3.14  Light and electron microscopy of archegonia embedded with epoxy resin

Takashi Murata and Naomi Sumikawa


In chapter 3.4, we described confocal microscopy of eggs and young embryos in cleared archegonia. Although the method is excellent for observation of tissue organization, subcellular structures are lost during clearing of tissues. Sectioning followed by light and electron microscopy is a useful method to visualize subcellular structures when good molecular probes for such structures are not available. In this section, we describe methods for light and electron microscopy of archegonia embedded with a low-viscosity epoxy resin. This method should be applicable for protonemata and other tissue with minor modification of fixative.


1 Fixation and Embedding

1)   Preparation of chemicals

WARNING: The chemicals below are hazardous. Use gloves. For detail of chemicals, see appendix.



5 % glutaraldehyde in 0.05 M sodium phosphate buffer, pH6.8.  Prepare just before use.



2 % osmium tetroxide in dH2O.  Handle in a hood because the vapor is extremely toxic. You can purchase the solution in ampoules. Use immediately after opening the ampoule. Instead, you can prepare the solution from osmium tetroxide and dH2O. The fresh solution should be clear (light yellow). The exhausted solution is black. Do not use the exhausted solution!


Epoxy resin mixture (Quetol 651 mixture by Nissin EM, Tokyo)

Prepare the mixture without accelerator. Add accelerator upon needed. Pot life of the mixture with accelerator is about 1 day.


Recipe for 50 ml mixture (without accelerator)

Quetol 651 (Nissin EM) 18.87 g

NSA 30.18 g

MNA 3.31 g


For 50 ml mixture, add 850 ml accelerator (DMP-30).  Mix thoroughly after adding the accelerator.



2) Protocols

I.     Pore the fixative to a plastic Petri dish.  Transfer a colony of P. patens to the dish.  Cut the colony with a razor blade or a surgical knife into small pieces (less than 5 mm square: as small as possible).  After that, transfer the tissues into a glass vial with fixative.

II.  Incubate the vial at room temperature for 5 h.

III.                     Wash the tissues with glutaraldehyde-free sodium phosphate buffer by three changes of solutions.  An interval of each change is 10 min.

IV.                     Incubate with post-fixative (2 % OsO4 in dH2O) at room temperature for 2 h.  Handle in a hood and use gloves.

V.  Rinse with 0.05 M sodium phosphate buffer.

VI.                     Dehydration with graded acetone series.

25% acetone     15-30 min, on ice

50% acetone     15-30 min, on ice

75% acetone     4ºC, overnight

99.5% acetone    15-30 min, on ice

100% acetone (dehydrated acetone)   15 min at room temperature, twice.


VII. Infiltration with resin (gently shake on rotator).

12.5 % resin (no accelerator) in deh. acetone      R.T. for 2 h

25 % resin (no accelerator) in deh. acetone       R.T. for 2 h

50 % resin (no accelerator) in deh. acetone       R.T. for 2 h

75 % resin (no accelerator) in deh. acetone       R.T. for 2 h

100 % resin (no accelerator)                  R.T. overnight

100 % resin (plus accelerator)                 R.T. for 4 h, twice


VIII. Transfer the infiltrated tissue to a mold.  Polymerize the resin at 60ºC for more than 1 day in an oven. Store the polymerized resin blocks until sectioning at room temperature under dehydrated condition.



(Continue to the next page)

2 Sectioning and light microscopy

Overview of sectioning is shown in the drawing below (in Japanese).


























1) Preparation of grids with formvar membrane (for electron microscopy only)

(Because formvar membrane becomes hydrophobic with time, the grids should be prepared on the day or one day before the sectioning.)


Wipe a glass slide with a dry lens paper.  Dip the slide into 0.5 % formvar in chloroform (or ethylene dichloride).  Bring the glass 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.  Dip slowly the glass into dH2O, which is filled in a 500 ml beaker or equivalent, so that a formvar layer is removed from the glass on the surface of water as a membrane.  Because some glass slides are sticky (depends on a lot of glasses), choose good glass slides before preparation.  Color of 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.  Faster up-speed from formvar solution will result thicker formvar membrane.  Because color of membrane varies even in a single membrane, put the grids onto better (that is, silver) areas of a membrane.  After putting grids, put a piece of Parafilm onto the membrane and remove the membrane from a surface of water.  Dry the membrane with grids on Parafilm and store until use in a clean Petri dish.

テキスト ボックス: Types of grids:
Single slot: A big (usually 2 x 1 mm) single hole is made on a grid.  All area on a section can be seen.  Formvar membrane is easily broken.
Mesh type:  Many small holes are made on a grid.  Only restricted area on a section can be seen.  Membrane is stable during observation and the grid is easily handled.






2)   Sectioning and light microscopy

I.                    Trim a block head into a 0.5 mm square.  Use a file for rough cut (first step).  For finishing use, use a razor blade under binocular microscope.

II.                 Cut the block head with a glass knife on a ultramicrotome until surface of a tissue appears.

III.               Upon encountering with the surface of a tissue, change the used glass knife to a diamond knife (for light microscopic sections) or a new glass knife with a boat.  Make 0.5 mm sections by the knife.  Transfer the sections from a boat to a APS (silane)-coated glass slide with a wire loop.  Observe the sections after staining with toluidine blue.  Repeat sectioning and observation, and stop sectioning when desired area will appear after a few sections.

- Toluidine blue staining -

Stain: 1 % toluidine blue in 0.1 M sodium phosphate buffer (pH7.0)


0.5 % toluidine blue in 2 % sodium borate

Drop the stain onto the sections, and incubate at room temperature for a few minutes followed by dH2O wash.  If staining is weak, heat the preparation on a heat plate for several ten seconds.

The staining will be fade after mounting with organic solvent-dissolved mounting medium.  Stained sections should be stored before mounting.


3) Sectioning for electron microscopy.

I. Change the knife to a diamond knife for electron microscopic sections (or freshly prepared good glass knife with a boat).  Make 70 nm sections.  Expand sections by chloroform vapor (by a toothpick dipped into chloroform solution).

II. Pick up the sections onto the grids with formvar membrane.  Store in a grid case until staining.


How to make a glass knife by glass knife maker (Messer type).















Cut the square glass as numbered.  Do not touch the gray area in the left and center schemes, because the area is used as a boat.  Finally, make a boat with an adhesive tape (right).


3. Uranyl acetate and lead citrate double staining

1)   Chemicals

1 % uranyl acetate in dH2O or in 70% methanol

Aqueous solution is the most commonly used for uranyl acetate staining.  However, the staining is sometimes weak for some low-viscosity resin or for thick sections. In such case, 70 % methanol solution gives stronger staining, but the solution easily precipitate (especially under illumination).  The methanol solution should be stored in the dark at –20ºC.


Laynold’s lead citrate

Dissolve 1.33 g of lead nitrate and 1.76 g of sodium citrate into 30 ml of distilled water.  Shake vigorously for 1 minute, and then keep at room temperature with shaking at a few minutes intervals.  After 30 minutes, add 8 ml of 1 N NaOH.  Confirm that the solution become clear.  Add distill water to 50 ml of total volume.  Seal tightly with Parafilm (avoid contact with carbon dioxide in air) and store at 4ºC.


2)   Protocols

I.     If you have many grids, attach the grids onto a specially designed stick bar (Grid-stick).  For use of Grid-stick, sections should be upside of the grid.

II.  Drop uranyl acetate solution onto a sheet of parafilm spread in a Petri dish (covered with alminium foil).  Put the grids onto the uranyl acetate drops, with section side down.  Incubate the grids in the Petri dish for 3-10 min at room temperature.

III.          Wash the grids with distilled water.

IV.          Drop lead citrate solution onto a sheet of parafilm spread in a Petri dish (filled with NaOH).  Incubate the grids with lead citrate solution for 3-5 min.

V.  Wash with distilled water.

VI.          Air dry and store until microscopy.


4. Microscopy

Please contact members of EM facility of your institute or university.  Operation of electron microscope should be learned by skilled persons, or by manufacturer’s training course.


Appendix.  Chemicals needed.


For fixation and embedding

0.2 M sodium phosphate buffer, pH 6.8 and pH 7.0.  Prepare from NaH2PO4 and Na2HPO4.

25 % glutaraldehyde solution (EM grade)

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

Quetol 651 resin kit (Nissin EM, Tokyo)

Acetone (99.5 %)

Acetone, dehydrated (commercially available for organic chemical synthesis)


For sectioning

0.5 % formvar in chloroform (or ethylene dichloride)


Toluidine blue

Glass slides (APS coated and uncoated)

Mounting medium



For staining with heavy metals (for EM)

Lead nitrate

Sodium citrate

Uranyl acetate