Lab Techniques in Electron Microscopy, Fall 2008
TEM and SEM Specimen Preparation Supplementary Manual
Arkansas State University
Dr. Stanley Trauth
Table of Contents
Specimen Preparation for SEM……………………………………………….…………..2
Specimen Preparation for TEM……………………………………………….…………..3
Chemical Fixation/Postfixation…………………………………………….…...………...4
Fixatives/Buffers……………………………………………………………….………….5
Dehydration of Specimen…………………………………………………………………8
Infiltration of Specimen………………………………………………………...…………8
Embedding of Specimen…………………………………………………………………..8
Sectioning of Specimen………………………………………………………...…………9
Glass Knife Making……………………………………………………………………….9
Trimming of Specimen Mold…………………………………………………...………..11
Copper Grids……………………………………………………………………..………15
Staining of Specimens…………………………………………………………..………..15
TEM Viewing of Specimen…………………………………………………….………..10
TEM Photographing of Specimen……………………………………………….………10
Developing Negatives of Specimen………………………………………………...……17
Development of Prints…………………………………………………………….……..18
Printing Negatives of Specimen…………………………………………………….……18
1
SUMMARY OF MAJOR STEPS FOR PREPARATION OF SPECIMENS FOR
SEM
1.
2.
3.
4.
5.
6.
7.
8.
9.
Fix in glutarladehyde
Rinse in buffer
Postfix in osmium tetroxide (optional)
Rinse in buffer
Dehydrate in a graded series of Alcohol
Place in Amyl Acetate solutions (optional)
Critical point dry in liquid carbon dioxide
Coat with carbon, then gold or gold-palladium
View with Vega TS 5136 XM SEM
(steps prior to digital SEM)
10. Take photographs of specimens at desired magnifications on Polaroid 52 or 55 PN
film
11. Coat positive prints
12. Place negatives in sodium sulfate solution
13. Wash negatives in running tap water
14. Hang to dry
15. Interpret positive prints
16. Label and store Polaroid positives in sheet protectors
17. Use Polaroid positives for study purposes
18. Print negatives using standard darkroom procedures
19. Dry micrographs
20. Label and store micrographs in sheet protectors
21. Place sheet protectors in binder
SUMMARY OF MAJOR STEPS FOR PREPARATION OF SPECIMENS FOR
TEM
1. Fix in gluataraldehyde
2. Rinse in buffer
3. Postfix in osmium tetroxide
4. Rinse in buffer
5. Dehydrate in Alcohol series
6. Infiltrate in Propylene Oxide and Epon 812
7. Embed in Beem Capsules in full strength Epon 812
8. Polymerize in oven
9. Trim blocks
10. Cut thick sections for light microscopy
11. Cut ultra-thin sections for TEM
12. Pick up on 200 or 300 Copper mesh grids
13. Stain in uranyl acetate and lead citrate solutions
2
14.
15.
16.
17.
18.
19.
20.
21.
View grids in JEOL 100 CXII TEM
Take photographs of sections at desired magnifications
Develop plates or films
Make print from EM plates or films
Dry electron micrographs
Interpret electron micrographs
Label and store micrographs in sheet protectors
Place sheet protectors in binder.
Preparation steps up to TEM viewing:
Day
1
2
3
4
5
Schedule
Tissue Removal and
Fixation
Equilibrate in
Refrigerator
Washing in Buffer
Postfixation
Wash and place in
buffer
Dehydration
Rotation
Preliminary
embedding
Bell jar
Final embedding
Put in oven
Making stains
Trim and cut thick
sections
Trim and cut thin
sections
Time (constant)
1
Time (waiting)
2.5
1.5
2
1.5
3
Overnight
1
1
1
1-2 days
1
1
1
Ready to view on TEM!
07-19-02
MNM
3
SEM Specimen Preparation:
Fix tissue for 1-2 days in 10% buffered formalin if soft tissue.
Remove and place in 70% ethanol solution.
Dehydrate the tissue in the following manner:
1.
2.
3.
4.
5.
6.
70% ethanol
90% ethanol
Absolute ethanol
50/50 ethanol-amyl acetate
Absolute amyl acetate
Absolute amyl acetate
30-45 min.
same
same
same
same
same
Tissue is now dried in critical point dryer. The size of the specimen is limited according
to basket. Be sure to do more than one tissue since this takes time.
New copper boats should be made with double stick tape to put specimens upon after
critical point drying. Be sure to etch the bottom with a diamond knife to label specimen.
After coating the specimen with carbon or gold/palladium, the specimen is ready for
viewing.
TEM Tissue Preparation:
The tissue mass should be no larger than 3 mm2; the best size is 1-2 mm2. Once the
tissue is removed from the organism, it should be placed in a mincing dish with fixative
and cut in ½ to 1 mm cubes using two Weck razorblades. Long and short blades will cut
at different thicknesses. Put the minced cubes in vials containing fixative, which should
be at room temperature and buffered. The fixation process is described in the next
section.
Chemical Fixation/Postfixation:
Fixatives/Buffers:
Common fixative is 2-6% glutarladehyde concentration depending on tissue and
difficulty of penetration. Fixative is buffered, commonly in phosphate buffer; sodium
cacodylate is suggested.
The recipe will be stored in a stock and working solution. The amount of buffer depends
on the amount of glutaraldehyde used. The following recipes and mixes are suggested:
Stock Buffer of 0.2 M sodium cacodylate
10.7g sodium cacodylate
Dilute to 250 ml with distilled water.
Store refrigerated.
Adjust pH according to table.
4
The following table can be used to adjust the pH of the buffer. It is calculated based on
adding the designated 0.2 M HCl to 100 ml of 0.2 M sodium cacodylate buffer.
0.2 M HCl
ml
5.4
8.4
12.6
18.6
26.6
36.6
47.6
59.2
69.6
78.4
86.0
90.0
94.0
pH
7.4
7.2
7.0
6.8
6.6
6.4
6.2
6.0
5.8
5.6
5.4
5.2
5.0
This table can be interpolated if needed.
Usually it is desirable to make at least 1 Liter of buffer.
Stock glutaraldehyde solution comes in 25, 50, and 70%
Recipe 1:
4% glutaraldehyde in Cacodylate Fixative CAUTION: HIGHLY TOXIC
8 mL 25% glutaraldehyde
25 mL 0.2 M Sodium Cacodylate
Adjust pH to 7.2 - 7.4 with 1 M HCl solution if not already done
Dilute to 50 mL
For 2% only use 4 mL 25% glutaraldehyde. It is recommended to use 2% for animal and
4% for plants.
Recipe 2:
2% glutaraldehyde in Cacodylate buffer – pH 7.3
50 mL 0.2 M Sodium Cacodylate buffer
4 mL 50% Stock glutaraldehyde
46 mL distilled water
Recipe 3:
2% glutaraldehyde in Cacodylate buffer – pH 7.3
35 mL 0.2 M Sodium Cacodylate buffer
2 mL 70% Stock glutaraldehyde solution
33 mL distilled water
5
Fixation:
2% glutaraldehyde usually takes 2-4 hours (2 ½ best). Care must be taken not to
underfix. Fixation must be done in the refrigerator with no light and in a Wheaton vial.
Washing:
The specimen is now taken out of the refrigerator and carried through a four-step rinse in
buffer such as sodium cacodylate buffer or phosphate buffer. Buffer should be placed in
vial following removal from fixative solution. The process involves covering the
specimen with the buffer with a 2 mL Pasteur pipette. This may involve 2-5 mL of buffer
depending on the size of the specimen. Allow to set for 10-15 minutes and then remove
the buffer with the pipette. Add and remove the buffer three more times in 10-15 minute
cycles.
Postfixation:
Postfixation is carried out with a 1% solution of osmium tetroxide. The following recipe
is recommended:
1% Osmium Tetroxide mixture:
1g osmium tetroxide
Dissolve in 25 mL distilled water overnight
(This makes 4% osmium tetroxide solution as a stock or concentrated solution).
5 mL concentrated stock solution
5 mL distilled water
10 mL 0.2 M sodium cacodylate
This makes 20 mL solution of 1% osmium tetroxide solution.
This postfixative retains lipids well, however it will not retain the proteins as well as
glutaraldehyde. The osmium is toxic and must be kept in the refrigerator. Wear gloves at
all times and do not breathe fumes.
Postfixation takes 1 ½ to 2 hours and must be kept in the refrigerator with no light during
the postfixation process. Once completed, remove the post fixative while at the same
time adding buffer using two pipettes. Again, wash the specimen four times as shown in
the previous section. Some say specimens may stay in buffer indefinitely, however, it is
probably best to go on to the next step immediately.
6
Dehydration:
The specimen must be dehydrated in a solvent. Ethanol is the most common. Dehydrate
the specimen in:
50% ethanol for ½ hour
70% ethanol for ½ hour
90% ethanol for ½ hour
100% ethanol for ½ hour
100% ethanol for ½ hour
100% ethanol for ½ hour
Once completed, the specimen must be put in a transitional solvent which is usually
acetone in the following manner:
50/50 ethanol acetone mix for 10-15 min.
100% acetone for 10-15 min.
100% acetone for 10-15 min.
Infiltration:
This involves the infiltration of the epoxy into the tissue in the following steps:
1. Mix the epoxy—see instructions.
This plastic mix is good for plants and animal tissues as an all around
embedding medium.
NOTE: Always use gloves during mixing or handling the plastic mix. Do not
touch the plastic mixing agents. Read the published warnings about each
chemical. Be sure to always wipe off bottle necks with paper towel. You
may use acetone as a solvent for cleaning. Be sure to use a fume hood.
Mollenhauer’s Epon-Araldite Mixture #2
(Makes about 230 mL)
62 mL Epon 812
81 mL Araldite 502 or 506
100 mL DDSA Hardener (Dodecenylsuccinic Anhydride)
4-7 mL DBP Plastizer (Dibutylphthalate)
The Epon 812 and Araldite 502/506 should be thoroughly mixed. Add the DBP Plastizer
to the DDSA Hardener and then mix into the first solution. Bubbles will form as a result
of mixing, but these will disappear. This solution should be refrigerated until used.
DMP-30 Accelerator is added to the mixture at time of use.
DMP-30 Accelerator has a life span of approximately six months after opening.
(For infiltration, use 4 drops per 10 mL mixture; for embedding, use 6 drops per 10 mL
mixture)
7
2. Measure epoxy out in tri-cornered beakers – usually 50 - 100 mL.
3. Add 4 drops of DMP-30 accelerator per 10 mL of epoxy.
4. Mix thoroughly and then add acetone to make a 50/50 mixture of epoxy resin
with accelerator and acetone.
5. Place specimen and tissue into vials in 50/50 mixture.
6. Put on rotator and rotate overnight.
Embedding in Plastic:
1. After rotation overnight, take tissue out of vials with wood applicator sticks
(make sticks by trimming end of sticks with knife) and place first into Beam
capsules or gelatin capsules (half full of resin) in a Beam capsule holder. Next,
cover with full strength epoxy mixture using a disposable pipette. This mixture
should have 6 drops of DMP-30 accelerator added for every 10 mL epoxy resin.
2. Place the Beam capsule holder into a bell vacuum jar under mild vacuum for
about 1 hour. This will bring the air bubbles to the surface for removal.
3. Final infiltration is done with pure epoxy resin mixture with 6 drops of DMP-30
accelerator per 10 mL. The tissue should be transferred either to Beam capsules
or plastic embedding molds. The procedure is filling the molds half way, placing
the specimen inside the mold and then filling the mold the rest of the way with
epoxy mixture.
4. Place the mold or capsules in an embedding oven for 1-2 days, normally one day
minimum at 60C. This allows for proper polymerization of the resin.
The tissue should now be embedded in tissue blocks that contain a mass of tissue. These
can be snapped out and should be ready for sectioning.
Be sure to label the samples properly. If you use the Beam capsules you can write small
labels, or if blocks are used, you can use legible writing instruments and indelible ink on
the end that will not be trimmed.
8
Glass Knife Making:
Sectioning is done with glass knives or diamond knives. Glass knives are made with 5
mm thick glass cut in 25 X 25 mm sections. The glass is scored diagonally from end to
end and then broken. The right half is thrown away and the left half is used. The glass
should only be touched with the fingers on the wide flat areas, never on the cutting edges.
The upper left tip is used as the cutting edge. Observation of this edge shows a fracture
mark. This edge can be divided into three parts: the sharpest cutting edge, center
section, and the rough cutting edge from left to right. The rough cutting edge is used for
the initial thick sectioning. The sharp cutting edge is used for the thin sectioning.
Normally, two knives are made, one being used for each type of sectioning, however, the
sharp cutting edge may be used for both.
Next, a water boat is added to the knife, using tape or preformed plastic boats. To seal
the boats to the knife, use nail polish or dental wax.
NOTE: The diamond knives should be kept for the experienced user.
Glass knife preparation drawings:
Inspect the knife edge face on. These are examples of some of the types of edges most
commonly seen:
9
Instructions for making knives:
Using LKB-7800 Knifemaker
1. Put on gloves – so not to smudge glass.
2. Get a piece of glass (cabinet above glass maker).
3. Clean glass with soap and water using a fine brush.
4. Treat glass with 20% acetone.
5. Air-dry.
6. Place the glass (burred edge down) with alignment on knifemaker.
7. Return break knob (black knob – front) all the way to the left.
8. Turn small metal control (front part of white scoring apparatus) to diagonal (//).
9. Make sure the pronged holder is under the left side of the glass.
10. Pull the lever on the left side of the machine (long metal rod with black knob) up,
and then down tightly to secure the glass in place.
11. Score the glass by pulling out the white knob on front of scoring apparatus; leave
the knob at this position until glass is broken.
12. Turn the large black knob (machine right front) to the right slowly until the glass
breaks.
13. Release white scoring knob, return break knob in front to the left, and release
black bail lever on the left of the machine.
14. Take the glass holder and bring the square glass toward you until you can position
the glass diagonally. (counter-clock-wise)
15. Release the small black knob on the left of the knifemaker, which will secure the
glass in the diagonal position, adjust with the small metal lever in front.
16. Again secure the glass with the black ball lever on the left.
17. Place the small metal control 25mm on front of scoring apparatus.
18. Score the glass by pulling the white knob toward you and leave in this position.
19. Turn the large black break knob on the right front to the right until the glass
breaks.
10
20. Repeat step 13. Remove the glass with the holder. Keep the left side to use for
your knife and throw the right side away.
21. Take the knife to the trimming scope and check for a good cutting edge (fracture
mark).
22. Secure a plastic boat on the knife by using fingernail polish. Be sure the top edge
of the boat is even with the cutting edge of the knife. The purpose of the boat is
to hold water, so get a tight seal.
23. Let air dry at least 10 minutes before use. Knives should be made only when
ready to section, as exposure to the air will dull the cutting edge. When dry, put
knife in a petri dish with cover until ready to use.
Trimming/Sectioning Specimens in Plastic Block:
A Weck razorblade and a dissecting microscope must be used to properly trim the
specimen for sectioning. Cut the specimen block to the specimen is in the block as
demonstrated in the following picture:
Next, trim the face so that it appears as shown here:
Thick sectioning:
11
The first sectioning is called thick sections. They are 1-2 m and are placed on a
glass slide, stained with a multiple stain, and examined under a compound
microscope to see if the tissue meets the desired quality.
Thin sectioning:
Next, the block must be trimmed to smaller specifications as shown in the figure below:
Procedure:
1. Secure tightly a specimen block under the dissecting microscope.
2. Focus microscope and trim the specimen at a 45-degree angle according to
above specifications for either thick or thin sectioning.
3. You may need to adjust the position of the scope and the light to focus in
properly.
Sectioning:
Once trimmed properly, the ultramicrotome is used to cut sections 40 nm to 70 nm thick.
One should look for a gray to silver color, the gray being the thinnest and the silver being
best. Never use the blue, gold, brown, or green sections for these sections will be too
thick. A ribbon should be formed that is face to face flat, as shown in the figure below.
When a ribbon of sections is formed, take the eyelash devise to center the sections in the
trough of water. Use a Q-tip with xylene or acetone to flatten the sections. Never touch
the sections with the Q-tip, just wave them above the section. This will cause the plastic
to flatten. At least 12-16 sections should be sliced.
12
Once the sections are centered, a copper grid should be crimped with forceps and touched
to the slices. Adhesion and surface tension of the water should cause the slices to cling to
a bubble on the grid. Turn over and touch the side of the grid without the specimen to the
edge of a kim wipe or filter paper to remove excess water and allow to dry briefly.
Copper grids: Copper grids are available in different sizes from 200 to 400. Remember,
the larger the number, the smaller the grid space.
Ultramicrotome Instructions:
1. Turn on the power--the switch is located on the control module 4802A.There
should be a 20 minute warm-up time.
2. Assemble all needed equipment:
Thick sectioningdistilled water
Glass slides
Pasteur pipette with bulb
Q-tip
Beaker
Forceps
Absorbant sticks
Permanent marker
Filter paper
Multi-stain (Ladd)
Thin sectioning – all if the above plus grid washing kit with the eye last
devise and grids for the sections.
3. Place the trimmed block in the appropriate holder (the beam capsule has a special
round based holder), secure tightly in the holder, then place in the microtome and
secure the holder with the block with the adjusting screw located underneath.
4. Release the specimen holder arm on the left side of the microtome.
5. Place the glass knife in the microtome and use adjustment screws to get the proper
position. Fill the boat with water—meniscus next to the cutting edge.
6. Adjust the microscope, turn on the focus light on the control module and focus in
on the specimen block while slowly using the micro/macro feed knobs on the
right of the microtiome to bring the cutting arm and specimen adjacent to the
glass knife.
13
7. Control module 4802A: Select cutting speed, thickness, adjust the focus light,
turn on the blower, set on “manual” for thick sectioning or “continuous” for thin
sectioning.
8. Turn the heat block on low.
9. Checking all settings again, begin to section by manually lifting the cutting arm
on the microtome and at the same time slowly macro/micro feeding with the
knobs on the right. Make sure you lift the cutting arm all the way up and make a
full swing each time. Adjust the thickness on the control module as needed to get
the right colored sections.
10. Thick sections: Remove the sections with the loop and place in drops of water on
a glass slide, make several slides, then place on the heat block to dry. When
slides are dry, stain them (see staining instructions) and check the sections under
the compound microscope for quality of tissue. Be sure to paramount the stained
slides and label properly with glass marker or indelible ink.
11. Thin sections: Re-trim the block and replace in the microtome. Reset all
adjustments. Prepare grids (see grid cleaning instructions) and do thin sectioning
using the continuous mode and adjusting as necessary for the silver color (see
color chart).
12. When you have a ribbon of silver colored sections in the trough, center with the
eyelash devise, take a properly cleaned grid and crimp the outer edge with the
forceps and touch the dull side of the grid to the sections. Then while still
clamped in the forceps, turn the specimen up and touch the bottom or shiny side
to a kim wipe or filter paper to remove all excess water and put in a numbered
grid petri dish for Uranyl Acetate and Lead Citrate Staining.
13. Remove all settings on the control module, leaving only the blower on for a 20
minute cool down time.
14. Remove the specimen and holder from the ultromicrotome, then remove the
specimen block from the holder.
15. Remove the cutting knife from the microtome.
16. Reset the cutting arm lever on the left to the lock position. Clean then cover the
microtome.
14
17. At the end of the 20 minute period, turn off the power on the control module
4802A and put on the cover.
18. Put up all equipment and clean the working area.
Grid Cleaning Procedure:
NOTE: The larger the grid number, the smaller the grid size. Prior to picking up tissue,
always clean the grid as follows:
Pick up grid with forceps and dip in the following solutions:
5 seconds
95% acetone
blot dry
5 seconds
0.1 M HCl
blot dry
5 seconds
distilled water
blot dry
Multi-Stain Procedure – Ladd stain:
1. Take the slide off the heat block with a close pin.
2. Take immediately to the sink and place Ladd stain on the slide on top of the
sections. Some tissue takes longer to stain, so you may have to make an
adjustment for the time the stain is left on the slide, usually 5-20 seconds.
3. Rinse with distilled water. Let air dry or blot dry.
4. Under compound microscope check for tissue quality, then paramount.
Uranyl Acetate and Lead Citrate Staining:
Use uranyl acetate and lead citrate for 5-10 minutes for staining. This is done with a 16slot rubber mat made for staining put into a petri dish with sodium hydroxide pellets.
First a drop of uranyl acetate is placed in each slot. Then with the forceps, place the grid,
specimen side down, on the droplet. Follow the same procedure with lead citrate stain in
another petri dish with fresh sodium hydroxide pellets. After staining, use a third petri
dish with pellets for drying. Once dry, store grids in a labeled grid box.
Once a large sample size (5-10) grids have been accumulated, the grids are ready for
observation and study on the TEM. You should have at least 150 micrographs of
specimen in some cases for stereology or quantitative microscopy. TEM viewing of
specimen should be done as instructed in TEM operation instructions. See TEM
instruction section.
15
Material needed:
1. 3 petri dishes
2. Aluminum foil
3. Pipettes (Pasteur)
4. Rubber bulb
5. NaOH pellets
6. Uranyl acetatae – Refrigerator door in a flask
7. Lead Citrate – Refrigerator door in a brown bottle
8. Timer
9. Grid box
10. 2 beakers with distilled water (Nalgene bottle)
11. Forceps for rinsing
Procedure:
Place a sheet of aluminum foil on working area.
1. Place 3 petri dishes filled with NaOH pellets on the foil.
2. Put a 1 x 2” sheet of dental wax in two of the dishes.
3. Place 5 drops of Uranyl Acetate on the dental wax in dish #1.
4. Place 5 drops of Lead Acetate on dental wax in dish #2.
5. With forceps, take the grid with specimen side down and place on top of the
uranyl acetate drop; repeat doing 5 specimens at a time or more if space
allows.
6. Stain 5-10 minutes. Set timer.
7. Pick up grid with forceps (always stay close to the edge with the forceps) rinse
5 times in beaker of distilled water by dipping in and out.
8. Blot on filter paper with specimen side up so rinse filters through specimen to
paper.
9. Repeat in second beaker of distilled water.
10. Place in Lead Acetate drops for 5-10 minutes.
11. Rinse as above.
12. Place in third petri dish with NaOH pellets for drying.
13. Take out of petri dish and place in a grid box.
14. Be sure to label specimens as to the position in the grid box.
15. Put stains back in the refrigerator and clean the area.
16