3.10 　Light and electron microscopy of protonemata embedded with epoxy resin

Takashi Murata and Naomi Sumikawa

Sectioning followed by light and electron microscopy is a useful method to visualize subcellular structures. In this section, we describe methods for light and electron microscopy of protonemata embedded with an epoxy resin.

1 Culture

Suspend cut protonemata in a medium with 0.8% agar (melted and cooled), and spread onto an agar medium with a cellophane before agar becomes solidified.　 As a result, a thin layer of agar medium is overlaid onto an agar medium, separated with a sheet of cellophane.　

The agar medium with a sheet of the suspension should be cultured under illumination with white light. For observation of caulonemata, please use BCDAT medium. Under these conditions, development of caulonemata depends on the duration of culture and the volume of medium. We use 10 ml of medium and 6 days of culture, but the conditions may differ between laboratories.

2 Fixation and Embedding

1) Preparation of fixatives

WARNING: Fixatives below are hazardous. Use gloves.

Primary fixative (2.5 % glutaraldehyde, 0.5% formaldehyde, 0.001% Triton X-100 in 0.05 M sodium phosphate buffer, pH7.4)

8% glutaraldehyde	6.25 ml
1% Triton X-100 in H2O	0.02 ml
2% paraformaldehyde in 0.2 M sodium phosphate buffer (pH 7.4)	5 ml
	Fill up to 20 ml with H2O

Dissolve 2% paraformaldehyde in 0.2 M sodium phosphate buffer (pH 7.4) on a hotplate prior to mixing chemicals.

Post-fixative (1 % osmium tetroxide in dH₂O)

2% osmium tetroxide in H2O (in ampoule, TAAB laboratory)	2 ml
Distilled water	2 ml

Handle in a hood because osmium vapor is extremely toxic. Use immediately after opening the ampoule. The fresh solution should be clear (light yellow). The exhausted solution is black. Do not use the exhausted solution.

Epoxy resin mixture (TAAB EPON 812 kit, TAAB laboratory)

TAAB EPON 812	10.2 g
DDSA	5.4 g
MNA	5.4 g
DMP-30 (accelerator)	0.18 g

Prepare the mixture without accelerator (DMP-30). Add accelerator as needed. The pot life of the mixture with accelerator is about 1 day.

Protocols

1. Make sure that protonemata are growing well. Pour the primary fixative into an empty plastic Petri dish. Cut the top layer of agar medium into squares (ca. 10 x 10 mm²) with a razor blade or a surgical knife. Transfer the square sheets of agar medium with protonemata to the primary fixative.

2. Drop the primary fixative onto the sheets so that they sink into the fixative.

3. Incubate at room temperature for 1 h.

4. Wash the sheets with aldehyde-free sodium phosphate buffer using three changes of solution, for 10 minutes each wash.　

5. Incubate with post-fixative at room temperature for 2 h.　 Handle in a hood and use gloves.

6. Rinse with distilled water.

7. Transfer the sheets to a glass petri dish filled with distilled water.

8. Dehydration with acetone/water mixture.

25% acetone for 10 min.

50% acetone for 10 min.

75% acetone for 10 min.

99.5% acetone for 10 min.

100% acetone (dehydrated acetone for chemical synthesis) for 10 min, twice.

9. Infiltration with resin.

12.5 % epoxy resin mixture (no accelerator) in deh. acetone for 2-3 h.

25 % epoxy resin mixture (no accelerator) in deh. acetone for 2-3 h.

50 % epoxy resin mixture (no accelerator) in deh. acetone overnight.

75 % epoxy resin mixture (no accelerator) in deh. Acetone for 2-3 h.

100 % epoxy resin mixture (no accelerator) for 3-4 h.

100 % epoxy resin mixture (plus accelerator) overnight.

100 % epoxy resin mixture (plus accelerator) for 3-4 h.

10. Pour 100% epoxy resin mixture (plus accelarator) into an aluminum dish (for making cake), until the depth of resin is 2-3 mm. Transfer the sheet with protonemata into the dish.　 Spread the sheets on the bottom of the dish. Polymerize the resin at 60^oC for 2 day in an oven. Store the polymerized resin blocks under dehydrated conditions, at room temperature until ready for sectioning.

3 Mounting blocks, making semi-thin (0.5-1 mm) sections and light microscopy

1) Preparation of solutions

Toluidine blue stain

Toluidine blue N (Chroma-Gesellschaft)	0.5 g
2% sodium borate in dH2O	50 ml

Dissolve Toluidine blue with 2% sodium borate in a tube or a bottle. If undissolved powder remains, centrifuge the mixture and use the supernatant.

2) Protocols

1. Choose a protonema to be sectioned. The polymerized resin is transparent so that you can observe protonemata in the resin. Mark the region of the chosen protonema with a fine felt pen.

2. Excise the marked region of the resin block using a compass saw.

3. Attach the excised block onto the top of a mounting stub (plain cone-shape resin block) with epoxy glue (Cemedine super; Cemedine Co. Japan). The top of the mounting stub should be flattened with a file. Polymerize the glue at 40-60^oC for 2-3 h.

4. Trim excess resin with a file and a razor blade. Use a binocular microscope.

5. Trim further with a glass knife on an ultramicrotome (e.g., Leica Ultracut UCT). The width of the top part of the trimmed block should be less than 2 mm, and the shape should be trapezoid.

6. Make 0.5-1 mm sections with a diamond knife (for semi-thin sections) on an ultramicrotome.

7. Transfer sections from the boat of a diamond knife to an APS (aminosilane) coated glass slide using a wire loop.

8. Heat the glass slide on a hot-plate until water evaporates.

9. Add 1-2 drops of toluidine blue stain onto the attached sections on the slide. Heat on a hot-plate for 1-2 min.

10. Wash off excess toluidine blue stain with distilled water.

11. Dry on the hot-plate.

12. Observe under a light microscope.

4 Making ultra-thin (50-100 nm) sections and electron microscopy

1) Preparation of solutions

0.5% formvar in ethylene dioxide

Polyvinyl formal (polyvinyl formvar)	0.25 g
Ethylene dichloride	50 ml

Dissolve polyvinyl formal with ethylene dichloride at room temperature in a 50 ml falcon tube.

2% uranyl acetate in 70% methanol

Uranyl acetate (powder)	0.2 g
70% methanol in dH2O	10 ml

For use of uranyl acetate, please follow rules of your university or institute. Dissolve uranyl acetate in a 15 ml falcon tube. Store at -20^oC in the dark.

Raynold’s lead citrate

Lead nitrate	1.33 g
Sodium citrate	1.76 g
dH2O	30 ml
1 N NaOH	8 ml
	Fill up to 50 ml with dH2O

Dissolve lead nitrate and sodium citrate in 30 ml of distilled water.　 Shake vigorously for 1 minute, and then keep at room temperature with shaking at intervals every few minutes.　 After 30 minutes, add 8 ml of 1 N NaOH.　 Make sure that the solution becomes clear.　 Add distilled water to 50 ml total volume.　 Seal tightly with Parafilm (avoid contact with carbon dioxide in air) and store at 4ºC.

2) Protocols

1. Make sure the of region aimed to be observed by observing toluidine-blue stained sections.

2. Trim the top part of a block with a razor blade on an ultramicrotome.　 The width of the top part of the trimmed block should be less than 0.5 mm, and the shape should be trapezoid.

3. Making a formvar membrane from 0.5% formvar dissolved in ethylene dichloride and mounting the membrane onto grids.

　 Wipe a glass slide with a dry lens paper.　 Dip the slide into 0.5 % formvar in ethylene dichloride.　 Move the slide up slowly into the air.　 Dry the slide at room temperature for approx. 2 min.　 Cut a formvar layer with a razor blade on four sides of the glass.　 Slowly dip the slide into dH₂O, in a 500 ml beaker or equivalent filled to the top, so that the formvar layer is removed from the glass onto the surface of water as a membrane.　 Because some glass slides are sticky (depends on the batch of glass), choose good glass slides before preparation.　 Wiping a glass slide with acetone may improve release of formvar from a glass slide. The colour of the formvar membrane is correlated with thickness of the membrane (interference color).　 Silver membrane is best for use.　 Gold or purple membrane is too thick, and dark gray membrane is too thin.　 A faster “up-speed” from the formvar solution will result in a thicker formvar membrane.　 Because the colour of the membrane varies even in a single membrane, put the grids onto better (that is, silver) areas of a membrane.　 After placing the grids, put a piece of Parafilm onto the membrane and remove the membrane from the surface of the water.　 Dry the membrane with the grids on Parafilm and store until use in a clean Petri dish.

　 We use single slot grids (Synaptek, Notch and Dot type) for observation of serial sections or for taking low magnification pictures of a protonema, and mesh-type grids for conventional observation of a single section.

4. Make 50-100 nm sections with a diamond knife (for ultrathin sections) on an ultramicrotome.

5. Expand sections by chloroform vapor (by a toothpick dipped into chloroform solution).

6. Mount sections onto the grids with a formvar membrane.

7. Staining with uranyl acetate.　 Incubate grids with 2% uranyl acetate dissolved in 70% methanol for 5 min. Avoid illumination by fluorescent lamps.

8. Wash the grids with distilled water.

9. Staining with lead citrate.　 Incubate grids with Raynold’s lead citrate solution for 5 min in the presence of sodium hydroxide (for removal of carbon dioxide).

10. Wash the grids with distilled water.　 Dry grids at room temperature and store them until observation.

11. Observe the sections under an electron microscope. Acceleration voltage should be 80-100 kV for 50-100 nm sections.

Appendix.　 Chemicals needed.

For fixation and embedding

0.2 M sodium phosphate buffer, pH 7.4.　 Prepare from NaH₂PO₄ and Na₂HPO₄.

8 % glutaraldehyde solution (EM grade)

Paraformaldehyde

1% Triton X-100 in water

2 % osmium tetroxide in water (we recommend package in ampoules)

TAAB EPON 812 kit

Acetone (99.5 %)

Acetone, dehydrated (commercially available for organic chemical synthesis)

For semi-thin sectioning and light microscopy

Toluidine blue

Sodium borate

Glass slides (APS coated)

For ultrathin sectioning and electron microscopy

Lead nitrate

Sodium citrate

Sodium hydroxide

Uranyl acetate

Polyvinyl formal

ethylene dichloride

Chloroform

Grids