended for delicate work.

Raewyn Eager, Dept. of Zoology, Auckland University. EM News.
Newsletter of the New Zealand Society for Electron Microscopy. No. 24, Dec. 1992

Formaldehyde Update

What you need to know about the revised OSHA standard on Formaldehyde. Refer to: Vinnie Speranza. On Stage. A Quarterly Publication of the New York State Histotechnological Society. Vol 15, No. 1 Summer, 1992.

Toxicological Properties and Occupational Hazards of Acrylate Compounds

For a brief report of the toxicological properties and occupational hazards of acrylate compounds and a personalized summary of the authors own experiences with resins and protective devices. Also summarized is guidelines for safe embedding, knowing the possible risks, and how to minimize them.

M. Tobler and A. U. Freiburghaus (1990). Occupational risks of (Meth)acrylate Compounds In Embedding Media for Electron Microscopy. J. of Microscopy Vol. 160. pp. 291-298

13. Quetol and the Addition of Water:

The addition of 1% water to the epoxy resin Quetol increases the labeling intensity of the sample. The significant decrease of the curing temperature of the epoxy resin may assist in preservation of antigens. Water may also reduce the cross-linkage of the resin allowing more antigen to be available to the antibodies. The modified Quetol resin is an option for use in immunocytochemistry studies.

Refer to: Andre R. Abad, (1992). Medium Temperature Epoxy Resin for Immunocytochemistry: Quetol 651 With Water. Microsc. Res. and Techn. 20:274-280.

14. Pre-embedding, Post-embedding, Non-embedding method, Protein A-gold technique, Double Labeling:

This review deals with the use of major methods for ultrastructural immunohistochemistry including pre-, post-, and non-embedding methods, particularly focusing on their application to human pituitary pathology. Problems of tissue processing and a protocol for double labeling technique using the protein A-gold complex are also described.

Refer to: Toshiaki Sano, 1992. Use of Ultrastructural Immunohistochemistry in Human Pituitary Pathology. Microscopy Research and Technique 20:152-161.

15. Inspecting The SEM While In Operation:

It would be very convenient if the SEM could be inspectedwhile in operation, with the image of the vacuum chamber visible on the SEM TV screen.

Refer to: R. Eckert, 1992. Inspecting the SEM Chamber with a Charged Polystyrene Mirror. Scanning Vol. 14, 73-75

16. Depex (EMS Cat.#13514) is an Improved Mounting Medium:

Most mounting media cause fading of histological preparations over time. A study was conducted to find the most suitable mounting media that would not fade.

This study indicated that Depex as a Mounting Media appeared to have a conserving effect on the stain as compared to other available mounting media. The study was based on Araldite embedded semithin sections of Rabbit cerebral cortex stained with Toluidine Blue and Pyronin G. The study also indicated that the average optical density of sections prepared with Depex after one year was higher than the sections treated with other mounting techniques.

Cordula Schmolke(1993). Effects of Mounting Media on Fading of Toluidine Blue and Pyronin G Staining in Epoxy Sections. Biotech. & Histochem. 68/3, pg.132-136

17. Simple Method to Avoid Precipitation on Lead onto thin Sections:

Lead precipitate on thin sections has plagued electron microscopists for generations. There are many possible reasons for this occurrence, however one of the most common causes of precipitation of lead citrate staining solutions is an incorrect pH. According to Reynolds (1963), as the citrate solution approaches a pH of 12 , no staining results with the accompanying loss of contrast. Continued observations in this laboratory have shown that as the pH is lowered to 8.0, heavy precipitation occurs under all circumstances even if good chealation of the components has been accomplished by adequate shaking and inversion. The most common cause of faulty pH, is the use of sodium hydroxide pellets instead of a commercially produced 1.0N carbonate free sodium hydroxide solution. Since it is frequently not possible to weigh pellets accurately due to their size, and since the pellets may absorb water from the atmosphere, the actual pH of the final lead solution may be unknown. This variance may result in the same laboratory alternating between poor contrast of sections and precipitation of lead crystals.

Therefore, Reynolds lead citrate staining solution should be produced in the EM laboratory ONLY with a 1.0N carbonate free sodium hydroxide solution (EMS Cat. #21170), which has been commercially titrated.

Hildegard H. Crowley, Department of Biological Sciences, University of Denver, Denver, CO 80208

18. Acetonitrile is a Safer, Better Dehydrating Solvent for Transmission Electron Microscopy:

Propylene Oxide and Ethanol are commonly used dehydrating solvents for processing tissues for electron microscopy. But both solvents, however, have some undesirable properties: they are highly flammable, volatile, toxic and potentially carcinogenic. Acetonitrile is a direct substitution for ethanol and propylene oxide and it is safer to use and requires shorter dehydration times. It is freely miscible with water, alcohol, acetone and epoxy resin and it does not interfere with epoxy polymerization.

Harold H. Edwards, Yu-Yan Yeh, Betty I. Tarnowsky, and Gregory R. Schonbaum. (1992). Acetonitrile as a Substitute for Ethanol/Propylene Oxide in Tissue Processing for Transmission Electron Microscopy: Comparison of Fine Structure and Lipid Solubility in Mouse Liver, Kidney, and Intestine. Microsc. Res. and Technique Vol. 21, pg.39-50

19. A Vacuum Drying Technique may be an Effective Method to Eliminate or Reduce the Wrinkles in Semithin Epoxy Sections:

Wrinkled, folded, buckled or warped sections can be vacuum dried as follows: a vacuum desiccator was furnished with a layer of calcium sulfate, and with a Petri dish filled to a depth of 3 mm with Phosphorous pentoxide. A large filter paper (Whatman #1) secured under the tray of the desiccator prevented dust from contaminating slides during pumping. Slides, positioned on top of the tray, were kept in vacuum for 48 hours. Immediately after removal from the vacuum jar, and without immersion in any organic solvent, the slides are coverslipped with mounting media.

Hildegard H. Crowley and Ben H. Leightling(1989). Elimination or Reduction of Wrinkles in Semithin Epoxy Sections by Vacuum Drying. Stain Technology, Vol. 64, No 5, pg. 221-223

20. LR White Technical Tips:

If you are having difficulty locating your tissue in LR white after it polymerizes due to the fact that the color of your tissue is the same as is LR White once polymerized there is a solution. Dye the LR white using any oil soluble dye. Remember however, that the intensity of the dye needs to be very high if contrast is to be seen at high magnification. Dyes with phenol or tertiary amine groups should be avoided because they effect the rate of cure of the resin.
It is our experience that Sudan Black (cat # 210610)is the best dye to use due to its color intensity. To make up a stock solution add the dry stain to the resin until a saturated solution is obtained. Then proceed as usual.
If you require the pH of LR White to be neutral (the standard is 5) you can adjust it by adding small amounts of caustic soda or a primary amine. The best primary amine to use is Ethanolamine (absolute) for it will mix into the resin very easily. Small amounts are to be titrated into the resin and the pH should be constantly checked until 7 is reached. Do not add more than 50 ml.