Cryosectioning of Contiguous Regions of a Single Mouse Skeletal Muscle for Gene Expression and Histological Analyses

Summary

Consecutive cryo-sections are collected to enable histological applications and enrichment of RNA for gene expression measurements using adjacent regions from a single mouse skeletal muscle. High-quality RNA is obtained from 20 - 30 mg of pooled cryosections and measurements are directly compared across applications.

Abstract

With this method, consecutive cryosections are collected to enable both microscopy applications for tissue histology and enrichment of RNA for gene expression using adjacent regions from a single mouse skeletal muscle. Typically, it is challenging to achieve adequate homogenization of small skeletal muscle samples because buffer volumes may be too low for efficient grinding applications, yet without sufficient mechanical disruption, the dense tissue architecture of muscle limits penetration of buffer reagents, ultimately causing low RNA yield. By following the protocol reported here, 30 μm sections are collected and pooled allowing cryosectioning and subsequent needle homogenization to mechanically disrupt the muscle, increasing the surface area exposed for buffer penetration. The primary limitations of the technique are that it requires a cryostat, and it is relatively low throughput. However, high-quality RNA can be obtained from small samples of pooled muscle cryosections, making this method accessible for many different skeletal muscles and other tissues. Furthermore, this technique enables matched analyses (e.g., tissue histopathology and gene expression) from adjacent regions of a single skeletal muscle so that measurements can be directly compared across applications to reduce experimental uncertainty and to reduce replicative animal experiments necessary to source a small tissue for multiple applications.

Introduction

The goal of this technique is to make multiple experimental analyses by different modalities, such as histology and gene expression, accessible from a single small skeletal muscle source tissue. Microscopy applications are the most sensitive to sample preservation methods, which must be carefully controlled to limit the formation of ice crystal artifacts during cryopreservation. Thus, method development is based on the tibialis anterior (TA) muscle frozen partially covered with embedding resin in a -140 °C liquid nitrogen-cooled 2-methylbutane bath as the source material for both immunofluorescence microscopy and gene expression analyses.

The need to use the same source material for diverse technical approaches is particularly important for intramuscular injection-based experiments where the left and right muscles represent different conditions, one experimental and one control. For example, in muscle regeneration studies, one muscle is injected with a toxin to cause widespread tissue damage while the contralateral muscle serves as a vehicle-injected control¹. Similarly, gene therapy studies for muscle disorders typically begin with validation of the gene therapy vector by intramuscular injection to be compared with empty vector, unrelated vector or vehicle control on the contralateral side². Therefore, it is not possible to source each TA muscle to a different application.

Common strategies to deal with this issue are: i) to use a different muscle group for each application, ii) to use additional mice, or iii) to cut off a piece of the muscle for each application. However, substantial differences between muscle groups make it difficult to compare data from separate applications, and additional animals increase expense and are poorly justified if other alternatives exist. Dividing the muscle after dissection to source different applications is the best option in many cases. However, the muscle pieces are often too small to use pulverization under liquid nitrogen or mechanical grinding techniques for homogenization^2-5. As muscle is a highly structural tissue packed with extracellular matrix and contractile proteins, inadequate mechanical homogenization leads to a low yield of subsequent DNA, RNA or protein. The method detailed here allows small quantities of tissue from one source muscle for use in multiple applications, and the inclusion of cryosectioning and needle trituration improves mechanical homogenization for better RNA yield.

Protocol

All animal procedures were approved by the University of Georgia Institutional Animal Care and Use Committee under animal use protocol A2013 07-016 (Beedle).

1. Cryopreservation of Unfixed Skeletal Muscle

1. Preparation
   1. Cut cork into small squares (approximately 1 cm x 1 cm) with a razor blade, write on the cork with a fine tip marker that is resistant to 2-methylbutane to identify the source mouse and muscle, and make a very shallow cut (approximately 1 mm) across the top surface. Insert a plastic coverslip into the cut to use for orienting the tissue. Repeat until a cork is ready for every tissue to be cryopreserved.
   2. Obtain liquid nitrogen, 2-methylbutane, embedding resin, low-temperature thermometer, corks, dissection tools, and study mouse.
      CAUTION: Liquid nitrogen is a compressed gas that may explode if heated. Wear a lab coat, low-temperature gloves, and face protection when handling liquid nitrogen; contact with skin or eyes may cause burns or frostbite. 2-Methylbutane is a flammable, toxic, health and environmental hazard. Wear personal protective equipment (lab coat, gloves, safety glasses), open the stock container in a fume hood, transfer the small amount needed for freezing (typically 200 to 400 ml) to a separate container that can be tightly closed, and avoid inhalation.
   3. Euthanize mice with an approved method of euthanasia under anesthesia. Briefly, insert the mouse into an inhalation chamber with 2.5% isoflurane in oxygen from an isoflurane vaporizer, wait until 20 sec after the mouse stops moving to check for a pedal reflex. When the pedal reflex is negative, euthanize the study mouse by cervical dislocation⁶.
      NOTE: Euthanasia methods require approval by the institutional ethics committee.
   4. Remove skin overlying the distal hindlimb and cut the distal anterior tendons just above the ankle with fine-point dissection scissors. Grasp the distal tendons with fine forceps and gently pull up and towards the knee while cutting lateral fascia to release the muscle. Pull the muscle out perpendicular from the knee and make a final cut to excise the TA muscle⁷.
      NOTE: The TA muscle is used here as an example, but any mouse skeletal muscle or the heart can be substituted for the TA in this protocol if appropriate to the user's experimental goals. The only limitation is that a tissue must be small enough to achieve rapid cryopreservation throughout its depth; a maximum tissue size of 1 cm x 1 cm is recommended.
   5. Repeat on the opposite leg, and dissect out any other tissues to be collected. Perform dissection as quickly as possible to limit tissue degradation before cryopreservation.
   6. Orient each muscle for transverse sections on its pre-labeled cork. Stand the muscle perpendicular to the cork with the distal tendon touching the cork and the top of the muscle extending away, held upright by the coverslip.
   7. Cryopreserve all tissues for histological analysis as soon as possible after dissection, preferably within 5 min but definitely not more than 15 min time elapsed since euthanization of the source animal.
2. Cryopreservation
   1. Begin cooling the cryopreservation bath five min before the end of the dissection. Pour 2-methylbutane into an open metal beaker to a depth of approximately 3 cm. Pour liquid nitrogen into an insulated container to a depth of 2 to 4 cm. Set the beaker of 2-methylbutane into the liquid nitrogen; the nitrogen will begin to boil. Avoid nitrogen splashing into the 2-methylbutane beaker.
      CAUTION: Prepare freezing bath in a fume hood or a well-ventilated area.
   2. Insert a low-temperature thermometer into the 2-methylbutane to monitor temperature and stir frequently with a fork to ensure even cooling. Continue to stir and cool until 2-methylbutane reaches -140 °C, adding new liquid nitrogen to the outer bath as necessary.
      NOTE: -140 °C is the optimum temperature to minimize ice crystal artifacts in striated muscle; other tissues may cryopreserve best at different temperatures (e.g., brain, -90 °C).
   3. Apply embedding resin only to the lower 35 - 50% of the muscle where it meets the cork and immediately drop the cork into the 2-methylbutane at -140 °C. Rapidly repeat for up to 8 tissues per freeze batch. Stir for 30 sec, scraping the bottom of the beaker to ensure that tissues don't freeze into the solidifying 2-methylbutane.
   4. Use a fork, slotted spoon or large forceps to pull each tissue cork from the 2-methylbutane. Quickly remove the coverslip, dab off excess 2-methylbutane into the beaker, and then drop the tissue cork into the outer nitrogen bath. Repeat for all remaining corks in the beaker.
   5. Transfer samples in liquid nitrogen or on dry ice to a -80 °C freezer for storage.
   6. If any tissues remain for cryopreservation, repeat steps 1.2.3 to 1.2.4. Add additional 2-methylbutane to the beaker or liquid nitrogen to the outer bath as necessary and re-cool to -140 °C. Dispose of used 2-methylbutane as hazardous waste.

2. Collect Cryosections for Histology and RNA Applications

1. Preparation.
   1. Pre-weigh a DNase/RNase-free autoclaved tube on an analytical balance. Then, move it into the cryostat chamber to precool. Repeat until tubes are ready for all pooled cryosection samples to be collected.
   2. For skeletal muscle sectioning, confirm that the cryostat chamber temperature is -21 to -22 °C by its internal thermostat. Transfer any containers with tissues to be cut into the chamber and allow the container(s) to equilibrate to the cryostat temperature for at least 15 min before opening.
   3. Insert a new disposable cryostat blade. Alternately, remove the existing blade, spray with RNase decontamination solution, rinse with ddH₂O, and reinsert into the cryostat to cool. Spray a clean tissue with 70% ethanol and carefully wipe the blade and blade clamping platform.
      1. Spray a clean tissue with an RNase decontamination solution and wipe cryostat brushes. Spray a tissue with ddH₂O, wipe the brushes again and set them in the cryostat chamber on a clean surface.
         CAUTION: The cryostat blade should be covered by the knife guard when not in use. Also, RNase decontamination solution is toxic and will freeze to form a precipitate in the cold cryostat chamber. Therefore, handle with care and avoid its use in the cryostat chamber.
   4. Within the cryostat chamber, place a dime-sized drop (approximately 300 μl) of embedding resin onto a warm specimen chuck, set a tissue cork on top of the resin, press down, and then set the chuck in the freezing rail or onto a fast freeze element if available.
   5. After the embedding resin solidifies (white), add additional embedding resin on top of the cork, around the lower 35% of the tissue and press the heat extractor into the embedding resin for a fast freeze to better stabilize the tissue. Wait 5 min before sectioning to allow the resin to harden fully.
   6. Ensure that the specimen clamp is in its most retracted position, farthest from the blade. Insert the tissue specimen chuck into the specimen clamp.
2. Cryosection preparation.
   1. Loosen the blade carrier holder, set the clearance angle of the blade to 10° (or an angle appropriate for the blade carrier used), and re-tighten. Release the brake and turn the hand wheel to lower the muscle towards the blade. Estimate the closest distance between the tissue and blade, then move the tissue away from the blade and engage the hand wheel brake.
   2. Loosen the height adjustment lever and move the blade carrier forward or back, respectively, if the end of the tissue is more than approximately 2 mm from the blade or if the blade strikes the tissue. Tighten and lower the tissue again to check distance from the blade. Repeat adjustments until the blade is 1 to 2 mm from the end of the tissue by visual estimate.
   3. Lower the tissue towards the blade and assess tissue angle for transverse sections. Lock the hand wheel and release the specimen clamp lever. Push the specimen clamp left or right until the horizontal orientation of the tissue is perpendicular to the blade.
   4. Push the specimen clamp up or down to adjust the "y" orientation so that tissue sections will be perpendicular to the long axis of the muscle. Tighten the specimen clamp lever.
   5. Use the course and fine forward feed to advance the specimen until it just touches the blade. If seeking sections from a particular tissue depth, reset the sum of section thicknesses to zero (top of the tissue).
   6. Set the cryostat to the trim function with section thickness at 30 μm. Cut and discard sections until the preselected depth for tissue collection is reached (e.g., 400 μm from the top).
3. Collect cryosections for RNA extraction.
   1. Open the tube for collecting sections and place it near the blade carrier. Use a pre-cooled, clean brush to pick up each section as it is cut from the blade and transfer the cryosection into the tube. Repeat until the pooled sections weigh 30 mg or the desired tissue depth is reached.
      NOTE: For an adult mouse tibialis anterior, collection from tissue depth of approximately 400 to 4,000 μm typically yields 25 - 40 mg. Using metal forceps to transfer sections to the collection tube is not recommended as the sections tend to stick and clump on the metal surface.
   2. If embedding resin surrounds the muscle cryosection, lock the handbrake and use a razor blade to shave off small pieces of resin until there is only a thin layer around the top of the muscle. Always cut resin with the blade angled away from the muscle.
      NOTE: A thin layer of embedding resin does not substantially impair the downstream RNA preparation. If thicker embedding resin is present, use brushes to tease it away from the muscle before moving the cryosection into the collection tube.
   3. Alternatively, pool sections on the blade carrier and transfer in bulk to the collection tube.
      NOTE: However, this method tends to be slower, and sections are more likely to stick and clump together, which can reduce the efficiency of needle homogenization in later steps.
   4. Quickly place the pooled cryosection tube into an analytical balance and record tube weight. Immediately return the tube to the cold cryostat chamber to maintain section temperature near -20 °C. Calculate the weight of the pooled sections.
      NOTE: If RNA isolation will occur on a different day, store the pooled cryosection tube at -80 °C until use.
4. Collect cryosections for histology.
   1. Press the section thickness button for fine sectioning and use arrows to set the cryostat section thickness to 7 μm (or other appropriate section thickness, typically 6 to 10 μm).
      NOTE: Thinner sections (6 to 10 μm) should be used for histological applications to ensure that staining reagents can penetrate the depth of the tissue section. Thin sections can be taken from any depth during the cryosectioning, but deeper sections are preferred because embedding resin, which increases with tissue depth, does not impair histological staining.
   2. Cut and discard 4 to 7 sections to obtain a consistent, even tissue surface. Make note of the tissue depth.
   3. Cut a section and orient it on the surface of the blade carrier. Pick up the section by quickly and gently touching a warm (room temperature) microscope slide to the section on the blade carrier. Return the slide to room temperature. Continue until the number of desired slides is obtained. Make note of the final tissue depth.
      NOTE: Collecting a second (duplicate) section for each tissue is recommended.
   4. With sectioning complete, engage the hand wheel brake, return the specimen to the rear-most position, and remove the tissue chuck. Use the cryostat heat element to melt the embedding resin holding the tissue cork on the specimen chuck. Remove tissue cork, dry with tissue, and return to storage.
   5. Repeat from step 2.1.2 for each remaining tissue. Allow slides to dry for 20 min after the last tissue is mounted. Then, use slides for histological or immunofluorescent staining or freeze at -80 °C in a slide box until needed.

3. RNA Isolation from Pooled Cryosections

1. RNA extraction
   1. Move tube(s) of pooled cryosections to ice. Immediately add an organic RNA extraction reagent at a ratio of approximately 1 ml reagent per 50 mg cryosection weight, typically 600 μl per tube.
      CAUTION: RNA isolation reagents are toxic, corrosive, and irritating. Follow the manufacturer's instructions for safe handling.
   2. Using a 1 ml syringe with an 18 gauge needle, draw the RNA extraction liquid up and rinse the walls of the tube until all tissue is suspended in the solution. Try to minimize air bubbles during needle homogenization.
   3. Use the tip of the needle to mash and disperse any clumped cryosections and pieces adhering to the tube wall. Triturate to homogenize by passing the sample up and down through the needle for five strokes, and then return the sample to ice. Repeat both steps three to five times, or as many times as is necessary to disrupt all clumps and pass the sample easily through the needle.
   4. Remove the 18 gauge needle and replace it with a 22 gauge needle. Carefully triturate the sample for five strokes, and then return the sample to ice. Repeat the trituration three to four times to achieve a very finely dispersed tissue homogenate.
      NOTE: If tissue particles start to block the needle when drawing up the sample, expel all sample from the syringe and switch back to the 18 gauge needle for further homogenization. An additional trituration step with a 25 or 26 gauge needle can be added to obtain maximal yield. However, the needle can become clogged so there is a risk of sample spilling.
   5. Move the sample from ice to room temperature. Incubate for 5 min to disrupt molecular complexes.
   6. In a fume hood, add 0.1 ml of 1-bromo-3-chloropentane (BCP) per 1 ml of RNA isolation reagent used (step 3.1.1), typically 60 μl per tube. Shake the tube vigorously by hand for 15 sec (do not vortex). Incubate the sample at room temperature for 2 to 3 min. Then, centrifuge for 15 min at 12,000 x g and 4 °C.
      CAUTION: BCP is flammable and toxic. Handle in a fume hood and wear gloves, a lab coat, and safety glasses.
   7. Carefully collect the upper aqueous phase (colorless) containing RNA and transfer it to a clean tube. Add an equal volume of 70% ethanol (in DEPC water) and vortex to mix. Incubate at room temperature for up to 10 min.
      NOTE: the middle interphase and lower phenol-BCP phases can be processed for genomic DNA and protein, respectively, according to the manufacturer's instructions or collected in a phenol-BCP hazardous waste container for appropriate disposal.
2. RNA purification.
   1. Follow the manufacturer's instructions for sample binding, washing and elution with minor variations⁸. For example:
   2. Place an aliquot of DNase/RNase-free water into a 37 °C bath or incubator to pre-warm for step 3.2.4.
   3. Add the RNA sample from 3.1.8 above to a purification column and centrifuge the sample for 15 sec at 12,000 x g at room temperature. Pour the flow through back into the same column and re-centrifuge. Repeat one additional time to maximize RNA binding, and then discard the final flow through.
   4. Perform wash steps according to the manufacturer's instructions⁸. Apply 700 μl of wash buffer (no ethanol) to the column, spin for 15 sec at 12,000 x g, and discard the flow through.
   5. Add 500 μl of wash buffer (with ethanol) to the column, spin for 15 sec at 12,000 x g, and discard the flow through. Repeat this step one time.
   6. Spin the empty column for 1 min at 12,000 x g to dry the membrane.
   7. Transfer the spin column to a clean RNase-, DNase-free tube. Add 40 μl of pre-warmed DNase/RNase-free water (from 3.2.1) to the center of the column membrane. At room temperature, incubate for 2 min and then centrifuge for 2 min at 12,000 x g.
      NOTE: The 40 μl elution volume is approximately 1.3 μl per mg of original tissue for 30 mg of pooled cryosections. Adjust the elution volume as appropriate for different starting tissue weights, but do not reduce below 32 μl. A second elution, using approximately 0.7 μl per mg of original tissue, can be added to increase total RNA yield.
   8. Analyze the RNA (column elution) for concentration and purity by a spectrophotometer, electrophoresis, and/or a bioanalyzer. Store the RNA at -80 °C until needed for downstream applications, such as reverse transcription for quantitative PCR⁴.
      NOTE: A DNase treatment is recommended before using the RNA in downstream applications. This treatment can be performed on some RNA purification columns before the wash step, at this specific point following column elution, or on an aliquot of the RNA as the first step in any downstream application. The RNA should be stable for several years.

4. Histological Analysis by Immunofluorescent Staining of Muscle Cryosections

1. Simple immunofluorescence protocol.
   1. Use a hydrophobic pen to circle the group of sections on each slide. Gently drop PBS into the circled area (approximately 80 μl for a small surface area up to 500 μl for a large surface area), being careful not to touch the tissues. Incubate for 5 min at room temperature.
      NOTE: If using frozen slides, remove slides from -80 °C freezer and incubate at room temperature for 20 to 30 min to thaw and dry, then proceed as above. At least two slides are needed: one experimental slide to be incubated in primary and secondary antibody; and one control slide for which primary antibody is excluded, the "secondary only" control.
   2. Tip the slide to pour off PBS. Add 5% donkey serum in PBS (blocking solution) to the circled area; be careful to ensure that the muscle sections do not dry out. Incubate at room temperature for 30 min (or up to several hours in a humidity chamber).
   3. Prepare primary antibody solution to analyze muscle regeneration. Mix blocking solution, with an eMHC antibody (1:40) and a ColVI antibody (1:1,000). Prepare approximately 150 μl, 300 μl or 500 μl for small, medium, or large slide areas, respectively. Vortex for 5 sec, and then centrifuge for 2 min at 15,000 x g to pellet any precipitate.
   4. For experimental slides, tip the slide to pour off blocking solution and then add primary antibody solution from 4.1.3. For the secondary control slide, tip the slide to pour off blocking solution and then add fresh blocking solution (no antibody). Incubate slides in a humidity chamber at 4 °C overnight.
      NOTE: When pipetting antibody solutions onto slides, always pull liquid from the top of the primary antibody solution tube, do not disrupt any precipitate at the bottom of the tube.
   5. Tip slides to pour off solution. Add PBS dropwise to the circled region of each slide to wash, tip to pour off, then add more PBS and incubate for 3 to 5 min. Repeat for a total of three washes.
      NOTE: The wash time is quite flexible and can be lengthened up to 20 min if desired. Generally, the number of solution changes is more important than the time.
   6. Prepare secondary antibody solution for all slides (including the secondary control slide) using a 1:500 dilution of red and green fluorophore-coupled secondary antibodies to detect mouse IgG1 (eMHC) and rabbit IgG (ColVI) and 1:10,000 dilution of DAPI nuclear stain.
      1. For example, mix 1 μl of DAPI with 9 μl of ddH₂O to make a 1:10 dilution of DAPI. Then, for a 500 μl final volume, add 1.0 μl anti-mouse IgG1-red + 1.0 μl anti-rabbit IgG-green + 0.5 μl of 1:10 DAPI to 447.5 μl of blocking solution. Vortex to mix and centrifuge for 2 min at 15,000 x g to pellet any precipitate.
         NOTE: Ideally, all secondary antibodies should be from the same host species.
   7. Tip the slides to pour off the last PBS wash. Add secondary antibody solution to cover all tissues. Cover the slides to protect them from light and incubate at room temperature for 30 min.
      NOTE: All secondary antibodies should be validated to have minimal cross reactivity with other species in dual labelling experiments.
   8. Wash the slides as described in 4.1.5. After the last wash, tip the slide to pour off the PBS and set the slide on a tissue. Add 3 to 4 drops of an aqueous mounting media along one side.
   9. Set the edge of a glass coverslip just to the outer edge of the mounting media. Using forceps, slowly lower the coverslip towards the tissues, then, release the coverslip and gently tap it into position. Finally, gently press each corner of the coverslip to stabilize it.
   10. Protect the slides from light and store at 4 °C until use.
       NOTE: For long term storage, apply a thin layer of nail polish along the edge of the coverslip to help prevent the slide from drying.
2. Histological evaluation.
   1. Image the slides using an epifluorescent or confocal microscope with appropriate filters to detect eMHC (red); ColVI (green); and DAPI-stained nuclei (blue), typically using a 20X objective^1,5.
   2. Confirm that the fluorescent signal from experimental slides is different from the secondary control slide to demonstrate the specificity of the eMHC and ColVI detection⁴.
      NOTE: eMHC positive regenerating fibers should be visible from approximately 2 to 7 days after a muscle injury and variably in mice with muscular dystrophy. Collagen VI is present in the extracellular matrix surrounding muscle fibers, blood vessels, and nerves and in the larger connective tissue bundles of peri- and epimysium. DAPI nuclear stain is useful to identify muscle fibers with central nuclei versus peripheral nuclei indicating fiber regeneration, and the presence of infiltrating cells. Necrotic or injured fibers may stain weakly with the mouse IgG secondary antibody due to detection of endogenous IgG that penetrates the fibers through damaged muscle membranes.
   3. To quantify regeneration, take overlapping microscope pictures with each fluorescent filter across the entire image. Merge the eMHC, ColVI and DAPI images for each location and then align the pictures to reconstruct a map of the entire section^1,5.
   4. Using analysis software, count the number of eMHC positive fibers and the number of total fibers to calculate recent regeneration (100*(# eMHC+ fibers/# total fibers)). Also, count the number of centrally nucleated fibers out of the total fibers to measure regeneration over a longer period (100*(# CN+ fibers/# total fibers))^1,5.

Results

Muscle cryosection RNA is high in quality and provides sufficient yield for most applications

Analyses of sixteen skeletal muscle RNA preparations are shown in Table 1 using 19.4 to 41 mg of pooled tibialis anterior (TA) muscle from 8 control mice. Both left (L) and right (R) TA muscles were prepared in regeneration experiments with muscles collected 3 days after longitudinal intramuscular injection of 25 μl of sal...

Discussion

To achieve best results with this method, keep embedding resin restricted to the lower third or half of the muscle during tissue cryopreservation because excess resin will slow the collection of the pooled cryosections and may increase embedding resin contamination in the RNA isolation. Also, careful attention during needle homogenization is important to maximize yield and minimize the probability of clogging the needle. The protocol may be modified by using a Luer-Lok syringe to protect against sample loss if the needle...

Materials

List of materials used in this article

Name	Company	Catalog Number	Comments
Cork	VWR Scientific	23420-708	Cut into small squares with a sharp blade.
Plastic coverslip	Fisher Scientific	12-547	Used to orient the muscle during freezing.
Low temperature thermometer	VWR Scientific	89370-158
2-methylbutane	Sigma	M32631-4L	Caution: hazardous chemical. Store in flammable cabinet.
Embedding resin: "cryomatrix"	Thermo Fisher Scientific	6769006	Other embedding resins can be substituted for cryomatrix.
Cryostat	Thermo Fisher Scientific	microm HM550 with disposable blade carrier	Any working cryostat should be sufficient for the protocol.
Disposable cryostat blade	Thermo Fisher Scientific	3052835	Use an appropriate blade or knife for the cryostat to be used.
RNAse decontamination solution: "RNase Zap"	Thermo Fisher Scientific	AM9780
Analytical balance	Mettler Toledo	XS64
Paint brush	Daler Rowney	214900920	Use to handle cryosections. Can be found with in stores with simple art supplies.
Razor blade	VWR Scientific	55411-050
Microscope slide	VWR Scientific	48311600
RNA organic extraction reagent: TRIzol	Thermo Fisher Scientific	15596026	Caution: TRIzol is a hazardous chemical. Note: Only organic extraction reagents are recommended for RNA extraction from skeletal muscle.
18 gauge needle	VWR Scientific	BD305185
22 gauge needle	VWR Scientific	BD305155
26 gauge needle	VWR Scientific	BD305115	Optional. Can be used for a third round of sample trituration in the RNA extraction protocol.
1 ml syringe	VWR Scientific	BD309659	For very high value samples, a Luer-Lok syringe is recommended (e.g., VWR BD309628).
1-bromo-3-chloropentane (BCP)	Sigma	B9673
For 70% ethanol in DEPC water: 200 proof alcohol	Decon Laboratories, Inc.	+M18027161M	Mix 35 ml 200 proof alcohol + 15 ml DEPC water.
For 70% ethanol in DEPC water: DEPC-treated water	Thermo Fisher Scientific	AM9922	Mix 35 ml 200 proof alcohol + 15 ml DEPC water.
RNA purification kit: PureLink RNA minikit	Thermo Fisher Scientific	12183018A	Final steps of RNA preparation.
DNase/Rnase-free water	Gibco	10977	DEPC-treated water can also be used.
Spectrophotometer: Nanodrop 2000	Thermo Fisher Scientific	NanoDrop 2000
Dnase I	Thermo Fisher Scientific	AM2222	Treat purified RNA to remove any DNA contamination before downstream appications.
Hydrophobic pen	Thermo Fisher Scientific	8899
Dulbecco's PBS	Gibco	14190	PBS for immunofluorescence protocol.
Donkey serum	Jackson ImmunoResearch Laoratories, Inc	017-000-121	Rehydrate normal donkey serum stock according to the manufacturer's instructions, then dilute an aliquot to 5% for immunofluorescence.  Normal goat serum can also be used.
eMHC antibody	University of Iowa Developmental Studies Hybridoma Bank	F1.652
Collagen VI antibody	Fitzgerald Industries	#70R-CR009x
Donkey anti-rabbit AlexaFluor488	Thermo Fisher Scientific	A21206
Goat anti-mouse IgG1 AlexaFluor546	Thermo Fisher Scientific	A21123
DAPI (4',6-diamidino-2-phenylindole)	Thermo Fisher Scientific	D1306
Aqueous mounting media: Permafluor	Thermo Fisher Scientific	TA-030-FM	Only use mounting media designed for fluorescent applications with anti-fade properties.
Glass coverslip	VWR Scientific	16004-314	Use for mounting slides at the end of immunofluorescence protocl
Image analysis software: ImageProExpress	Media Cybernetics, Inc.	Image-Pro Express, or more advanced products	Freeware ImageJ should also work for manual counting. More advanced software with segmentation abiities may allow partial automation of the process; e.g., ImageProPremier.
Merge and map section images: Photoshop	Adobe	Photoshop
Cardiotoxin	Sigma	C9659	Sigma C9659 has been discontinued. Other options for cardiotoxin are EMD Millipore #217503; American Custom Chemicals Corp. # BIO0000618; or Ge Script # RP17303; but these have not been validated.
reverse transcription kit: Superscript III First-strand synthesis system	Thermo Fisher Scientific	18080051	Any validated, high quality reverse transcription reagents can be used.
Standard PCR: GoTaq Flexi polymerase system	Promega	M8298	Any validated, high quality Taq polymerase system can be used. If DNA sequencing is to be used for any application downstream of the PCR, then a high fidelity PCR system should be used instead.
SYBR green	Thermo Fisher Scientific	S7585	For use in qPCR when not using a dedicated qPCR master mix. Use with SuperROX (for Applied Biosystems instruments) and GoTaq Flexi polymerase and buffers.
ROX: SuperROX, 15 mM	BioResearch Technologies, Inc. Novato CA	SR-1000-10	SuperROX is more stable in the PCR reaction, so it is preferred for use as a qPCR passive reference dye over ROX (carboxy-X-rhodamine). For qPCR with Applied Biosystems instruments
Real-time PCR	Applied Biosystems	7900HT