High-Throughput, Multi-Image Cryohistology of Mineralized Tissues

Summary

In this manuscript, we present a high-throughput, semi-automated cryohistology platform to produce aligned composite images of multiple response measures from several rounds of fluorescent imaging on frozen sections of mineralized tissues.

Abstract

There is an increasing need for efficient phenotyping and histopathology of a variety of tissues. This phenotyping need is evident with the ambitious projects to disrupt every gene in the mouse genome. The research community needs rapid and inexpensive means to phenotype tissues via histology. Histological analyses of skeletal tissues are often time consuming and semi-quantitative at best, regularly requiring subjective interpretation of slides from trained individuals. Here, we present a cryohistological paradigm for efficient and inexpensive phenotyping of mineralized tissues. First, we present a novel method of tape-stabilized cryosectioning that preserves the morphology of mineralized tissues. These sections are then adhered rigidly to glass slides and imaged repeatedly over several rounds of staining. The resultant images are then aligned either manually or via computer software to yield composite stacks of several layered images. The protocol allows for co-localization of numerous molecular signals to specific cells within a given section. In addition, these fluorescent signals can be quantified objectively via computer software. This protocol overcomes many of the shortcomings associated with histology of mineralized tissues and can serve as a platform for high-throughput, high-content phenotyping of musculoskeletal tissues moving forward.

Introduction

Biological research often requires efficient phenotyping, which is frequently associated with some sort of histological analysis^1-3. This need is even more evident with the ambitious projects to disrupt each gene in the mouse genome⁴. These histological analyses can range from assessing cell morphology and/or anatomical features to mapping expression of specific genes or proteins to individual cells. In fact, one of the fundamental contributions of histology to the field of genomics is the ability to associate a specific molecular signal to a specific region or cell type.

Traditional methods of histology, especially for musculoskeletal tissues, are often time consuming and laborious, requiring sometimes weeks to fix, decalcify, section, stain, and image the specimen then analyze the images via human interpretation. Analyzing multiple molecular signals, whether via immunohistochemistry, in situ hybridization, or special stains, requires multiple sections and even multiple specimens to perform appropriately. In addition, these multiple responses cannot be co-localized to the same cell and sometimes cannot be co-localized to a specific region within a given specimen. As the genomics and epigenomics field moves into the digital age, the histological field must also follow suit to provide efficient, high-throughput, and automated analysis of a variety of molecular signals within a single histological section.

Indeed, there is a demand for improved histological techniques that can associate multiple molecular signals to specific cells within a given specimen. Recently, we have published a new high-throughput cryohistological method for assessing several response measures within a given section from mineralized tissue^5-14. The process involves stabilizing the cryosection with frozen cryotape, adhering the taped section rigidly to a microscope slide, and conducting several rounds of staining and imaging on each section. These rounds of images are then aligned manually or via computer automation prior to image analysis (Figure 1). Here, we present detailed protocols of this process and provide examples where these techniques have improved our understanding of different biological processes.

Protocol

The University of Connecticut Health Center institutional animal care and use committee approved all animal procedures.

1. Fixation and Embedding

1. Euthanize the animal via CO₂ asphyxiation or other approved methods.
2. Harvest the tissue of interest (e.g., limb, vertebrae, etc.) and place in 10% neutral buffered formalin at 4 °C until properly fixed. Take special care to maintain consistent anatomical placement prior to fixation. For instance, fix limbs with joints at consistent flexion, internal rotation, and external rotation angles¹¹. Typically, fix whole mouse limbs for 1 - 3 days.
   Caution: Formalin is toxic and should be handled in a fume hood while wearing appropriate personal protective equipment.
   Note: The timeframe of fixation depends on the size of the specimen. If the experiment allows, cutting open the bone will improve fixation of the marrow compartment. Some specimens may require perfusion fixation¹⁵ to minimize fluorescent background (e.g., faint fluorescent signals within bone marrow).
3. Transfer specimens from formalin to 30% sucrose made in 1x PBS and incubate for 12 - 24 hr at 4 °C.
   Note: Once incubated for 12 - 24 hr, specimens can then be placed at -80 °C for long-term storage. This method of storage is recommended for experiments in which researchers intend to embed multiple samples in the same block but may not harvest all of these samples at the same time (e.g., experiments with multiple time points).
4. Remove specimens from the sucrose and dissect away any excess tissue.
5. Fill a cryomold (size of mold is dependent on specimen size) part of the way with cryo embedding medium. Place sample in mold such that cutting plane for the region of interest is parallel with the bottom of the mold.
   Note: An example of specific tissue placement and orientation can be found in Figure 2A - B.
6. Place the cryomold onto a piece of dry ice until a thin layer of cryo embedding medium freezes and subsequently fixes the specimen in place. Fill the remaining volume of the cryomold with cryo embedding medium while maintaining the cryomold on the dry ice pellet.
7. Place the cryomold in a container containing 2-methyl-butane pre-chilled by dry ice. Once specimens are completely frozen, remove cryomolds and shake off excess 2-methyl-butane. Wrap cryomolds in cellophane and place in a -20 °C or -80 °C freezer for storage.
   Note: Samples can dry out over time when stored in cryo embedding medium, especially at -20 °C. We recommend storage of samples for longer than 1 - 2 months in cryo embedding medium at -80 °C or in 30% sucrose at -80 °C (see step 1.3).

2. Tape-stabilized Tissue Sectioning

1. Remove specimen blocks from freezer and place in cryostat with the temperature set between -20 and -25 °C.
2. Remove the block from the cryomold and trim away excess cryo embedding medium with a razor blade.
3. Place some cryo embedding medium onto the specimen disc and then align the block such that the surface of the specimen disc is parallel to the bottom surface of the block thereby establishing the proper cutting plane for the region of interest.
4. Place the specimen disc in the cryostat and allow for the cryo embedding medium to freeze.
5. Place the specimen disc onto the specimen head and adjust the head such that the blade cuts parallel to the surface of the specimen block.
6. Trim the block down to a level within the region of interest and brush off any shavings from the surface of the block.
7. Cut a piece of the cryotape large enough to cover the region of interest and pre-chill in the cryostat.
   Note: Multiple pieces can be cut of consistent sizes and stored within the cryostat during sectioning.
8. Remove non-adherent backing from the cryotape by grasping the tape by the non-sticky silver/gold tabs using forceps then place the tape onto the block sticky-side down.
9. Apply pressure to the cryotape using the roller (Figure 2C).
10. Make a section (5 - 8 μm) using either the automatic motor or manual cutting wheel of the cryostat.
    Note: It is good practice to hold the edge of the cryotape as it comes onto the stage such that the section does not fall off the stage during sectioning (Figure 2D).
11. Place the section tissue side up on a plastic microscope slide within the cryostat.
12. Remove the plastic slide from the cryostat and allow the cryo embedding medium to melt such that the section adheres to the surface of the slide.
13. Repeat sectioning for serial sections or other regions of interest.
14. Once finished, store the sections in a slide box at 4, -20 or -80 °C depending on the length of storage required and the downstream response measures. For instance, use slides that are going to be stained for enzymatic activity (see 5.2 - 5.3) within a month if stored at 4 °C.

3. Adhesion of Sections to Microscope Slides

1. UV-curable adhesive method.
   1. Label the microscope slide.
      Note: For increased automation, samples and slides can be labeled with barcodes.
   2. Apply a drop of UV-activated optical adhesive to two glass slides and use the edge of a plastic slide to spread a thin layer of adhesive across the surface of the glass slides.
   3. Cut off the silver/gold tab and lay the taped section tissue side up on the adhesive of the first slide. Apply the section in a rolling motion from one edge to the other in order to minimize the formation of entrapped bubbles underneath the tape.
      NOTE: The first slide is used to pre-wet the underside of the tape prior to placing it on the second (permanent) slide (step 3.1.4).
   4. Remove the taped section from the first slide and place it on the adhesive layer of the second slide (Figure 3A). Again, take care to avoid entrapment of bubbles.
      NOTE: This step takes practice to master.
   5. Once the appropriate number of sections for the given experiment is placed on each slide, wipe off the excess adhesive from the surrounding areas.
   6. Inspect each section closely for bubbles or fibers underneath the tape. Perform this inspection under a microscope or magnifying glass. If there are obstructions, remove the individual section, remove the obstructions, and then reapply it to the glass slide.
   7. Once it is determined that there are no obstructions under the taped sections, place the microscope slides under the UV black light for 5 - 10 min to crosslink the adhesive.
2. Chitosan adhesive method.
   1. Prepare the 1% chitosan adhesive. Prepare acetic acid solution by dissolving 0.25 ml of concentrated acetic acid in 100 ml DI water (0.25% v/v). Dissolve 1 g of chitosan powder in 100 ml of acetic acid solution and stir the solution O/N or until all the chitosan powder is dissolved.
      NOTE: The solution is stable at RT.
   2. Deposit a drop of chitosan solution for each section that will be placed on the slide.
   3. Cut off the silver/gold tab of the tape and place each taped section tissue side up onto the adhesive (Figure 3A). Use great care to avoid entrapment of bubbles.
   4. Once all sections are placed on the slide, tilt the slide up on one of its long edges and drag excessive chitosan to the bottom edge using forceps.
   5. Place slides in this orientation on top of a paper towel in a slide box. Gravity will then cause the excess chitosan to fall down onto the towel, yielding a flat and uniform layer of chitosan.
   6. Place the slide box with its lid propped open in the refrigerator O/N to allow the chitosan to dry.
      NOTE: See Table 1 for a comparison between the two adhesive methods.

4. Application of Reference Markers to Slides

1. Prepare reference marker solution by dissolving 50 μl of green and 50 μl of red microspheres in 100 μl of water. Store the solution in the dark at 4 °C.
2. Place a 10 μl or 20 μl pipet tip into the microsphere solution. Add a small drop of the microsphere solution to each section adjacent to the region of interest (Figure 3C). Be careful to not get the microspheres within the region of interest.
3. Dry the slides at RT (protected from light) for 30 min if processing the slides on that day. If not, place the slides in a slide box at 4 °C.
   Note: As long as the microsphere solution dries prior to hydrating the slides, the microspheres will remain adhered to the slides during multiple rounds of staining/imaging.

5. Multiple Rounds of Staining

NOTE: By choosing a compatible sequence of imaging, staining, and reimaging steps, it is possible to detect and co-localize many biological signals on the same tissue section. Each round of imaging/staining/reimaging has to be developed for the particular histological question. The imaging/staining/reimaging sequence typically involves acquiring the endogenous fluorescent signals (e.g., cellular GFP, mineralization dyes, in vivo imaging probes) on the first round of imaging followed by fluorescent multiplexed immunostaining and multiple rounds of enzymatic activity stains. Lastly, the section can be stained using chromogenic dyes (e.g., H&E, toluidine blue, safranin O, etc.) to highlight the tissue architecture. Presented in this section are custom methods adapted from commercially available protocols.

1. Calcein blue staining for accumulated mineral
   NOTE: Calcein blue staining yields a consistent mineral surface that is amenable to automated image analysis unlike darkfield or differential interference contrast (DIC) imaging. The problem with calcein blue staining is that the fluorescent spectrum overlaps with tetracycline and demeclocycline labeling. Therefore, if the sample contains these mineral labels, image the labels in the first round of imaging prior to staining with calcein blue.
   1. If the endogenous signals within the tissue are imaged prior to this staining step, submerge the slides from the previous imaging round in a coplin jar filled with 1xPBS until the cover slips fall away from the slides. Remove and dry the cover slips.
   2. Prepare 30 mg/ml of calcein blue solution in 2% NaHCO₃ (made by dissolving 2 g NaHCO₃ in 100 ml H₂O and adjusting to pH of 7.4 with HCl). Aliquot and store the solution at -20°C.
   3. Immerse the slides in a coplin jar containing 1x PBS for 15 min to hydrate the sections, if not already hydrated from the previous round.
   4. Remove the slides and dry the back and edges with a paper towel.
   5. Apply 100 - 500 μl of calcein blue solution per slide and incubate for 10 min in a humidity chamber at RT.
   6. Rinse the slides in 1x PBS for 10 min with 3 changes.
   7. Apply ~ 200 μl of 50% glycerol in 1x PBS to each slide and mount the cover slip.
   8. Remove excess glycerol and dry the bottom of the slides.
2. Tartrate-resistant acid phosphatase (TRAP) enzymatic activity staining
   NOTE: TRAP is expressed by multiple cell types in the hematopoietic lineage including osteoclasts. The TRAP staining presented here uses a yellow fluorescent acid phosphatase substrate. Certain fluorescent signals will overlap with the custom yellow filter set including tetracycline/demeclocycline mineralization labels, calcein blue stain, and DAPI counterstain.
   1. Submerge slides from the previous imaging round in a coplin jar filled with 1x PBS until the cover slips fall away from the slides. Remove and dry the cover slips.
   2. Prepare Buffer 1 by dissolving 9.2 g of sodium acetate anhydrous and 11.4 g of sodium tartrate dibasic dihydrate in water and bringing the final volume to 1,000 ml. Adjust the pH to 4.2 with concentrated glacial acetic acid (~ 1.5 - 2 ml). Store the solution at 4 °C for up to 12 months.
   3. Prepare Buffer 2 by dissolving 40 mg of sodium nitrite in water and bringing the final volume to 1 ml. Store the solution at 4 °C for up to 1 week.
   4. On the day of staining, prepare the reaction buffer by mixing 7.5 ml of buffer 1 and 150 µl of buffer 2.
   5. Apply the reaction buffer without the substrate to the slides for 10 - 15 min to equilibrate the tissue at the lower pH.
      NOTE: The typical volume is ~ 200 μl but needs to be large enough to cover all sections.
   6. Dilute the yellow fluorescent acid phosphatase substrate between 1:50 and 1:100 in the reaction buffer.
      NOTE: Dilution will be dictated by TRAP activity within the sample.
   7. Pour the reaction buffer off of the slides and apply the reaction buffer with the yellow fluorescent substrate to the slides.
   8. Place the slides under the UV black light for ~ 5 min to activate the substrate.
      NOTE: Incubation time may vary depending on TRAP activity within the sample.
   9. Rinse the slides in 1x PBS for 10 min with 3 changes.
   10. Apply ~ 200 μl of 50% glycerol in 1x PBS to each slide and mount the cover slip.
   11. Remove excess glycerol and dry the bottom of the slides.
       NOTE: The acidic TRAP buffer will decalcify the sections. The added benefit of the decalcification is that certain colors will be available again for downstream staining (e.g., mineralization labels). For instance, the calcein blue staining will be removed during the TRAP staining. Therefore, DAPI counterstain can be imaged in this channel during a subsequent round.
3. Alkaline phosphatase (AP) enzymatic activity staining
   NOTE: Multiple cell types involved with mineralization including osteoblasts and mineralizing chondrocytes express AP. The Fast Red substrate used in this protocol elicits both a fluorescent red and chromogenic red signal. Because of this, the chromogenic substrate will quench fluorescent signals in the regions where the substrate precipitates. Therefore, it is best to do this stain after imaging the fluorescent signals that may be quenched by the AP substrate.
   1. Submerge slides from the previous imaging round in a coplin jar filled with 1x PBS until the cover slips fall away from the slides. Remove and dry the cover slips.
   2. Prepare 1 M Tris by dissolving 12.1 g of Tris in 100 ml of water. Adjust pH to 9.5 with 1 M NaOH.
   3. Prepare 1 M MgCl₂ hexahydrate by dissolving 20.33 g of MgCl₂ in 100 ml of deionized water.
   4. Prepare 2 M NaCl by dissolving 11.68 g NaCl in 100 ml of deionized water.
   5. Prepare 100x (20 mg/ml) stock solution of Fast Red TR Salt by dissolving 1 g powder in 50 ml of deionized water. Make 100 μl aliquots and store at -20°C.
   6. Prepare 100x (10 mg/ml) Naphthol AS-MX phosphate stock solution by dissolving 500 mg powder in 50 ml of N,N dimethylformamide. Make 100 μl aliquots and store at -20 °C.
   7. On the day of staining, prepare the AP buffer by adding 1 ml of 1 M Tris, 0.5 ml of 1 M MgCl₂, 0.5 ml of 2 M NaCl and bring the final volume to 10 ml using deionized water (final concentrations: 100 mM Tris, 50 mM MgCl₂, 100 mM NaCl).
   8. Place AP buffer on each slide and incubate in a humidity chamber for 10 min at RT.
   9. Prepare AP substrate buffer by diluting 100x stocks of Fast Red TR Salt and Naphthol AS-MX to 1x in the AP buffer.
   10. Pour AP buffer off of slides and replace with AP substrate buffer. Incubate slides for 5 - 10 min in a humidity chamber at RT.
       NOTE: Incubation time will be dictated by AP activity of the sample.
   11. Wash slides 3x in 1x PBS for 5 min each.
   12. Mount cover slips with DAPI counterstaining solution (1:1,000 dilution of DAPI in 50% glycerol in 1x PBS).
4. Toluidine blue O (TB) chromogenic staining
   NOTE: Because all previous steps are imaged under temporary aqueous mounting in 50% glycerol/PBS solution, the chromogenic step is also imaged under aqueous conditions. However, the staining solution may tend to leach out over time. Therefore, it is suggested to image the toluidine blue as soon as possible after staining.
   1. Submerge slides from previous imaging round in a coplin jar filled with deionized water until the cover slips fall away from the slides. Remove and dry the cover slips.
   2. After removing the cover slips, replace with fresh water and incubate the slides for at least 10 min so that salts from PBS are sufficiently removed from the tissue.
   3. Prepare 0.025% TB solution in deionized water.
   4. Place the slides in the TB solution for 1 - 5 min.
      NOTE: Incubation time is dependent on user preference but should be kept consistent across all samples.
   5. Place slides into coplin jar with deionized water and wash slides 3x for 5 min each.
   6. Mount cover slip with 30% glycerol dissolved in deionized water (NOT 1x PBS as this will cause the stain to leach out from the tissue). NOTE: Glycerol mounting medium can be substituted with fructose syrup, which helps prevent diffusion of the stain from the tissue. To prepare the fructose syrup, dissolve 30 g of fructose in 10 ml of deionized water and heat to 60 °C until dissolved.

6. Multiple Rounds of Imaging

1. Load the slides into the microscope trays (Figure 3D).
   Note: The trays are spring loaded.
2. Insert the trays into the tray stack of the slide-scanning microscope (Figure 3E).
3. Click on the profile list to load a profile for each slide with appropriate exposure times for each fluorophore. Click on the slide name to name each slide manually or have the software read the barcode provided on the slide label. Click the start preview scan button to take a preview image of the slide.
4. Set the regions of interest in the tissue detection wizard and save them for subsequent rounds of imaging. Once each slide is setup, start the scan process by clicking the start scan button.
   Note: The system can hold up to 100 slides at a time.
5. Once the imaging has finished, export the images from each individual channel and imaging round as .tif or .jpg files for image assembly and analysis.

7. Image Assembly

1. Manual method using Photoshop (or similar image editing software capable of producing layered images).
   1. Open each individual channel image from each imaging round.
   2. Assemble the individual images as layers within one composite image. To do so, click on the layer within the layer palette from one image. Then drag the layer onto another image. This drag and drop operation will create a new layer in the image. Do this for all other images. Alternatively, use a script (File>Scripts>Load Files into Stack…) to automate this process by loading multiple image files as layers in an image stack.
   3. Once each image is listed as an individual layer in the composite image, double click on the layer name to rename the layers as needed to discern the different channels.
   4. In order to see through each layer and create a truly composite image, change the blend mode for each layer from "Normal" to "Screen" within the layer palette.
      Note: To speed up this step, change a single layer to screen, then right-click on the layer, select "copy layer style", then highlight all other layers and select "paste layer style" to apply the screen style to the other layers.
   5. If desired, decrease the opacity (change value to 10 - 40%) of the chromogenic layer (i.e., toluidine blue) to better visualize the fluorescent signals through the chromogenic layer.
      Note: The tiled scanning microscope used in this protocol holds the slides on 3 edges, thereby minimizing the amount of rotation that can occur to the slide during each round of imaging. Therefore, it is much easier for the user to manually align the images without having to rotate images derived from different rounds of imaging.

Results

A General Workflow for the High-Throughput, Multi-Image Cryohistology

Figure 1 represents the general workflow used for this technique. It includes several steps from fixation through several rounds of imaging and finally image alignment/analysis. The process can take as little as a week to go from sample fixation through 4 rounds of imaging, which is less time than it takes to decalcify these t...

Discussion

Here we have presented a detailed cryohistology protocol to co-localize and quantify several biological measures by aligning images from multiple rounds of staining/imaging on a single section. The method outlined using the cryotape is especially useful as it maintains the morphology of difficult to section tissue (e.g., mineralized bone and cartilage). In addition, the sectioned tissue is adhered firmly to the glass slide, allowing for multiple rounds of staining/imaging of the same section; unlike traditional ...

Materials

Name	Company	Catalog Number	Comments
10% neutral buffered formalin	Sigma Aldrich	HT501128-4L	Multiple suppliers available. Toxic. Can be substituted with 4% paraformaldehyde.
Sucrose	Sigma Aldrich	S9378	Multiple suppliers available.
PBS	Sigma Aldrich	P5368	Multiple suppliers available.
Cryomolds	Fisher Scientific	Fisherbrand #22-363-554	Different sizes can be used depending on tissue
Cryomatrix	Thermo Scientific	6769006	Can be substitituted with other cryomatrices. However, PVA/PEG-based resins have worked best in our hands.
2-methyl-butane	Sigma Aldrich	M32631	Multiple suppliers available.
Cryostat	Leica Biosystems	3050s	Can be substituted with other brands/models.
Specimen disc	Leica Biosystems	14037008587	Can be substituted with other brands/models.
Cryostat blades	Thermo Scientific	3051835	Can be substituted with other brands/models.
Cryotape	Section Lab	Cryofilm 2C
Roller	Electron Microscopy Sciences	62800-46	Can be substituted with other brands/models.
Plastic microscope slides	Electron Microscopy Sciences	71890-01	Can be substituted with other brands/models.
Glass microscope slides	Thermo Scientific	3051	Can be substituted with other brands/models.
Norland Optical Adhesive, 61	Norland Optical	Norland Optical Adhesive, 61
UV Black Light	General Electric	F15T8-BLB
Glacial acetic acid	Sigma Aldrich	ARK2183	Multiple suppliers available.
Chitosan	Sigma Aldrich	C3646	Multiple suppliers available.
InSpeck red microscopheres	ThermoFisher Scientific	I-14787
InSpeck green microspheres	ThermoFisher Scientific	I-14785
Calcein Blue	Sigma Aldrich	M1255	Multiple suppliers available.
Calcein	Sigma Aldrich	C0875	Multiple suppliers available.
Alizarin complexone	Sigma Aldrich	A3882	Multiple suppliers available.
Demeclocycline	Sigma Aldrich	D6140	Multiple suppliers available.
NaHCO3	Sigma Aldrich	S5761	Multiple suppliers available.
Glycerol	Sigma Aldrich	G5516	Multiple suppliers available.
ELF 97 yellow fluorescent acid phosphatase substrate	ThermoFisher Scientific	E-6588
DAPI	ThermoFisher Scientific	62247	Multiple suppliers available. Can be substituted with Hoechst 33342 or other nuclear dyes.
TO-PRO-3 (Cy5 nuclear counterstain)	ThermoFisher Scientific	T3605
Propidium Iodide	ThermoFisher Scientific	R37108	Multiple suppliers available.
Sodium acetate anhydrous	Sigma Aldrich	S2889	Multiple suppliers available.
sodium tartrate dibasic dihydrate	Sigma Aldrich	T6521	Multiple suppliers available.
Sodium nitrite	Sigma Aldrich	S2252	Multiple suppliers available.
Tris	Sigma Aldrich	15504	Multiple suppliers available.
MgCl2 hexahydrate	Sigma Aldrich	M9272	Multiple suppliers available.
NaCl	Sigma Aldrich	S7653	Multiple suppliers available.
Fast Red TR Salt	Sigma Aldrich	F8764	Multiple suppliers available. Can also be substituted with other substrate kits such as Vector Blue (Vector Laboratories, Cat# SK-5300)
Naphthol AS-MX	Sigma Aldrich	N4875	Multiple suppliers available.
N,N dimethylformamide	Sigma Aldrich	D158550	Multiple suppliers available.
Toluidine blue O	Sigma Aldrich	T3260	Multiple suppliers available.
Axio Scan.Z1	Carl Zeiss AG	Axio Scan.Z1	Other linear or tiled scanners may also be used.
DAPI Filter Set	Chroma Technology Corp.	49000
CFP Filter Set	Chroma Technology Corp.	49001
GFP Filter Set	Chroma Technology Corp.	49020
YFP Filter Set	Chroma Technology Corp.	49003
Custom yellow (ELF 97) Filter Set	Chroma Technology Corp.	custom; HQ409sp, HQ555/30m, 425dxcr
TRITC Filter Set	Chroma Technology Corp.	49004
Cy5 Filter Set	Chroma Technology Corp.	49009
CryoJane Tape Transfer System	Electron Microscopy Sciences	62800-10	Multiple suppliers available.
CryoJane Tape Windows	Electron Microscopy Sciences	62800-72	Multiple suppliers available.
CryoJane Adhesive Slides	Electron Microscopy Sciences	62800-4X	Multiple suppliers available.