Flat Embedding of Worms (Slam Freeze) Fixation
by Steve Fields (Rand Lab)

Overview

Flat Embedding: This is a very simple concept, in which the dehydrated, resin-embedded animals are flattened between two sheets of aclar plastic film in a thin layer of fresh resin. If your resin is oxygen-sensitive, the edges of the aclar should be sealed to exclude air. If your resin is more forgiving, there is no need to seal the edges. Then the aclar/worm sandwich can be cured in the oven, or by UV in the cold. Aclar has marvelous release properties (it won't stick to most substrates), but it can also be thin sectioned. This flat embedding method yields worms in a thin, transparent layer of resin. Worms can be examined under LM, trimmed out and glued onto a new block in any orientation for thin sectioning. A variation of this method is used below for the slam freezing. Aclar is available in 8x11 inch sheets from several EM suppliers, including Ted Pella.

Slam Freezing/ Freeze Substitution: This method has potential for improved high resolution TEM. It provides rapid fixation of that portion of the worm which hits the metal mirror; best fixation propagates for a few microns into the specimen before ice damage occurs. Briefly, after live worms are fast frozen, they are freeze substituted with osmium tetroxide or an osmium/aldehyde mixture, infiltrated with resin, and cured for thin sectioning. Below we provide Steve Field's recipe, his notes on alternate freeze substitution media, and some references on the general method.

Protocol

  1. Cool the slam-freezing unit with LN2 and place a copper metal mirror in position on the freezing stage.
  2. Wash worms off of 60 mm plate with 5 ml M9 buffer. (Make sure worms are not starving.)
  3. Centrifuge 5 min at low speed (i.e. setting 2 on an IEC tabletop centrifuge).
  4. Resuspend pellet in 0.5 to 1.0 ml M9 buffer.
  5. Cut 0.45 µl Millipore filters (type HA) into 7 mm x 10 mm rectangles (these are soluble in acetone).
  6. Put a dry Millipore rectangle on a glass fiber filtration unit and apply a house vacuum.
  7. Slowly add 200 µl of worm suspension - do not allow surface tension of meniscus to break or worms will be lost off the edge of the filter. (The dense population of worms should be fairly evenly distributed on the millipore filter.)
  8. To avoid drying the worms, remove the Millipore filter from the filtration unit as the last of the liquid is pulled through. Immediately place it on a wet Whatman filter paper (9mm x 11 mm) that is positioned on a styrofoam freezing disk supplied with the Reichert MM80E slam freezer.
  9. Attach the freezing disk to the magnet on the end of the plunging arm, then hit the release button allowing the arm to plunge the worms onto the cooled copper block.
  10. Detach the freezing disk from the arm and quickly transfer the freezing disk and worms to a dewar of nitrogen.
  11. Working in a -80°C ultralow freezer, use forceps to remove the Whatman/Millipore filters from the freezing disk while it is submerged in LN2 and place the filters into a mesh transfer basket in a container of the primary freeze substitution medium* at -80°C. (The Whatman and Millipore filters are frozen together and cannot be separated at this point.)
  12. Allow 3-5 days for freeze substitution in the primary medium if the sample is not being agitated. This time can be decreased to 2 days if samples are agitated on dry ice.
  13. Rinse 3x10 min in -80°C solvent (i.e. acetone) minus fixatives.
  14. Transfer basket with sample to -80°C secondary freeze substitution medium containing 1% OsO4. (This is only necessary for studies employing standard ultrastructural analyses).
  15. Warm freeze substitution medium slowly by placing the container on a lead brick at -80°C and transferring this to a -20°C freezer for 8-16 h.
  16. Then transfer to a 4°C chamber for 8-16 h. (Make sure medium is above 0°C before proceeding.)
  17. Rinse 3x10 min in 4°C solvent minus fixatives if a secondary freeze substitution medium was employed. In the final rinse, remove the Whatman filters from the basket with forceps (the Millipore filters should have been dissolved) and transfer the individuals and clumps of worms to a 15 ml polypropylene centrifuge tube. Centrifuge one time, resuspend in 1 ml of solvent, transfer to a microfuge tube, and warm to room temp.
  18. Add 0.5 ml resin to make 1/3 resin (i.e. Spurr's or Epon 812) and rotate for 1-2 h at rt.
  19. Infiltrate in 2/3 resin for 1-2 h at rt.
  20. Infiltrate in 100% resin for 24-48 h, making at least 5 changes if fresh resin. To change resin, spin briefly in microfuge 6 times at 10 sec each. Turn tube 180° between each spin so that worms are moved down to the tip by the final spin.
  21. For the final resin change, make up a fresh batch and infiltrate for 1 h. Put a drop (100µl) of worm suspension on a sheet of Aclar and cover with a Thermanox coverslip.
  22. Place in a 65°C oven and polymerize for 16 h.
  23. After polymerization, peel off Aclar film and use a dissecting microscope to select worms to be sectioned.
  24. Cut our selected worms with a razor blade and use a 5 min epoxy to glue the resin square to a blank resin block. (Glue the resin side of the square to the block. The Thermanox coverslip will pop off the resin when trimming the block.)
  25. Trim and section block, picking up sections on formvar-coated slot grids or on mesh grids.
  26. Stain with the double lead stain method.

Notes of freeze substitution media

There are a variety of solvents used to substitute water out of biological samples. The two most commonly used are methanol and acetone. In my (Steve Fields) experience, acetone has proven more useful, in that samples seem to be less extracted and membranes appear to be better preserved. However, this is highly dependent on the type of tissue being freeze substituted. Some advantages of methanol are that salts found in some buffers are more soluble and a higher percentage of water can be tolerated before the methanol solution is saturated.

While the solvent itself acts as a mild fixative during the freeze substitution step, other fixatives may be added to the primary freeze substitution medium depending on the goal of the study. If the goal is to do immunogold labeling, then other fixatives are not added. If ultrastructural analysis is the primary goal, then other fixatives are added. In the latter cases, I usually include 1% anhydrous glutaraldehyde (from EMS) and 1% tannic acid. Tannic acid seems to improve membrane preservation and contrast. I avoid warming the samples while they are exposed to these fixatives to avoid adverse osmotic effects, esp for samples exposed to glutaraldehyde (try to stay below -20C).

For ultrastructural studies, a secondary freeze substitution medium that includes 1% OsO4 further improves membrane preservation and contrast. This solution begins at -80°C and is made from a stock of 4% OsO4. I have found that methanolic OsO4 stock solutions last longer. A yellow color indicates that the solution is good; whereas a gray to black color indicates it has been reduced. Adding a methanolic OsO4 stock to acetone freeze substitution medium does not adversely affect ultrastructure. Samples should be warmed all the way to 4°C in this solution. At the higher temperatures OsO4 turns black along with the sample.

References

Fields, S.D., Strout, G.W. and Russell, S.D. 1997. Spray-freezing freeze substitution (SFFS) of cell suspensions for improved preservation of ultrastructure. Microsc. Res. Tech. 38: 315-328. Abstract

Gilkey, J.C. and Staehelin, L.A. 1986. Advances in ultrarapid freezing for the preservation of cellular ultrastructure. J. Electron Microsc. Tech. 3: 177-210. Abstract

Ryan, K.P. 1992. Cryofixation of tissues for electron microscopy. A review of plunge cooling methods. Scanning Electron Miscrosc. 6: 715-743.

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