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All procedures involving animal samples have been reviewed and approved by the appropriate animal ethical review committee.
1. Preparation of Brain Tissue and SBFSEM Imaging
- Anesthetize a young (~2 - 4 months old) C57 black mouse (C57BL/6J strain) using 4 - 5% isoflurane following the institutional guidelines. Confirm anesthetization by monitoring the loss of muscle tone, lack of voluntary movements, and responses to aversive stimuli like a tail pinch.
- Pin the mouse on a dissection tray and make an incision on the skin along the ventral midline. Make further incisions in the skin to expose the rib cage of the mouse. Incise the diaphragm, carefully dissect the chest cavity along the periphery to expose the beating heart, and then make an incision on the right atrium.
- Using a butterfly cannula, cannulate the left ventricle. Perfuse the mouse transcardially using 10 - 20 ml of phosphate buffer saline (PBS) pH 7.2, until exsanguination is confirmed by a change in the color of the liver.
Note: A change in the color of the liver is used as a guide to determine the extent of exsanguination. Confirm that the liver color changes from reddish brown to pale pink. - Perfuse the mouse transcardially using 10 - 20 ml of 2% glutaraldehyde and 4% paraformaldehyde made in 0.10 M cacodylate buffer (pH 7.2), to fix the brain tissue rapidly from within. Fixation is monitored by observing the tail stiffening.
Note: Prepare 0.10 M cacodylate buffer (pH 7.2) by dissolving 2.14 g of sodium cacodylate in 80 ml water, add hydrochloric acid to adjust the pH, and make the volume to 100 ml with water. - Following perfusion, decapitate the mouse dissect the brain, followed by fixation in 0.10 M sodium cacodylate buffer (pH 7.2) containing 2% glutaraldehyde and 4% paraformaldehyde for 48 hr at 4 oC.
- After 48 hr make 400 µm coronal sections using a vibratome and then carefully dissect the region of interest in the brain (for example, the hippocampus) under the dissection microscope. Carefully trim the tissue and take a picture for preserving orientation.
Note: Post processing methods of tissue staining in SBFSEM combines a variety of heavy metal staining methods in order to improve resolution. . - Wash the glutaraldehyde-fixed tissues 3 times, 5 min each, in 0.1 M cacodylate buffer (pH 7.2).
- Prepare 0.1% tannic acid solution by dissolving tannic acid in 0.1 M sodium cacodylate buffer (pH 7.2). Swirl until dissolved and filter through a 0.45 µm filter, if necessary.
- Postfix the tissues with cacodylate buffered 0.1% tannic acid by incubating in 1 ml reagent for 30 - 60 min at RT.
Note: The incubation time is dependent on tissue size, but 30 min works best for most tissues. - Wash tissues 3 times, 5 min each, in cacodylate buffer (pH 7.2).
- Dissolve 0.3 g potassium ferrocyanide and 0.86 g sodium cacodylate in 10 ml distilled H2O (dH2O). Keep the potassium ferrocyanide solution on ice. Just before use, add 10 ml of 4% osmium tetroxide (OsO4).
- Stain the tissues with 2% osmium-ferrocyanide solution for 90 min, on ice. Wash 3 times, 5 min each in dH2O.
- Prepare 1% thiocarbohydrazide (TCH) solution by dissolving 0.1 g TCH in 10 ml dH2O. Dissolve at 60 °C by swirling every 10 min until fully dissolved. Observe safety precautions while handling TCH, particularly taking care to avoid the use of metals, heating to high temperatures, or allowing the solution to dry out.
- Treat the samples with freshly prepared 1% TCH for 20 min at RT. Wash 3 times, 5 min each, in dH2O.
- Dilute 4% OsO4 to 2% with dH2O, and stain tissues with 2% aqueous osmium tetroxide by incubating for 1 hr. Wash tissues 3 times, 5 min each, with dH2O.
- Incubate the samples O/N in 1% uranyl acetate in dH2O at 4 °C.
- Prepare Walton's lead aspartate solution.
- Dissolve 0.998 g L-Aspartate in 250 ml dH2O and then add 10 N potassium hydroxide (KOH) in a dropwise fashion until the pH reaches 5.5. After pH adjustment, add 0.066 g of lead nitrate in 10 ml aspartic acid stock and heat to 60 °C for 30 min.
- Rinse tissues in dH2O and incubate with Walton's lead aspartate stain for 30 min in a 60 °C oven. Wash 3 times, 5 min each, in dH2O.
- Dehydrate samples through a graded series of alcohol using chilled solutions of 20%, 50%, 75%, 85,% and 95% ethanol for 5 min each, followed by 100% ethanol 3 times, 10 min each.
Note: Use 100% ethanol from freshly opened bottles, as opened ethanol absorbs water from the air and causes embedding to fail. Longer incubations will be necessary with larger tissue samples. - Wash samples 2 times, 15 min each, in propylene oxide.
- Make plastic embedding resin using 25 ml resin, 10.5 ml DDSA (dodecenyl succinic anhydride), 15.5 ml NMA (nadic methyl anhydride), and 1 ml DMP-30 (2,4,6-Tris dimethylaminomethyl phenol). Mix the resin by shaking. Spin and allow the resin to stand until bubbles resolve.
Note: This is the standard medium hardness recipe. Other types of electron microscopy (EM) plastic resin may be used but should be tested with a non-essential control sample in advance as not all resins work for SBFSEM. - Incubate the tissues O/N in a 50: 50 mix of embedding resin and propylene oxide in a vial that is capped initially and uncapped after 2 hr so that the propylene oxide evaporates over a period of about 8 - 10 hr.
- Transfer the tissues to 100% fresh embedding resin in clean vials for 2 hr.
- Embed samples in fresh embedding resin, in flat molds containing printed paper labels, and cured them in an oven at 60 °C for 48 hr. After about 1 hr, check the tissue placement and alignment again and adjust if necessary.
- Trim samples to the area of interest and mount them on an aluminum pin using gelling cyanoacrylate superglue or a conductive epoxy resin. Then, coat the sides of the block with colloidal silver paste to provide a conductive path to the aluminum pin.
- Examine tissue specimens using a scanning electron microscope system equipped with an in-chamber ultramicrotome stage and low kV backscattered electron detector.
Note: Obtain instrument- and site-training in scanning electron microscopy (SEM) use and become an authorized user. Alternatively, collaboration with a researcher or core facility for small projects may be possible. Radiation training may also be required as SEMs generate x-rays. - To image the samples, use the following settings: 2.25 kV, at 5 - 10 nm/pixel resolution, with field sizes between 80 - 250 µm in x,y (other field sizes possible), and slice thickness of 50 - 100 nm, with a total of 250 - 600 slices in a 16 - 20 hr time period.
Note: Settings vary substantially between different microscopes, individual samples, and desired resolution. These settings should produce images that are readily interpretable for many samples.
2. Analyzing the Imaging Dataset
Note: The Image J/Fiji software is used to analyze the dataset and relies upon the TrakEM2 plugin. Preprocessing steps may be performed using a variety of software and may be extensive or minor depending on experience level and the stacks obtained. The main transformations using the open-source software (ImageJ ver 1.50b, FIJI download Oct 1, 2015) are described here.
- Convert images to 8-bit tiff format from the original proprietary 16-bit images by opening the software and selecting menu items Image→Type→8 bit.
- If automatic contrast/brightness conversion during this step is not ideal for images, reopen the 16-bit images, and press Image→Adjust→Brightness/Contrast. Select a range that works for all images, and press Apply. Then perform conversion. Note: On some SEMs, this step may require microscope manufacturer software.
- If required, due to unacceptable image movement between slices (e.g. drift due to charging), register/align the image stacks (menu items Plugins→Registration→Linear StackAlignmentWithSIFT). In most registration software, set for "translation-only" mode rather than "rigid body".
Note: Many approaches and software may work: the SIFT registration plug-in works for many applications and there is a virtual stack version.- If required, enlarge the canvas size prior to registration (Image→Adjust→CanvasSize) or reduce it to an area of interest (Image→Crop).
Note: Some drift or splaying may occur, and trying other plugins or software may produce better results. Manual options are also available (e.g. ImageJ/FIJI, Plugins→Registration→ManualLandmarkSelection).
- If desired, scale images to a smaller, more manageable size (e.g. 25%) using ImageJ (Image→Scale).