3.10 Light and electron microscopy of protonemata embedded with epoxy resin
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.
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.
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 60oC
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 40-60oC 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 -20oC 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.
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)
Toluidine
blue
Sodium
borate
Glass
slides (APS coated)
Lead
nitrate
Sodium
citrate
Sodium
hydroxide
Uranyl
acetate
Polyvinyl
formal
ethylene
dichloride
Chloroform
Grids