Serial Block-Face Scanning Electron Microscopy (SBF-SEM) of Biological Tissue Samples

Summary

This protocol outlines a routine method for using serial block-face scanning electron microscopy (SBF-SEM), a powerful 3D imaging technique. Successful application of SBF-SEM hinges on proper fixation and tissue staining techniques, as well as careful consideration of imaging settings. This protocol contains practical considerations for the entirety of this process.

Abstract

Serial block-face scanning electron microscopy (SBF-SEM) allows for the collection of hundreds to thousands of serially-registered ultrastructural images, offering an unprecedented three-dimensional view of tissue microanatomy. While SBF-SEM has seen an exponential increase in use in recent years, technical aspects such as proper tissue preparation and imaging parameters are paramount for the success of this imaging modality. This imaging system benefits from the automated nature of the device, allowing one to leave the microscope unattended during the imaging process, with the automated collection of hundreds of images possible in a single day. However, without appropriate tissue preparation cellular ultrastructure can be altered in such a way that incorrect or misleading conclusions might be drawn. Additionally, images are generated by scanning the block-face of a resin-embedded biological sample and this often presents challenges and considerations that must be addressed. The accumulation of electrons within the block during imaging, known as "tissue charging," can lead to a loss of contrast and an inability to appreciate cellular structure. Moreover, while increasing electron beam intensity/voltage or decreasing beam-scanning speed can increase image resolution, this can also have the unfortunate side effect of damaging the resin block and distorting subsequent images in the imaging series. Here we present a routine protocol for the preparation of biological tissue samples that preserves cellular ultrastructure and diminishes tissue charging. We also provide imaging considerations for the rapid acquisition of high-quality serial-images with minimal damage to the tissue block.

Introduction

Serial block face scanning electron microscopy (SBF-SEM) was first described by Leighton in 1981 where he fashioned a scanning electron microscope augmented with an in-built microtome which could cut and image thin sections of tissue embedded in resin. Unfortunately, technical limitations restricted its use to conductive samples, as non-conductive samples such as biological tissue accumulated unacceptable levels of charging (electron buildup within the tissue sample)¹. While coating the block-face between cuts with evaporated carbon reduced tissue charging, this greatly increased imaging acquisition time and image storage remained a problem as computer technology at the time was insufficient to manage the large file sizes created by the device. This methodology was revisited by Denk and Horstmann in 2004 using a SBF-SEM equipped with a variable pressure chamber². This allowed for the introduction of water vapor to the imaging chamber which reduces charging within the sample, making imaging of non-conductive samples viable albeit with a loss of image resolution. Further improvements in tissue preparation and imaging methods now allow for imaging using high vacuum, and SBF-SEM imaging no longer relies on water vapor to dissipate charging^{3,4,5,6,7,8,9}. While SBF-SEM has seen an exponential increase in use in recent years, technical aspects such as proper tissue preparation and imaging parameters are paramount for the success of this imaging modality.

SBF-SEM allows for the automated collection of thousands of serially-registered electron microscopy images, with planar resolution as small as 3-5 nm^10,11. Tissue, impregnated with heavy metals and embedded in resin, is placed within a scanning electron microscope (SEM) containing an ultramicrotome fitted with a diamond knife. A flat surface is cut with the diamond knife, the knife is retracted, and the surface of the block is scanned in a raster pattern with an electron beam to create an image of tissue ultrastructure. The block is then raised a specified amount (e.g., 100 nm) in the z-axis, known as a "z-step," and a new surface is cut before the process is repeated. In this way a 3-dimensional (3D) block of images is produced as the tissue is cut away. This imaging system further benefits from the automated nature of the device, allowing one to leave the microscope unattended during the imaging process, with the automated collection of hundreds of images possible in a single day.

While SBF-SEM imaging primarily uses backscattered electrons to form an image of the block-face, secondary electrons are generated during the imaging process¹². Secondary electrons can accumulate, alongside backscattered and primary-beam electrons that do not escape the block, and produce "tissue charging," which can lead to a localized electrostatic field at the block-face. This electron accumulation can distort the image or cause electrons to be ejected from the block and contribute to the signal collected by the backscatter detector, decreasing the signal-to-noise ratio¹³. While the level of tissue charging can be decreased by reducing the electron beam voltage or intensity, or reducing beam dwell time, this results in a diminished signal-to-noise ratio¹⁴. When an electron beam of lower voltage or intensity is used, or the beam is only allowed to dwell within each pixel space for a shorter period of time, less backscattered electrons are ejected from the tissue and captured by the electron detector resulting in a weaker signal. Denk and Horstmann dealt with this problem by introducing water vapor into the chamber, thereby reducing charge in the chamber and on the block face at the cost of image resolution. With a chamber pressure of 10-100 Pa, a portion of the electron beam is scattered contributing to image noise and a loss of resolution, however this also produces ions in the specimen chamber which neutralizes charge within the sample block². More recent methods for neutralizing charge within the sample block use focal gas injection of nitrogen over the block-face during imaging, or introducing negative voltage to the SBF-SEM stage to decrease probe-beam-lading energy and increase signal collected^6,7,15. Rather than introducing stage bias, chamber pressure or localized nitrogen injection to decrease charge buildup on the block surface, it is also possible to increase the conductivity of the resin by introducing carbon to the resin mix allowing for more aggressive imaging settings¹⁶. The following general protocol is an adaptation of the Deerinck et al. protocol published in 2010 and covers modifications to tissue preparation and imaging methodologies we found useful for minimizing tissue charging while maintaining high resolution image acquisition^3,17,18,19. While the previously mentioned protocol focused on tissue processing and heavy metal impregnation, this protocol provides insight into the imaging, data analysis, and reconstruction workflow inherent to SBF-SEM studies. In our laboratory, this protocol has been successfully and reproducibly applied to a wide variety of tissues including cornea and anterior segment structures, eyelid, lacrimal and harderian gland, retina and optic nerve, heart, lung and airway, kidney, liver, cremaster muscle, and cerebral cortex/medulla, and in a variety of species including mouse, rat, rabbit, guinea pig, fish, monolayer and stratified cell cultures, pig, non-human primate, as well as human^20,21,22,23. While small changes may be worthwhile for specific tissues and applications, this general protocol has proven highly reproducible and useful in the context of our core imaging facility.

Protocol

All animals were handled according to the guidelines described in the Association for Research in Vision and Ophthalmology Statement for the Use of Animals in Vision and Ophthalmic Research and the University of Houston College of Optometry animal handling guidelines. All animal procedures were approved by the institutions in which they were handled: Mouse, rat, rabbit, guinea pig, and non-human primate procedures were approved by the University of Houston Animal Care and Use Committee, zebrafish procedures were approved by the DePauw University Animal Care and Use Committee, and pig procedures were approved by the Baylor College of Medicine Animal Care and Use Committee. All human tissue was handled in accordance with the Declaration of Helsinki regarding research on human tissue and appropriate institutional review board approval was obtained.

1. Tissue Processing

1. Prepare a stock solution of 0.4 M sodium cacodylate buffer by mixing sodium cacodylate powder in ddH₂O. Thoroughly mix buffer and pH adjust the solution to 7.3. This buffer is used to make fixative (composition described below in step 1.3), washing buffer, as well as osmium and potassium ferrocyanide solutions.
   NOTE: Perfusion fixation is often the best method of fixation for SBF-SEM studies, as fixation occurs rapidly and throughout the body. If perfusion fixation is not possible within your study design, skip to step 1.3.
2. Perform perfusion fixation with the appropriate physiological pressure for the animal model^24,25,26. This is done via transcardial sequential perfusion with heparinized saline followed by fixative, each placed at a specific height (e.g., 100 cm) above the organism (appropriate to the physiological pressure of the vascular system in the animal model), with fixative flowing into the left ventricle, and exiting out of an incision made in the right atrium. Tissue of interest will become pale as blood is replaced with fixative, if all or a portion of your tissue does not blanche then tissue may not be appropriately fixed and ultrastructure may not be preserved.
3. Use a razor blade or sharp scalpel to trim tissue samples into blocks no larger than 2 mm x 2 mm x 2 mm. If step 1.2 was skipped, do this swiftly so that tissue can be immersion fixed as quickly as possible.
   1. Alternatively, dissect tissue under fixative and transfer to fresh fixative to complete the immersion process. The final composition of the fixative consists of 0.1 M sodium cacodylate buffer containing 2.5% glutaraldehyde and 2 mM calcium chloride. Allow fixation to proceed for a minimum of 2 hours at room temperature and a maximum of overnight at 4 °C. If possible, use a rocker/tilt plate to gently agitate samples while fixing.
   2. Alternatively, if an inverter microwave is available, fix tissue in aforementioned fixative under vacuum at 150 watts for 4 cycles of 1 minute on, 1 minute off. Microwave fixation is the preferred method for step 1.3 as it rapidly fixes tissue and preserves tissue ultrastructure²⁷.
      NOTE: Tissue should never be allowed to dry during this protocol, care should be taken to transfer tissue quickly from one solution to the next.
4. Wash fixed tissue 5x for 3 minutes each (15 minutes total) at room temperature in 0.1 M sodium cacodylate buffer containing 2 mM calcium chloride.
5. Make the following osmium ferrocyanide solution fresh, preferably during the previous wash steps. Combine a 4% osmium tetroxide solution (prepared in ddH₂O) with an equal volume of 3% potassium ferrocyanide in 0.2 M cacodylate buffer with 4 mM calcium chloride. After the previous wash step, place the tissue in this solution for 1 hour on ice in the dark, and in the fume hood.
   NOTE: Osmium tetroxide is a yellow crystalline substance that comes in an ampule. To create the osmium tetroxide solution, crack open the ampule, add ddH₂O, and sonicate for 3-4 hours in the dark until the crystals are completely dissolved. Osmium tetroxide solution is a clear yellow solution, if the solution is black the osmium has been reduced and should no longer be used.
6. While the tissue is incubating in the osmium ferrocyanide solution, begin preparing the thiocarbohydrazide (TCH) solution. Prepare this solution fresh and have it readily available at the end of the 1 hour osmium ferrocyanide fixation period. Combine 0.1 g of thiocarbohydrazide with 10 mL of ddH₂O and place this solution into a 60 °C oven for 1 hour. To ensure the solution is dissolved, gently swirl every 10 minutes. Prior to use, filter this solution through a 0.22 µm syringe filter.
7. Prior to incubating in TCH, wash the tissue with room temperature ddH₂O 5x for 3 minutes each (15 minutes total).
8. Place the tissue in the filtered TCH solution for a total of 20 minutes at room temperature (Figure 1A-C).
9. Following incubation in TCH, wash the tissue 5x for 3 minutes each (15 minutes total) in room temperature ddH₂O.
10. Place tissue in ddH₂O containing 2% osmium tetroxide (not osmium reduced with potassium ferrocyanide) for 30 minutes at room temperature. This should be done in the fume hood and in the dark as osmium can be reduced by light (e.g., under aluminum foil) (Figure 1D-F).
11. Following osmium tetroxide incubation, wash tissue 5x for 3 minutes each (15 minutes total) in room temperature ddH₂O.
12. Place tissue in 1% aqueous uranyl acetate (uranyl acetate powder mixed in ddH₂O) overnight in a refrigerator at 4 °C.
13. Just before removing tissue from the refrigerator, prepare fresh Walton's lead aspartate solution. Begin by dissolving 0.066 g of lead nitrate in 10 mL of 0.03 M aspartic acid solution (0.04 g aspartic acid in 10 mL of distilled water) and adjust pH to 5.5 with 1 N KOH (0.5611 g in 10 mL of distilled water).
    CAUTION: A precipitate can form when adjusting the pH. This is not acceptable.
    1. Using a stir bar, slowly add the 1 N KOH dropwise while monitoring pH. Pre-heat the finished clear lead aspartate solution in a 60 °C oven for 30 minutes. If a precipitate forms the solution cannot be used and another solution must be prepared.
14. Remove the tissue from the refrigerator and wash 5x for 3 minutes each (15 minutes total) in room temperature ddH₂O.
15. After washing, place the tissue in the warmed Walton's lead aspartate solution for 30 minutes while maintaining the temperature at 60 °C.
16. After incubation in Walton's lead aspartate, wash the tissue 5x for 3 minutes each (15 minutes total) in room temperature ddH₂O (Figure 1G-I).
17. Dehydrate the tissue through an ice-cold acetone series (30%, 50%, 70%, 95%, 95%, 100%, 100%, and 100% acetone (in ddH₂O where applicable) allowing 10 minutes for each step in the series.
18. Following the ice-cold dehydration series, place tissue in room temperature acetone for 10 minutes.
    1. During this time, formulate Embed 812 ACM resin. Use the "hard mix" recipe as it is more resistant to beam damage. Mix the resin thoroughly, and place the tissue into Embed 812:acetone (1:3 mix) for 4 hours, followed by Embed 812:acetone (1:1 mix) for 8 hours or overnight, and finally Embed 812:acetone (3:1 mix) overnight. Perform these resin-infiltration steps at room temperature.
19. The next day, place the tissue in 100% Embed 812 for 4-8 hours, then in fresh 100% Embed 812 overnight, and finally into fresh 100% Embed 812 for 4 hours. Perform these resin-infiltration steps at room temperature.
    1. Just before embedding, place a small amount of resin into a mixing container and slowly mix (a wooden stick can be used for stirring) in carbon black powder until the resin is saturated with the powder but is still fluid and does not become grainy. It should resemble thick ink and be able to slowly drip from the wooden stick without visible clumps.
20. Orient the tissue samples in a silicone rubber mold and take a picture so that sample orientation within the resin block is recorded and can be referenced. Cover the samples in carbon black saturated resin at the tip of the silicone mold and place the mold in an oven for ~1 hour at 65 °C.
    1. Place the mold at an incline to contain the resin at the tip of the mold where it covers the tissue sample. Place a label with an experiment/tissue sample identifier in the mold at the opposite end of the resin (Figure 2A).
21. Remove the silicone mold from the oven and fill the remainder of the mold with clear resin (no carbon black) making sure that the label remains visible. Cure the resin infused with carbon black enough as to not readily mix with the clear resin.
    1. Prepare an extra well within the mold that does not contain tissue. Beginning with the extra well, fill the remainder of the mold with clear resin.
    2. If the carbon black infused resin begins to bleed into the clear resin, place the silicone mold back into the oven for additional time (e.g., 15 minutes).
    3. Once all of the tissue samples have been topped off with clear resin, place the silicone mold back into the oven (flat, no incline) at 65 °C for 48 hours to complete the curing process.

2. Block Preparation

NOTE: The method will depend on how the sample is oriented in the block and how the sectioning is to take place. However, the most common tissue orientation finds the tissue centered in the tip of the resin block, perpendicular to the long end of the resin block.

1. In most cases, first trim the end of the block to locate the tissue by placing the specimen block in the microtome chuck with the tapered end sticking up approximately 5-6 mm out of the chuck. Lock it in place with the set screw and place it under a heat lamp.
2. After several minutes the block will be malleable and easy to trim. Place the chuck in the stereomicroscope holder and use a new double-edge razor blade to make thin sections parallel with the block face until the tissue is visible. This is best seen by angling light across the block face, the tissue sample will be less reflective and granular compared to those portions of the resin that are devoid of tissue. Consult the photograph taken of tissue samples prior to introduction of carbon black saturated resin for an idea of how and where the tissue is located.
3. Set one specimen pin holder aside for trimming purposes. This pin holder is never placed into the SEM chamber and can therefore be handled without gloves, this will be referred to as the trimming pin holder. Any specimen holder destined to be placed into the imaging chamber should never be touched without gloves. This avoids introducing grease and oil into the microscope chamber.
4. Place an aluminum specimen pin in the trimming pin holder and slightly tighten the set screw with the face (flat surface) of the pin held 3-4mm above the pin holder.
5. Make several deep, crisscrossing scratches in the face of the pin to provide a larger surface area for the glue used to hold the specimen in place. If an aluminum pin is used, a small steel flathead screwdriver is recommended for this step (Figure 2B).
6. Place the chuck containing the tissue sample back under the heat lamp until the resin becomes soft and malleable, then place it into the chuck-receptacle under the stereomicroscope.
7. Using a double-edged razor blade to trim away excess resin from the portion of the resin block containing the tissue sample. Ultimately the size of tissue block attached to the pin will be approximately 3 mm in diameter and 2-3 mm in height.
   1. Carefully push the razor straight down into the resin block roughly 1-2 mm, then carefully push the razor horizontally into the resin block at a depth equal to the previous cut. Do this slowly and with great care, as it is possible to damage or cut away the portion of the block containing the tissue sample. As the two cuts meet, the excess resin will separate from the block. Continue to remove resin until only a 3 mm x 3 mm raised area remains.
8. After this initial trimming, place the block (still in the chuck) under the heat lamp for several minutes.
9. Once the resin becomes soft and malleable, place the block back under the stereomicroscope. Using a new double-edge razor blade, cut off the top of the resin block, roughly 1 mm below the trimmed portion, with a single smooth cut. A flat surface is preferable as this will be glued to the specimen pin. Be careful not to allow the sample to become lost, as this step requires some force which can transfer to the removed portion of the block and cause it to fly away. Place the cut and trimmed sample aside.
10. Place the trimming pin holder containing the cut aluminum pin in the stereomicroscope receptacle. Apply a thin layer of cyanoacrylate glue to the pin face such that it completely covers the pin without forming a visible meniscus. Pick up the trimmed piece of the tissue block with forceps and place in on the pin face. Center the tissue sample on the specimen pin. Push it down and hold it for several seconds. Allow the glue to set for several minutes.
11. When the glue is thoroughly dry, place the trimming pin holder back under the stereomicroscope. Using a fine file, file away excess resin so that no resin is overhanging the pin. The resin shape should resemble the circular pin head.
12. Locate the tissue on the raised portion of your resin block, oblique lighting is useful for this. Using a double-edge razor, the raised portion of the resin containing the tissue sample must be trimmed to an area no larger than 1 mm². If possible, the block-face can be trimmed even smaller, this will reduce stress on the diamond knife and improve its longevity.
    1. Remove as much excess resin as possible, leaving the block slightly longer in one dimension. This is done slowly and with care, as it is possible for the resin containing the tissue sample to break away if too much force is applied. While a razor is recommended, a fine metal file can be used for this step.
13. With a fine metal file angle the excess resin, in the area outside the raised portion containing the tissue sample, down towards the edge of the pin (Figure 2C).
14. Remove resin particles and dust from the prepared sample before applying silver paint followed by gold sputtering. Mix silver with acetone so that it is an easily spreadable liquid, akin to nail polish (but not so thin that it drips off of the applicator) and apply a thin coat to the entire sample block surface. Acetone evaporates rapidly, so it may be necessary to add additional acetone as the silver paint begins to thicken.
    1. Allow the silver paint to dry overnight before loading into the microscope.
       ​NOTE: This silver layer must be thin in order to avoid expanding the block-face beyond 1 mm x 1 mm, and while the silver paint has never damaged the diamond knife, smaller block-faces are still recommended to preserve the longevity of the diamond knife. The acetone mixed in with the silver must evaporate completely before gold-sputtering or loading the sample into the microscope to avoid introducing acetone vapor into the imaging chamber.
    2. Following application of silver paint, apply a thin layer of gold to the sample block. Using a standard vacuum sputtering device equipped with a standard gold foil target, a chamber pressure of 200 milliTorr (Argon gas) and 40 milliamps running for 2 minutes will result in a 20 nm thick gold coating.
15. After coating, place the mounted and trimmed block in a tube with the appropriate experiment label attached. Create custom tubes using disposable transfer pipettes.
    1. Cut the transfer pipette just below the bulb, leaving a short portion of the transfer pipette tube attached below the bulbous end. Shorten the tubular portion that was cut away, and cut the pipette tip back enough so that the aluminum specimen pin can be pushed snuggly inside of it.
    2. Place the end containing the aluminum specimen pin within the bulbous end of the modified transfer pipette.
16. Before loading a prepared tissue block, carefully trim away excess silver paint from the surface of the block face.

3. SEM Settings for Imaging the Block Face

NOTE: The imaging settings that follow were produced on the device used by the authors, which is listed in the Table of Materials provided. While this device is capable of variable pressure imaging, best results were captured under high vacuum.

1. Dwell Time: Use 12 µs/px during serial sectioning. When a region of interest has been identified, a higher resolution image can be acquired at 32 µs/px.
2. Vacuum Settings: Use a gun pressure of 9e-008 Pa, a column pressure of 1.1e-004 Pa, and a chamber pressure of 9.5e-002 Pa.
3. Capture Time: With the above settings, capture a 2048x2048 px image stack at a rate of 50 seconds per image. Higher resolution images of regions of interest can be captured at 4096x4096 px at just under 9 minutes per image.
4. Section Thickness: Use 100-200 nm. Less is possible, but may require lower beam voltage, intensity, or dwell time.
5. High Voltage (HV): Use 7-12 kV. While increasing the voltage reduces the spot size and increases resolution, it introduces more possibility for beam damage. Higher kV increases the beam penetration which results in loss of details. However, lowering the kV degrades the signal to noise ratio (Figure 3)¹⁴.
6. Beam Intensity (BI): The author's SBF-SEM device offers a beam intensity scale ranging from 1-20. On this scale, values of 5-7 give quality images without excessive charging and beam damage. The higher the BI the greater the resolution however, there is more chance of charging and beam damage¹⁴.
7. Spot Size and Image Magnification: Determine the spot size by the beam intensity and voltage level. Ideally, the spot size should not be larger than the pixel size used. The pixel size is determined by dividing the field of view (FOV) by the number of pixels. For example, a 25 µm FOV with an image size of 2048x2048 px would give 12.2 nm per pixel. Therefore spot size should be no greater than 12.2 nm. Figure 4 shows how HV, BI and spot size are related.
8. Working Distance (WD) - With block face imaging the working distance is not adjustable. It is simply a factor of focus. It will be nearly identical for all blocks imaged. While the working distance is not adjustable, it plays a critical role in the resolution of the image captured. As working distance decreases, the resolution limit on images captured increases. In some cases it may be possible to decrease the working distance by making modifications within the imaging chamber, however these modifications must be made at the user's discretion. In order to decrease the working distance and increase image resolution, we loosened the door mount microtome screws and repositioned the microtome so that it rested ~2 mm closer to the beam detector after retightening the screws.
9. Resolution - Using the above settings, x & y resolution as high as 3.8 nm is possible. It is important to note that resolution is limited by beam spot size as well as the pixel resolution of the image captures (e.g., a 20 µm field of view captured in a 2048x2048 pixel image has a pixel resolution of 9.8 nm, even if a 3.8 nm spot size was used). Image resolution in the z-plane is dependent on sectioning thickness, we find that 100-200 nm works well with this protocol.

Results

Mouse Cornea
This protocol has been applied extensively to the mouse cornea. Using SBF-SEM imaging a network of elastin-free microfibril bundles (EFMBs) were shown to be present within the adult mouse cornea. It was previously believed that this network was only present during embryonic and early postnatal development. SBF-SEM revealed an extensive EFMB network throughout the cornea, with individual fibers found to be 100-200 nm in diameter when measured in cross-section. It was also found that thi...

Discussion

The purpose of this methods paper is to highlight the tissue preparation and imaging methodology that has allowed our lab to reliably capture high-resolution serial electron microscopy images, and to point out critical steps that lead to this outcome as well as potential pitfalls that can occur when conducting SBF-SEM imaging. Success using this protocol requires proper fixation of tissue, impregnation of heavy metals into the sample, modifications of the embedding resin to reduce charging, as well as an understanding of...

Materials

Name	Company	Catalog Number	Comments
1/16 x 3/8 Aluminum Rivets	Industrial Rivet & Fastener Co.	6N37RFLAP/1100	Used as specimen pins.
2.5mm Flathead Screwdriver	Wiha Quality Tools	27225
Acetone	Electron Microscopy Sciences	RT 10000	Used to dilute silver paint.
Aspartic Acid	Sigma-Aldrich	A8949
Calcium Chloride	FisherScientific	C79-500
Conductive Silver Paint	Ted Pella	16062
Denton Desk-II Vacuum Sputtering Device equipped with standard gold foil target	Denton Vacuum	N/A	This is the gold-sputtering device used by the authors, alternates are acceptable.
Double-edged Razors	Fisher Scientific	50-949-411
Embed 812	Electron Microscopy Sciences	14120
Gatan 3View2 mounted in a Tescan Mira3 Field emission SEM	Gatan & Tescan	N/A	This is the SBF-SEM device used by the authors, alternates are acceptable.
Glass Shell Vials, 0.5 DRAM (1.8 ml)	Electron Microscopy Sciences	72630-05
Gluteraldehyde	Electron Microscopy Sciences	16320
Gorilla Super Glue - Impact Tough	NA	NA	Refered to as cyanoacrylate glue in text.
Ketjen Black	HM Royal	EC-600JD	Refered to as carbon black in text.
KOH	FisherScientific	18-605-593
Lead Nitrate	Fisher Scientific	L62-100
Microwave	Pelco	BioWave Pro	This is the microwave used by the authors, alternates are acceptable.
Osmium Tetroxide	Sigma-Aldrich	201030
Potassium Ferrocyanide	Sigma-Aldrich	P9387
Silicone Embedding Mold	Ted Pella	10504
Sodium Cacodylate Trihydrate	Electron Microscopy Sciences	12300
Samco Transfer Pipette	ThermoFisher Scientific	202	Used to make specimen pin storage tubes.
Swiss Pattern Needle Files	Electron Microscopy Sciences	62115
Thiocarbohydrazide	Sigma-Aldrich	223220
Uranyl Acetate	Polysciences, Inc.	21447-25
Reconstruction Software
Amira Software	Thermo Scientific	N/A	Used to create the reconstructions found in figures 5-7 and 9.
Fiji (Fiji is Just ImageJ)	ImageJ.net	N/A	TrakEM2 can be added to Fiji to asist in manual segmentation.
Microscopy Image Browser (MIB)	University of Helsinki, Institute of Biotechnology	N/A
Reconstuct Software	Neural Systems Lab	N/A
SuRVoS Workbench	Diamond Light Source & The University of Nottingham	N/A
SyGlass	IstoVisio, Inc.	N/A	Allows for reconstruction in virtual reality and histogram-based reconstruction methods.