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Dec 2010

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Transmission Electron Microscopy for Analysis of Mitochondria in Mouse Skeletal Muscle
使用透射电子显微镜技术分析小鼠骨骼肌线粒体   

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Abstract

Skeletal muscle is the most abundant tissue in the human body and regulates a variety of functions including locomotion and whole-body metabolism. Skeletal muscle has a plethora of mitochondria, the organelles that are essential for aerobic generation of ATP which provides the chemical energy to fuel vital functions such as contraction. The number of mitochondria in skeletal muscle and their function decline with normal aging and in various neuromuscular diseases and in catabolic conditions such as cancer, starvation, denervation, and immobilization. Moreover, compromised mitochondrial function is also associated with metabolic disorders including type 2 diabetes mellitus. It is now clear that maintaining mitochondrial content and function in skeletal muscle is vital for sustained health throughout the lifespan. While a number of staining methods are available to study mitochondria, transmission electron microscopy (TEM) is still the most important method to study mitochondrial structure and health in skeletal muscle. It provides critical information about mitochondrial content, cristae density, organization, formation of autophagosomes, and any other abnormalities commonly observed in various disease conditions. In this article, we describe a detailed protocol for sample preparation and analysis of mouse skeletal muscle mitochondria by TEM.

Keywords: Transmission electron microscopy (透射电子显微镜技术), Skeletal muscle (骨骼肌), Mitochondria (线粒体), Autophagy (自噬), Myopathy (肌病), Atrophy (萎缩), Oxidative metabolism (氧化代谢)

Background

Skeletal muscle is a highly plastic tissue that undergoes morphological and metabolic adaptations in response to a number of extracellular cues. A number of perturbations including resistance or endurance exercise stimulates mitochondrial biogenesis leading to increased metabolic capacity and resistance to fatigue (Li et al., 2008; Sandri, 2008). By contrast, during aging, inactivity, and in many catabolic disease states, skeletal muscle mitochondrial number and function decline, leading to increased fatigability and insulin resistance (Sandri, 2008). An accumulation of dysfunctional mitochondria may also result in progressive reactive oxygen species-induced damage, producing a further impairment of oxidative capacity in skeletal muscle (Bonnard et al., 2008).

Mitochondria exist as a reticular membrane network that is located in different subcellular compartments in skeletal muscle. The subsarcolemmal (SS) mitochondria, account for 10-15% of the mitochondrial volume and lie directly beneath the sarcolemmal membrane, whereas the intermyofibrillar (IMF) mitochondria are located in close contact with the myofibril (Takahashi and Hood, 1996). Mitochondria are double membrane structures containing an intermembrane space between the outer and inner membranes as well as the inner matrix compartment, where most of the metabolic processes take place. The inner membrane is highly folded, forming so-called cristae, to accommodate its large surface area. The five complexes that make up the respiratory chain where oxidative phosphorylation takes place are embedded within the inner mitochondrial membrane. In this process, a proton gradient across the inner membrane is coupled to ATP synthesis at complex V (Peterson et al., 2012). In addition to producing ATP for cross-bridge cycling between actin and myosin, mitochondria are a source of free radicals which regulate skeletal muscle physiology (Peterson et al., 2012).

Transmission electron microscopy (TEM) is a powerful technique for ultrastructural studies (Watson, 1958). TEM has been very useful in studying mitochondrial structure in skeletal muscle in both physiological and pathological conditions (Picard et al., 2013). For example, TEM can provide information about mitochondrial content, organization, cristae structure, and vacuolization as observed in some neuromuscular disorders such as Amyotrophic lateral sclerosis (Picard et al., 2013). In many muscle wasting conditions, mitochondrial content is reduced through autophagy, also known as mitophagy. In this regard, TEM has been found to be an important approach to study autophagosome formation (Sandri, 2008). We have developed an efficient protocol that can be easily adapted in any laboratory to study the ultrastructure of mouse mitochondria in skeletal muscle by TEM (Paul et al., 2010; Hindi et al., 2014 and 2018). In the following sections, we provide a step-wise protocol for sample preparation and analysis of SS and IMF mitochondria in skeletal muscle. A similar protocol can be used for studying other organelles in skeletal muscle by TEM as well.

Materials and Reagents

  1. Glass specimen vials (Electron Microscopy Sciences, catalog number: 72630-05 )
  2. Razor blades, Double Edge Coated, Washed Version (Electron Microscopy Sciences, Personna, catalog number: 72000-WA )
  3. Transfer pipette (Fisher Scientific, catalog number: 13-711-9BM )
  4. Nitrile gloves
  5. Glass strips, ultramicrotomy grade, 6.4 x 25 x 400 mm (Electron microscopy sciences, catalog number: 71012 )
  6. Glass slides (Fisher Scientific, catalog number: 12-550-15 )
  7. Syringes 1 ml (BD, catalog number: 329652 )
  8. Syringes 30 ml (BD, catalog number: 302833 )
  9. 0.22 μm syringe filter (Merck, catalog number: SLGV033RS )
  10. Conical centrifuge tube, 50 ml (VWR, catalog number: 21008-169 )
  11. Conical centrifuge tube, 15 ml (VWR, catalog number: 21008-089 )
  12. Non-sterile urine specimen container (Electron Microscopy Sciences, catalog number: 64231-10 )
  13. Filter paper, Qualitative Grade 1 Circles (GE Healthcare, Whatman, catalog number: 1001-090 )
  14. Glass stirring rod (United Scientific Supplies, catalog number: GSR012 )
  15. Wood applicators (Electron microscopy Sciences, catalog number: 72300 )
  16. Flat, silicone embedding mold (Electron Microscopy Sciences, catalog number: 70900 )
  17. Glass knife boat, 6.4 mm (Electron Microscopy Sciences, catalog number: 71007 )
  18. Glass knife box (Electron Microscopy Sciences, catalog number: 71010 )
  19. N95 respirator, with valve (VWR, catalog number: 89201-510 )
  20. Metal loop, perfect loop (Electron Microscopy Sciences, catalog number: 70944 )
  21. Grids, tabbed, copper, 200 mesh (Ted Pella, catalog number: 3HGC200 )
  22. Grid storage box, tabbed (Ted Pella, catalog number: 161 )
  23. Petri dish, glass, 100 x 20 mm (Corning, catalog number: 70165-102 )
  24. Glutaraldehyde, EM grade, 8% (Polysciences, catalog number: 00216-30 )
  25. Sodium phosphate monobasic monohydrate, NaH2PO4·H2O (Sigma-Aldrich, catalog number: S9638 )
  26. Sodium phosphate dibasic anhydrous, Na2HPO4 (Sigma-Aldrich, catalog number: S9763 )
  27. Osmium tetroxide, 10 x 1 g (Electron Microscopy Sciences, catalog number: 19110 )
  28. Ethanol, 200 Proof (Decon Labs, catalog number: 2701 )
  29. EMbed-812 kit, includes: EMbed-812, DDSA, NMA, and DMP-30 (Electron Microscopy Sciences, catalog number: 14120 )
  30. Sodium borate (MP Biomedicals, catalog number: 0219030980 )
  31. Toluidine blue O (Amresco, catalog number: 0672-25G )
  32. Uranyl acetate dihydrate powder (depleted) (Electron Microscopy Sciences, catalog number: 22400 )
  33. NaOH pellets (Amresco, catalog number: 0583-500G )
  34. Lead Nitrate, Pb(NO3)2 (Electron Microscopy Sciences, catalog number: 17900 )
  35. Sodium citrate, Na3(C6H5O7)·2H2O (Electron Microscopy Sciences, catalog number: 21140 )
  36. Propylene oxide, EM grade (Electron Microscopy Sciences, catalog number: 20401 )
  37. Dental wax (Electron Microscopy Sciences, catalog number: 72660 )
  38. 3% glutaraldehyde (see Recipe 1)
  39. 0.1 M sodium phosphate buffer pH 7.4 (see Recipe 2)
  40. 1% osmium tetroxide (see Recipe 3)
  41. Ethanol dilutions (see Recipe 4)
  42. Embedding media (see Recipe 5)
  43. 1% toluidine blue stain (see Recipe 6)
  44. 4% uranyl acetate stock (aq) (see Recipe 7)
  45. 1 N-NaOH (see Recipe 8)
  46. Reynold’s Lead Citrate (see Recipe 9)

Precautions/Hazards: As with any chemicals and reagents handled in the lab, users should be aware of how to use and manipulate them safely. Please refer to each chemical’s Material Safety Data Sheet (MSDS) for detailed information about precautions and hazards. Electron microscopy uses quite a few hazardous chemicals, such as: glutaraldehyde, osmium tetroxide, propylene oxide, uranyl acetate, lead citrate, and others. Please handle these chemicals using the proper personal protective equipment (PPE), ventilation conditions, and dispose of these chemicals in accordance with your institution’s Department of Environmental Health and Safety.


Equipment

  1. Amber, wide-mouth glass bottle, 125 ml (VWR, catalog number: 10861-846 )
  2. Clear, media bottle, 1 L (Corning, PYREX®, catalog number: 1399-1L )
  3. Graduated cylinder, 1,000 ml (VWR, catalog number: 65000-012 )
  4. Graduated cylinder, 25 ml (VWR, catalog number: 65000-002 )
  5. Magnetic stirring bar (VWR, catalog number: 58948-025 )
  6. General-Purpose Liquid-In-Glass Thermometer (VWR, catalog number: 89095-626 )
  7. Negative-action, curved self-closing tweezers (Electron Microscopy Sciences, catalog number: 72864-D )
  8. Eyelash manipulator (Electron Microscopy Sciences, catalog number: 71182 )
  9. 1,000 ml Glass Griffin beaker (VWR, catalog number: 10754-960 )
  10. 50 ml Glass Griffin beakers (VWR, catalog number: 10754-946 )
  11. Glass funnel, 100 mm (VWR, catalog number: 10546-048 )
  12. 50 ml volumetric flask (VWR, catalog number: 10123-996 )
  13. 100 ml volumetric flask, amber (VWR, catalog number: 10124-022 )
  14. -20 °C Freezer (VWR, catalog number: 97014-903 )
  15. 4 °C Refrigerator (VWR, catalog number: 14236-525 )
  16. Clinical rotator, variable speed tube rotator (Cole-Parmer, Stuart, catalog number: SB3 )
  17. Culture tube holder for clinical rotator, variable speed tube rotator, 12 mm offering a rolling action for tubes (Cole-Parmer, Stuart, catalog number: SB3/3 )
  18. pH meter, SymPhony B10P (VWR, catalog number: 89231-662 )
  19. pH probe, refillable, glass (VWR, catalog number: 89231-580 )
  20. Precision Balance, AV212C (Ohaus, out of production)
  21. Precision Balance (OHAUS, catalog number: 30122632 )
  22. Hotplate stirrer (Fisher Scientific, catalog number: SP88857200P )
  23. Vacuum oven (Electron Microscopy Sciences, catalog number: 63235-10 )
  24. Ultramicrotome (Reichert-Jung, model: Ultracut E , out of production, eBay or other second-hand markets)
  25. New ultramicrotome (Leica Microsystems, model: Leica EM UC7 )
  26. Glass knifemaker (LKB, model: LKB Type 7801B , out of production, eBay or other second-hand markets)
  27. New glass knifemaker, Leica EM KMR3 (Leica Microsystems,, catalog number: Leica EM KMR3 )
  28. Light microscope, Olympus CX31 (Olympus, catalog number: CX31 )
  29. Diamond knife, Diatome wet ultra 45°, 3.5 mm (Electron Microscopy Sciences, model: Diatome Ultra )
  30. Phillips CM10 transmission electron microscope retrofitted with a new digital camera (Phillips, catalog number: CM10 )
  31. High Definition CCD Camera for TEM retrofitted to Phillips CM10 scope (Advanced Microscopy Techniques, catalog number: BioSprint )

Procedure

The basic steps for using TEM to analyze mitochondrial ultrastructure in skeletal muscle are presented in Figure 1.


Figure 1. The schematic view of general procedures for TEM of skeletal muscle from start to finish

  1. Primary fixation
    The goal of primary fixation with glutaraldehyde is to preserve the ultrastructure of the skeletal muscle tissue in a physiologically relevant state by cross-linking proteins, primarily reacting with nucleophiles and other amines.
    1. Following Recipe 1, prepare the 3% glutaraldehyde solution.
    2. Remove the muscle tissue and immediately submerge it in 3% glutaraldehyde in a glass specimen vial and label accordingly (see Note 1).
    3. Place the specimen vials on a clinical rotator for 30 min at room temperature.
    4. After 30 min, quickly transfer the muscle to filter paper and cut it into the desired size (for mouse soleus, the muscle is cut in half for transverse and longitudinal sections, respectively) using a razor blade or scalpel with a light pulling motion to slice, do not crush tissue (see Note 2).
    5. As quickly as possible, place each new section of tissue into its own specimen vial filled with 3% glutaraldehyde and label the vials appropriately.
    6. Place the specimen vials on a clinical rotator and leave for 24 h at room temperature.

  2. Secondary fixation
    The purpose of secondary fixation with osmium tetroxide is two-fold, it cross-links lipids preserving their structure and it effectively stains the tissue by adding electron-dense material, enhancing contrast.
    1. Following Recipe 2, prepare 0.1 M sodium phosphate buffer pH 7.4.
    2. Remove the 3% glutaraldehyde from the specimen vials with a transfer pipette and transfer the solution to a proper waste container.
    3. Add phosphate buffer to the specimen vials and return the samples to the clinical rotator for 5 min.
    4. Dispose of the phosphate buffer in the proper waste container and repeat Step B3 twice more, on the third wash before disposing of the phosphate buffer, have your 1% osmium solution ready so that your sample does not dry out.
    5. Following Recipe 3, prepare 1% osmium tetroxide (see Note 3).
    6. Under a chemical fume hood with complete attention and care, dispense 1% osmium tetroxide into the specimen vials, cap the vials, and then return to the clinical rotator for 1 h.
    7. The specimens should appear black.
    8. Under a fume hood, remove the 1% osmium tetroxide solution from the specimen vials and dispose of the solution into the appropriate waste container.
    9. Dispose of the plastic caps in an appropriate waste container as well and then use new plastic caps on your specimen vials for subsequent steps.
    10. Add the phosphate buffer to the specimen vials and return the samples to the clinical rotator for 5 min.
    11. Dispose of the phosphate buffer in the proper waste container and repeat Step B10 twice more.

  3. Dehydration
    Water must be removed from the tissue through graded alcohol dehydration to allow for the infiltration of embedding media.
    1. Following Recipe 4, prepare all ethanol dilutions.
    2. Transfer 10% ethanol solution to the specimen vials and return them to the clinical rotator for 5 min.
    3. Remove 10% ethanol and repeat Step C2 with 25%, 50%, 75%, and 95% ethanol solutions.
    4. Transfer 100% ethanol to the specimen vials and return them to the clinical rotator for 5 min.
    5. Remove 100% ethanol and repeat Step C4 twice more.
    6. Under a chemical fume hood, transfer 100% propylene oxide to the specimen vials and cap the lids tightly (lids may pop off if not secured).
    7. Return the samples to the clinical rotator for 5 min.
    8. Remove the 100% propylene oxide and dispose of it in the proper waste container.
    9. Repeat Steps C6, C7, and C8 twice more.

  4. Infiltration and embedding
    Propylene oxide is equally miscible in both alcohol and embedding media, thereby clearing the alcohol and facilitating infiltration of the embedding media into the depths of the tissue.
    1. Following Recipe 5, prepare the embedding media.
    2. In a chemical fume hood, prepare a 2:1 (propylene oxide:embedding media) solution, the total volume you make will depend on how many samples you have (you want enough solution to completely cover your sample, even when rotating).
    3. Transfer 2:1 (propylene oxide:embedding media) solution to the specimen vials, cap them, and place them on the clinical rotator for 1 h (see Note 4).
    4. Remove 2:1 solution from the specimen vials and dispose of it in a plastic waste container.
    5. In a chemical fume hood, prepare a 1:1 (propylene oxide:embedding media) solution, the same total volume you used with the 2:1 solution can be used here as well.
    6. Transfer 1:1 (propylene oxide:embedding media) solution to the specimen vials, cap them, and return the vials to the clinical rotator. Leave the samples on the clinical rotator overnight and store the remaining embedding media in the -20 °C freezer.
    7. The next day, dispose of the 1:1 solution in the same waste container and bring the embedding media to room temperature.
    8. In a chemical fume hood, prepare a 1:2 (propylene oxide:embedding media) solution, using the same total volume as above.
    9. Transfer 1:2 (propylene oxide:embedding media) solution to the specimen vials cap them, and place them on the clinical rotator for 1 h.
    10. Remove 1:2 solution from the specimen vials and dispose of it in the designated waste container.
    11. Transfer 100% embedding media to the specimen vials and return them to the rotator for 1 h.
    12. Remove 100% embedding media and dispose of it in the designated waste container.
    13. Transfer 100% embedding media to the specimen vials, leave the lids off, and place in a vacuum oven pressurized to 20 PSI for 1 h set to room temperature.
    14. Specimens are now ready for curing and there is no need to remove the remaining 100% media from the vials, as the specimens will be transferred directly from the vials to the mold.

  5. Curing
    The viscous embedding media must be cured so that the block containing the tissue becomes hard enough for ultrathin sectioning.
    1. Set a vacuum oven to 60 °C and 0 PSI, then confirm the temperature with a thermometer before proceeding.
    2. With a pencil, write sample information on a small piece of paper that can fit into the base of the mold to be cured with the sample (will be visible through the cured plastic).
    3. Fill the individual wells of the plastic mold halfway with 100% embedding media using a wood applicator to drizzle the media into the wells of the mold.
    4. With fine negative-action tweezers, transfer the piece of paper with sample information into the bottom portion of the mold (the part that is farthest away from where the sample will rest).
    5. Next, with fine negative-action tweezers, transfer the blackened, osmicated (hard) sample from the specimen vial to the mold (see Note 5).
    6. Orient the sample accordingly to ensure that the sample can be sectioned in a cross-section or in a longitudinal section with ease (see Figure 6 for schematic on how it should look in the block).
    7. Repeat Steps E4, E5, and E6 with the remaining samples.
    8. Transfer enough 100% embedding media to fully fill the wells.
    9. Ensure that the samples are correctly oriented one last time, as adding media can distort their position slightly.
    10. Carefully, place the plastic mold into the oven to cure for at least 24 h (see Note 6).

  6. Glass knife fabrication
    Fabricating your own glass knives is a cost-saving technique, allowing you to preserve and extend the life of your expensive diamond knife for only cutting ultrathin sections. Pre-fabricated glass knives can be purchased if this equipment is not available at your institution.
    1. Wearing nitrile gloves, clean your glass strip (6.4 x 25 x 400 mm) with detergent and a brush and then rinse it thoroughly with tap water (see Note 7).
    2. Using a squirt bottle, rinse another time with distilled water.
    3. Leave the glass strip on paper towels to dry in a relatively clean, dust-free environment.
    4. Retract the Rear Glass Holder (Figure 2A) by rotating the Rear Glass Holder Knob (Figure 2A) and pull out the knob to lock it in place.
    5. Ensure that the Breaking Knob (Figure 2B) on the front is turned fully counterclockwise.
    6. Rotate the Locking Lever (Figure 2A) back to the rear position, fully raising the scorer.
    7. Push the Scoring Shaft (Figure 2B) all the way in.
    8. Make sure that the Score Selector (Figure 2B) above the Scoring Knob is set to.
    9. Transfer a cleaned, dry glass strip to the glass knife maker and place it flush against the White Guide Plate (Figure 2B) and then push the strip against the first Lateral Arresting Stud (Figure 2C) (used for 25 mm wide strips).
    10. While holding the glass strip in place, swing the Locking Lever gently forward until the Vertical Support Studs (Figure 2C) come into contact with the glass strip.
    11. Remove your hands and then press the Locking Lever down further until there is significant resistance.


      Figure 2. Glass knife maker with labeled parts. A. Angled side view with labeled parts. B. Angled frontal view with labeled parts. C. Close-up of angled side view with labeled parts.

    12. Place the Glass Fork (Figure 2A) under the left side of the glass strip to catch your glass square.
    13. Pull the glass Scoring Shaft Knob in a swift fluid, smooth movement across the face of the glass strip until it stops to score the glass (Figure 3A).
    14. Rotate the Breaking Knob clockwise until the glass breaks, you will be able to visually and audibly sense this.
    15. Reset the Breaking Knob fully counterclockwise.
    16. Rotate the Locking Lever back to the rear position, fully raising the scorer.
    17. Push the Scoring Shaft Knob back in completely.
    18. Clean off the glass break area with a brush.
    19. Place the Dampening Pressure Adjustment Lever (Figure 2C) on the dot.
    20. Adjust the Forward and Rear Glass Holder adjustment knobs to fit the glass square.
    21. Place the square piece of glass that you just made at a 45° angle onto the Breaking Pins and leave the Glass Fork underneath the square (Figure 3B).
    22. Disengage the Rear Glass Holder Knob from Step F4 by pressing it in and then rotating it clockwise to bring the Rear Glass Holder in contact with the corner of the glass square.
    23. The glass square should sit with its corners between the forward and rear glass holders.
    24. Gently pull the Locking Lever to the forward position until the Support Studs come into contact with the glass square.
    25. Press the Locking Lever down further until there is significant resistance.
    26. Pull the Scoring Shaft Knob in a swift fluid, smooth movement across the face of the glass square until it stops to score the glass.
    27. Rotate the Dampening Pressure Adjustment Lever until the damping pad contacts the corner of glass in contact with the Front Glass Holder.
    28. Rotate the Breaking Knob clockwise until the glass breaks; you will be able to visually and audibly hear this.
    29. Reset the Breaking Knob fully counterclockwise.
    30. Rotate the Locking Lever back to the rear position, fully raising the scorer.
    31. Push the Scoring Shaft Knob back in completely.
    32. Reset the Dampening Pressure Adjustment Lever back to the dot.
    33. Rotate the Rear Glass Holder Knob counter-clockwise to retract the Rear Glass Holder away from the glass triangles and pull the knob out to lock it in place.
    34. Use the Glass Fork to lift the left and right glass knives up and out of the knife maker and place the knives in a glass knife holder.
    35. Clean off the glass break area with a brush.
    36. Repeat Steps F1-F35 to make more glass knives.
    37. To attach a plastic boat to the knife, heat some dental wax in a glass beaker on a hot plate to set to 100 °C.
    38. Once the wax has fully melted, use a wood applicator to apply wax to the edges of the plastic boat and immediately align the plastic boat onto the back of your glass knife (see Note 8).
    39. With the wood applicator, transfer more melted wax to the edges of the attached boat to seal completely (Figure 3D).


      Figure 3. Fabrication of glass knives. A. Glass strip after being scored and broken into a glass square; B. Glass square aligned between the rear and forward glass holders with the dampening pressure adjustment lever adjusted and the glass fork underneath to catch the glass; C. Glass square, prior to scoring with the scorer above, being pressed onto the breaking pins with the locking lever pressed down fully; D. Several glass knives without boats and one with a plastic boat attached with dental wax in the glass knife holder.

    40. Test the wax seal by transferring distilled water with a transfer pipette into the plastic boat, creating a convex meniscus of water.
    41. Leave the glass knife with the plastic boat for 5 min and then check the water level to determine if there are leaks.
    42. If leak-free, empty water and store it in your glass knife holder for future use.

  7. Trim block
    Trimming the block into the trapezoidal pyramid shape enhances the thick and thin sections you are able to produce by reducing the frictional and resistive forces on your knives.
    1. Remove the plastic mold from the oven (from Step E10) and let it cool for at least 30 min.
    2. Once cool, place the plastic block with your tissue sample into the Specimen Holder (Figure 4A) that is set into the Specimen Holder Adapter (Figures 4A and 5A).


      Figure 4. Reichert-Jung Ultramicrotome with specimen holder in position to be trimmed and knife holder in position to cut specimens. A. Specimen holder clamped into the adaptor and guide track for trimming of the embedded specimen. B. This is what the specimen block looks like while looking through the binocular eye pieces. C. The knife holder with a diamond knife is clamped into place and adjusted for cutting thin sections. D. This is what the diamond knife edge looks like while looking through the binocular eye pieces.

    3. Clamp the Specimen Holder and Adapter (Figure 4A) to ensure that the specimen block does not move.
    4. Press the green On-Off Main Switch (Figure 5A) on the power supply to the ultramicrotome and switch on the lights using the Multi-Position Illumination Switch (Figure 5C), both the overhead light and the light under the stage will turn on.
    5. Bring the specimen into focus by turning the Focus Knob (Figure 5D) while looking through the Wide-Field Binocular Eyepieces.


      Figure 5. Reichert-Jung Ultramicrotome with labeled parts. A. Power supply; B. Front view of the ultramicrotome; C. Angled left side view; D. Angled right side view.

    6. The objective is to trim the specimen block into a trapezoidal pyramid that is as small as possible while including your specimen in the center (see Note 9) (Figure 6).


      Figure 6. The schematic view of how to trim the blocks prior to sectioning. A. A fully cured block with an osmicated muscle sample aligned to be cut in cross-section should appear like this. From the top, looking down the osmicated muscle will look relatively less dark due to the layer of plastic between the tissue and the top of the block. Using a razor blade, the top of the block face needs to be trimmed down to expose the tissue sample. B. Initially, the trimmed layers will feature plastic only and eventually you will begin to see the black tissue in the trimmed layer with surrounding plastic once the tissue is exposed. Next, trim the plastic around the sample block to create a trapezoidal pyramid. The sample block is now ready to be faced using a glass knife.

    7. While looking through the Wide-Field Binocular Eyepieces (Figure 5B) and using a razor blade, trim the face of the block until you bring your sample into plane. You should see the lighter-colored resin and then the dark shadow of your sample in the trimmed slices that you are creating.
    8. Next, while looking through the Wide-Field Binocular Eyepieces, use a razor blade to remove excess resin from all four sides of the block around your sample to sculpt your pyramid.
    9. Unclamp the Specimen Holder from the Adapter, place the Specimen Holder onto the Specimen Arm (Figures 5C and 5D) of the ultramicrotome and use the screw to secure the Specimen Holder tightly into place.
    10. Unclamp and remove the Specimen Holder Adapter, place your Knife Holder (Figures 4C and 5B) onto the ultramicrotome stage, and clamp the Knife Holder into place.
    11. Make sure that the Clearance Angle Adjustment Knob (Figure 4C) is adjusted to the proper clearance angle, which should correspond to the angle written on the box that came with your diamond knife (6°).
    12. Use the Knife Clamping Screw (Figure 4C) to clamp a glass knife onto the Knife Holder.
    13. Use the East-West Stage Control Knob (Figure 5C) and the North-South Knife Feed Knob (Figure 5B) to bring the knife holder with your glass knife close to your block, without touching.
    14. Adjust your Specimen Holder so that the leading edge of your sample (the part that is cut first and the base of your pyramid) is parallel with the knife edge.
    15. Using the North-South Knife Feed Knob on the ultramicrotome, slowly bring the glass knife closer to the sample block. Your ultramicrotome is equipped with a light that shines up from the base, illuminating the gap between the knife and the block face. As you bring the knife closer to the block face, this illuminated area will dim more and more, signifying that the two surfaces are almost touching.
    16. Set the Manual Fine-Feed Knife Control Knob (Figure 5C) to 1 μm.
    17. While looking through the Wide-Field Binocular Eyepieces, rotate the Manual Fine-Feed Knife Control Knob to advance the sample to the knife edge while rotating the Handwheel (Figure 5D) for the Specimen Arm for a full cycle.
    18. Repeat Step G17 until you begin to see sections of the sample face coming off onto the edge of the glass knife and then continue until the full face of your sample is being sectioned.
    19. Now, the face is trimmed, smooth, and ready for thick sections.
    20. Use the North-South Knife Feed Knob to move the Knife Holder away from your sample.
    21. Unscrew the Knife Clamping Screw to loosen your glass knife and remove from the Knife Holder.

  8. Cut thick sections
    The purpose of cutting thick sections and then staining them is to confirm the orientation of the tissue sample before proceeding.
    1. Following Recipe 6, prepare the 1% toluidine blue stain.
    2. Turn on a hot plate to approximately 100 °C.
    3. Place a glass knife with a plastic boat and tighten the Knife Clamping Screw (Figure 4C) to clamp the knife into place.
    4. Use the East-West Stage Control Knob (Figure 5C) and the North-South Knife Feed Knob (Figure 5B) to bring the knife holder with your glass knife close to your block, without touching.
    5. Adjust your Specimen Holder (Figure 5D) so that the leading edge of your sample (the part that is cut first and the base of your pyramid) is parallel with the knife edge.
    6. Using the North-South Knife Feed Knob on the ultramicrotome, slowly bring the glass knife closer to the sample block. Your ultramicrotome is equipped with a light that shines up from the base, illuminating the gap between the knife and the block face. As you bring the knife closer to the block face, this illuminated area will dim more and more, signifying that the two surfaces are almost touching.
    7. Fill the plastic boat with distilled water until the water is convex.
    8. Place the Suction Syringe (Figure 5D) into the boat beneath the water line.
    9. While looking through the Wide-Field Binocular Eyepieces (Figure 5B), use the Suction Knob (Figure 5D) to adjust the water level until you see an even silver reflection over the surface of the water (see Notes 10 and 11) (Figure 7).


      Figure 7. Views through binocular lens showing proper water level in the diamond knife prior to sectioning. The convex surface of water overfilling the diamond knife boat (A). Using the ultramicrotome syringe, the water level is lowered until there is a silvery, reflective surface as shown (B). After lowering the water and attempting to cut sections, if any water collects on the face of the block, the water may need to be lowered slightly further as shown (B), where you can see a slight shadow forming at the knife edge. However, if the water is lowered too much, it will pull away from the knife edge and will need to be refilled.

    10. Set the Manual Fine-Feed Knife Control Knob (Figure 5C) to 1 μm.
    11. While looking through the Wide-Field Binocular Eyepieces, rotate the Manual Fine-Feed Knife Control Knob to advance the sample to the knife edge while rotating the Handwheel for the Specimen Arm (Figure 5D) for a full cycle.
    12. Repeat Step H17 until you begin to see sections of the sample face coming off the edge of the glass knife onto the surface of the water and then continue until the full face of your sample is being sectioned (see Note 12).
    13. Once you get a full-faced section, use a metal loop to fish out the part-sections and the full section you just cut and discard them by rotating the metal loop in a beaker of distilled water.
    14. Set the Manual Fine-Feed Knife Control Knob to 0.5 μm.
    15. While looking through the Wide-Field Binocular Eyepieces, rotate the Manual Fine-Feed Knife Control Knob to advance the sample to the knife edge while rotating the Handwheel for the Specimen Arm for a full cycle.
    16. Discard the first couple of sections into the beaker of water.
    17. Cut 5-10 more 0.5 μm sections.
    18. Label two glass slides with the pertinent sample information and apply a drop (~2 cm diameter) of distilled water to the slide.
    19. Use the metal loop to transfer individual sections to the drop of water on the glass slide and rotate the loop to transfer.
    20. After you finish distributing the sections between the two glass slides, place the glass slides onto the hot plate (see Note 13).
    21. After the water has fully evaporated, the sections are ready to be stained.
    22. While the slides are still on the hot plate, add 1% toluidine blue stain drop-wise to completely cover the dried sections and leave for 2 min (see Note 14).
    23. Wash the slides with running water to remove as much excess stain from the slides as possible and allow them to dry.
    24. After allowing the slides to dry, view the sections with a light microscope (see Notes 15 and 16).
    25. If satisfied with the orientation of the sections, then no adjustment is necessary and you can move on to cutting thin sections.
    26. Use the North-South Knife Feed Knob to move the Knife Holder away from your sample.
    27. Unscrew the Knife Clamping Screw to loosen your glass knife and remove it from the Knife Holder.

  9. Cut thin sections
    In order to form a high-resolution image, the electron beam must be able to penetrate the section without a significant loss of speed, requiring an ultrathin section.
    1. Place a diamond knife onto the Knife Holder (Figure 5B) and tighten the Knife Clamping Screw (Figure 4C) to clamp the knife into place.
    2. Use the East-West Stage Control Knob (Figure 5C) and the North-South Knife Feed Knob (Figure 5B) to bring the knife holder with your glass knife close to your block, without touching.
    3. Adjust the Specimen Holder (Figure 5D) so that the leading edge of your sample (the part that is cut first and the base of your pyramid) is parallel with the knife edge.
    4. Using the North-South Knife Feed Knob on the ultramicrotome, slowly bring the glass knife closer to the sample block. Your ultramicrotome is equipped with a light that shines up from the base, illuminating the gap between the knife and the block face. As you bring the knife closer to the block face, this illuminated area will dim more and more, signifying that the two surfaces are almost touching.
    5. Fill the diamond knife boat with distilled water until the water is convex.
    6. Place the Suction Syringe (Figure 5D) into the boat beneath the water line.
    7. While looking through the Wide-Field Binocular Eyepieces (Figure 5B), use the Suction Knob (Figure 5D) to adjust the water level until you see an even silver reflection over the surface of the water (Figure 7).
    8. Set the Manual Fine-Feed Knife Control Knob (Figure 5C) to 0.5 µm.
    9. While looking through the Wide-Field Binocular Eyepieces, rotate the Manual Fine-Feed Knife Control Knob to advance the sample to the knife edge while rotating the Handwheel for the Specimen Arm (Figure 5D) for a full cycle.
    10. Repeat Step I9 until you begin to see sections of the sample face coming off the edge of the glass knife onto the surface of the water and then continue until the full face of your sample is being sectioned (see Note 17).
    11. Once you get a full-faced section, use a metal loop to fish out the part-sections and the full-faced section you just cut and discard by rotating the metal loop in a beaker of distilled water.
    12. Next, on the power supply, set the value on the Semi-Thin Sectioning Control (Figure 5A) to 0.50 μm.
    13. On the power supply, adjust the Cutting Speed Control (Figure 5A) to 1.0 mm/sec (this corresponds to the speed that the ultramicrotome arm uses on the down stroke).
    14. Press down on the Motor Drive Control Lever (Figure 5D) that begins the automated advancement and rotation of the Specimen Arm.
    15. Look through the Wide-Field Binocular Eyepieces and observe the full-faced sections coming off the knife (see Note 18).
    16. After 3-4 sections, look through the Wide-Field Binocular Eyepieces and once the Specimen Arm completes its down stroke sending a new section into the boat, adjust the Semi-Thin Sectioning Control to 0.40 µm.
    17. Use a metal loop to fish out the sections you just cut and discard them by rotating the metal loop in a beaker of distilled water.
    18. After 3-4 sections, look through the Wide-Field Binocular Eyepieces and once the Specimen Arm completes its down stroke sending a new section into the boat, adjust the Semi-Thin Sectioning Control to 0.30 µm.
    19. Use a metal loop to fish out the sections you just cut and discard them by rotating the metal loop in a beaker of distilled water.
    20. When observed in the binocular lenses, sections after this adjustment should begin to display colors that correspond to the approximate thickness of the section and the refractive index of the embedding material. At this point, they will most likely be yellow.
    21. After 3-4 sections, look through the Wide-Field Binocular Eyepieces and once the Specimen Arm completes its down stroke sending a new section into the boat, adjust the Semi-Thin Sectioning Control to 0.25 µm.
    22. Use a metal loop to fish out the sections you just cut and discard them by rotating the metal loop in a beaker of distilled water.
    23. When observed in the Wide-Field Binocular Eyepieces, sections after this adjustment should begin to look yellow to blue.
    24. After 3-4 sections, look through the Wide-Field Binocular Eyepieces and once the Specimen Arm completes its down stroke sending a new section into the boat, adjust the Semi-Thin Sectioning Control to 0.20 µm.
    25. Use a metal loop to fish out the sections you just cut and discard them by rotating the metal loop in a beaker of distilled water.
    26. When observed in the Wide-Field Binocular Eyepieces, sections after this adjustment should begin to look blue to purple.
    27. After 3-4 sections, look through the Wide-Field Binocular Eyepieces and once the Specimen Arm completes its down stroke sending a new section into the boat, adjust the Semi-Thin Sectioning Control to 0.17 µm.
    28. Use a metal loop to fish out the sections you just cut and discard them by rotating the metal loop in a beaker of distilled water.
    29. When observed in the Wide-Field Binocular Eyepieces, sections after this adjustment should begin to look purple.
    30. After 3-4 sections, look through the Wide-Field Binocular Eyepieces and once the Specimen Arm completes its down stroke sending a new section into the boat, adjust the Semi-Thin Sectioning Control to 0.14 µm.
    31. Use a metal loop to fish out the sections you just cut and discard them by rotating the metal loop in a beaker of distilled water.
    32. When observed in the Wide-Field Binocular Eyepieces, sections after this adjustment should begin to look purple to gold.
    33. After 3-4 sections, look through the Wide-Field Binocular Eyepieces and once the Specimen Arm completes its down stroke sending a new section into the boat, adjust the Semi-Thin Sectioning Control to 0.12 µm.
    34. Use a metal loop to fish out the sections you just cut and discard them by rotating the metal loop in a beaker of distilled water.
    35. When observed in the Wide-Field Binocular Eyepieces, sections after this adjustment should begin to look gold.
    36. After 3-4 sections, look through the Wide-Field Binocular Eyepieces and once the Specimen Arm completes its down stroke sending a new section into the boat, adjust the Semi-Thin Sectioning Control to 0.11 µm.
    37. Use a metal loop to fish out the sections you just cut and discard them by rotating the metal loop in a beaker of distilled water.
    38. When observed in the Wide-Field Binocular Eyepieces, sections after this adjustment should begin to look purely gold (see Note 19).
    39. If satisfied, then continue cutting these sections until you have your desired quantity and continue to Step I47. If not satisfied, then continue to Step I40.
    40. After 3-4 sections, look through the Wide-Field Binocular Eyepieces and once the Specimen Arm completes its down stroke sending a new section into the boat, adjust the Semi-Thin Sectioning Control to 0.10 µm.
    41. Use a metal loop to fish out the sections you just cut and discard them by rotating the metal loop in a beaker of distilled water.
    42. When observed in the Wide-Field Binocular Eyepieces, sections after this adjustment should begin to look gold to silver.
    43. After 3-4 sections, look through the Wide-Field Binocular Eyepieces and once the Specimen Arm completes its down stroke sending a new section into the boat, adjust the Semi-Thin Sectioning Control to 0.09 µm (90 nm).
    44. Use a metal loop to fish out the sections you just cut and discard them by rotating the metal loop in a beaker of distilled water.
    45. When observed in the Wide-Field Binocular Eyepieces, sections after this adjustment should begin to look silver.
    46. Continue cutting sections until you are satisfied, typically 15-20 homogenously colored sections are what to aim for.
    47. Use the North-South Knife Feed Knob to move the Knife Holder away from your sample.
    48. Use the negative-action forceps to pick up a copper grid with your dominant hand and place the eyelash manipulator tool in your non-dominant hand.
    49. While looking through the Wide-Field Binocular Eyepieces, bring these tools into view.
    50. Submerge the grid under the surface of the water and begin to advance it directly under your section of choice (see Note 20).
    51. At the same time, slightly submerge your eyelash manipulator tool next to your section of choice and gently guide the section over your grid.
    52. Hold the section in place as best as you can and slowly raise the grid beneath the section until the section rests relatively centered on top of the grid and remove from the water.
    53. With your section-mounted grid still resting in the forceps, take a triangle of filter paper and gently lower it next to the outer rim of the grid. Water should begin transferring from the grid to the filter paper (see Note 21).
    54. After the water is removed as best that you can, place the grid into the grid box.
    55. Continue collecting sections from the boat of your diamond knife, typically 10 grids for each sample is plenty.
    56. Allow grids to fully dry in your grid box overnight prior to staining.

  10. Stain grids
    Staining the sections with uranyl acetate and lead citrate enhances the contrast for countless cellular structures.
    1. Turn on a hot plate/stirrer and adjust the temperature to ~200 °C.
    2. Place a 1 L beaker filled with distilled water and a magnetic stir bar on top of the hot plate (Figure 8).


      Figure 8. Setup for staining grids. A. General equipment for staining grids. B. A close-up showing the use of sodium hydroxide pellets with two grids being stained with lead citrate, the lid is closed immediately after floating the grids on drops of lead citrate to prevent precipitation of lead citrate.

    3. Bring the water to a boil and then turn off the hot plate while leaving the stirrer on (see Note 22).
    4. Distribute the water into four 50 ml beakers and allow to cool to ~25 °C (see Note 23).
    5. Following Recipe 7, prepare 1% uranyl acetate solution.
    6. Following Recipe 8, prepare 1 N-NaOH solution.
    7. Following Recipe 9, prepare Reynold’s lead citrate solution.
    8. Use a 1 ml syringe, with the needle removed and a 0.22 μm syringe filter attached, to distribute a drop of uranyl acetate into a glass Petri dish, with the bottom filled with dental wax (see Note 24).
    9. With negative-action forceps, place your grid, section-side down, onto the drop of uranyl acetate.
    10. Set a timer for 30 min and cover the Petri dish with something that blocks light (see Note 25).
    11. After 30 min, use negative-action forceps to pick up the grid and then dunk the grid into one of the 50 ml beakers filled with previously boiled, distilled water. Dunk the grid into and out of the water ~30 times.
    12. Advance to the next 50 ml beaker filled with previously boiled, distilled water and dunk the grid into and out of the water ~30 times (use these same two beakers for subsequent uranyl acetate washes).
    13. Set the negative-action forceps onto the benchtop and apply a triangle of filter paper gently to the outer rim of the copper grid to remove water from the surface of the grid.
    14. Use a second glass Petri dish, with dental wax in the bottom, and place pellets of sodium hydroxide around the area of the dish where you will add drops of lead citrate. Close the lid immediately while preparing the lead citrate.
    15. Use a 1 ml syringe, with the needle removed and a 0.22 µm syringe filter attached, to distribute a drop of lead citrate onto the bottom of the glass Petri dish. Cover immediately.
    16. Transfer the grid that was resting in the negative-action forceps section-side down onto the drop of lead citrate. Close immediately and set a timer for 5 min.
    17. After 5 min, use negative-action forceps to pick up the grid and then dunk the grid into one of the 50 ml beakers filled with previously boiled, distilled water. Dunk the grid into and out of the water ~30 times.
    18. Advance to the next 50 ml beaker filled with boiled, distilled water and dunk the grid into and out of the water ~30 times (use these same two beakers for subsequent lead citrate washes).
    19. Set the negative-action forceps onto the benchtop and apply a triangle of filter paper gently to the outer rim of the copper grid to remove water from the surface of the grid.
    20. Transfer the grid back into the grid box and allow it to dry for at least a couple of hours prior to viewing in the electron microscope.

  11. Capturing images with the transmission electron microscope
    The goal of imaging with the transmission electron microscope is to provide high-resolution images of skeletal muscle and mitochondrial ultrastructure in an unbiased manner, taking enough images to get a true sense of the microanatomy.
    1. Press the Panel On-Off Knob (Figure 9D) to illuminate the screen displaying the microscope settings.
    2. Press the HT (High Tension) button (Figure 9D) where a green light above the button will illuminate and check the emission gauge to ensure that this returns closer to ~0 before proceeding.
    3. Turn the Filament Knob (Figure 9D) clockwise ~23 steps with associated clicks as you turn the knob to reach saturation. This will take approximately 2 min and the scope will beep when this occurs. The viewing screen of the microscope should be illuminated in bright green light (see Note 26).
    4. Remove the Specimen Holder (Figure 9B) from the microscope by rotating the holder approximately a quarter turn clockwise and then pull it out completely.
    5. Place the Specimen Holder onto the Specimen Holder Stand (Figure 9F) and use the clip tool to raise the specimen clip at the end of the Specimen Holder (see Note 27).


      Figure 9. Phillips CM10 Transmission Electron Microscope. A and B. A general overview of the electron microscope and its parts. C. The left control panel is mainly used to control the intensity of the electron beam and for adjusting the screens for focusing images with the objective lenses or capturing images by lifting the fluorescent screen. D. The right control panel houses most of the functions needed for adjusting magnification, focusing, and the screen for monitoring the electron microscope system as a whole. E. The vacuum screen shows a schematic of the vacuum system with the various pumps, valves, and pressure gauges used to monitor the performance of the system. F. The specimen holder on the specimen holder stand with the clip opening tool inserted and the grid clip lifted.

    6. Use the negative-action forceps to pick up and transfer your grid from the grid box to the circular grid holder at the bottom of the Specimen Holder, ensuring that the grid rests completely within the area and is not resting slightly outside.
    7. Use the clip tool to lower the clip down onto your grid, holding the grid in place.
    8. Insert the Specimen Holder halfway into the scope and stop. A red Pre-vacuum Indicator Light (Figure 9B) will turn on and you will hear the pre-vacuum click on.
    9. Wait for the red Pre-vacuum Indicator Light to go out and then continue to insert the Specimen Holder fully into the high vacuum chamber by gently rotating the holder counter-clockwise and the vacuum will literally pull the specimen holder in (see Note 28).
    10. The grid will now be in view on the viewing screen with a low magnification.
    11. Move around your viewing area to find your sample area of interest by rotating the two grips (-x and -y) (Figure 9C).
    12. Once in the desired area, increase the magnification with the Magnification Knob (Figure 9D). The screen will display your current level of magnification.
    13. As you zoom in, you will have to adjust the viewing area with the two grips to keep it centered as much as possible.
    14. After the magnification is increased to ~500x, there will be an audible beep prompting you to flip the Condenser Aperture Lever (Figure 9B), switching from the low magnification aperture to high magnification.
    15. As you continue to increase the magnification, the muscle section will begin to come into view; however, the screen will get darker and darker because you have not increased the intensity of the electron beam (see Note 29).
    16. As you zoom in, gently adjust the electron beam intensity by rotating the Intensity Knob (Figure 9C) so that you can see the muscle ultrastructure clearly.
    17. Bring the muscle ultrastructure into focus with the Focus Knob (Figure 9D).
    18. Turn on the computer monitor and click on the AMT software icon.
    19. Before switching on the software and lifting the viewing screen, as a rule of thumb, adjust the intensity of the electron beam to 0.500, which is displayed on the screen next to ‘meter’ (see Note 30).
    20. Lift the Fluorescent Screen (Figure 9C) using the lever and click on ‘Click for Live Image’, a gray-scale image of your sample will appear on the monitor (See Note 31).
    21. The software is automatically set to ‘QualityLive’ and is in ‘Survey’ mode.
    22. If you want to adjust the focus slightly better, click on the ‘Focus’ button that is under the ‘Survey’ button in the software, which will zoom in the image allowing you to get a finer focus.
    23. Click the ‘Survey’ button to return to the preview screen.
    24. If satisfied, click on ‘Final Image’.
    25. Name your image with the relevant information and save the image in a designated folder.
    26. Once finished taking images, close the software and lower the Fluorescent Screen using the lever.
    27. Decrease your zoom with the magnification knob, while rotating the electron beam intensity counter-clockwise to compensate.
    28. An audible beep ~500x will prompt you to flip the Condenser Aperture Lever from the high magnification aperture back to the low magnification one.
    29. Continue to decrease the magnification to ~75x and then re-center the viewing area with the -x and -y axis grips.
    30. Remove the Specimen Holder from the high vacuum chamber and remove your grid.
    31. Place the specimen holder back into the high vacuum chamber exactly as was described above, waiting for the pre-vacuum to kick off before inserting fully.
    32. Rotate the filament knob counter-clockwise until the viewing screen is no longer illuminated and there will be an audible beep.
    33. Depress the high-tension button and the green light will go off.
    34. Pull out the panel dim knob to turn off the screen.

Data analysis

For most of our studies focused on the effect of knocking out or knocking in a signaling molecule and subjecting the muscle to various conditions that induce hypertrophy or atrophy, we perform TEM analysis on a minimum of n = 3 samples per group. We subjected TRAF6ff and TRAF6mko mice to denervation of the sciatic nerve for 10 days and then utilized TEM to study the ultrastructure of the TA muscle between these groups (Paul et al., 2010). With TEM, we were able to complement our molecular biological experimental results by showing a reduction in atrophy and autophagy in the denervated TRAF6mko mice when compared to control TRAF6ff mice. Ultrastructurally, it was noted that the denervated muscle of TRAF6ff mice showed disorganization of SS and IMF mitochondria and an increase in autophagic vacuoles with the fusion of mitochondria to autophagosomes; whereas, this phenotype was largely absent in the denervated muscle of TRAF6mko mice (Paul et al., 2010). In another study, we looked at the role of transgenic overexpression of the TWEAK cytokine in mice and used TEM to evaluate the ultrastructure of the soleus muscle in the naïve condition. It was found that there was a drastic decrease in the size and number of mitochondria in the TWEAK-TG mice when compared to wild-type littermates at 6 months of age (Hindi et al., 2014). This was quantified by counting the SS mitochondria in three samples per group utilizing at least 10 pictures per sample magnified at 2,200x. This was repeated with the IMF mitochondria (Hindi et al., 2014). We present the data as mean ± standard deviation (SD) and use a paired Student’s t-test to determine statistical differences among the different groups with a P < 0.05 being considered as statistically significant. Most recently, we used TEM to qualitatively compare the mitochondrial ultrastructure in the soleus muscle of Tak1fl/fl mice to those of muscle-specific knockout or Tak1mKO mice. We found that the Tak1mKO mice had an abundance of vacuolated mitochondria and an increased proportion of enlarged mitochondria with disturbances in the structure of their cristae when compared to the Tak1fl/fl mice (Hindi et al., 2018). Our TEM observations were combined with biochemical analyses conducted between these groups to conclude that TAK1 is required for mitochondrial homeostasis in the skeletal muscle of these mice. Please see Figure 10 for examples of electron micrographs in the subsarcolemmal and intermyofibrillar regions of mouse skeletal muscle.


Figure 10. TEM micrographs of mouse skeletal muscle mitochondria. A. Intermyofibrillar mitochondria from sagittal sections (Scale bar = 1 μm); B. Subsarcolemmal mitochondria from sagittal sections (Scale bar = 1 μm); C. Healthy subsarcolemmal mitochondria with well-defined, uniform cristae (Scale bar = 500 nm); D. Several enlarged subsarcolemmal mitochondria with non-uniform cristae (Scale bar = 500 nm).

Notes

  1. Any mouse skeletal muscle tissue can be used for this analysis. Typically, in the skeletal muscle field, the mouse hind limb muscles are isolated for study, each muscle with its own characteristic fiber type composition. We use the mouse tibialis anterior (TA) and mouse soleus for TEM analyses due to their distinct differences in fiber type composition. The TA muscle has been characterized as a fast-glycolytic muscle, while the soleus has been characterized as a slow-oxidative muscle (Schiaffino et al., 2011; Kammoun et al., 2014). Therefore, it is necessary to compare the same muscle between experimental groups. For a helpful video on how to isolate hind limb muscles from mice, please see the video in Hindi et al., 2017. After isolating the muscle, it is imperative that you submerge the whole muscle into the glutaraldehyde as quickly as possible to preserve the muscle in its physiological state.
  2. It must be emphasized that you don’t need very much of your tissue sample for TEM. The mouse soleus muscle is a relatively small muscle when compared to the mouse TA muscle. For the soleus, we typically cut the muscle in half mid-belly and use one half for our cross-section analysis and the other half for our sagittal section analysis without the need to further trim the tissue down. On the other hand, the mouse TA muscle is much larger and needs to be trimmed down to a thin strip, similar in size to the diameter of the soleus. If you need help with this visualization, use your mouse soleus muscle as a comparison. From there, divide your strip into two parts, one for cross-section analysis and one for sagittal section analysis.
  3. Many of the chemicals in TEM tissue processing are highly toxic, especially osmium tetroxide. Osmium tetroxide is capable of fixing your corneas, so make sure that these steps are done in a properly ventilated chemical fume hood and personal protective equipment is worn.
  4. Ensure that the black samples remain mobile in the various dilutions and that they aren’t stuck to the side of the vial. This can be helped by adjusting the rotator speed to rotate more appropriately with the increasingly viscous solutions. After adding the next media solution, briefly shake or flick samples to dislodge them from the bottom or side.
  5. Try to avoid introducing bubbles into the block, especially around the sample itself. If there is a bubble at the bottom, then it’s not a big deal. If there is a bubble close to the sample, use a syringe needle to move and then lift it out of the mold.
  6. If not given enough time to cure, the plastic won’t be hard enough to yield proper thin sections. This will be realized immediately when trimming the block with your razor blade. If the plastic appears to be giving too much to the pressure of the blade or there is significant resistance when trimming as if the blade is sticking (not simply because the block is hard), then this is a good indication that the block could use additional time in the oven to finish curing. Simply add the blocks back to the oven and give them some more time. Alternatively, if the oven maintains the temperature well without fluctuation, there is no problem leaving the tissue blocks in for longer periods of time. Often, I have placed the blocks into the oven to cure over the weekend and then returned to trim them on Monday without issue.
  7. Do not touch the glass with your bare hands, the oils on your hands will ruin the quality of the glass when sectioning. In addition, if you feel that you need a supplement to go along with the instructions and figures, search for ‘LKB Type 7801B videos’ and there are a couple that may help illustrate these steps.
  8. Attaching the plastic boats to the glass knives with wax can be a challenge at first. Practice is important. Transferring enough wax to the edges of the plastic boat and then attaching the boat to the glass as fast as possible before the wax begins to harden is the difficult part.
  9. A trapezoidal pyramid is desired so that as the block encounters the diamond knife, there is a decrease in the surface area encountered by the blade as the blade cuts the full section. This relieves pressure/friction and ensures a smoother section with as little sectioning artifact as possible. This also preserves your knife edge for as long as possible. One video that does a pretty good job of demonstrating the trimming process that can be applied here is in Jenny, 2011.
  10. Adjusting the water level to the point where there is a silver reflection is difficult for the first time, but is noticeable. Practice moving the water up and down as you go from convex to silver reflection and then to concave as you see the water begin to pull away from the knife edge. Please refer to the video by Soplop et al., 2009 that is helpful for both thick and thin sectioning and shows how the sections float on the surface of the water in the knife boat as they are sectioned.
  11. If water has pulled away from the knife edge, the water level is too low and needs to be increased. Sometimes there is only a small portion of the knife where this has happened and in which case you can use a tool with an eye-lash adhered to the end to brush water up onto the edge.
  12. When you begin trying to cut sections, you may notice that water has pulled up onto the face of your sample block most likely via surface tension. If this happens, then you need to stop, dry off the face with a piece of filter paper and then adjust your water level a little lower. Sometimes it needs to be a little lower closer to the knife edge to where you see a shadow closer to the knife edge, but not to the point where the water has pulled off the edge.
  13. You don’t want the water on the hot plate to evaporate too quickly or else the sections will dry unevenly on the glass plate and will be bunch up in a wrinkly manner. If you notice this occurring, adjust the temperature so that they dry relatively slowly.
  14. Similarly, you don’t want the hot plate to be hot enough to where it boils on contact, which sends toluidine blue stain bubbling everywhere. Adjust accordingly. You want a ring of yellowish gold, dried stain to develop, ~2 min. At this point, you wash.
  15. With the light microscope, the main objective is to determine that your sections are oriented appropriately. You want to see that the section is free from obvious defects and that it is a true cross-section or longitudinal section.
  16. If the sections look more oblique in the light microscope, then you can adjust the chuck that is holding the block in the way that you think necessary to correct the sectioning angles. At this point, depending on how drastic your changes are, you may need to re-face the block with the original glass knife in the previous ‘trimming step’.
  17. When manual advancing the ultramicrotome arm and then rotating the arm to cut a section, ensure that this is a slow, smooth rotation. You don’t want to rotate the ultramicrotome arm to quickly or aggressively, else you may decrease the life of your diamond knife.
  18. If the sample pyramid is small enough and the face was aligned parallel to the edge of the diamond knife, the sections will come off the knife and form a ribbon of sections that don’t float away from one another. This can be helpful when you collect the sections with a grid. In addition, you may be wondering why we are using the Semi-thin Sectioning Control rather than the Ultra-thin Sectioning Control. Our ultramicrotome model is relatively old and the Ultra-thin Sectioning Control feature relies on a thermal advance feature that we found to be inaccurate, most likely due to the age of the device. While it may take longer as you slowly decrease the section thickness from 0.5 µm to 0.100 µm (100 nm), the quality and consistency of our sections improved. So feel free to try out your Ultra-thin Sectioning Control feature, which may save some time if it is working properly.
  19. Gold sections provide higher contrast, but not as much resolution. Silver sections provide high resolution, but with less contrast. Most of the time, gold sections are desired for skeletal muscle unless you’re trying to resolve difficult structures like the sarcoplasmic reticulum, T-tubules, etc. where silver sections may be more preferable.
  20. The ultrathin sections are extremely fragile and care should be taken with the hair tools to gently guide and not destroy your sections. It may take some time to get the fine dexterity to comfortably mount your thin sections on grids without frustration. Practice is the best way.
  21. When attempting to dry the grids with the triangle of filter paper, don’t place the filter paper directly onto your section. There is typically an outer ring of thicker copper where you should aim to rest the filter paper so that your section doesn’t get damaged.
  22. Boiling the distilled water removes carbon dioxide from the water. The goal here is to prevent the precipitation of our heavy metal stains that can leave our grids with aggregated clumps of metal that interfere with the image quality. This can be quite frustrating after getting this far into the process. Take every precaution.
  23. If you don’t allow your water to cool enough, you will notice that the contrast and staining looks relatively ‘flat’ when looking at the sections in the scope. Be patient and let it cool.
  24. Filtering your heavy metal stains directly before use will hopefully remove any aggregates that are in the solution and lead to better staining.
  25. Uranyl acetate is photosensitive and should be protected from light when in use and when not in use in dark colored bottles. Lead citrate is sensitive to carbon dioxide and should be stored in a bottle with a tight seal and parafilm to prevent aggregation. During the staining process, adding sodium hydroxide is a necessary step to remove as much carbon dioxide from the staining dish as possible while the reaction is taking place.
  26. This scope uses a tungsten filament, which is why the filament knob is turned that many steps and in quick succession. If there were a LaB6 filament, this would be a much slower, more gradual process. The biggest difference between these two is the longevity and the price of the filaments. Tungsten filaments are good for ~100 h of use, whereas the LaB6 filaments yield ~1,000+ h of use and are considerably more expensive.
  27. Be careful with the specimen holder and especially the specimen holder tool, which is very fragile and will break if too much force is applied. Be careful not to let your fingers come into contact with the lower, shiny half of the specimen holder. Oil from your hands will ruin the device if this is not done.
  28. After the red light goes off, you want to gently rotate the specimen holder with constant pressure and rotation. If this is done too quickly and aggressively, the pressure in this chamber may increase and the high tension may kick off as a safety mechanism. Then you would need to start over. If you want to monitor the pressure of the chamber when inserting the specimen holder, press the button next to where it says vacuum on the ready screen, where you will see a schematic of the vacuum systems and pressure values on the bottom. The Ion Getter Pump (IGP) values will correspond to the pressure in the chamber that you are inserting the specimen holder. Try to keep this as low as possible (< 50).
  29. Do not adjust the beam intensity too quickly or you may burn a hole in your sample grid. Always work on the right side (or clockwise) of crossover. If you rotate the intensity knob counter-clockwise, you will decrease the intensity and at a certain point you will be able to visualize the filament itself. This is known as crossover. If you continue to rotate the intensity knob counter-clockwise, it will get brighter and brighter. This would be the ‘left’ or ‘counter-clockwise’ side of crossover. We want to work on the right side where increasing the intensity is a result of rotating clockwise.
  30. This CM10 Phillips Electron Microscope has been retrofitted with a new camera and uses AMT software to acquire images. Adjusting the electron beam intensity to 0.500 ensures that the camera won’t be saturated or burned by a beam that is too intense. One can adjust the intensity accordingly once the image is being looked at in the software and it will indicate when saturation or low signal occurs by the histogram in the software.
  31. There is a fluorescent screen where your EM image is projected for manual adjustment by looking at it in the viewing screen, like looking through the objectives of a light microscope. This is an older electron microscope retrofitted with a computer and digital camera for imaging, allowing high resolution images. The camera is located at the bottom of the microscope and the beam is coming from above. The fluorescent screen needs to be flipped out of the way so that the digital camera can capture the final image.

Recipes

  1. 3% glutaraldehyde
    1. In the fume hood, crack open the scored 10 ml vial of 8% glutaraldehyde and empty contents into a 50 ml conical test tube
    2. Dilute to 3% glutaraldehyde by adding 16.6 ml of phosphate buffer
    3. Invert several times to mix and it’s ready for immediate use
  2. 0.1 M sodium phosphate buffer pH 7.4
    1. Measure 800 ml of distilled water in a graduated cylinder and add a magnetic stir bar
    2. Turn on the stirrer
    3. Weigh 3.1 g of sodium phosphate monobasic monohydrate (NaH2PO4·H2O) and add to the graduated cylinder
    4. Weigh 10.9 g sodium phosphate dibasic anhydrous (Na2HPO4) and add to the graduated cylinder
    5. Check the pH using a pH meter and ensure that it has a pH of 7.4
    6. Volume up to 1,000 ml and store in a glass bottle
  3. 1% osmium tetroxide
    1. Open the container containing the scored ampoules with 1 g of osmium tetroxide under the fume hood
    2. Prepare 100 ml of 0.1 M phosphate buffer in a clean tight-sealing, amber glass bottle
    3. Drop the scored ampoule into the glass bottle and using a clean glass rod, break open the ampoule into several pieces
    4. Seal the glass bottle immediately with a cap and swirl several times
    5. Allow the osmium crystals to dissolve completely at room temperature (can be left in the hood overnight)
    6. Once dissolved, store your amber bottle inside its own air-tight container (a metal can with lid or any air-tight container large enough to fit the amber bottle inside) and then place in a 4 °C refrigerator until ready to use
  4. Ethanol dilutions
    Prepare ethanol dilutions (10%, 25%, 50%, 75%, 95%, 100%):
    1. For 100%, pour 50 ml of 200 Proof ethanol into a 50 ml conical centrifuge tube and label accordingly
    2. For 95%, measure 47.5 ml of 200 Proof ethanol in a graduated cylinder and then transfer to a 50 ml conical centrifuge tube. Volume up to the 50 ml graduation with distilled water and label accordingly
    3. For 75%, measure 37.5 ml of 200 Proof ethanol in a graduated cylinder and then transfer to a 50 ml conical centrifuge tube. Volume up to the 50 ml graduation with distilled water and label accordingly
    4. For 50%, measure 25.0 ml of 200 Proof ethanol in a graduated cylinder and then transfer to a 50 ml conical centrifuge tube. Volume up to the 50 ml graduation with distilled water and label accordingly
    5. For 25%, measure 12.5 ml of 200 Proof ethanol in a graduated cylinder and then transfer to a 50 ml conical centrifuge tube. Volume up to the 50 ml graduation with distilled water and label accordingly
    6. For 10%, measure 5.0 ml of 200 Proof ethanol in a graduated cylinder and then transfer to a 50 ml conical centrifuge tube. Volume up to the 50 ml graduation with distilled water and label accordingly
  5. Embedding media
    1. Turn on the hot plate to 60 °C and place the two anhydrides, DDSA and NMA, on top to reduce their viscosity
    2. Measure 40 ml of EMbed 812 in a 50 ml conical centrifuge tube and then pour into a disposable urine specimen container
    3. Using a new 50 ml conical centrifuge tube, measure 17 ml of warmed DDSA and then pour into the same disposable urine specimen container
    4. Using a new 50 ml conical centrifuge tube, measure 26 ml of warmed NMA and pour into the same disposable urine specimen container
    5. Using a graduated transfer pipette, transfer 1 ml of DMP-30 to the urine specimen container
    6. Using the same transfer pipette, ensure that any remaining volume of EMbed 812, DDSA, and NMA is removed and transferred to the mixture in the urine specimen container
    7. Cap the 50 ml conical centrifuge tubes and discard
    8. Thoroughly stir the embedding mixture in the urine specimen container with a wooden applicator until homogenous
    9. The embedding media can be stored in the -20 °C freezer until ready to use
    10. If you’re ready, simply prepare propylene oxide solutions as needed and with respect to how many samples you are processing
  6. 1% toluidine blue stain
    1. Weigh 2 g of sodium borate and dissolve in 100 ml of distilled water
    2. Weigh 1 g of toluidine blue powder and dissolve in sodium borate solution
    3. Filter the stain solution using a syringe filter and ready for use 
  7. 4% uranyl acetate stock (aq)
    1. Wear appropriate personal protective equipment including at least N-95 respirator
    2. Boil 500 ml of distilled water in a beaker on a hot plate to remove CO2 and then turn off the hot plate
    3. In the fume hood, weigh 4 g of uranyl acetate dihydrate (depleted) and transfer to 100 ml volumetric flask
    4. Transfer 100 ml of near-boiling water to the amber glass bottle and place on a stirrer with magnetic stir bar to stir overnight
    5. Filter solution with a Whatman #1 filter into a clean, amber glass bottle
    6. Cap and label the bottle appropriately
    7. This solution can be stored in a 4 °C refrigerator for months.
    8. Make 1% working solution by diluting 2 ml of 4% uranyl acetate solution into 6 ml of previously boiled distilled water, CO2-free
    9. Filter the 1% uranyl acetate solution immediately prior to use using a 0.22 μm syringe filter placing filtered drops directly into your staining dish
  8. 1 N-NaOH
    1. Measure 0.40 g of sodium hydroxide
    2. Dissolve in 8 ml of water
    3. Volume up to 10 ml of solution
  9. Reynold’s lead citrate (Reynold’s, 1963)
    1. Wear appropriate personal protective equipment including at least an N-95 respirator
    2. Boil 500 ml of distilled water in a beaker on a hot plate to remove CO2 and then let cool
    3. In the fume hood, weigh 1.33 g of lead nitrate, Pb(NO3)2 and transfer to a 50 ml volumetric flask
    4. In the fume hood, weigh 1.76 g of sodium citrate, Na3(C6H5O7)·2H2O and transfer to the same volumetric flask
    5. Add 30 ml of the cooled CO2-free distilled water from above to the volumetric flask and cap
    6. Shake the suspension vigorously for 1 min and then let it stand for 30 min with intermittent shaking to ensure complete conversion of lead nitrate to lead citrate. The solution will remain cloudy
    7. Carefully add 1 N-NaOH 1 ml at a time (~8 ml total) swirl and check the pH using a pH meter. The pH should be 12.0 ± 0.1. If the measurement is above 12.1, then start over
    8. The solution will go from cloudy to clear and should not have any turbidity
    9. Volume up to 50 ml total and store in a tightly-sealed glass bottle

Acknowledgments

The authors would like to thank Yann S. Gallot for his thorough reading of the manuscript and the helpful comments. The authors would also like to declare no conflicts of interest or competing interests.

References

  1. Bonnard, C., Durand, A., Peyrol, S., Chanseaume, E., Chauvin, M. A., Morio, B., Vidal, H. and Rieusset, J. (2008). Mitochondrial dysfunction results from oxidative stress in the skeletal muscle of diet-induced insulin-resistant mice. J Clin Invest 118(2): 789-800.
  2. Hindi, L., McMillan, J. D., Afroze, D., Hindi, S. M. and Kumar, A. (2017). Isolation, culturing, and differentiation of primary myoblasts from skeletal muscle of adult mice. Bio-protocol 7(9): e2248.
  3. Hindi, S. M., Mishra, V., Bhatnagar, S., Tajrishi, M. M., Ogura, Y., Yan, Z., Burkly, L. C., Zheng, T. S. and Kumar, A. (2014). Regulatory circuitry of TWEAK-Fn14 system and PGC-1alpha in skeletal muscle atrophy program. FASEB J 28(3): 1398-1411.
  4. Hindi, S. M., Sato, S., Xiong, G., Bohnert, K. R., Gibb, A. A., Gallot, Y. S., McMillan, J. D., Hill, B. G., Uchida, S. and Kumar, A. (2018). TAK1 regulates skeletal muscle mass and mitochondrial function. JCI Insight 3(3).
  5. Jenny, A. (2011). Preparation of adult Drosophila eyes for thin sectioning and microscopic analysis. J Vis Exp (54): 2959.
  6. Kammoun, M., Cassar-Malek, I., Meunier, B. and Picard, B. (2014). A simplified immunohistochemical classification of skeletal muscle fibres in mouse. Eur J Histochem 58(2): 2254.
  7. Li, H., Malhotra, S. and Kumar, A. (2008). Nuclear factor-kappa B signaling in skeletal muscle atrophy. J Mol Med (Berl) 86(10): 1113-1126.
  8. Paul, P. K., Gupta, S. K., Bhatnagar, S., Panguluri, S. K., Darnay, B. G., Choi, Y. and Kumar, A. (2010). Targeted ablation of TRAF6 inhibits skeletal muscle wasting in mice. J Cell Biol 191(7): 1395-1411.
  9. Peterson, C. M., Johannsen, D. L. and Ravussin, E. (2012). Skeletal muscle mitochondria and aging: a review. J Aging Res 2012: 194821.
  10. Picard, M., White, K. and Turnbull, D. M. (2013). Mitochondrial morphology, topology, and membrane interactions in skeletal muscle: a quantitative three-dimensional electron microscopy study. J Appl Physiol (1985) 114(2): 161-171.
  11. Reynolds, E. S. (1963). The use of lead citrate at high pH as an electron-opaque stain in electron microscopy. The Journal of Cell Biology 17(1), 208-212.
  12. Sandri, M. (2008). Signaling in muscle atrophy and hypertrophy. Physiology (Bethesda) 23: 160-170.
  13. Schiaffino, S. and Reggiani, C. (2011) Fiber types in mammalian skeletal muscles. Physiol Rev 91(4): 1447-531.
  14. Soplop, N. H., Patel, R. and Kramer, S. G. (2009). Preparation of embryos for Electron Microscopy of the Drosophila embryonic heart tube. J Vis Exp (34): 1630.
  15. Takahashi, M. and Hood, D. A. (1996). Protein import into subsarcolemmal and intermyofibrillar skeletal muscle mitochondria. Differential import regulation in distinct subcellular regions. J Biol Chem 271(44): 27285-27291.
  16. Watson, M. L. (1958). Staining of tissue sections for electron microscopy with heavy metals. J Biophys Biochem Cytol 4(4): 475-478.

简介

骨骼肌是人体中含量最丰富的组织,可调节各种功能,包括运动和全身代谢。骨骼肌有很多线粒体,这是ATP好氧生成所必需的细胞器,它提供化学能量来促进收缩等重要功能。骨骼肌中线粒体的数量及其功能随着正常衰老和各种神经肌肉疾病以及癌症,饥饿,去神经支配和固定等分解代谢条件而下降。此外,受损的线粒体功能也与包括2型糖尿病在内的代谢紊乱有关。现在清楚的是维持骨骼肌中的线粒体含量和功能对于整个寿命期间的持续健康是至关重要的。虽然有许多染色方法可用于研究线粒体,但透射电子显微镜(TEM)仍然是研究骨骼肌中线粒体结构和健康的最重要方法。它提供关于线粒体含量,嵴密度,组织,自噬体形成以及在各种疾病状况中经常观察到的任何其他异常的关键信息。在本文中,我们描述了一个详细的协议样本制备和透射电镜分析小鼠骨骼肌线粒体。

【背景】骨骼肌是一种高度塑性的组织,经过响应一些细胞外信号的形态和代谢适应性。包括抵抗或耐力运动在内的许多干扰刺激线粒体生物发生,导致增加的代谢能力和抵抗疲劳(Li等人,2008; Sandri,2008)。相反,在衰老期间,不活动,以及在许多分解代谢疾病状态下,骨骼肌线粒体数量和功能下降,导致易疲劳性和胰岛素抵抗增加(Sandri,2008)。功能失调的线粒体的累积也可能导致进行性活性氧物质诱导的损伤,从而进一步损害骨骼肌中的氧化能力(Bonnard等人,2008)。

线粒体作为网状膜网络存在,位于骨骼肌中的不同亚细胞区室中。线粒体亚线粒体(线粒体)占线粒体体积的10-15%,直接位于肌膜上,而肌间纤维细胞(IMF)线粒体与肌原纤维密切接触(Takahashi和Hood,1996)。线粒体是双膜结构,其在外膜和内膜之间以及内部基质隔室之间包含膜间隙,其中大部分代谢过程发生。内膜高度折叠,形成所谓的嵴,以适应其大表面积。构成氧化磷酸化发生的呼吸链的五个复合物嵌入内线粒体膜内。在这个过程中,穿过内膜的质子梯度与复合物V处的ATP合成偶联(Peterson等人,2012)。除了产生ATP用于肌动蛋白和肌球蛋白之间的跨桥循环之外,线粒体是调节骨骼肌生理学的自由基的来源(Peterson等人,2012)。

透射电子显微镜(TEM)是超微结构研究的强大技术(Watson,1958)。在生理和病理条件下,TEM在研究骨骼肌线粒体结构方面非常有用(Picard等人,2013)。例如,TEM可以提供关于线粒体内容,组织,嵴结构和空泡形成的信息,如在一些神经肌肉疾病例如肌萎缩性侧索硬化中观察到的(Picard等人,2013)。在许多肌肉萎缩症状中,线粒体含量通过自噬(也称为线粒体自噬)而降低。在这方面,TEM被发现是研究自噬体形成的重要方法(Sandri,2008)。我们开发了一种有效的方案,可以很容易地在任何实验室中使用,以通过TEM研究骨骼肌中的小鼠线粒体的超微结构(Paul等人,2010; Hindi等人, 2014年和2018年)。在下面的章节中,我们提供了一个逐步协议,用于样品制备和分析骨骼肌中的SS和IMF线粒体。通过TEM也可以使用类似的方案来研究骨骼肌中的其他细胞器。

关键字:透射电子显微镜技术, 骨骼肌, 线粒体, 自噬, 肌病, 萎缩, 氧化代谢

材料和试剂

  1. 玻璃标本瓶(电子显微镜科学,目录号:72630-05)
  2. 剃刀刀片,双面涂层,洗版(电子显微镜科学,Personna,目录号:72000-WA)
  3. 移液器(Fisher Scientific,目录号:13-711-9BM)
  4. 丁腈手套
  5. 玻璃条,超微切割级,6.4×25×400毫米(电子显微镜科学,目录号:71012)
  6. 玻璃载玻片(Fisher Scientific,目录号:12-550-15)
  7. 注射器1毫升(BD,目录号:329652)
  8. 注射器30毫升(BD,目录号:302833)
  9. 0.22μm注射器过滤器(Merck,产品目录号:SLGV033RS)
  10. 锥形离心管,50毫升(VWR,目录号:21008-169)
  11. 锥形离心管,15毫升(VWR,目录号:21008-089)
  12. 非无菌尿液标本容器(Electron Microscopy Sciences,目录号:64231-10)
  13. 滤纸,定性1级圆圈(GE Healthcare,Whatman,目录号:1001-090)
  14. 玻璃搅拌棒(United Scientific Supplies,目录号:GSR012)
  15. 木器(电子显微镜科学,目录号:72300)
  16. 平的硅树脂嵌入模具(电子显微镜科学,目录号:70900)

  17. 玻璃刀舟,6.4毫米(电子显微镜科学,目录号:71007)
  18. 玻璃刀盒(电子显微镜科学,目录号:71010)
  19. N95呼吸器,带阀门(VWR,目录号:89201-510)
  20. 金属环,完美的环(电子显微镜科学,目录号:70944)
  21. 网格,标签,铜,200目(泰德佩拉,目录号:3HGC200)
  22. 网格储物盒,标签(Ted Pella,目录编号:161)
  23. 培养皿,玻璃,100×20毫米(Corning,目录号:70165-102)
  24. 戊二醛,EM等级,8%(Polysciences,目录号:00216-30)
  25. 磷酸二氢钠一水合物,NaH2PO4·2H2O(Sigma-Aldrich,目录号:S9638 )
  26. 磷酸氢二钠无水Na 2 HPO 4(Sigma-Aldrich,目录号:S9763)
  27. 四氧化锇,10×1克(电子显微镜科学,目录号:19110)
  28. 乙醇,200证明(Decon Labs,目录号:2701)
  29. EMbed-812试剂盒包括:EMbed-812,DDSA,NMA和DMP-30(电子显微镜科学,目录号:14120)
  30. 硼酸钠(MP Biomedicals,目录号:0219030980)
  31. 甲苯胺蓝O(Amresco,目录号:0672-25G)
  32. 乙酸双氧铀粉末(耗尽)(Electron Microscopy Sciences,目录号:22400)
  33. NaOH颗粒(Amresco,目录号:0583-500G)
  34. 硝酸铅,Pb(NO 3)2(电子显微镜科学,目录号:17900)
  35. 柠檬酸钠,Na 3(C 6 H 5 O 7)2·2H 2/2 > O(电子显微镜科学,目录号:21140)
  36. 环氧丙烷,EM级(电子显微镜科学,目录号:20401)
  37. 牙科蜡(Electron Microscopy Sciences,目录号:72660)
  38. 3%戊二醛(见配方1)
  39. 0.1M磷酸钠缓冲液(pH 7.4)(见配方2)
  40. 1%四氧化锇(见配方3)
  41. 乙醇稀释(见第4部分)
  42. 嵌入媒体(见第5章)
  43. 1%甲苯胺蓝染色(见第6部分)
  44. 4%醋酸双氧铀(aq)(见第7部分)
  45. 1 N-NaOH(见方法8)
  46. 雷诺的铅柠檬酸盐(见第9条)
    预防措施/危害:与在实验室中处理的任何化学品和试剂一样,用户应该注意如何安全地使用和操作它们。请参阅每种化学品的材料安全数据表(MSDS),了解有关预防措施和危害的详细信息。电子显微镜使用相当多的有害化学物质,如:戊二醛,四氧化锇,环氧丙烷,乙酸铀酰,柠檬酸铅等。请使用适当的个人防护设备(PPE),通风条件处理这些化学品,并根据您所在机构的环境健康和安全部处理这些化学品。

设备


  1. 琥珀色,广口玻璃瓶,125毫升(VWR,目录号:10861-846)
  2. 清洁介质瓶,1升(Corning,PYREX ®,产品目录号:1399-1L)
  3. 刻度量筒,1,000毫升(VWR,目录号:65000-012)
  4. 刻度量筒,25毫升(VWR,目录号:65000-002)
  5. 磁力搅拌棒(VWR,目录号:58948-025)
  6. 通用液体玻璃钢温度计(VWR,目录号:89095-626)
  7. 负动作弯曲自闭镊子(Electron Microscopy Sciences,目录号:72864-D)
  8. 睫毛操纵器(电子显微镜科学,目录号:71182)
  9. 1000毫升玻璃格里芬烧杯(VWR,目录号:10754-960)
  10. 50毫升玻璃格里芬烧杯(VWR,目录号:10754-946)
  11. 玻璃漏斗,100毫米(VWR,目录号:10546-048)
  12. 50毫升容量瓶(VWR,目录号:10123-996)

  13. 100毫升容量瓶,琥珀色(VWR,目录号:10124-022)
  14. -20°C冷冻机(VWR,目录号:97014-903)
  15. 4°C冰箱(VWR,目录号:14236-525)
  16. 临床旋转器,变速管旋转器(Cole-Parmer,Stuart,目录号:SB3)
  17. 用于临床旋转器,变速管旋转器的培养管支架,12毫米为管材提供滚动作用(Cole-Parmer,Stuart,目录号:SB3 / 3)
  18. pH计,SymPhony B10P(VWR,目录号:89231-662)
  19. pH探头,可再填充玻璃(VWR,目录号:89231-580)
  20. 精密天平,AV212C(奥豪斯,停产)
  21. 精确平衡(OHAUS,目录号:30122632)
  22. 加热板搅拌器(Fisher Scientific,目录号:SP88857200P)
  23. 真空烘箱(Electron Microscopy Sciences,目录号:63235-10)
  24. 超薄切片机(Reichert-Jung,型号:Ultracut E,停产,eBay或其他二手市场)
  25. 新的超薄切片机(徕卡显微系统,型号:Leica EM UC7)
  26. 玻璃刀匠(LKB,型号:LKB 7801B型,停产,eBay或其他二手市场)
  27. 新的玻璃刀匠Leica EM KMR3(Leica Microsystems,目录号:Leica EM KMR3)
  28. 光学显微镜,奥林巴斯CX31(奥林巴斯,产品目录号:CX31)
  29. 钻石刀,Diatome Wet超45°,3.5毫米(电子显微镜科学,型号:Diatome Ultra)
  30. 菲利普斯CM10透射电子显微镜改装一台新的数码相机(菲利普斯,目录号:CM10)
  31. 用于TEM的高分辨率CCD摄像机改装为Phillips CM10示波器(高级显微镜技术,目录号:BioSprint)

程序

使用TEM分析骨骼肌线粒体超微结构的基本步骤如图1所示。


图1.从始至终
骨骼肌透射电镜的一般程序示意图

  1. 主要固定
    戊二醛主要固定的目标是通过交联蛋白质来保持生理相关状态下骨骼肌组织的超微结构,主要与亲核试剂和其他胺反应。
    1. 遵循配方1,制备3%戊二醛溶液。
    2. 取出肌肉组织并立即将其浸入玻璃样品瓶中的3%戊二醛中并相应标记(参见注释1)。
    3. 在室温下将标本小瓶置于临床旋转器上30分钟。
    4. 30分钟后,迅速将肌肉转移到滤纸上,并将其切成所需尺寸(对于小鼠比目鱼,分别将横向和纵向切片切成两半),用剃刀刀片或解剖刀轻轻拉动,请勿挤压组织(见注2)。
    5. 尽快将每个新的组织切片放入装有3%戊二醛的标本瓶中,并适当标记小瓶。
    6. 将标本小瓶置于临床旋转器上并在室温下放置24小时。

  2. 二次固定
    使用四氧化锇进行二次固定的目的是双重的,它交联脂质以保持其结构,并且通过添加电子致密材料来有效地染色组织,增强对比度。
    1. 遵循配方2,制备0.1M磷酸钠缓冲液(pH 7.4)。

    2. 使用移液管从样品瓶中取出3%戊二醛并将溶液转移到合适的废液容器中。
    3. 将磷酸盐缓冲液加入样品瓶中,并将样品送回临床旋转器5分钟。
    4. 将磷酸盐缓冲液倒入适当的废液容器中,重复步骤B3两次,第三次清洗,然后弃置磷酸盐缓冲液,准备好1%的锇溶液,以便样品不会变干。
    5. 遵循配方3,准备1%四氧化锇(见注3)。
    6. 在化学通风橱中,注意并小心,将1%四氧化锇注入标本瓶中,盖上瓶子,然后返回临床旋转器1小时。
    7. 标本应显示为黑色。
    8. 在通风橱下,从样品瓶中取出1%四氧化锇溶液,并将溶液倒入适当的废物容器中。
    9. 将塑料盖丢弃到适当的废物容器中,然后在样品瓶上使用新塑料盖进行后续步骤。
    10. 将磷酸盐缓冲液加入样品瓶中,并将样品送回临床旋转器5分钟。
    11. 将磷酸盐缓冲液置于适当的废物容器中,并重复步骤B10两次以上。

  3. 脱水
    水必须通过分级酒精脱水从组织中去除,以允许包埋介质的渗透。
    1. 遵循配方4,准备所有乙醇稀释液。
    2. 将10%乙醇溶液转移到样品瓶中,并将它们返回到临床旋转器5分钟。
    3. 取出10%乙醇,用25%,50%,75%和95%乙醇溶液重复步骤C2。
    4. 将100%乙醇转移到样品瓶中,并将其返回到临床旋转器5分钟。
    5. 去除100%乙醇并重复步骤C4两次以上。
    6. 在化学通风橱下,将100%氧化丙烯转移到样品瓶中并密封盖子(如果不安全,盖子可能会弹出)。

    7. 将样品返回临床旋转器5分钟。
    8. 除去100%环氧丙烷并将其处理在适当的废物容器中。
    9. 重复步骤C6,C7和C8两次以上。

  4. 渗透和嵌入
    环氧丙烷同样可溶于酒精和包埋介质,从而清除酒精并促进包埋介质渗入组织深处。
    1. 遵循配方5,准备嵌入介质。
    2. 在化学通风橱中,准备2:1(环氧丙烷:包埋介质)溶液,您制造的总体积取决于您有多少样品(即使在旋转时,您想要足够的溶液以完全覆盖样品)。< br />
    3. 将2:1(环氧丙烷:包埋介质)溶液转移到样品瓶中,盖上盖子,并将它们放置在临床旋转器上1 h(见注4)。
    4. 从样品瓶中取出2:1的溶液,并将其丢弃在塑料废物容器中。
    5. 在化学通风橱中,准备1:1(环氧丙烷:包埋介质)溶液,这里也可以使用与2:1溶液相同的总体积。
    6. 将1:1(环氧丙烷:包埋介质)溶液转移到样品瓶中,盖上它们,然后将样品瓶返回临床旋转器。将样品留在临床旋转器上过夜,并将剩余的包埋介质存放在-20°C的冷冻箱中。
    7. 第二天,将1:1溶液置于同一废物容器中,并将包埋介质置于室温。
    8. 在化学通风橱中,使用与上述相同的总体积制备1:2(环氧丙烷:包埋介质)溶液。
    9. 将1:2(环氧丙烷:包埋介质)溶液转移到样品瓶盖上,并将它们放置在临床旋转器上1小时。

    10. 从样品瓶中取出1:2溶液并将其处理到指定的废物容器中。
    11. 将100%包埋介质转移到样品瓶中,并将它们返回到转子1小时。
    12. 删除100%的包埋介质并将其处理到指定的废物容器中。
    13. 将100%包埋介质转移到样品瓶中,取下盖子,并放入加压至20 PSI的真空箱中1 h,使其达到室温。
    14. 标本现在可以进行固化了,不需要从小瓶中取出剩余的100%培养基,因为标本将直接从小瓶转移到模具中。

  5. 固化
    粘稠的包埋介质必须固化,以使包含组织的块变得足够硬以进行超薄切片。
    1. 将真空烘箱设置为60°C和0 PSI,然后在继续操作前用温度计确认温度。
    2. 使用铅笔,将样品信息写在一张小纸片上,这些小纸片可以放入模具底部,与样品一起固化(可通过固化塑料看到)。

    3. 使用木材涂抹器将100%包埋介质中的塑料模具的各个孔填充到中途,以将介质淋入模具的孔中。
    4. 用精细的负片镊子,将带有样品信息的纸张转移到模具的底部(离样品最远的部分)。
    5. 接下来,使用精细的负动作镊子,将黑化的渗透(硬)样品从样品瓶转移到模具(参见注释5)。
    6. 相应地确定样品的位置,以确保样品可以轻松地在横截面或纵向切片中切片(参见图6,了解它在模块中的外观示意图)。

    7. 对剩余的样品重复步骤E4,E5和E6。
    8. 转移足够的100%嵌入介质以充分填满井。
    9. 确保样品最后一次正确定位,因为添加介质会稍微扭曲它们的位置。
    10. 小心地将塑料模具放入烤箱中固化至少24小时(参见注释6)。

  6. 玻璃刀制造
    制作自己的玻璃刀是一种节约成本的技术,可以让您保留和延长昂贵的钻石刀的寿命,以便仅切割超薄切片。如果您的机构不提供该设备,可以购买预制玻璃刀。
    1. 戴丁腈手套,用洗涤剂和毛刷清洁玻璃条(6.4 x 25 x 400毫米),然后用自来水彻底冲洗(见注7)。
    2. 用喷射瓶,用蒸馏水冲洗另一次。
    3. 将纸巾上的玻璃条在相对干净无尘的环境中晾干。
    4. 通过旋转后部玻璃固定器旋钮(图2A)缩回后部玻璃固定器(图2A),然后拉出旋钮将其锁定到位。
    5. 确保前部的分离旋钮(图2B)逆时针旋转完全。

    6. 转动锁定杆(图2A)回到后面位置,完全抬高记录器。

    7. 将刻痕轴(图2B)推入。
    8. 确保评分旋钮上方的分数选择器(图2B)设置为。
    9. 将清洁干燥的玻璃条转移到玻璃切割刀制造商处,并将其与白色导向板对齐(图2B),然后将条带推向第一个侧向锁定螺柱(图2C)(用于25mm宽的条)。< br />
    10. 将玻璃条固定到位时,轻轻摇动锁定杆,直至垂直支撑柱(图2C)与玻璃条接触。

    11. 。移开你的手,然后再按下锁定杆,直到有明显阻力。


      图2.具有标记部件的玻璃刀制造机 A.带有标记部件的倾斜侧视图。 B.带标签部件的正面视图。 C.带有标记部分的倾斜侧视图特写。

    12. 将玻璃叉(图2A)放在玻璃条的左侧下方,以抓住您的玻璃方块。
    13. 将玻璃刻痕轴旋钮以迅速流畅的方式拉过玻璃条表面,平滑移动,直至玻璃刻痕为止(图3A)。

    14. 。顺时针旋转分解开关,直到玻璃破碎,您将可以在视觉上和听觉上感觉到这一点。

    15. 。将分离旋钮完全逆时针方向复位

    16. 。将锁定杆转回到后部位置,完全抬高记分器。

    17. 完全推入计分轴旋钮。
    18. 用刷子清理玻璃破碎区域。
    19. 将润版压力调节杆(图2C)放在圆点上。
    20. 调整前后玻璃座调节旋钮以适应玻璃方块。

    21. 将刚制成的方形玻璃片以45°角放置在破坏销上并将玻璃叉放在方形下方(图3B)。
    22. 将步骤F4中的后玻璃座固定器旋钮按下,然后顺时针旋转,使后玻璃固定器与玻璃正方形的角部接触。

    23. 玻璃正方形应该坐在前后玻璃支架之间
    24. 轻轻地将锁定杆拉至前方位置,直至支撑螺栓与玻璃方块接触。

    25. 进一步向下按锁定杆,直至出现明显阻力。

    26. 在玻璃方块面上迅速流畅地移动刻痕轴旋钮,直至其停止对玻璃打分。
    27. 旋转减震压力调节杆,直到减震垫接触与前玻璃座接触的玻璃角。
    28. 顺时针旋转分离旋钮,直至玻璃破碎;您将能够在视觉和听觉上听到这一点。

    29. 。将分离旋钮完全逆时针方向复位

    30. 。将锁定杆转回到后部位置,完全抬高记分器。

    31. 完全推入计分轴旋钮。
    32. 将调湿压力调节杆重新放回点。
    33. 逆时针旋转后部玻璃固定器旋钮,将后部玻璃固定器从玻璃三角形处缩回,然后将旋钮拉出,将其锁定到位。
    34. 使用玻璃叉将左侧和右侧玻璃刀抬起并从刀具制造机中取出,并将刀放入玻璃刀架中。
    35. 用刷子清理玻璃破碎区域。
    36. 重复步骤F1-F35来制作更多的玻璃刀。
    37. 要将塑料小船连接到小刀上,请将热盘上的玻璃烧杯中的某些牙科蜡加热至100°C。
    38. 一旦蜡完全熔化后,使用木材涂抹器将蜡涂在塑料船的边缘,并立即将塑料船对准玻璃刀背面(见注8)。
    39. 使用木材涂抹器,将更多熔化的蜡转移到附属船的边缘以完全密封(图3D)。


      图3.玻璃刀的制造A.玻璃条在被打分并分解成玻璃方块后, B.玻璃正方形在后部和前部玻璃支架之间对齐,调节减震压力调节杆并在下面安装玻璃叉以抓住玻璃; C.玻璃方形,在用上述记分器打分之前,将锁定杆完全压下,将其压在断针上; D.一些没有船的玻璃刀和一个玻璃刀架上装有牙蜡的塑料船。

    40. 通过用移液管将蒸馏水转移到塑料船中测试蜡封,从而形成一个凸形弯月面的水。

    41. 。将玻璃刀放在塑料舟中5分钟,然后检查水位以确定是否有泄漏。
    42. 如果无泄漏,请将空水倒入玻璃刀架以备将来使用。

  7. 修剪块
    将块体修剪成梯形金字塔形状,可以减少刀具上的摩擦力和阻力,从而增强您能够生产的厚薄部分。

    1. 从烤箱中取出塑料模具(来自步骤E10)并让其冷却至少30分钟。
    2. 一旦冷却后,将带有组织样本的塑料块放入样本支架(图4A和5A)中的样本支架(图4A)。


      图4. Reichert-Jung超薄切片机,样本架位置可以修剪,刀架位于切割样本的位置。A.样本架夹在适配器和导轨中以修剪嵌入的样本。 B.这是标本块在通过双目眼睛片时的样子。 C.带有钻石刀的刀架被夹紧到位并调整为切割薄片。 D.这就是在钻透双目眼镜时钻石刀刃的样子。

    3. 夹紧试样夹具和适配器(图4A)以确保试样块不移动。
    4. 按下超薄切片机电源上的绿色开关主开关(图5A),并使用多位照明开关(图5C)打开照明灯,同时舞台灯光和舞台下的灯都会亮起。
    5. 通过转动聚焦旋钮(图5D),同时通过宽视场望远镜目镜观察时,使样本聚焦。


      图5.带有标记部件的Reichert-Jung超薄切片机A.电源; B.超薄切片机的正视图; C.倾斜的左侧视图; D.有角度的右侧视图。

    6. 目标是将样本块修剪成尽可能小的梯形金字塔,同时将样本包括在中心(参见注释9)(图6)。


      图6.如何在切片前修剪块的示意图 A.完全固化的块与渗透的肌肉样本对齐切割横截面应该是这样的。从顶部看,由于组织和块顶部之间的塑料层,向下看渗透肌将看起来相对较暗。使用剃刀刀片,需要修剪块面的顶部以暴露组织样本。 B.最初,修剪后的图层将仅包含塑料,并且一旦组织暴露出来,最终您将开始用包围塑料看到修剪层中的黑色组织。接下来,修剪样品块周围的塑料以创建一个梯形金字塔。现在可以使用玻璃刀对样品块进行处理。

    7. 在通过宽视野双目镜(图5B)和使用剃刀刀片观察时,修整块的表面,直到将样品带入平面。您应该看到颜色较浅的树脂,然后看到您正在创建的修剪过的切片中样品的黑影。
    8. 接下来,在通过Wide-Field双目目镜观察时,使用剃刀刀片从样品四周的四周除去多余树脂,以雕刻您的金字塔。
    9. 从适配器上松开标本固定器,将标本固定器放在超薄切片机的标本臂(图5C和5D)上,并使用螺丝将标本固定器紧紧固定到位。
    10. 松开并取下样品支架适配器,将您的刀架(图4C和5B)放在超薄切片台上,并将刀架夹紧到位。
    11. 确保间隙角度调节旋钮(图4C)调节到合适的后角,该角度应与钻石刀(6°)随附的箱子上写下的角度一致。
    12. 使用刀具夹紧螺钉(图4C)将玻璃刀夹在刀架上。
    13. 使用东西方舞台控制旋钮(图5C)和南北刀进给旋钮(图5B)将玻璃刀的刀架靠近您的挡块,不要触碰。
    14. 调整您的样本架,使您的样本的前缘(首先切割的部分和您的金字塔底部)与刀刃平行。
    15. 在超薄切片机上使用南北切刀进刀旋钮,慢慢地将玻璃切刀靠近样品块。您的超薄切片机配备有从底座照射的光线,照亮刀和块体表面之间的缝隙。当您将刀子靠近块体表面时,此照明区域将变得越来越暗淡,表明两个表面几乎是相互接触的。
    16. 将手动精饲刀控制旋钮(图5C)设置为1μm。
    17. 在通过宽视场望远镜目镜观察时,旋转手动精进刀控制旋钮,将样品前进至刀尖,同时旋转样品臂的手轮(图5D)一个完整周期。
    18. 重复步骤G17,直到开始看到样品表面的部分脱落到玻璃刀的边缘上,然后继续,直到您的样品的整个表面被切割为止。
    19. 现在,脸部被修剪,平滑,并准备厚厚的部分。
    20. 使用南北刀进给旋钮将刀架移离样品。
    21. 拧下刀夹紧螺钉松开玻璃刀并从刀架上取下。

  8. 切厚切片
    切割厚切片然后染上它们的目的是在继续之前确认组织样本的取向。
    1. 遵循配方6,准备1%甲苯胺蓝染色。
    2. 打开一个热板到约100°C。
    3. 放置一把玻璃刀和一个塑料舟,然后拧紧刀夹紧螺钉(图4C),将刀夹紧到位。
    4. 使用东西方舞台控制旋钮(图5C)和南北刀进给旋钮(图5B)将玻璃刀的刀架靠近您的挡块,不要触碰。
    5. 调整您的试样支架(图5D),使样品前缘(首先切割的部分和金字塔底部)与刀刃平行。
    6. 在超薄切片机上使用南北切刀进刀旋钮,慢慢地将玻璃切刀靠近样品块。您的超薄切片机配备有从底座照射的光线,照亮刀和块体表面之间的缝隙。当您将刀子靠近块体表面时,此照明区域将变得越来越暗淡,表明两个表面几乎是相互接触的。
    7. 用蒸馏水填充塑料船,直到水凸起。
    8. 将吸入注射器(图5D)放入水管下方的船内。
    9. 在观察宽视野双目镜(图5B)时,使用吸水旋钮(图5D)调整水位,直到水面上出现均匀的银反射(见注10和11)(图7 )。


      图7.通过双目镜头的视图显示切片前钻石刀中的适当水位。金刚石刀舟(A)的水面凸出表面。如图(B)所示,使用超薄切片注射器,降低水位直至出现银色反光表面。如图(B)所示,在放下水并尝试切割部分后,如果任何水收集在块的表面上,则可能需要稍稍降低水位,在此处可以看到在刀口处形成的轻微阴影。但是,如果水降得太多,它会从刀刃上脱落,需要重新加注。

    10. 将手动精饲刀控制旋钮(图5C)设置为1μm。
    11. 在通过宽视野双目镜看时,转动手动精饲刀控制旋钮,将样品推到刀口,同时将样品臂的手轮(图5D)旋转一个完整周期。
    12. 重复步骤H17,直到您开始看到样品表面的部分从玻璃刀边缘脱落到水面上,然后继续,直到您的样品的整个表面被切开(见注12)。
    13. 一旦你得到一个全面的部分,用一个金属环去掉部分部分和你刚刚切割的整个部分,并通过在蒸馏水烧杯中旋转金属环来丢弃它们。
    14. 将手动进料刀控制旋钮设置为0.5μm。
    15. 在通过宽视场望远镜目镜观察时,转动手动精进刀控制旋钮,将样品推到刀口,同时将样品臂的手轮旋转一个完整周期。
    16. 丢弃第一个几节到水烧杯。

    17. 切5-10个0.5μm的部分
    18. 在相应的样品信息上标注两张载玻片,并在载玻片上滴上一滴(〜2厘米直径)的蒸馏水。
    19. 使用金属环将各个部分转移到载玻片上的水滴上,然后旋转环以进行转移。
    20. 在完成分配两个载玻片之间的部分后,将载玻片放在热板上(见注13)。
    21. 水完全蒸发后,这些部分已准备好被染色。
    22. 当载玻片仍在热板上时,滴加1%甲苯胺蓝染色剂以完全覆盖干燥的切片并保持2分钟(参见注释14)。

    23. 使用自来水清洗玻片,尽可能多地去除玻片上多余的污渍,并让玻璃干燥。
    24. 让幻灯片干燥后,用光学显微镜观察各部分(见注15和16)。
    25. 如果对部分的方向感到满意,则不需要调整,您可以继续切割薄部分。
    26. 使用南北刀进给旋钮将刀架移离样品。
    27. 拧下刀夹紧螺钉松开玻璃刀并将其从刀架上取下。

  9. 切薄片
    为了形成高分辨率的图像,电子束必须能够穿透该部分而不会显着降低速度,需要超薄部分。
    1. 将一把钻石刀放在刀架上(图5B),然后拧紧刀夹紧螺钉(图4C),将刀夹紧到位。
    2. 使用东西方舞台控制旋钮(图5C)和南北刀进给旋钮(图5B)将玻璃刀的刀架靠近您的挡块,不要触碰。
    3. 调整样品架(图5D),使样品前缘(首先切割的部分和金字塔底部)与刀口平行。
    4. 在超薄切片机上使用南北切刀进刀旋钮,慢慢地将玻璃切刀靠近样品块。您的超薄切片机配备有从底座照射的光线,照亮刀和块体表面之间的缝隙。当你把刀子靠近块面时,这个照明区域会变得越来越暗,这表明两个表面几乎是接触的。
    5. 用蒸馏水填充钻石刀船,直到水凸出。
    6. 将吸入注射器(图5D)放入水管下方的船内。
    7. 在透视宽视野双目镜(图5B)时,使用吸力旋钮(图5D)调整水位,直到水面上出现均匀的银反射(图7)。
    8. 将手动上料刀控制旋钮(图5C)设置为0.5μm。
    9. 在通过宽视野双目镜看时,转动手动精饲刀控制旋钮,将样品推到刀口,同时将样品臂的手轮(图5D)旋转一个完整周期。
    10. 重复步骤I9,直到您开始看到样品表面的部分从玻璃刀边缘脱落到水面上,然后继续,直到样品的整个表面被切割(参见注释17)。
    11. 一旦你得到一个全面的部分,用一个金属环去掉部分部分和你刚切割的全面部分,然后在蒸馏水烧杯中旋转金属环。
    12. 接下来,在电源上,将Semi-Thin Sectioning Control(图5A)上的值设置为0.50μm。
    13. 在电源上,将切割速度控制(图5A)调整到1.0毫米/秒(这对应于超薄切片机手臂在下冲程中使用的速度)。
    14. 按下马达驱动控制杆(图5D),开始自动前进和旋转试样臂。
    15. 通过宽视场望远镜观察并观察从刀上脱落的全脸部分(见注18)。
    16. 3-4节之后,通过宽视野双目镜观察,一旦试样臂完成向下的行程,将新的截面发送到船上,将半薄切片控制调整为0.40μm。
    17. 使用一个金属环去掉你刚刚切割的部分,并通过在蒸馏水烧杯中旋转金属环来丢弃它们。
    18. 3-4节之后,通过宽视野双目镜观察,一旦试样臂完成向下的行程,将一个新的部分放入船中,将半薄切片控制调节至0.30μm。

    19. 使用金属环来捞出刚刚切割的部分,并通过在蒸馏水烧杯中旋转金属环来丢弃它们。
    20. 在双目镜片中观察时,调整后的部分应开始显示对应于部分的近似厚度和包埋材料的折射率的颜色。此时,它们很可能会变成黄色。
    21. 3-4节之后,通过宽视野双目镜观察,一旦试样臂完成向下的行程,将新的截面发送到船上,将半薄切片控制调整为0.25μm。

    22. 使用金属环来捞出刚刚切割的部分,并通过在蒸馏水烧杯中旋转金属环来丢弃它们。
    23. 在宽视场双筒望远镜中观察时,调整后的部分应开始看起来黄色变为蓝色。
    24. 3-4节之后,通过宽视场望远镜目镜观察,一旦试样臂完成向下的行程,将新的截面发送到船上,将半薄切片控制调整为0.20μm。

    25. 使用金属环来捞出刚刚切割的部分,并通过在蒸馏水烧杯中旋转金属环来丢弃它们。
    26. 在宽视野双目目镜中观察时,调整后的部分应开始看起来蓝至紫色。
    27. 3-4节之后,通过宽视场望远镜目镜观察,一旦标本臂完成向下的行程,将新的剖面发送到船上,将半薄切片控制调整到0.17μm。

    28. 使用金属环来捞出刚刚切割的部分,并通过在蒸馏水烧杯中旋转金属环来丢弃它们。
    29. 在宽视场双筒望远镜中观察时,调整后的部分应开始变为紫色。
    30. 3-4节之后,通过宽视场望远镜目镜观察,一旦样本臂完成向下的行程,将新的剖面发送到船上,将半薄切片控制调整为0.14μm。

    31. 使用金属环来捞出刚刚切割的部分,并通过在蒸馏水烧杯中旋转金属环来丢弃它们。
    32. 在宽视野双目目镜中观察时,调整后的部分应开始看起来像紫色。
    33. 3-4节之后,通过宽视野双目镜观察,一旦试样臂完成其向下的行程,将新的截面发送到船上,将半薄切片控制调整为0.12μm。

    34. 使用金属环来捞出刚刚切割的部分,并通过在蒸馏水烧杯中旋转金属环来丢弃它们。
    35. 在宽视野双目目镜中观察时,调整后的部分应开始看起来像金。
    36. 3-4节之后,通过宽视野双目镜观察,一旦试样臂完成向下的行程,将新的截面发送到船上,将半薄切片控制调整为0.11μm。

    37. 使用金属环来捞出刚刚切割的部分,并通过在蒸馏水烧杯中旋转金属环来丢弃它们。
    38. 在宽视场双筒望远镜中观察时,调整后的部分应开始看纯金(参见注19)。
    39. 如果满意,则继续切割这些部分,直至获得所需数量并继续步骤I47。如果不满意,则继续步骤I40。
    40. 3-4节之后,通过宽视场望远镜目镜观察,一旦样本臂完成向下的行程,将新的剖面发送到船上,将半薄切片控制调整为0.10μm。

    41. 使用金属环来捞出刚刚切割的部分,并通过在蒸馏水烧杯中旋转金属环来丢弃它们。
    42. 在宽视野双目目镜中观察时,调整后的部分应开始看起来像金银。
    43. 3-4节之后,通过宽视场望远镜目镜观察,一旦样本臂完成向下的行程,将新的剖面发送到船上,将半薄切片控制调整为0.09μm(90 nm)。

    44. 使用金属环来捞出刚刚切割的部分,并通过在蒸馏水烧杯中旋转金属环来丢弃它们。
    45. 在宽视场双目镜中观察时,调整后的部分应开始看起来银色。
    46. 继续切割部分,直到您满意为止,通常15-20个均匀颜色的部分是目标。
    47. 使用南北刀进给旋钮将刀架移离样品。
    48. 使用负动作钳用您的优势手拿起铜网,并将睫毛操纵器工具置于您的非优势手中。
    49. 在通过宽视野双目目镜观察时,请将这些工具放入视野。
    50. 将网格浸没在水面下并开始直接在您选择的部分下推进(参见注释20)。
    51. 同时,将您的睫毛操纵器工具稍微浸入您选择的部分旁边,并轻轻地将该部分引导至您的网格上方。
    52. 尽可能最好地保持该部分的位置,然后缓慢地升高该部分下方的格栅,直至该部分搁置在格栅的顶部相对中心并从水中移出。
    53. 使用安装在支架上的网格仍然放在钳子中,取一个三角形滤纸,轻轻将其放在网格外缘旁边。水应该开始从网格转移到滤纸上(参见注释21)。
    54. 尽可能地去除水后,将网格放入网格框中。
    55. 继续从你的钻石刀的船上收集部分,通常每个样本10格很多。
    56. 在染色之前,让网格在网格盒中充分干燥过夜。

  10. 染色网格
    用醋酸铀和柠檬酸铅染色切片可增强无数细胞结构的对比度。
    1. 打开热板/搅拌器并将温度调节至〜200°C。

    2. 在热板上放置一个装满蒸馏水和磁力搅拌棒的1升烧杯(图8)。


      图8.用于染色网格的设置 A.用于染色网格的通用设备。 B.特写显示使用含有柠檬酸铅染色的两个格栅的氢氧化钠颗粒,在将格栅漂浮在柠檬酸铅滴上之后立即关闭盖子以防止柠檬酸铅沉淀。

    3. 将水烧开,然后关闭电热板,同时使搅拌器继续运转(参见注释22)。
    4. 将水分配到四个50毫升的烧杯中,让其冷却到约25°C(见注23)。
    5. 遵循配方7,制备1%醋酸双氧铀溶液。
    6. 遵循配方8,制备1N-NaOH溶液。
    7. 遵循配方9,准备雷诺的柠檬酸铅溶液。
    8. 使用1 ml注射器,取下针头并连接0.22μm注射器过滤器,将一滴醋酸铀酰分配到玻璃培养皿中,底部填充牙蜡(参见注释24)。
    9. 用负面作用的钳子,把你的网格,截面朝下,放在醋酸铀酰滴上。
    10. 设置一个计时器30分钟,用挡住光线的东西覆盖培养皿(参见注释25)。
    11. 30分钟后,使用负动作钳取出网格,然后将网格浸入装满预先煮沸的蒸馏水的50ml烧杯中的一个中。
      。将网格进出水面〜30次。
    12. 进入下一个50毫升的烧杯,烧杯中装满以前煮过的蒸馏水,并将网格进出水中约30次(使用相同的两个烧杯进行醋酸铀酰洗涤)。
    13. 将负作用钳放在台面上,并轻轻将一个三角形滤纸涂在铜网的外缘,以除去网格表面的水分。
    14. 使用第二块玻璃培养皿,在底部涂上牙科蜡,然后在培养皿周围放置氢氧化钠丸,在那里加入柠檬酸铅滴剂。
      在准备柠檬酸铅时立即关闭盖子。
    15. 使用1毫升注射器,取下针头并连接0.22微米注射器过滤器,以将一滴柠檬酸铅分配到玻璃培养皿的底部。立即盖上。
    16. 将在负动作钳部分侧静止的栅格转移到柠檬酸铅滴上。立即关闭并设置一个计时器5分钟。
    17. 5分钟后,使用负反应钳取出电网,然后将电网扣入装满预先煮沸的蒸馏水的50ml烧杯中的一个中。
      。将网格进出水面〜30次。
    18. 前进到下一个装满煮沸蒸馏水的50毫升烧杯中,并将网格浸入水中和水中约30次(用同样的两个烧杯进行后续的柠檬酸铅清洗)。
    19. 将负作用钳放在台面上,并轻轻将一个三角形滤纸涂在铜网的外缘,以除去网格表面的水分。
    20. 将电网转移回电网盒并在电子显微镜观察前至少让其干燥至少几个小时。

  11. 用透射电子显微镜捕捉图像
    透射电子显微镜成像的目标是以公正的方式提供骨骼肌和线粒体超微结构的高分辨率图像,并获取足够的图像以获得真正意义上的显微解剖。
    1. 按面板上的开关旋钮(图9D)来点亮显示显微镜设置的屏幕。
    2. 按下HT(高张力)按钮(图9D),按钮上方的绿色指示灯将点亮,并在继续操作前检查排放计以确保返回接近0。
    3. 顺时针旋转长丝旋钮(图9D)〜23步,同时旋转旋钮达到饱和。这将需要大约2分钟,当发生这种情况时,示波器会发出蜂鸣声。显微镜的显示屏应亮绿灯(见注26)。

    4. 通过将支架顺时针旋转约四分之一圈然后将其完全拉出,从显微镜上取下试样支架(图9B)。
    5. 将样品架放在样品架支架上(图9F),并使用夹具升高样品架末端的样品夹(见注释27)。


      图9.菲利普斯CM10透射电子显微镜 A和B.电子显微镜及其零件的总体概述。 C.左控制面板主要用于控制电子束的强度,并用于调整用物镜聚焦图像或通过抬起荧光屏捕捉图像的屏幕。 D.正确的控制面板具有调节放大倍数,聚焦和整个电子显微镜系统监控屏幕所需的大部分功能。 E.真空屏幕显示了真空系统的示意图,其中包含用于监测系统性能的各种泵,阀门和压力计。 F.将样品架放在样品架支架上,并将夹子打开工具插入并提起网格夹。

    6. 使用负动作镊子将电网从电网箱中取出并转移到样本架底部的圆形电网支架上,确保电网完全停留在该区域内,并且不会稍微偏离外部。
    7. 使用剪辑工具将剪辑放下到您的网格上,将网格固定到位。
    8. 将样品架插入仪器并停下。一个红色预真空指示灯(图9B)将亮起,您将听到预真空点击。
    9. 等待红色的预真空指示灯熄灭,然后继续轻轻地逆时针旋转固定器将样品架完全插入高真空室中,真空将逐字拉出样品架(见注28)。
    10. 网格现在将以低倍率显示在屏幕上。
    11. 通过旋转两个夹点(-x和-y)来移动您的查看区域以找到您感兴趣的样本区域(图9C)。
    12. 一旦进入所需区域,用放大旋钮增加放大倍数(图9D)。屏幕将显示当前的放大倍数。
    13. 放大时,必须用两个手柄调整观看区域,以尽可能保持其居中。
    14. 放大倍率增加到〜500倍后,会发出哔哔声,提示您翻转聚光器光圈控制杆(图9B),从低倍光圈切换到高倍率。
    15. 随着您继续增加放大倍数,肌肉部分将开始进入视野;然而,由于没有增加电子束的强度,屏幕会变得越来越黑(见注29)。
    16. 在放大时,通过旋转强度旋钮轻轻调整电子束强度(图9C),以便清晰地看到肌肉超微结构。
    17. 用Focus Knob将肌肉超微结构带入焦点(图9D)。
    18. 打开电脑显示器并点击AMT软件图标。
    19. 根据经验,在打开软件和抬起观察屏幕之前,将电子束的强度调整为0.500,这将显示在'meter'旁边的屏幕上(参见注释30)。
    20. 用杠杆抬起荧光屏(图9C)并点击'Click for Live Image',样品的灰度图像将出现在显示器上(参见注释31)。
    21. 该软件自动设置为“QualityLive”并处于“调查”模式。
    22. 如果您想稍微调整焦点,请单击软件中“测量”按钮下的“焦点”按钮,这将放大图像,使您可以获得更好的焦点。
    23. 点击“调查”按钮返回到预览屏幕。
    24. 如果满意,请点击“最终图像”。
    25. 用相关信息命名图像并将图像保存在指定的文件夹中。
    26. 完成拍摄图像后,关闭软件并使用杠杆降低荧光屏。
    27. 使用放大旋钮减小您的缩放比例,同时逆时针旋转电子束强度以进行补偿。
    28. 一声〜500x的声音会提示您将聚光镜光圈杆从高倍率光圈翻转回低倍率。
    29. 继续将放大倍数降至〜75x,然后用-x和-y轴夹点重新对准观看区域。
    30. 从高真空室中取出样品架并取出您的网格。
    31. 完全按照上述方法将试样架放回高真空室,等待预真空完全插入后再启动。
    32. 逆时针旋转灯丝旋钮,直到观看屏幕不再亮起,并会发出哔哔声。
    33. 按下高压按钮,绿灯熄灭。
    34. 拉出面板调光旋钮关闭屏幕。

数据分析

对于我们大多数研究的重点是敲除或敲入信号分子并使肌肉受到诱导肥大或萎缩的各种条件的影响,我们对每组最少n = 3个样本进行TEM分析。我们使TRAF6 ff和TRAF6 mko小鼠对坐骨神经进行10天的去神经支配,然后利用透射电镜研究这些组之间TA肌肉的超微结构(Paul >等人,,2010)。使用TEM,与对照TRAF6 ff小鼠相比,我们能够通过显示失神经TRAF6 mko小鼠的萎缩和自噬减少来补充我们的分子生物学实验结果。在超微结构上,注意到TRAF6ff小鼠的失神经肌肉显示出SS和IMF线粒体的解体和线粒体与自噬体融合时自噬泡的增加;而这种表型在TRAF6 mko小鼠的失神经肌肉中大部分不存在(Paul等人,2010)。在另一项研究中,我们研究了TWEAK细胞因子在小鼠中的转基因过表达的作用,并使用TEM评估了幼稚状态下比目鱼肌的超微结构。发现TWEAK-TG小鼠的线粒体大小和数量与6个月龄时的野生型同窝出生仔猪相比显着减少(Hindi等人,2014年)。这通过计数每组三个样品中的SS线粒体来量化,每个样品至少使用10张图片在2200x放大。这与国际货币基金组织线粒体重复进行(印地语等人,,2014年)。我们以平均值±标准偏差(SD)呈现数据,并使用配对学生t检验来确定不同组间的统计学差异,其中p 0.05被认为具有统计学意义。最近,我们使用TEM来定性比较Tak1 fl / fl小鼠的比目鱼肌的线粒体超微结构与肌肉特异性敲除或Tak1 mKO小鼠的线粒体超微结构。我们发现Tak1 mKO小鼠具有丰富的空泡化线粒体,并且与Tak1 fl / fl相比,其嵴结构紊乱的扩大线粒体比例增加,小鼠(Hindi等人,2018)。我们的TEM观察与这些组之间进行的生物化学分析相结合,得出TAK1是这些小鼠的骨骼肌线粒体体内平衡所需的。
请参见图10,了解小鼠骨骼肌的子宫下和肌间纤维区域的电子显微照片示例。


图10.小鼠骨骼肌线粒体的TEM显微照片A.来自矢状切片的肌间纤维细胞线粒体(比例尺=1μm); B.来自矢状切片的亚线粒体线粒体(比例尺=1μm); C.具有明确定义的均匀嵴的健康亚线粒体线粒体(比例尺= 500nm); D.具有非均匀嵴的几个增大的亚线粒体线粒体(比例尺= 500nm)。

笔记

  1. 任何小鼠骨骼肌肉组织均可用于此分析。通常,在骨骼肌场中,将小鼠后肢肌肉分离出来进行研究,每个肌肉具有其自身的特征纤维类型组成。由于它们在纤维类型组成上的明显差异,我们使用小鼠胫骨前部(TA)和小鼠比目鱼进行TEM分析。 TA肌肉已被表征为快速糖酵解肌肉,而比目鱼肌已被表征为慢氧化肌肉(Schiaffino等人,2011; Kammoun等人 >,2014)。因此,有必要比较实验组之间相同的肌肉。有关如何从小鼠中分离后肢肌肉的有用视频,请参阅2017年印地语 et al。中的视频。隔离肌肉后,必须将整个肌肉浸入戊二醛尽可能快地保持肌肉处于其生理状态。
  2. 必须强调的是,您不需要太多的组织样本来进行TEM。与小鼠TA肌肉相比,小鼠比目鱼肌是相对较小的肌肉。对于比目鱼,我们通常将腹部肌肉切成一半,并使用一半用于我们的横截面分析,另一半用于我们的矢状切面分析,而不需要进一步修剪组织。另一方面,小鼠TA肌肉更大,需要修剪成一个细条,尺寸与比目鱼的直径相似。如果您需要此图表的帮助,请使用您的小鼠比目鱼肌作为对比。从那里,将你的钢带分成两部分,一部分用于横截面分析,另一部分用于矢状剖面分析。
  3. TEM组织处理中的许多化学物质都是剧毒的,特别是四氧化锇。四氧化锇能够固定角膜,因此请确保这些步骤是在通风良好的化学通风橱中完成的,并且佩戴个人防护装备。
  4. 确保黑色样品在各种稀释液中保持活动状态,并且不会粘在样品瓶的侧面。这可以通过调整转子转速来更好地适应日益粘稠的解决方案。添加下一个媒体解决方案后,请简单摇动或轻弹样本,以从底部或侧面移除它们。
  5. 尽量避免在块中引入气泡,特别是在样品本身周围。如果底部出现泡沫,那么这不是什么大问题。如果靠近样品有气泡,请使用注射器针头移动,然后将其从模具中取出。
  6. 如果没有足够的时间来治疗,塑料不会很难产生适当的薄切片。用剃刀刀片修剪模块时会立即实现此功能。如果塑料看起来对刀片的压力太大或者在修剪时存在明显的阻力,就好像刀片粘在一起(不仅仅是因为刀块很硬),那么这就很好地表明刀具块可以使用额外的在烤箱中完成固化。只需将块添加回烤箱,给他们更多的时间。或者,如果烤箱保持良好的温度而没有波动,将组织块留在较长的时间内没有问题。通常情况下,我已经把块放入烤箱,在周末治好,然后在周一回来修剪它们,没有问题。
  7. 切勿用裸手触摸玻璃杯,切片时手上的油会破坏玻璃杯的质量。此外,如果您觉得您需要补充说明和图表,请搜索'LKB Type 7801B视频',并且有一些可能有助于说明这些步骤。
  8. 首先,将塑料船用蜡附着在玻璃刀上可能是一个挑战。实践很重要。将足够的蜡转移到塑料舟的边缘,然后在蜡开始变硬之前尽可能快地将舟附着到玻璃上是困难的部分。
  9. 梯形金字塔是理想的,因此当块体遇到金刚石刀时,当刀片切割完整部分时,刀片遇到的表面积减小。这减轻了压力/摩擦,并确保了尽可能小的切片伪影的平滑切面。这也尽可能地保留了你的刀刃。其中一个视频在展示可应用于此的修剪过程方面做得相当不错,在2011年的Jenny中。
  10. 第一次很难调整水位到有银色反射的地方,但是很明显。当你从凸面转向银反射时,练习将水上下移动,然后在看到水开始从刀刃上拉开时凹入。请参阅Soplop et。 2009年的视频,这对厚和薄的切片有帮助,并显示切片在切割时如何浮在刀架水面上。
  11. 如果水从刀刃上脱落,则水位太低并且需要增加。有时只有一小部分刀发生这种情况,在这种情况下,您可以使用附着在端部的眼睫毛的工具将水冲刷到边缘上。
  12. 当您开始尝试切割切片时,您可能会注意到水很可能通过表面张力拉到了样品块的表面。如果发生这种情况,则需要停下来,用一张滤纸擦干脸部,然后调低水位。有时它需要靠近刀刃的一点,以便看到靠近刀刃的阴影,但不能到达水从边缘脱落的位置。
  13. 您不希望热板上的水蒸发得太快,否则这些部分会在玻璃板上不均匀地干燥,并且会以皱折的方式堆积起来。如果您发现这种情况发生,请调整温度以使其干燥得相对较慢。
  14. 同样,你不希望热板足够热到接触沸腾的地方,这会使甲苯胺蓝染色剂在各处冒出泡沫。相应地调整。你需要一个黄金色的环,干燥的污点发展,约2分钟。此时,你洗。
  15. 使用光学显微镜,主要目标是确定您的部分是否适当定位。您希望看到该部分没有明显缺陷,并且它是一个真正的横截面或纵向部分。
  16. 如果这些部分在光学显微镜中看起来更倾斜,那么您可以按照您认为有必要纠正切片角度的方式调整夹持块的卡盘。此时,根据您的改变的剧烈程度,您可能需要在前一个“修剪步骤”中用原始玻璃刀重新对着该块。
  17. 当手动推进超薄切片机手臂,然后旋转手臂切割切片时,确保这是一个缓慢而平稳的旋转。你不想旋转超薄切片机的手臂快速或积极,否则你可能会减少你的钻石刀的寿命。
  18. 如果样品金字塔足够小并且表面与钻石刀的边缘平行对齐,那么这些部分将从刀上脱落并形成不会彼此浮动的部分。当您使用网格收集部分时,这会很有帮助。另外,您可能想知道为什么我们使用半薄切片控件而不是超薄切片控件。我们的超薄切片机型号相对较旧,超薄切片控制功能依赖于我们发现不准确的热提前功能,这很可能是由于设备的使用年限所致。虽然可能需要更长的时间,因为您将截面厚度从0.5μm慢慢减小到0.100μm(100 nm),但我们各部分的质量和一致性得到了改善。因此,请随时尝试您的超薄切片控制功能,如果它工作正常,可能会节省一些时间。
  19. 黄金部分提供更高的对比度,但没有那么多的分辨率。银色部分提供高分辨率,但对比度较低。大多数情况下,除非您尝试解决肌浆网,T小管,等这样的困难结构,否则骨骼肌需要黄金切片。其中银部分可能是更优选的。
  20. 超薄部分非常脆弱,应该小心使用头发工具轻轻地引导并且不会破坏您的部分。可能需要一些时间才能获得良好的灵巧性,以便轻松地将您的薄切片安装在网格上而不会感到沮丧。实践是最好的方法。
  21. 尝试用滤纸三角形烘干网格时,请勿将滤纸直接放在您的部分上。通常有一个较厚的铜外环,您应该瞄准静止滤纸,以便您的部分不会被损坏。
  22. 沸腾蒸馏水可以去除水中的二氧化碳。这里的目标是防止我们的重金属污渍沉淀,这些污渍可能会使我们的网格聚集成块金属,干扰图像质量。在将这个过程纳入这个过程之后,这可能会令人非常沮丧。采取一切预防措施。
  23. 如果你不允许你的水足够冷,你会注意到当观察示波器中的部分时,对比度和染色看起来相对'平坦'。要有耐心,让它凉快。
  24. 在使用前直接过滤掉重金属污渍,有望去除溶液中的任何聚集物并导致更好的染色。
  25. 醋酸铀酰是光敏性的,在使用时和不使用时应避光避光。铅柠檬酸盐对二氧化碳很敏感,应该储存在有密封和封口膜的瓶子里,以防止聚集。在染色过程中,添加氢氧化钠是在反应发生时从染色皿中尽可能多地除去二氧化碳的必要步骤。
  26. 这个示波器使用钨丝,这就是为什么灯丝旋钮多次转动并快速连续转动的原因。如果有LaB <6>长丝,这将是一个更慢,更渐进的过程。这两者之间最大的区别是长丝的使用寿命和价格。钨丝适用于约100小时的使用,而LaB6细丝产生约1,000 + h的使用并且相当昂贵。
  27. 小心试样架,尤其是试样架工具,这种工具很脆弱,如果施加的力太大,会损坏。小心不要让手指接触到样品架下半部有光泽的部分。如果没有完成,手中的油将毁坏设备。
  28. 红灯熄灭后,您需要以恒定的压力和旋转轻轻旋转样品架。如果这样做过快而且积极,这个腔室内的压力可能会增加,高张力可能会作为一种安全机制启动。那么你需要重新开始。如果要在插入样品支架时监测腔室的压力,请按下就绪屏幕上真空的位置旁边的按钮,您将在其中看到底部真空系统和压力值的示意图。离子吸气泵(IGP)值将与您插入样品架的腔室中的压力相对应。尽量保持尽可能低(<50)。
  29. 不要太快调整光束强度,否则可能会在采样网格中烧伤一个孔。始终在交叉的右侧(或顺时针方向)工作。如果逆时针旋转强度旋钮,则会降低强度,并且在某个点您可以看到灯丝本身。这被称为交叉。如果您继续逆时针旋转强度旋钮,它会变得更亮更明亮。这将是交叉的'左'或'逆时针'侧。我们希望在右侧工作,增加强度是顺时针旋转的结果。
  30. 这款CM10菲利普斯电子显微镜采用新相机进行改造,并使用AMT软件获取图像。将电子束强度调整为0.500可确保相机不会被太强烈的光束饱和或烧毁。一旦软件查看图像,人们可以相应地调整强度,并通过软件中的直方图指示何时发生饱和或低信号。
  31. 有一个荧光屏幕,通过在观看屏幕中查看您的EM图像进行手动调整,例如通过光学显微镜的目标进行观察。这是一台更旧的电子显微镜,改装了一台计算机和数码相机以进行成像,从而获得高分辨率图像。相机位于显微镜的底部,光束从上方进入。
    荧光屏需要翻转,以便数码相机可以拍摄最终图像。

食谱

  1. 3%戊二醛
    1. 在通风橱中,打开打开10毫升8%戊二醛小瓶,并将内容物倒入50毫升锥形试管中。

    2. 加入16.6ml磷酸盐缓冲液稀释至3%戊二醛
    3. 多次翻转混合,可立即使用
  2. 0.1M磷酸钠缓冲液pH7.4
    1. 用量筒量取800毫升蒸馏水并加入磁力搅拌棒
    2. 打开搅拌器
    3. 称量3.1克磷酸二氢钠一水合物(NaH 2 PO 4·2H 2 O)并加入量筒中。
    4. 称量10.9克无水磷酸氢二钠(Na 2 HPO 4 4)并加入量筒中。
    5. 使用pH计检查pH值并确保其pH值为7.4
    6. 体积可达1000毫升,存放在玻璃瓶中
  3. 1%四氧化锇

    1. 在通风橱下用1克四氧化锇打开盛有安瓿的容器

    2. 准备100毫升的0.1 M磷酸盐缓冲液在一个干净的密封琥珀色玻璃瓶中
    3. 将打开的安瓿放入玻璃瓶中,使用干净的玻璃棒,将安瓿打开成几块。
    4. 立即用一个盖子密封玻璃瓶并旋转几次

    5. 允许锇晶体在室温下完全溶解(可以留在通风橱中过夜)
    6. 一旦溶解后,将琥珀色的瓶子存放在自己的密封容器内(带有盖子的金属罐或任何足够大的密封容器,以便将琥珀色的瓶子放入里面),然后放入4°C的冰箱中直至可以使用。 />
  4. 乙醇稀释
    准备乙醇稀释液(10%,25%,50%,75%,95%,100%):
    1. 100%时,将50毫升200毫升乙醇倒入50毫升圆锥形离心管中并相应标签。
    2. 对于95%,在量筒中量取47.5ml的200号乙醇,然后转移到50ml锥形离心管中。
      用蒸馏水将体积调至50毫升刻度并标记
    3. 对于75%,在量筒中量取37.5ml的200份Proof乙醇,然后转移到50ml锥形离心管中。
      用蒸馏水将体积调至50毫升刻度并标记
    4. 对于50%,在量筒中量取25.0ml的200 Proof乙醇,然后转移到50ml锥形离心管中。
      用蒸馏水将体积调至50毫升刻度并标记
    5. 对于25%,在量筒中量取12.5毫升的200毫升乙醇,然后转移到50毫升锥形离心管中。
      用蒸馏水将体积调至50毫升刻度并标记
    6. 10%时,量取刻度量筒中的5.0毫升200毫升乙醇,然后转移至50毫升锥形离心管中。用蒸馏水定容至50ml刻度并相应标签
  5. 嵌入媒体
    1. 将热板打开至60°C,将两种酸酐DDSA和NMA置于顶部以降低其粘度
    2. 在50毫升圆锥形离心管中量取40毫升EMbed 812,然后倒入一次性尿液标本容器。
    3. 使用新的50毫升锥形离心管,测量17毫升加热的DDSA,然后倒入同一个一次性尿液标本容器中。
    4. 使用新的50毫升锥形离心管,测量26毫升温热的NMA,并倒入同一个一次性尿液标本容器中。
    5. 使用分级移液器,将1毫升DMP-30转移到尿液标本容器中。
    6. 使用相同的移液管,确保除去任何剩余体积的EMbed 812,DDSA和NMA,并将其转移到尿液标本容器中的混合物中。
    7. 盖上50毫升圆锥形离心管并丢弃
    8. 用木制的涂药器彻底搅拌尿液标本容器中的嵌入混合物,直至均匀
    9. 嵌入介质可以储存在-20°C冷冻箱中,直到可以使用
    10. 如果您准备好了,只需根据需要准备环氧丙烷溶液,并根据您要加工的样品数量来决定
  6. 1%甲苯胺蓝染色
    1. 称取2克硼酸钠,溶于100毫升蒸馏水中
    2. 称取1克甲苯胺蓝粉末并溶于硼酸钠溶液中。

    3. 使用注射器过滤器过滤染色溶液并准备使用
  7. 4%醋酸铀酰(aq)
    1. 佩戴适当的个人防护装备,包括至少N-95呼吸器
    2. 在热板上的烧杯中煮沸500毫升蒸馏水以除去CO 2,然后关闭热板。
    3. 在通风橱中,称取4 g醋酸铀酰二水合物(耗尽)并转移至100 ml容量瓶中。
    4. 将100毫升近沸水转移到琥珀色玻璃瓶中,置于带磁力搅拌棒的搅拌器上搅拌过夜。
    5. 用Whatman#1过滤器将溶液过滤到干净的琥珀色玻璃瓶中。
    6. 适当地盖上瓶子并贴上标签

    7. 该解决方案可以在4°C的冰箱中储存数月。
    8. 通过将2毫升4%乙酸双氧铀溶液稀释到6毫升先前煮沸的蒸馏水中制备1%工作溶液,不含二氧化碳。
    9. 立即使用0.22μm注射器过滤器过滤1%醋酸双氧铀溶液,将过滤后的液滴直接放入染色皿中。
  8. 1N-NaOH
    1. 测量0.40克氢氧化钠

    2. 溶于8毫升水中
    3. 体积达10毫升的解决方案
  9. 雷诺的铅柠檬酸盐(Reynold's,1963)
    1. 佩戴适当的个人防护装备,包括至少一个N-95呼吸器
    2. 在热板上的烧杯中煮沸500毫升蒸馏水以除去CO 2,然后冷却。
    3. 在通风橱中称量1.33g硝酸铅Pb(NO 3)2并转移到50ml容量瓶中。
    4. 在通风橱中,称重1.76g柠檬酸钠Na 3(C 6 H 5 O 7) ·2H 2 O并转移到相同的容量瓶中
    5. 从上面将30毫升冷却的不含CO 2的蒸馏水加入到容量瓶和盖子中。
    6. 剧烈摇动悬浮液1分钟,然后静置30分钟,间歇摇动以确保硝酸铅完全转化为柠檬酸铅。该解决方案将保持阴天
    7. 每次小心添加1毫升1毫升NaOH(总共约8毫升)涡流并使用pH计检查pH值。 pH应该是12.0±0.1。如果测量结果高于12.1,则重新开始
    8. 解决方案将从多云转为清晰,不应该有任何混浊
    9. 总体积达50毫升,储存在密封的玻璃瓶中

致谢

作者要感谢Yann S. Gallot对他的手稿和有用的评论的全面阅读。作者还想声明不存在利益冲突或利益冲突。

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引用: Readers should cite both the Bio-protocol article and the original research article where this protocol was used:
  1. McMillan, J. D. and Eisenback, M. A. (2018). Transmission Electron Microscopy for Analysis of Mitochondria in Mouse Skeletal Muscle. Bio-protocol 8(10): e2455. DOI: 10.21769/BioProtoc.2455.
  2. Paul, P. K., Gupta, S. K., Bhatnagar, S., Panguluri, S. K., Darnay, B. G., Choi, Y. and Kumar, A. (2010). Targeted ablation of TRAF6 inhibits skeletal muscle wasting in mice. J Cell Biol 191(7): 1395-1411.
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