Whole-mount Staining Methods for TEM


There are several investigators at the University of Florida College of Medicine involved in the extraction of cellular inclusions, including exosomes, mitochondria, viruses, etc. In addition to biochemical assays to determine the presence and/or purity of the extracts, electron microscopy is a frequently used and often more rewarding evidence of successful extraction. Typical methods used for transmission electron microscopy (TEM) include resin-embedding and sectioning the sample, a long and involved method taking several days to accomplish, or on-grid negative staining. Bacteria, viruses, or organelles from cells can often be visualized on the TEM, without the tedious dehydration and embedment in an epoxy, by whole-mount or negative staining, which easily allows one to see the results within hours of extraction. Most of the time, the whole-mount technique simply entails attaching the extracts to the surface of a 3mm formvar-coated grid, applying a stain such as uranyl acetate or methyl tungstate to the specimen-covered surface of the grid, drying, then viewing the results on a TEM. This method takes only minutes instead of days and may answer questions such as size, shape, or purity of sample; exterior features, such as flagella; and concentration of the sample in the preparation. It is even possible to label the sample with immunogold particles in order to unquestionably identify it. We have presented here two protocols, one a quick-and-easy technique known simply as “negative staining”, and another, a more involved method utilizing methyl cellulose.

The first method is for the TEM preparation of negatively stained AAV virus capsids and results (Figures1a and 1b) as prepared and imaged by Prasad Trivedi, a doctoral student in Dr. Nathalie Clement’s lab in the UF COM Powell Gene Therapy Center. This procedure takes about 30 minutes and can be done in most labs, or the EM Core staff can prepare both the solutions and the stained grids for you to view on our Hitachi 7600 Transmission Electron Microscope.

However, we consider the methyl cellulose method, taken from a report entitled “Isolation and Characterization of Exosomes from Cell Culture Supernatants and Biological Fluids” by Thery et al. (Current Protocols in Cell Biology, Supplement 30, 3.22.13) to produce superior evidence of exosomal extraction by TEM. Therefore, our second protocol is for the TEM preparation of methyl-cellulose negatively stained exosomes and results (Figures 2a and 2b) as prepared and imaged by Regina Oshins in Dr. Mark Brantly’s Alpha-1 research lab in the UF COM Pulmonary Department. Once the solutions are prepared, this procedure takes about 2 hours, and can be done in most labs; or the EM Core staff can prepare both the solutions and the stained grids for you to view on our Hitachi 7600 TEM.

Negative Staining of AAV Capsids

Figure 1a


Figure 1b



  • Eppendorf tube(s) containing pre-fixed sample(s) in solution (provided by investigator)
  • Small petri dishes equal to the number of samples to be stained
  • Filter paper in each petri dish with sample ID written in pencil
  • 1mL syringe and syringe filter
  • A container of Distilled Water for rinses
  • 3mm Formvar-Carbon coated TEM grids (Ladd Research Industries, Inc. Williston, VT)
  • Timer
  • Large petri dish lined with parafilm
  • Fine forceps, #3 or #5
  • Pipets and Pipet tips (a separate one for each sample to prevent cross-contamination)
  • A container of rinse solutions, usually the same as the carrier for stain used
  • Stain solution (1% UAc, aqueous, in EtOH, or in MetOH; 1% Methyl Tungstate, aqueous; Other?)
    • The EM Core uses 1% aqueous UAc, as follows: Place 10mL distilled water in a 20mL scintillation vial. Using a disposable spatula, weigh out and add 0.1g depleted uranyl acetate (Ladd Research Industries, Inc., Williston, VT). Mix vigorously for 30 sec., then place on a shaker overnight – dispose of the contaminated spatula and weigh paper in the properly labeled waste container. Remove the vial from shaker and store under staining hood; allow to settle before using. Using a 1mL syringe, carefully draw staining solution from the top; filter through a syringe filter onto the parafilm-lined petri dish. This solution will last indefinitely.

(Caution: Uranyl acetate, made from ‘depleted’ uranium, is still radioactive. Handle with care under a hood, wear gloves, and dispose of all contaminated waste in a properly labeled waste container.)


Set up droplets (~10-20mL) on parafilm in a large petri dish for no more than 4 samples, as follows:

◍ Sample 1         ◍ Sample 2         ◍ Sample 3         ◍ Sample 4         (2 minutes ea)

◍ DW rinse         ◍ DW rinse         ◍ DW rinse         ◍ DW rinse         (1 minute ea)

◍ Stain                ◍ Stain                ◍ Stain                ◍ Stain                (2 minutes ea)

◍ Rinse               ◍ Rinse              ◍ Rinse               ◍ Rinse              (Briefly)

  1. Choose “Count UP” selection on a timer.
  2. Place formvar-coated grid face down on Sample 1 droplet and start timer
  3. Place grids on Samples 2, 3, & 4 at 20-second intervals
  4. After 2 minutes on the sample, begin moving grids at 20-second intervals to DW rinse
  5. After 1 minute on the rinse, begin moving grids at 20-second intervals to stain
  6. After 2 minutes on the stain, begin moving grids at 20-second intervals to rinse.
  7. Dip each grid briefly in rinse solution, then edge-dry on filter paper and allow to air dry for 60 min in small petri dish ID’d for that sample
  8. Allow samples to dry for at least 60 min before viewing on TEM.  Samples properly stored should last indefinitely.
  9. Dispose of all waste contaminated with uranyl acetate in appropriate radioactive waste container.

Negative Staining of Exosome with Methyl Cellulose – UA

Figure 2a

exosome methyl cellulose1

Figure 2b


  • Exosome samples suspended in buffered formaldehyde, provided by the investigator
  • Large Glass petri dish
  • Parafilm
  • Formvar-Carbon coated grids, 3mm (Ladd Research Industries, Inc., Williston, VT)
  • Forceps, fine-pointed, #3 or #5
  • 3+mm wire loops
  • Syringe filters, 0.22mm
  • 2-50mL disposable centrifuge tubes
  • 3-10mL disposable syringes
  • 1mL disposable syringe
  • Whatman #1 filter paper, cut into small wedges
  • Timer


Methyl Cellulose, 2% (w/v) Make this solution ahead, at least 4 days before needed.

Heat 98mL of distilled water to 90oC; while stirring, add 2g methyl cellulose (Millipore-Sigma, Burlington, MA, cat. #M-6385).  When completely dissolved, rapidly cool on ice (in a larger beaker with ice water) while stirring until the solution reaches 10oC.  Let the solution continue stirring overnight at 4oC.  Stop the stirring and allow the solution to “ripen” at 4oC for 3 days.  Bring the final volume to 100mL with d-water.  Pour into 2-50mL tubes and centrifuge at 100,000g (29,000 rpm in Nephrology’s ultra-centrifuge, rm CG-92), 4oC, for 95 min.  (It will take 30 min to an hour for the centrifuge to come to a stop).  Collect the supernatant and store it for up to 3 months at 4oC.

Uranyl Acetate (4% w/v)

Dissolve 0.4g depleted uranyl acetate (Ladd Research Industries, Inc., Williston, VT) in 10mL distilled water.  Place in a 10ml disposable syringe, wrapped in foil to protect from light, and store up to 4 months at 4oC.  Just before use, filter the needed amount through a 0.22mm filter.

(Caution:  Uranyl acetate, made from ‘depleted’ uranium, is still radioactive.  Handle with care under a hood, wear gloves, and dispose of all contaminated waste in a properly labeled waste container.) 

Oxalic Acid (0.15 M)

Mix 19mg of oxalic acid in 10mL distilled water.  Store at 4oC in a 10-ml disposable syringe.

Uranyl-Oxalate, pH 7

Mix filtered 4% UA with 0.15 M oxalic acid in a 1:1 ratio (5mL of each).  Adjust the pH to 7 by adding 25% NH4OH in drops (for small amounts, may need to reduce to 2.5%).  Store in a 10mL disposable syringe in the dark up to 1 month at 4oC (Wrap in foil).

Methyl Cellulose-UA

Just before use, mix 9 parts of 2% methyl cellulose with 1 part of 4% filtered uranyl acetate

Glutaraldehyde, 1% (v/v)

Dilute 0.2mL of 50% glutaraldehyde, EM grade, (Ladd Research Industries, Inc.) in 10mL PBS.  Store up to 6 months at -20oC or 1 week at 4oC.

Distilled Water



  1. Exosome pellets should be suspended with an equal volume of 4% paraformaldehyde in PBS.  The resulting exosomes in 2% PFA can be stored up to 1 week at 4oC.
  2. Place 10-20mL drops of the resuspended exosomes on clean Parafilm in a glass petri dish, and float 1-2 Formvar-carbon coated grids (coated membrane down) on top of the drops.  Cover and let the membranes adsorb for 20 min.
  3. Place 50mL drops of  PBS on the Parafilm.  Transfer the grids (membrane side down) to the drops of PBS to wash.  Always try to keep the opposite side of the grid dry.
  4. Transfer the grids to 50mL drops of 1% glutaraldehyde in PBS for 5 min.
  5. Transfer the grids to 50mL drops of distilled water  8 x 2 min.
  6. Transfer the grids to a filtered 50mL drop of uranyl-oxalate solution, pH 7. Cover  5 min
  7. Add 1 part filtered 4% aqueous uranyl acetate to 9 parts methyl cellulose.
  8. Place a glass petri dish lined with Parafilm on a cold plate, put 50mL drops of the methyl cellulose-UA on the Parafilm, and place the grids on the drops.  Cover.    10 min.
  9. Remove the grids, one at a time, with a stainless steel loop, and blot excess fluid by gently touching the edge of the loop with wedges of Whatman no. 1 filter paper, so that a thin film is left on the exosome side of the grid.  (The faster you remove the excess fluid, the thicker the film will be, so try to remove the fluid slowly for a thin film.)
  10. Air-dry the grids 5 to 10 min. while still on the loop.  (After drying, a properly coated grid will have a blue-gold interference color.)
  11. The grids may be viewed immediately on an electron microscope and may be stored for many years in a grid storage box.