PRIMARY FIXATIVES GLUTARALDEHYDE (glutaric acid dialdehyde)
NOTES:
a. Most widely used primary fixative for EM
b. To minimize extraction of cellular components by autolysis, fix at low temperatures (e.g. 4 C in fridge).
c. Concentration for usage: below 2%, extraction may occur. above 4%, shrinkage may occur. Usually
prepared at 2.5% for biological specimens.
d. Purified GA is good at around pH 3-6. If below 3% then discard. You can purchase in ampoules of 25 or
8% from EM vendors which will last several months unopened). Biological grade GA contains impurities
such as glutaric acid, acrolein glutaradozamine, ethanol, methanol and various polymers and products of
oxidation and photochemical degradation.
e. Osmolarity: purified around 300 mosmols; biological around 500-600 mosmols
g. Glutaraldehyde/formaldehyde mixtures provide faster penetration and good crosslinking of proteins.
Structures are stabilized with formalin, then crosslinked with GA. Basic EM primary fixation To mix 2 - 2.5% Glutaraldehyde:
2.5% buffered GA - 10 ml 25% GA into 90 ml buffer.
2% buffered GA - 1 ml 8% GA into 2 ml double strength buffer and 1 ml water (or 1 part GA, 2 parts buffer, 1
part water) GLUTARALDEHYDE/PARAFORMALDEHYDE MIXTURES
Note: Lower concentrations of GA (0.02 0.2%) mixed with formaldehyde are preferred when preparing
samples for immunolocalization. Karnovskys (1965)
Extremely hypertonic (2010 mosmols). Not often used. 5% glutaraldehyde, 4% formaldehyde in cacoldylate buffer (cacodylic acid). 2.5% GA, 2% paraformaldehyde
-Most widely used fixative for EM Cacodylate (0.2M) 25 ml pForm (10%) 10 ml GA (25%) 5 ml distilled H2O to 50 ml The mixture results in 0.1M buffered solution. McDowell and Trumps (1976)
-Used mainly in pathological labs where both EM and light microscopy are required from the same tissue. GA (50%) 2 ml Formaldehyde (40%) 10 ml NaH2PO4 H2O 1.16 g NaOH 0.27g dH2O 88 ml GA/FA mix
- The high concentration of formaldehyde in this mix is effective in localizing amines by formation of
fluorescent products and for fixing CNS for EM. Fluorescent intesity is enhanced if dried, but lost if
rehydrated. GA 0.5-1%
FA 4%
0.1M Cacodylate or PBS (pH 7.0) SPECIALIZED FIXATION GA- Ruthenium red
Preserves and defines membranes and myofilaments. Will stain acidic mucopolysaccharides.
CAUTION! Ruthenium red is toxic. Soln A: GA (4% aqueous) 5 ml Soln B: OsO4 (5% aqueous) 5 ml cacodylate (0.2M, pH 7.3) 5 ml cacodylate buffer 5 ml ruthenium red (stock) 5 ml ruthenium red (stock) 5 ml (1,500 ppm in H2O) Fix in solution A for 1 hr at RT
Rinse 3x10 min in cacodylate buffer
Fix in solution B for 3 hr at RT
No post staining necessary GA Alcian blue
To obtain staining of a cell coat and intercellular substances. Its behavior is identical to reuthenium red except
alcian blue is not toxic. The GA-mucosubstance-alcian blue complex formed is osmiphilic. Fix in 4% GA, 1% alcian blue in buffer for 1-18 hours.
wash
post-fix in OsO4
buffer both GA and OsO4 to pH 6.5 GA- digitonin (Okros 1968)
Helpful in retaining fine structural localization of free cholesterol and cholesterol esters. The complex is
osmiphilic and insoluble in lipid solvents.
Fix at RT and dehydrate only 70 and 95% ethanols. Infiltrate with 95% ethanol/Epon mix. Crystal artifacts
are not uncommon. The 1% digitonin causes complete disruption of lysosomal membranes in intact tissues
but has little effect on peroxisomes. GA (2.5%) 5.0 ml fix at RT for 2 hr FA (2%) 5.0 ml The mixture is stable for two days digitonin in buffer (0.2%) 5.0 ml GA hydrogen peroxide
Used to enhance quality of fixation. The possible increase in oxygen during fixation is thought to be needed
for irreversible protein crosslinking.
CAUTION! Hydrogen peroxide cannot be mixed with formaldehyde/formalin as explosive compounds may
result. Cacodylate buffer (0.1M) 25 ml GA (25%) 5 ml Hydrogen peroxide (15%) 5-25 drops Add drops while continuously stirring. The final concentration of GA will be between 3 and 6%.
Fix for 1-2 hr at RT or 3-4 hr at 4 C. If you fix at the cooler temp, fix for one hour in 3-5% GA after the initial
fix. GA- Lead acetate
Used to preserve soluble inorganic phosphate (nucleolar orthophosphate). Osmium and UA are not used in
this prep as they will remove the lead precipitate.
Rxn: GA-lead acetate and inorganic phosphate will form lead hydroxyapatite. GA 2%
Lead acetate 2%
0.1 M Cacodylate fix at pH 7.0 at RT for 3 hr GA-malachite green
Used to preserve lipids containing granules in mammalian sperm. Enhances lipid opacity under the big eye.
The overall ultrastructural preservation when using this combination is not satisfactory. GA 2% Malachite green 0.1% Fix as usual. GA trinitro compounds (Picric acid, trinitrocresol.)
Preserves smooth ER in testicular interstitial cells and other steroid secreting cells. Preserves peroxidase
activity. CAUTION! Trinitro compounds are unstable when crystallized and can explode. Buffer (0.2M, pH 7.2) 45 ml GA (25%) 5 ml FA (4%) 50 ml trinitrocresol (2%) 50 ml fix for 2 hr at RT GA potassium dichromate
For visualizing biogenic amines. Useful for demonstrating norepinephrine and argentaffin cells. a. Fix tissue in 3% GA in Cacodylate buffer (pH 7.2) for 4 hr followed by incubation in 2.5% potassium
dichromate in Caco buffer (pH 4.1) for 4 hr. b. Mix: GA 1% FA 0.4% 0.01 M Na chromate-potassium dichromate buffer (pH 7.2) Fix tissue in above formulation for 1 to 10 minutes. Store in Na chromate-K dichromate buffer (pH 6.0) for
18 hr at 4 C. Do not stain with UA. Champys fluid
3% potassium dichromate 7 parts 1% chromic acid 7 parts 2% OSO4 4 parts GA- potassium ferrocyanide OsO4 (Elbers 1965)
Preserves labile lipids, surfactants in the lung and increases visualization of lipids. Used for the
demonstration of liposome particles. Fix in 1% acrolein and 2.5% GA in 0.067M cacodylate (pH 7.4) containing 1mM CaCl 2 for 24 hr at RT. Store overnight in buffer.
Postfix in 1% OsO 4 in same buffer containing K 3 Fe(CN) 6 (0.05M) and CaCl 2 (0.05M) for 3-4 hr in dark at 4 C. GA Tannic acid
Tannic acid is a mordant for heavy metal staining of mucins and complex carbohydrates. Treatment prior to
OsO4: Provides enhanced cyto-membranes and cytoskeletal features. Prevents disruption of MTs by OsO4.
Reacts with phospholipids and phosphatidyl cholines. Treatment after OsO4: High density staining of
secretory bodies.
Preserves intercellular glycosaminoglycans (Singer & Solursh 1980). Prepare all solutions in buffer
Treatment 1: Treatment 2: GA (5 %) 10 ml GA (5%) 20 ml tannic acid (8%) 10 ml FA (16%) 10 ml Adjust pH with NaOH tannic acid (16%) 10 ml sucrose 0.05 M Postfix in OsO4 GA- Uranyl acetate
Recommended for fixing bacteria containing intracellular phages (Sechaud and Kellenberger 1972). Gels the
DNA. See the article for concentrations. Acrolein
Allows for increased penetration with large or impermeable tissues. This is a component or impurity found in
biological grade glutaraldehyde. CAUTION! Acrolein is highly volatile and toxic. Very dangerous and
should only be used when absolutely necessary. Acrolein formulations: 1% acrolein 2.25% GA in 0.1 M buffer
cacodylate buffer (0.2 M) 50 ml Acrolein 1 ml GA (25%) 10 ml dH2O to make 100 ml GA (25%) 12 ml acrolein 3 ml Millonigs Phosphate buffer 85 ml Either formulation; Fix for 1-3 hr, start in cold and allow to come to RT.
Wash
Postfix in 2% buffered OsO4 at 4 C for 2 hr. Osmium Vapor fix
Primarily used with specimens that are in an aqueous solution or very delicate and addition of one of the
above fixatives would disrupt or be osmotically detrimental.
Take a lid or cover and tape a filter paper to it. Put a drop of 4% aqueous osmium on the filter paper and
cover the sample for at least 1 hr. Another drop of 4% osmium into the sample directly can be done if
required for good fixation. Wash completely. Do not use glutaraldehyde after as precipitation will occur and
you will have a mess. SECONDARY FIXATION OsO4 (osmium tetroxide, osmic acid) OsO4 acts as both a stain and a fixative. Most commonly used post-fixative. Has a high molecular weight (254.2) in a water soluble crystalline form that will not change the pH. It is a noncoagulant fix that
stabilizes proteins further. It is a non-polar compound which oxidizes aliphatic double bonds. It is thought that
one molecule of osmium reacts with one double bond in the lipid molecule and forms glycol osmates
stabilizing the molecule. Fixation in OsO4 should be kept to a minimum to avoid leaching and damage to cellular components. Also, longer times causes the lipids to over stain decreasing the information gained from staining. Optimal
concentration is between 1-2%. Higher concentrations cause cleavage of protein molecules resulting in loss of
peptide fragments. Usually obtained in ampoules of 4% aqueous or in solid form. If solid, make aqueous
stock solution of 4% that can be frozen until used. Tissue size should not exceed 0.5 mm for optimal fixation
and penetration of sample. One method used by Dr. Farmer and others is a method of using the osmium vapors to initially fix aquatic specimens (e.g Euglena and other protists). See above in Primary fix for details. Crystalline OsO4 can be mixed in acetone or methanol for freeze substitution protocols. If used in this manner, extra precautions are needed to keep the solution at 80 C as it will oxidize rapidly. If either
aqueous or other osmium solution is dark brown to black then discard and prepare fresh. Fresh solution
should appear light tan or weak tea colored.
Cryofixation and freeze substitution are covered in detail in Bozzola EM text. Briefly, one rapidly freezes the
sample by plunging into liquid propane at liquid nitorgen teperature and then transferring the sample to the
osmium/acetone or methanol substitution fluid which is kept at 80 C. This substitution fluid will remove the
water over time, usually 48 hrs while kept at 80. The sample in substitution fluid is then warmed very slowly
by peroidically moving the sample from 80 to 20 freezer for several hours, then to the 4 C fridge and
finally washing the sample with several changes of fresh acetone or methanol. Infiltration can begin
immediately. CAUTION! Osmium vapors will fix tissue! ALWAYS use in hood and with appropriate safety
precautions. BUFFERS Most buffers used in electron microscopy are effective in the physiological range: pH 7.2 - 7.4 PHOSPHATE BUFFERS Non-toxic to cells in culture and the pH is stable at various temperatures. They can be stored for several
weeks in the refrigerator and is the most widely used buffer for many cellular methods and EM. Forms a
precipitant when contaminated and has a tendency to decrease the nuclear mass. Will extract non-
chromosomal proteins in nucleus and can cause swelling on organelles. Will precipitate polyvalent cations,
lead and uranium salts. Not good for negative staining.
May inhibit certain enzymes. Phosphate ions are thought to precipitate in concentrations of ethanol above
50%, adhere to cellular structures and attract uranyl and lead ions. Rinsing in lower concentratins of ethanol
reduces much of the phosphate.
Osmolarity is around 290 mosmols. Millonigs 1. Phosphate buffer (1961) Soln A: 2.26% monobasic sodium phosphate in water Soln B: 2.52% NaOH in water Buffer final concentration: 0.13 M Mix 41.5 ml of Soln A with 8.5 ml Soln B
Remove 5 ml of mix and add 5 ml of 4% sucrose solution.
Add 25g of MgCl2 or CaCl2 to each 100 ml of buffer
Adjust pH to 7.3 with Soln B
Stable for several weeks at 4 C 2. Phosphate buffer (1964) In 500 ml dH2O:
Monobasic sodium phosphate 1.8 g Dibasic sodium phosphate 23.25 g NaCl 5 g Ph to desired value then add d H2O to make 1,000 ml Karlsson and Schultz Phosphate (1965) Monobasic sodium phosphate 3.31 g Dibasic sodium phosphate 33.77 g dH2O to make 1.0 L
pH 7.4, Osmolarity 320 mosmols (equivalent to cerebrospinal fluid in rats). Maunsbach Phosphate (1966) Monobasic sodium phosphate 2.98 g Dibasic sodium phosphate 30.40 g dH2O to 1.0 L Sorensons Phosphate (0.1 M) Solution A: 0.2 M Na 2 HPO 4 . 2H2O 35.61 g or Na 2 HPO 4 . 7H2O 53.65 g or Na 2 HPO 4 . 12H2O 71.64 g then dH2O to make 1.0 L Solution B: 0.2 M Na 2 HPO 4 . H2O 27.6 g or Na 2 HPO 4 . 2H2O 31.21 g Then dH2O to make 1.0 L Prepare by mixing two solutions as given here and diluting to 100 ml with distilled water. pH at 25 C Soln A Soln B 6.4 13.25 36.75 6.6 18.75 31.25 6.8 24.5 25.5 7.0 30.5 19.5 7.2 36.0 14.0 7.4 40.5 9.5 7.6 43.5 6.5 7.8 43.75 4.25 8.0 47.35 2.65 Osmolarity of 0.1 M buffer (pH 7.2) is 226 mosmols. Addition of 0.18 M sucrose raises it to 425 mosmols. Phosphate Buffered Saline (PBS)
For immunofluorescence, immunocytochemistry pH 7.0 For 1.0 L
NaCl 6.8 g Na 2 HPO 4 (dibasic) 1.5 g NaH 2 PO 4 (monobasic) 0.43 g Dulbeccos Phosphate buffered saline In 800 ml water:
Soln A: 8.0g NaCl 0.2 g KCl 1.15 g Na2HPO4 (dibasic)
0.2 g KH2PO4 (monobasic) In 100 ml water each:
Soln B: 0.1 g CaCl2 Soln C: 0.1 g MgCl2 Dissolve separately, then combine. CACODYLATE BUFFER (cacodylic acid)
Made at 0.2 M and then diluted accordingly. Effective in pH range of 6.4 to 7.4. Avoids interference of
extraneous phosphates in cytochemical localization. Does not increase nuclear mass, little removal of acid
soluble proteins from nuclei. Desirable for auto-radiography and enzyme localizations. Resistant to bacterial
contamination. Calcium can be added without precipitation. The buffer is incompatible with UA, so enbloc
staining is not recommended. Membrane permeability may be altered due to toxicity of the buffer leading to a
redistribution along the osmotic gradient of changed chemical activity. This impairment will affect the quality
of fixation.
CAUTION! Contains arsenic contact with acid produces arsenic gas. Cacodylate formulation: (0.05M)
Prepare in fume hood and wear gloves. Solution A: Sodium cacodylate trihydrate 42.8 g Add dH2O to 1.0 L Solution B: 0.2 M HCl
Concentrated HCl (36-38%) 10 ml dH2O 603 ml The desired pH can be obtained by adding soln B to 50 nl of soln A and diluting to a total volume of 200 ml
with dH2O, according to the following schedule: Soln B pH 0.2 M Cacodylate 18.3 6.4 Molecular Wt in grams into 1.0L H2O 13.3 6.6 Adjust pH accordingly. 9.3 6.8 6.3 7.0 4.2 7.2 2.7 7.4 COLLIDINE BUFFER Efficient at pH 7.4 when half neutralized with HCl (range of buffer 6.0 8.0)
Does not react with OsO4 and is stable at RT indefinitely. Great for lung tissue but collidine isa pyridine
derivative and therefore extracts phospholipids. Tissue sections very easily, proibably due to this extraction.
Extraction facilitates penetration of fixative into larger specimens. It is not recommended for EM.
CAUTION! TOXIC Stock: s-collidine (pure) 2.67 ml dH2O to make 50 ml Buffer: Stock soln 50 ml 1.0 M HCl 9.0 ml dH2O to make 100 ml adjust pH to 7.4 with HCl TRIS BUFFERS As a primary amine, it reacts with GA. It has poor buffering capacity below 7.5. Biological inhibitor and used
mostly in enzyme localizations. Tris (0.05 M) STAINS THICK SECTION STAINING Procedure
1. Transfer sections to small drop of water on slide.
2. Heat slide gently until section adheres to slide evaporate all water but do not boil.
3. Cover section with drop of staining solution and heat gently for thirty sections to one minute.
4. Drain off excess stain, and wash well in two changes of dH2O
5. Dry with heat. STAINS Toluidine blue
Most commonly used stain for epoxy sections 1 part dH2O
1 part 5% toluidine blue
1 part 2% sodium borate
Mix well, then filter. Store at RT Methylene Blue
1% Azure blue or Azure II in 1% borax Paragon stain (Martin et al. 1966) 1% aqueous p-phenylene diamine (Estable-Puig 1965)
for black and white photography. THIN SECTION STAINING Usually thin sections for EM are sequentially stained with 4% uranyl acetate followed by Reynolds lead
citrate. URANYL ACETATE
Most widely used stain for thin sections. Provides high contrast by staining nucleic acids, proteins, free amino
groups. Solutions are photolabile and should be kept dark. Alcohol solutions produce better contrast and
require shorter staining times. Aqueous solutions must be used when working with supportive films or when
enzymes have been localized. Concentrations my be varied so it is suggested that you use the one that
provides your material with best contrast.
Common formulations
1-4% aqueous 2% in methanol or ethanol saturated aqueous saturated in 50% methanol saturated in methanol Staining procedure: 1. Staining should be performed at RT. Filter solution.
2. Place one drop onto parafilm in Petri dish and float grids section-side on solution. Do not allow drop to evaporate or precipitated UA will result. 3. Stain for 15 to 45 minutes in dark. This can be done by covering the Petri cover with aluminum foil. 30
minutes is typical.
4. wash grids in three changes of dH2O or rinse with a continuous flow over the grid. If alcohol based, wash
with progressively lower concentrations of alcohol. En Bloc Staining with UA When UA is used en bloc, it has a fixative effect giving the fine structural preservation of DNA filaments,
membranous structures and cell junctions. But is can also cause extraction of cellular components. Procedure:
Apply stain to tissue either before or after OsO4. It is thought that application of UA after OsO4 leads to
better retention of phopholipids. 1% UA in buffer or 50% ethanol. CAUTION! UA is an uranium salt and is slightly radioactive. It is not advisable to come in contact with UA.
Dispose of waste in appropriate receptacles. LEAD STAINING Stains most intensely at high pH, more intense staining occurs when preceded by OsO4 fixation especially
membranes because phosphate, sulfhydryl, tyrosyl and carboxyl groups become more ionized after osmium
resulting in increased binding of lead. Stains glycogen, membranes and the ground substance of cells.
Insoluble crystals of lead may form while staining by precipitating in the presence of CO2. NaOH pellets
around the staining chamber will chelate lead hydroxide and prevent precipitation.
Precautions:
Maintain a clean work area.
Use NaOH around grids and keep lid on Petri dish.
Hold your breath when transferring or washing
Filter stain before using
Wash initially with 0.02M NaOH before washing well with dH2O. Some prefer to boil water to drive off
CO2, allowing the water to cool before using for wash. Reynolds Lead Citrate (1963) Boil 100 ml of dH2O to remove dissolved CO2. Let cool to RT.
Lead Nitrate 1.33 g Sodium citrate 1.76 g
dH2O 30 ml in 50 ml volumetric flask Shake vigorously for 1 minute and then every 5 minutes or so over 30 minutes time period.
Add 8 ml of 1N NaOH to clear solution and dilute to 50 ml.
Dispense into syringes with filter and store at 4 C. Can be stored up to 1 month. Stain grids in similar fashion as described above for UA. It is advisable to place NaOH pellets around the area
of stain in the Petri dish. Discard first few drops from syringe and then use drops for grids.
Stain for 1 minute. Longer periods will decrease UA staining, as the UA bleaches from the high pH.
First wash in 0.02N NaOH, then three thorough washes in water.
Dry grid by wicking water away with filter paper. Allow to dry completely before viewing in scope (approx.
1 hr minimum) Venable and Cogglesall Lead Citrate (1965) dH2O 100 ml Lead citrate 0.4 g 10 N NaOH 1.0 ml Shake vigorously in screw-capped vial. Do not expose to atmospheric CO2. Stain as described above. Satos Lead Citrate (1967) Lead nitrate 1.5 g Lead acetate 1.5 g Lead citrate 1.5 g dH2O 90 ml Heat to 40 C while stirring for one minute.
Add 3 g Sodium citrate to this mixture and stir for 1 min
Add 24 ml 1N NaOH and 24 ml dH2O to this mix
This solution can be stored for one year at RT. 1. Stain sections by immersion with concentrated solution or dilute with dH2O 1:7 Stain for 10 minutes 2. Wash with water and air dry Lever Lead Hydroxide (1960) Add 1 g lead hydroxide to 100 ml dH2O and bring to boil.
Cool and filter
Add drop by drop, 2N Potassium hydroxide until the solution clears completely.
Stain the sections by floatation for 5 min
Wash in 1% aqueous potassium hydroxide followed by thorough water washes. Potassium Permanganate Stains membranes, myelin sheaths, tonofibrils, glycogen and desmosomes. Reactive with NMA, a component
of EPON 812 resin.
Used aqueous at 1% for section staining
For en bloc, 1% in 100% acetone. Bismuth
Has a strong affinity for nucleic acids. Great for chromosome studies and is a general enhancer of contrast.
An alkaine bismuth subnitrate solution is used
See Ainsworth & Karnovsky (1972) J.Histochem. Cytoschem. 20:995. Thorium
Specific for mucosubstances. Radioactive, used at low pH (2 2.5). Stains ourside of plasma membrane. Colloidal thorium 0.5 g 3% caetic acid (pH2.5) 50 ml Wash glutaraldehyde osmium fixed tissue with dH2O. Immerse the tissue in staining solution for 24 hr at
RT.
Rinse the blocks in 3% acetic acid
Dehydrate and embed as usual. Indium trichloride
For staining of nucleic acids. The staining is attributed to binding of indium by phosphate groups in the
nucleic acids. Other groups that may react are first blocked by acetylation and borohydride reduction. The
method is not completely specific as keratohyalin granules, mammalian sperm tails and mast cell granules
stain as well. Produces good contrast and ultrastructural staining. Fix tissue in buffered aldehyde. Do not use osmium at any stage.
Dehydrate in an acetone series at 0-5 C.
Once tissue in absolute acetone, add pyridine gradually and bring tissue to pure pyridine in 15 minutes.
wash in pyridine 3x 10 min at 4 C.
Reduce for 2 hr with pyridine saturated with lithium borohydride prepared just before.
Wash 3x 10 min with cold pyridine.
Acetylate overnight at RT with freshly prepared mix of 6 parts pyridine// 4 parts acetic anhydride saturated
with sodium acetate.
Wash 3x 10 min with 100% acetone at RT
Stain en bloc for 2 hr at 4 C in 25 mg anhydrous indium trichloride in 1 ml acetone.
Wash 2x 10 min in acetone at RT
Embed in methacrylate or epoxy. Do not use araldite. Phosphotungstic acid (PTA)
Has an affinity for polysaccharides, used as a precipitation agent in the localization of amino acids. Reacts
with serotonin, histamine, epinephrine; proteins rich in lysine, histadine, and arginine. Stains lysosomes and
plasma membrane.
Used in acidic environment (pH 1-3). Very popular as a negative stain, has a finer grain than UA. Oxidize sections for 20 min in 1% periodic acid
stain in 5% PTA for 1-2 hr Always use PTA in an ethanolic solution. Store the reagent in freezer. NEGATIVE STAINING The method of negative staining described by Brenner & Horne (1959) is essentially a simple one. Its original
purpose was to enable large numbers of specimens prepared by other physical and chemical methods to be
examined by EM. Their original procedure was to prepare a 2% solution of phosphotungstic acid (PTA) in
water or ammonium acetate and adjust the pH to a neutral value between 6.4 7.4, by adding small drops of
N-KOH. The potassium phosphotunstate (KPT) was then added to the virus suspension and the mixture
sprayed onto carbon filmed supports by one of the methods described. Droplet patterns were formed by the
electron-dense KPT enclosing or surrounding the virus particles, thus producing a reversal in the contrast
seen in the final image. A method also used is to dry the sample suspension (bacteria, flagellar suspension, viral) onto the coated grid
and then put a drop of either PTA or 4% UA onto the grid, then wait a period of time determined previously to
provide best results. Wick of the excess stain and allow to dry before viewing in the TEM. PHOTOGRAPHY SOLUTIONS FORMULATIONS For Film Development: Microdol-X:
Gives the finest grain with a minimum speed loss. Produces a very low fog level. When possible use a 1:3
dilution over full strength the grain will be finer. HC-110:
Rapid developer for processing most B&W films. Produces sharp negs with normal contrast. Great for TRI-X
film. Use dilution B for better grain. Stored at full strength until diluted. D-11
High contrast developer used in graphic arts. use this developer for Ortho film. 2.5 min at full strength at RT. D-19
Rapid developer that yields high contrast negs. Used to develop EM negs. Use 1:1 with H2O
Change after approx. 200 negs Dektol
Yields high contrast negs and used to develop cold-tone papers. Used to develop LPD IV and B&W papers. D-76
Produces maximum emulsion speed and great shadow detail. Produces a normal contrast. Common dilutions: B&W 35 mm film
Microdol-X 1:3 Dektol 1:2 D-11 Straight HC-110 1:7 (dil.B) B&W paper:
Ektaflo type I: 1:9 for cold tone papers
Dektol: 1:2 MISCELLANEOUS PROCEDURES Preparing mixtures and volumes When preparing mixtures, use the following calculations to minimize solutions used and decrease waste
production. (original concentration of starting solution) x (amount of original solution to add) = (the final amount
required) x (the working concentration). For example: 8% glut (X) = 20 ml (2% glut) 8% is the original concentration of glut in ampoule 2% is the working concentration 8X = 40 X= 40/8 X=5 so 5 ml of 8% glut added to 15 ml buffer (20ml - 5ml) to provide 2% final. Preparing solutions from dry ingredients When preparing any solutions, such as a buffer, it is important to read the instructions and know the molecular weight (or formula weight) of the reagent and the final pH of the solution. A 1 molar solution (1.0
M) is equivalent to adding the m.w. in grams to a liter.
A normal solution is usually used for pH reagents and is the same as molar as long as there is only 1
hydrogen involved in the formula (e.g. HCl or NaOH). e.g. 1N HCl = 1M HCl Always begin with less water than the final volume, as adding the dry ingredients and pH reagents will add volume. Usually about