Introduction:
TEM sample preparation can be completed using many methods. The purpose of this preparation is to 1) preserve the structure of the sample as close to the natural state as possible, 2) make the sample compatible with the environment of the electron microscope, and 3) make it able to keep its structural integrity while being cut into ultra thin sections. This protocol utilizes the method of chemical fixation, dehydration, and embedding for sample preservation. It should be noted that there is no one best method for preserving all biological materials as each type of tissue has its own requirements for best preservation. Considerations for pH, osmolarity, and physical/chemical characteristics need to be addressed when selecting specific chemicals and processing times for a specific tissue and desired outcome. Because of the nature of what we are trying to accomplish with this process you need to be aware that most of the chemicals used are very toxic or carcinogenic and need to be handled with great care. Most of the steps should be performed in a functional fume hood. Refer to the MSDS sheets for each of the chemicals for further information. The following protocol will work with most mammalian tissues and provide adequate results for morphologic analysis. A flow chart for the procedure follows.
Procedure:
Sample Acquisition:
Samples need to be as fresh as possible prior to placing in the primary fixative. Ideally the fixative should be used to kill the tissue being used. If the entire organism is being used the perfusion of the anesthetized animal is desired if the animal is small enough. Otherwise perfusion of the organ being studied is appropriate. If the animal is not going to be euthanized as part of the sample acquisition then removing the tissue prior to fixation is necessary. In this case, or in the case of not being able to perfuse the sample then plunging the sample into the primary fixative as quickly as possible is necessary. This may necessitate dissection of the desired tissue prior to putting into fixative if the organ is too large to put into fixative immediately.
Once the tissue has been acquired it needs to be put into the primary fixative and cut down to appropriately sized tissue blocks and then allowed to continue fixing for at least 2 hours in the primary fixative. Tissue blocks need to be quite small. One axis of the block needs to be no more than ½ mm in thickness to allow for complete fixation with the secondary fixative. The other axes can be larger to allow for orientation if needed. Still, no one dimension of the tissue block should be more than about 2 mm.
Primary Fixation:
We typically use a primary fixative of 2% glutaraldehyde in 0.06 M sodium cacodylate buffer. The fixative itself can be quite harsh on the tissue so rather than just using an aqueous solution of the fixative we mix it in a buffer to help control pH and osmolarity. The buffer helps keep the cell components intact while the fixative is reacting with the tissue and keeps the cell from swelling or shrinking during processing. There are many buffers available that will work well for sample fixation. Sodium cacodylate is used in this lab because it is stable over a long time and doesn’t promote microbial growth. It is, however, quite toxic due to the As component in it so it must be handled with care and disposed of properly. Other common buffers that are used are various phosphate buffers, zwitter ionic buffers, and others that can be found in the literature. The different buffers will affect the appearance of the tissue so choosing one that emphasizes the structures of interest could be important. It should be noted that PBS is not suitable for carrying the fixative as it has insufficient buffering capacity to keep the pH in the proper range. It can be used for washing the tissue between steps if desired.
There are several types of primary fixatives available, most of which use some type of aldehyde as the fixative. As mentioned above, we typically use 2% glutaraldehyde as our primary fixative. Glutaraldehyde is typically used in concentrations of 2% to 4%. It is often paired with freshly made formaldehyde (derived from paraformaldehyde) in concentrations of 2% to 4%. Aldehyde fixatives act primarily to crosslink protein found in the sample helping to stabilize the sample and hold everything in place. The sample should be in the primary fixative for at least 2 hours unless the sample is very small such as a cell culture, in which case the sample will be adequately fixed after about 1 hour. If using small particulates, such as a cell culture, it is important to wash the cells first in a buffer prior to fixing as the fixative will crosslink any protein in the supernatant and will make further processing difficult.
Wash:
The next step is to wash the sample. This wash is used to remove all unreacted primary fixative from the system so that it doesn’t adversely affect the following steps. Washing is accomplished by removing the primary fixative and then adding a buffer to the sample vials and soaking for a period of time. We typically recommend 10 minutes of soaking time prior to the next step. This should be repeated 5 to 6 times to make sure all of the unreacted fixative has been removed from the tissue. It is important to remember that all of these steps require diffusion of the chemicals through the tissue and this takes time. It is better to go longer than it is to rush any of these steps. In fact, the times recommended in this protocol should be considered minimums rather than absolutes.
Post or Secondary fixation:
Following the washing step we again fix the sample with another fixation step. In this step we use 1% osmium tetroxide (OsO4) in the buffer. This should go for at least 1.5 hours but not more than 2 hours unless the specific protocol being used indicates differently. It is important that you realize that OsO4 is highly toxic and must be used in a functional fume hood. OsO4 primarily fixes unsaturated lipids. It does this by breaking the double bond in the lipid and inserting itself into the bond of two adjacent lipid molecules. This crosslinks the lipid stabilizing it and adding a heavy metal to the sample. This heavy metal helps add contrast to the sample when imaging. It also provides a bonding site for other metallic stains which also provides for better contrast. Be aware that OsO4 is highly reactive so make sure your sample is ready for the treatment prior to applying it and that all of it gets washed away before continuing with further processing. Once the secondary fixation step is complete, make sure you dispose of the waste OsO4 in the proper container. This container is located in the fume hood and contains vegetable oil that will help reduce the OsO4 and make it safer to handle.
Wash:
After the OsO4 post fixation it is very important to wash the unreacted OsO4 out of the tissue. If this does not happen then OsO4 precipitate is likely to occur in the tissue. This wash is performed as the previous wash with the exception of using distilled water instead of buffer. At this point in the processing the tissue is stable. Concern with pH and osmolarity is no longer a problem. The waste water from this stem should be disposed of in the cup sink in the back of the fume hood.
In Block Staining:
This step is optional. After completing the second washing step, contrast can be added to the sample by allowing it to soak in 0.5% uranyl acetate (UA) overnight. This step adds uranium to the sample enhancing contrast in the images. It is important to note that UA can extract glycogen so if that is a component that is expected in the sample then this may not be a good place to add this contrast. UA can also be used post-sectioning to add contrast as well. If for some reason you will not be able to continue the sample processing the next day it is best to skip this step to avoid over-staining.
Dehydration:
Depending on the resin you are planning on using, this step may need to be modified. This protocol assumes embedding in Spurr’s low viscosity resin.
Dehydration is needed to remove water from the sample and substitute it with a solvent that is suitable for the resin being used. This is accomplished by using either a graded series of ethanol or acetone. The Spurr’s resin that we typically use in this lab requires acetone as the final solvent so if ethanol is used then a final series with 100% acetone will be necessary. Dehydration is carried out very similarly to the washes by going through a series of increasing dehydrant concentrations until 100% dehydrant us used. We use the following series: 10%, 30%, 50%, 70%, 95%, 3X100%. Again if ethanol is used then we will also need to complete the series with 3X100% acetone. Any water remaining in the tissue will cause voids or result in unpolymerized resin. It is therefore critical that the final steps of dehydration be completed with absolutely pure acetone.
Infiltration:
From this step forward you will be working with embedding Spurr’s low viscosity embedding resin. If you are using another resin and need help then consult with the lab manager. This resin is carcinogenic and is quite messy and sticky (like honey but won’t clean up with water). For this reason all of the remaining steps need to be carried out in the fume hood and there should be ample (as in a lot) of paper towels on the work surface being used to catch any spills and drips. Also, the waste will be put in a special container in the fume hood instead of the usual waste containers. All of the other containers and tools used with this process needs to be disposable so we don’t ruin our glassware.
Mix the Spurr’s resin as demonstrated by Mike. The mixed resin will stay usable for at least 24 hours at room temperature. Remove the acetone from the vial leaving enough to cover the samples. Add 100% acetone to the vial until it fills the vial approximately ⅔ full. Do this for all of the sample vials you are working with. Then fill the vial to the neck, leaving a little air at the top, with the prepared resin mixture. Shake the vial to combine the two ingredients until it is homogenous. Cap the vial tightly and place on the shaker. Do this with all of the sample vials. Leave the vials on the shaker for at least 1 hour. If the sample pieces are large or particularly difficult to infiltrate then it may need to go longer.
Remove the first dilution of resin from the vial(s) leaving the samples covered with a little of the mixture. Add 100% resin mixture to the vial(s) until approximately ⅔ full. Fill to the neck with 100% acetone. Shake as before and put on the shaker for at least another hour. This step can go overnight if necessary.
Remove the second dilution of resin from the vial(s) removing as much of the mixture as reasonably possible. Fill the vial(s) with 100% resin and sake for at least another hour.
Embedding:
Select the proper embedding mold for your sample. If orientation is not an issue then use the Beem® capsules. If orientation is an issue then use the bottle caps or other suitable embedding mold. Make labels for each of the molds by creating a unique identifier on a piece of paper using a pencil. Ink will dissolve in the resin so pencil is required. Put the label in each of the molds. Fill each of the molds most of the way with fresh Spurr’s resin.
Remove some of the resin from the sample vial(s). Carefully remove one of the sample pieces using an applicator stick and then place the sample into the embedding mold. If using Beem® capsules then only put one piece of sample in each one unless you have lots of samples then you may put more than one sample in the capsule. Generally you will probably trim away excess samples so more than one piece isn’t really helpful. If you are using bottle caps then feel free to put multiple pieces in them as they will be removed and remounted for correct orientation.
Once the samples are in the embedding molds, finish filling the molds with fresh resin until they are completely full. Don’t fill them until they overflow but do make sure they are filled to the top. You need to make sure there is enough resin around the sample to work with. It is very difficult to work with samples that don’t have enough resin.
After the samples have all been put into the embedding molds, put the molds into the polymerization oven at 70 degrees C at least overnight.
This completes the sample preparation process. The rest of the preparation includes sectioning and staining which will be covered elsewhere.
TEM Sample Processing Outline
Acquire Sample
- Sample must be small - < 0.5 mm in one axis and no more than 2mm in other axes.
- Put in fixative as quickly as possible - ideally fixative should kill the sample.
Primary Fixation (2% glutaraldehyde in buffer)
- Need to be in fixative for at least 2 hours for tissue blocks indicated above.
- Can store tissue in primary fixative for several weeks if necessary but best results will be obtained when processed as soon as possible.
- Small particulates that need to be centrifuged need to be washed prior to fixation to remove any protein in the supernatant.
- Dispose of chemical waste into appropriate waste container
Wash samples in buffer
- 5 to 6 times for 5 to 10 minutes each wash
- Dispose of chemical waste into appropriate waste container
Post or Secondary fixation (1% OsO4 in buffer)
- Fix for no more than 2 hours unless specific protocol indicates otherwise
- Handle this fixative very carefully as it is highly toxic.
- Dispose of this fixative in the appropriate waste receptacle in the fume hood.
Wash with distilled water using same procedure as before
- Waste water goes down the cup sink in the fume hood
- Samples can be held at this step for a few days if necessary
In-block stain with 0.5% uranyl acetate (UA) overnight
- Dispose of waste UA down the sink with plenty of water
- May extract glycogen so use with caution
- This step is optional - exclude it if processing cannot be continued the next morning
Dehydrate in ethanol or acetone series
- Use the series 10%, 30%, 50%, 70%, 95%, 3 X 100%
- Must finish with 3 X 100% acetone
- Time in each concentration should be 5 to 10 minutes
- Samples can be held for extended periods of time in 70% ethanol
- Dispose of waste in appropriate waste container
Infiltrate sample with Spurr’s Low Viscosity resin
- Remove most of the last 100% acetone soak
- Put fresh 100% acetone into vial until approximately ⅔ full
- Finish filling vial with 100% Spurr’s resin
- Shake to mix thoroughly
- Put on shaker for at least 1 hour
- Repeat the previous steps exchanging Spurr’s resin with the 100% acetone
- Put on shaker for at least 1 hour
- Remove resin mixture and replace with fresh 100% Spurr’s resin
- Put on shaker for at least 1 hour
Embedding
- Make paper labels for each of the selected embedding molds using pencil to write on the paper
- Place labels into the molds
- Fill molds most of the way with 100% Spurr’s resin
- Carefully put samples into the molds with the resin
- If necessary push samples down to the tip of the embedding mold
- Finish filling molds with 100% Spurr’s resin
- Put embedding molds in a 70 degrees C polymerization oven overnight to polymerize.
- Check resin to make sure it has polymerized. Samples are ready when pressing a fingernail into the resin does not leave a mark.