This work was supported by the Swedish Olympic Committee.

The present protocol using human biopsied skeletal muscle samples was approved by The Regional Committees on Health Research Ethics for Southern Denmark (S-20170198). Muscle biopsies were obtained through an incision in the skin from the vastus lateralis muscle using a Bergstr&#246;m needle with suction after local anesthesia was given subcutaneously (1-3 mL of Lidocaine 2% per incision). If isolated whole rat muscles were used, the animals were sacrificed by cervical dislocation before the muscle biopsies were obtained, in accordance with the guidelines of the animal ethics committee at Odense University Hospital, Denmark.
1. Primary fixation, post-fixation, embedding, sectioning, and contrasting
<ol>
	<li>Prepare 1.6 mL of primary fixative solution (2.5% glutaraldehyde in 0.1 M sodium cacodylate buffer (pH 7.3)) in a 2 mL micro centrifugation tube. Store it at 5 &#176;C for a maximum of 14 days.</li>
	<li>From the muscle biopsy or whole muscle, isolate a small specimen, which has a maximum diameter of 1 mm in any direction and is a bit longer in the longitudinal fiber direction than cross-sectionally (for orientation purposes).</li>
	<li>Place the specimen in the tube containing the cold primary fixation solution. Store it at 5 &#176;C for 24 h.</li>
	<li>Wash the specimen four times (15 min between each wash) in 0.1 M sodium cacodylate buffer (pH 7.3). Using transfer pipettes, remove the used buffer from the tube leaving the specimen untouched, and subsequently add the fresh buffer. 
	NOTE: Following the final wash, the specimen can be stored in the 0.1 M sodium cacodylate buffer at 5 &#176;C for several months11. The protocol can be paused here.</li>
	<li>Postfix with 1% osmium tetroxide (OsO4) and 1.5% potassium ferrocyanide (K4Fe(CN)6) in 0.1 M sodium cacodylate buffer (pH 7.3) for 120 min at 4 &#176;C. 
	NOTE: The use of 1.5% potassium ferrocyanide (K4Fe(CN)6) is essential for an optimal contrast of glycogen particles11,12,13.</li>
	<li>Rinse twice in double-distilled water at room temperature (RT).</li>
	<li>Dehydrate by submerging in a graded series of alcohol (ethanol) at RT using the following concentrations: 70% (10 min), 70% (10 min), 95% (10 min), 100% (10 min), and 100% (10 min). 
	NOTE: In each step, the specimen is submerged in ethanol, which is subsequently only partly removed to avoid drying of the specimen. Finally, the left-over ethanol is discarded.</li>
	<li>Infiltrate with graded mixtures of propylene oxide and epossidic resin at RT using the following volume ratios (propylene oxide/epossidic resin): 1/0 (10 min), 1/0 (10 min), 3/1 (45 min), 1/1 (45 min), 1/3 (45 min), 0/1 (overnight). The following day, embed specimens in 100% fresh epossidic resin in molds and polymerize at 60 &#176;C for 48 h. 
	NOTE: This graded method is per the previous protocols11,12. The protocol can be paused here.</li>
	<li>Cut ultra-thin (60-70 nm) sections of longitudinally oriented fibers and collect them on one-hole copper grids as follows.
	<ol>
		<li>Mount the block of a specimen on the ultramicrotome holder.</li>
		<li>Trim the block on the surface with a razor blade in order to reach the level of the tissue.</li>
		<li>Mount a diamond knife (ultracut 45) in front of the sample and align the sample surface parallel to the knife.</li>
		<li>Produce a semi-thin (1 &#181;m) section with the diamond knife to check the orientation of the sample. Stain the semi-thin section with toluidine blue for observation with light microscopy.</li>
		<li>Trim the block further to reduce the area of interest in order to get proper ultrathin sections.</li>
		<li>Cut ultrathin (60-70 nm) sections with a second diamond knife (ultracut 45).</li>
		<li>Collect 1-2 sections on one-hole copper grids using a Perfect Loop. 
		NOTE: One-hole copper grid has a single hole in the middle with Formvar supporting membrane.</li>
	</ol>
	</li>
	<li>Contrast sections with uranyl acetate and lead citrate by immersing the above grids in uranyl acetate solution (0.5% in double-distilled water) for 20 min, and then in lead citrate solution (1% in double-distilled water) for 15 min. Wash the grids in double-distilled water between and after the two stains. 
	NOTE: The protocol can be paused here.</li>
</ol>
2. Imaging
<ol>
	<li>Turn on the transmission electron microscope (operated at an accelerating voltage of 80 kV), computer, and image recording software. Record digital images with a digital slow-scan 2 k x 2 k CCD camera and the associated imaging software.</li>
	<li>Insert the grid with multiple sections in the microscope stage.</li>
	<li>Screen the grid initially at low magnification (e.g., x100) to determine the quality of sections (i.e., holes in the supporting membrane, debris, etc.) and choose the best quality sections. At low magnification, determine the direction of the muscle fibers.</li>
	<li>Next, increase the magnification with the beam centered on a peripheral fiber in the section. Focus the image at magnification above 30 k to ensure sufficient fine details in the image, guided by a Real-Time Fast Fourier Transformation, if available. Finally, record images with 1 s exposure time at the desired magnification.</li>
	<li>Acquire a total of 24 images of a randomly selected fiber, i.e., 12 images of the myofibrillar space and 12 images of the subsarcolemmal space, at a magnification between 10 k and 40 k. Ensure that the images are distributed across the length and width of the fiber in a randomized but systematic order to obtain unbiased results (Figure 1A). 
	NOTE: The optimal magnification depends on the available camera resolution and the size of the micrographs. The goal is to achieve a final resolution, where glycogen particle diameters can be measured within 1 nm steps, and to include a total area of the myofibrillar region of at least 70 &#181;m2 and a total length of the fiber of at least 25 &#181;m distributed into 12 images of the myofibrillar space and 12 images of the subsarcolemmal space per fiber, respectively. The 24 images per fiber will most likely give a precision (coefficient of error) of the volumetric content of the different pools of glycogen between 0.1 and 0.2 in individual fibers from human, rat, and mice skeletal muscles20,21,24 (Figure 2E).</li>
	<li>Repeat steps 2.4 and 2.5 until a total of 6-10 fibers are imaged. If needed, cut additional sections (separated by at least 150 &#181;m to avoid overlap of already imaged fibers) and repeat steps 1.9-2.5.</li>
</ol>
3. Image analyses
<ol>
	<li>Import images to ImageJ by clicking on File &#62; Open.</li>
	<li>Set global scale to match the original size of the image by clicking on Analyze &#62; Set Scale.</li>
	<li>Zoom in 100% by clicking on Image &#62; Zoom &#62; In.</li>
	<li>Measure the thickness of one Z-disc per image of the myofibrillar space (12 per fiber) using the Straight Line tool from the Tools menu (Figure 1D). Calculate the average Z-disc thickness of each of the 6-10 fibers.</li>
	<li>Define 2-3 fibers with the thickest average Z-disc as type 1 fibers and 2-3 fibers with the thinnest average Z-disc as type 2 fibers. Disregard the intermediate 2-4 fibers for further analyses (Figure 1E). 
	NOTE: The following steps are repeated for each of the 4-6 fibers from the sample. The glycogen volume fractions are estimated by point counting as described elsewhere25,26. The size of the grids is chosen to obtain a satisfactory high precision of the estimates. This is often obtained by achieving 250 hits, which then dictates the total number of points needed and, in turn, the area per point.</li>
	<li>Use the Segmented Line tool to measure the length of the outermost myofibril visible just below the subsarcolemmal region (Figure 2A). 
	NOTE: This length is used to express subsarcolemmal glycogen per surface area (i.e., length of the outermost myofibril multiplied by the thickness of the section (60 nm); see step 4.5). Therefore, only the subsarcolemmal region, which is represented by this length, is included in the analysis.</li>
	<li>Insert a grid by clicking on Analyze &#62; Tools &#62; Grid and set Area Per Point at 32,400 nm2. Count the number of hits within the available length in the 12 subsarcolemmal images, where a cross hits the subsarcolemmal glycogen (Figure 2A). A hit is defined as a glycogen particle being present in the upper-right corner of a cross.</li>
	<li>Insert a grid by clicking on Analyze &#62; Tools &#62; Grid and set Area Per Point at 160,000 nm2. Count the number of hits in the 12 myofibrillar images, where a cross hits the intramyofibrillar space (Figure 2B).</li>
	<li>Insert a grid by clicking on Analyze &#62; Tools &#62; Grid and set Area Per Point at 3,600 nm2. Count the number of hits in the 12 myofibrillar images, where a cross hits the intramyofibrillar glycogen (Figure 2C).</li>
	<li>Insert a grid by clicking on Analyze &#62; Tools &#62; Grid and set Area Per Point at 32,400 nm2. Count the number of hits in the 12 myofibrillar images, where a cross hits the intermyofibrillar glycogen (Figure 2D).</li>
	<li>Using the Straight Line tool, measure the diameter of five randomly chosen glycogen particles of each pool for each of the 12 images to obtain an average of 60 particles per pool per fiber. 
	NOTE: The average of 60 particles largely covers the variation within the fiber (Figure 2F).</li>
</ol>
4. Calculations
<ol>
	<li>Calculate the apparent area fraction (AA) of the intramyofibrillar space per myofibrillar space as the sum of all hits divided by the sum of all points from the 12 images (from step 3.8).</li>
	<li>Calculate the apparent area fraction of intramyofibrillar glycogen per myofibrillar area, intermyofibrillar glycogen per myofibrillar area, and subsarcolemmal glycogen per image area as the sum of all hits divided by the sum of all points from the 12 images (from steps 3.7, 3.9, and 3.10).</li>
	<li>Calculate the volume fraction (VV) of intramyofibrillar, intermyofibrillar, and subsarcolemmal glycogen, respectively, as the apparent area fraction (AA) minus the product of surface density (SV) with section thickness (t), where surface density is the numerical density of particles multiplied by the mean particle surface: 
	Vv = AA - (1 / 4) &#183; Sv &#183; t 
	where 
	Sv (&#181;m-1) = AA / ( (&#960; &#183; (((1 / 2) &#183; H)2)) &#183; (t + H))&#160;&#183; (&#960; &#183; H2) 
	t = 0.06 &#181;m 
	H = mean diameter of the particles (&#181;m) 
	NOTE: The volume fraction is smaller than the apparent area fraction due to the contribution of caps from particles with their center outside the slice25.</li>
	<li>To express intramyofibrillar glycogen per intramyofibrillar space, divide the area fraction of intramyofibrillar glycogen (step 4.2) by the area fraction of the intramyofibrillar space (step 4.1). The intermyofibrillar glycogen is expressed per myofibrillar space as calculated in the previous step (step 4.3).</li>
	<li>To express subsarcolemmal glycogen per surface area of the fiber (VS) (outermost myofibril), convert the volume fraction of glycogen to an absolute amount by multiplying with the volume of the image (product of area and section thickness) and dividing by the product of mean available length (from step 3.6) with section thickness (t).</li>
	<li>Estimate the total volumetric glycogen content using the values from steps 4.1, 4.4, and 4.5, as follows: 
	Myofibrillar glycogen = Intermyofibrillar glycogen + (intramyofibrillar glycogen &#183; area fraction of intramyofibrillar space) 
	By assuming an average fiber radius of 40 &#181;m27, the volume to surface ratio is 20:1, so total glycogen is: 
	Total glycogen (VV) = Myofibrillar glycogen + (subsarcolemmal glycogen (VS) / 20) 
	NOTE: The volume to surface ratio of 20:1 can vary from fiber to fiber depending on the actual fiber size and the size of the subsarcolemmal region. This is not taken into account with the present protocol.</li>
	<li>From this, the relative contribution from each pool is calculated as fractions of total glycogen: 
	Intermyofibrillar glycogen / Total glycogen =&#160;Intermyofibrillar glycogen / Total glycogen 
	Intramyofibrillar glycogen / Total glycogen = (Intramyofibrillar glycogen &#183; area fraction of intramyofibrillar space) / Total glycogen 
	Subsarcolemmal glycogen / Total glycogen = Subsarcolemmal glycogen / 20 / Total glycogen</li>
	<li>For each glycogen pool, calculate the coefficient of error (CE), which expresses the uncertainty of the glycogen estimate on a fiber level, based on the number of images (n), the total number of crosses in each image (x), and the number of crosses hitting glycogen in the relevant pool in each image (y) as follows28: 
	CE = n-1 &#183; &#8721;x2 &#183; (&#8721;x)-2 + &#8721;y2 &#183; (&#8721;y)-2 - 2&#8721;(xy) &#183; &#8721;x-1 &#183; &#8721;y-1</li>
</ol>

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K., de Paoli, F. V., Vissing, K. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Enhanced+glycogen+storage+of+a+subcellular+hot+spot+in+human+skeletal+muscle+during+early+recovery+from+eccentric+contractions.">Enhanced glycogen storage of a subcellular hot spot in human skeletal muscle during early recovery from eccentric contractions.</a> PLoS One. 10 (5), 0127808 (2015).</li><li>Sj&#246;str&#246;m, M., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Morphometric+analyses+of+human+muscle+fiber+types.">Morphometric analyses of human muscle fiber types.</a> Muscle Nerve. 5 (7), 538-553 (1982).</li><li>Gejl, K. D., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Local+depletion+of+glycogen+with+supramaximal+exercise+in+human+skeletal+muscle+fibres.">Local depletion of glycogen with supramaximal exercise in human skeletal muscle fibres.</a> Journal of Physiology. 595 (9), 2809-2821 (2017).</li><li>Stanley, W. C., Recchia, F. A., Lopaschuk, G. D. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Myocardial+substrate+metabolism+in+the+normal+and+failing+heart.">Myocardial substrate metabolism in the normal and failing heart.</a> Physiological Reviews. 85 (3), 1093-1129 (2005).</li></ol>

The authors declare no competing interests.

The critical step of the method is the use of reduced osmium by potassium ferrocyanide during post-fixation. The selectivity of this modified fixative for glycogen detection cannot be fully explained by chemistry, but also includes experimental findings demonstrating no detection of such particles in tissues known to be free of glycogen or in the extracellular space11.
Critical parameters are the precision of the estimates and the fiber-to-fiber variation. By following ...

Glycogen particles are composed of branched polymers of glucose and various associated proteins1 and constitute an important fuel during high metabolic demands2. Although not widely recognized, glycogen particles also constitute a local fuel, where some subcellular processes preferentially utilize glycogen despite the availability of other and more long-lasting fuels as plasma glucose and fatty acids3,4.
The importance of storing glycogen as a subcellular specific localized fuel has been discussed in several reviews5,6 mainly based on some of the earliest documentations of the subcellular distribution of glycogen by transmission electron microscopy (TEM)7,8. The first studies used different protocols to increase the contrast of glycogen from histochemical staining techniques to negative and positive stainings9,10. An important methodological development was the refined post-fixation protocol with the potassium ferrocyanide-reduced osmium11,12,13,14, which significantly improved the contrast of glycogen particles. This refined protocol was not used in some of the pioneering work on exercise-induced glycogen depletion15 but was re-introduced by Graham and colleagues16,17.
Based on the 2-dimensional images, the subcellular distribution of glycogen is most often described as glycogen particles located in three pools: subsarcolemmal (just beneath the surface membrane), intermyofibrillar (between the myofibrils), or intramyofibrillar (within the myofibrils). However, glycogen particles could also be described as associated with, for example, sarcoplasmic reticulum7 or nuclei18. In addition to the subcellular distribution, the advantage of TEM-estimated glycogen content is also that quantification can be conducted at the single fiber level. This allows investigation of fiber-to-fiber variability and correlative analyses with fiber types and cellular components as mitochondria and lipid droplets.
Here, the protocol for the TEM-estimated fiber type-specific volumetric content of the three common subcellular pools of glycogen (subsarcolemmal, intermyofibrillar, and intramyofibrillar) in skeletal muscle fibers is described. The method has been applied to skeletal muscles from humans19, rats20, and mice21; as well as birds and fish22; and cardiomyocytes from rats23.

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			<td>Taab</td>
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			<td>Leica</td>
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			<td>Ultrostain lead citrate 3%, stabilised solution</td>
			<td>Leica</td>
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			<td>Uranyl acetate dihydrate</td>
			<td>Polysciences</td>
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quantification of subcellular glycogen distribution in skeletal muscle fibers using transmission electron microscopy

With the use of transmission electron microscopy, high-resolution images of fixed samples containing individual muscle fibers can be obtained. This enables quantifications of ultrastructural aspects such as volume fractions, surface area to volume ratios, morphometry, and physical contact sites of different subcellular structures. In the 1970s, a protocol for enhanced staining of glycogen in cells was developed and paved the way for a string of studies on the subcellular localization of glycogen and glycogen particle size using transmission electron microscopy. While most analyses interpret glycogen as if it is homogeneously distributed within the muscle fibers, providing only a single value (e.g., an average concentration), transmission electron microscopy has revealed that glycogen is stored as discrete glycogen particles located in distinct subcellular compartments. Here, the step-by-step protocol from tissue collection to the quantitative determination of the volume fraction and particle diameter of glycogen in the distinct subcellular compartments of individual skeletal muscle fibers is described. Considerations on how to 1) collect and stain tissue specimens, 2) perform image analyses and data handling, 3) evaluate the precision of estimates, 4) discriminate between muscle fiber types, and 5) methodological pitfalls and limitations are included.

Using this protocol, glycogen particles appear black and distinct (Figures 1 and Figure 2). The normal values of glycogen are depicted in Figure 3. These data are based on a total of 362 fibers from 41 healthy young men as collected in different previous studies19,24,29,30,31</...

Watch this Scientific Journal Video about Quantification of Subcellular Glycogen Distribution in Skeletal Muscle Fibers using Transmission Electron Microscopy at JoVE.com

Quantification of Subcellular Glycogen Distribution in Skeletal Muscle Fibers using Transmission Electron Microscopy

A modified post-fixation procedure increases the contrast of glycogen particles in tissue. This paper provides a step-by-step protocol describing how to handle the tissue, conduct the imaging, and use stereological methods to obtain unbiased and quantitative data on fiber type-specific subcellular glycogen distribution in skeletal muscle.

With the use of transmission electron microscopy, high-resolution images of fixed samples containing individual muscle fibers can be obtained. This ...

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Biochemistry

3.9K Views.  University College South Denmark. A modified post-fixation procedure increases the contrast of glycogen particles in tissue. This paper provides a step-by-step protocol describing how to handle the tissue, conduct the imaging, and use stereological methods to obtain unbiased and quantitative data on fiber type-specific subcellular glycogen distribution in skeletal muscle.

Video: Quantification of Subcellular Glycogen Distribution in Skeletal Muscle Fibers using Transmission Electron Microscopy

Method to Visualize and Analyze Membrane Interacting Proteins by Transmission Electron Microscopy

Monotopic proteins exert their function when attached to a membrane surface, and such interactions depend on the specific lipid composition and on the availability of enough area to perform the function. Nanodiscs are used to provide a membrane surface of controlled size and lipid content. In the absence of bound extrinsic proteins, sodium phosphotungstate-stained nanodiscs appear as stacks of coins when viewed from the side by transmission electron microscopy (TEM). This protocol is therefore designed to intentionally promote stacking; consequently, the prevention of stacking can be interpreted as the binding of the membrane-binding protein to the nanodisc. In a further step, the TEM images of the protein-nanodisc complexes can be processed with standard single-particle methods to yield low-resolution structures as a basis for higher resolution cryoEM work. Furthermore, the nanodiscs provide samples suitable for either TEM or non-denaturing gel electrophoresis. To illustrate the method, Ca2+-induced binding of 5-lipoxygenase on nanodiscs is presented.

Many proteins perform their function when attached to membrane surfaces. The binding of extrinsic proteins on nanodisc membranes can be indirectly imaged by transmission electron microscopy. We show that the characteristic stacking (rouleau) of nanodiscs induced by the negative stain sodium phosphotungstate is prevented by the binding of extrinsic protein.

Monotopic proteins exert their function when attached to a membrane surface, and such interactions depend on the specific lipid composition and on the ...

Improved Protocol for Chromatin Immunoprecipitation from Mouse Skeletal Muscle

We describe an efficient and reproducible protocol for the preparation of chromatin from adult mouse skeletal muscle, a physically resistant tissue with a high content of structural proteins. Dissected limb muscles from adult mice are physically disrupted by mechanical homogenisation, or a combination of mincing and douncing, in a hypotonic buffer before formaldehyde fixation of the cell lysate. The fixed nuclei are purified by further cycles of mechanical homogenisation or douncing and sequential filtrations to remove cell debris. The purified nuclei can be sonicated immediately or at a later stage after freezing. The chromatin can be efficiently sonicated and is suitable for chromatin immunoprecipitation experiments, as illustrated by the profiles obtained for transcription factors, RNA polymerase II, and covalent histone modifications. The binding events detected using chromatin prepared by this protocol are predominantly those taking place in the muscle fiber nuclei despite the presence of chromatin from other fiber-associated satellite and endothelial cells. This protocol is therefore adapted to study gene regulation in the adult mouse skeletal muscle.

A novel protocol for the preparation of chromatin from adult mouse skeletal muscle adapted to the study of gene regulation in muscle fibers by chromatin immunoprecipitation is presented.

We describe an efficient and reproducible protocol for the preparation of chromatin from adult mouse skeletal muscle, a physically resistant tissue with a ...

Analysis of &#946;-Amyloid-induced Abnormalities on Fibrin Clot Structure by Spectroscopy and Scanning Electron Microscopy

This article presents methods for generating in vitro fibrin clots and analyzing the effect of beta-amyloid (A&#946;) protein on clot formation and structure by spectrometry and scanning electron microscopy (SEM). A&#946;, which forms neurotoxic amyloid aggregates in Alzheimer&#39;s disease (AD), has been shown to interact with fibrinogen. This A&#946;-fibrinogen interaction makes the fibrin clot structurally abnormal and resistant to fibrinolysis. A&#946;-induced abnormalities in fibrin clotting may also contribute to cerebrovascular aspects of the AD pathology such as microinfarcts, inflammation, as well as, cerebral amyloid angiopathy (CAA). Given the potentially critical role of neurovascular deficits in AD pathology, developing compounds which can inhibit or lessen the A&#946;-fibrinogen interaction has promising therapeutic value. In vitro methods by which fibrin clot formation can be easily and systematically assessed are potentially useful tools for developing therapeutic compounds. Presented here is an optimized protocol for in vitro generation of the fibrin clot, as well as analysis of the effect of A&#946; and A&#946;-fibrinogen interaction inhibitors. The clot turbidity assay is rapid, highly reproducible and can be used to test multiple conditions simultaneously, allowing for the screening of large numbers of A&#946;-fibrinogen inhibitors. Hit compounds from this screening can be further evaluated for their ability to ameliorate A&#946;-induced structural abnormalities of the fibrin clot architecture using SEM. The effectiveness of these optimized protocols is demonstrated here using TDI-2760, a recently identified A&#946;-fibrinogen interaction inhibitor.

Analysis of &amp;#946;-Amyloid-induced Abnormalities on Fibrin Clot Structure by Spectroscopy and Scanning Electron Microscopy

Presented here are two methods that can be used individually or in combination to analyze the effect of beta-amyloid on fibrin clot structure. Included is a protocol for creating an in vitro fibrin clot, followed by clot turbidity and scanning electron microscopy methods.

This article presents methods for generating in vitro fibrin clots and analyzing the effect of beta-amyloid (A&#946;) protein on clot formation and ...

Preparation of Sample Support Films in Transmission Electron Microscopy using a Support Floatation Block

Structure determination by cryo-electron microscopy (cryo-EM) has rapidly grown in the last decade; however, sample preparation remains a significant bottleneck. Macromolecular samples are ideally imaged directly from random orientations in a thin layer of vitreous ice. However, many samples are refractory to this, and protein denaturation at the air-water interface is a common problem. To overcome such issues, support films-including amorphous carbon, graphene, and graphene oxide-can be applied to the grid to provide a surface which samples can populate, reducing the probability of particles experiencing the deleterious effects of the air-water interface. The application of these delicate supports to grids, however, requires careful handling to prevent breakage, airborne contamination, or extensive washing and cleaning steps. A recent report describes the development of an easy-to-use floatation block that facilitates wetted transfer of support films directly to the sample. Use of the block minimizes the number of manual handling steps required, preserving the physical integrity of the support film, and the time over which hydrophobic contamination can accrue, ensuring that a thin film of ice can still be generated. This paper provides step-by-step protocols for the preparation of carbon, graphene, and graphene oxide supports for EM studies.

Sample preparation for cryo-electron microscopy (cryo-EM) is a significant bottleneck in the structure determination workflow of this method. Here, we provide detailed methods for using an easy-to-use, three-dimensionally printed block for the preparation of support films to stabilize samples for transmission EM studies.

Structure determination by cryo-electron microscopy (cryo-EM) has rapidly grown in the last decade; however, sample preparation remains a significant ...

Micropatterning Transmission Electron Microscopy Grids to Direct Cell Positioning within Whole-Cell Cryo-Electron Tomography Workflows

Whole-cell cryo-electron tomography (cryo-ET) is a powerful technology that is used to produce nanometer-level resolution structures of macromolecules present in the cellular context and preserved in a near-native frozen-hydrated state. However, there are challenges associated with culturing and/or adhering cells onto TEM grids in a manner that is suitable for tomography while retaining the cells in their physiological state. Here, a detailed step-by-step protocol is presented on the use of micropatterning to direct and promote eukaryotic cell growth on TEM grids. During micropatterning, cell growth is directed by depositing extra-cellular matrix (ECM) proteins within specified patterns and positions on the foil of the TEM grid while the other areas remain coated with an anti-fouling layer. Flexibility in the choice of surface coating and pattern design makes micropatterning broadly applicable for a wide range of cell types. Micropatterning is useful for studies of structures within individual cells as well as more complex experimental systems such as host-pathogen interactions or differentiated multi-cellular communities. Micropatterning may also be integrated into many downstream whole-cell cryo-ET workflows, including correlative light and electron microscopy (cryo-CLEM) and focused-ion beam milling (cryo-FIB).

The goal of this protocol is to direct cell adhesion and growth to targeted areas of grids for cryo-electron microscopy. This is achieved by applying an anti-fouling layer that is ablated in user-specified patterns followed by deposition of extra-cellular matrix proteins in the patterned areas prior to cell seeding.

Whole-cell cryo-electron tomography (cryo-ET) is a powerful technology that is used to produce nanometer-level resolution structures of macromolecules ...

Measurement of Protein Import Capacity of Skeletal Muscle Mitochondria

Mitochondria are key metabolic and regulatory organelles that determine the energy supply as well as the overall health of the cell. In skeletal muscle, mitochondria exist in a series of complex morphologies, ranging from small oval organelles to a broad, reticulum-like network. Understanding how the mitochondrial reticulum expands and develops in response to diverse stimuli such as alterations in energy demand has long been a topic of research. A key aspect of this growth, or biogenesis, is the import of precursor proteins, originally encoded by the nuclear genome, synthesized in the cytosol, and translocated into various mitochondrial sub-compartments. Mitochondria have developed a sophisticated mechanism for this import process, involving many selective inner and outer membrane channels, known as the protein import machinery (PIM). Import into the mitochondrion is dependent on viable membrane potential and the availability of organelle-derived ATP through oxidative phosphorylation. Therefore its measurement can serve as a measure of organelle health. The PIM also exhibits a high level of adaptive plasticity in skeletal muscle that is tightly coupled to the energy status of the cell. For example, exercise training has been shown to increase import capacity, while muscle disuse reduces it, coincident with changes in markers of mitochondrial content. Although protein import is a critical step in the biogenesis and expansion of mitochondria, the process is not widely studied in skeletal muscle. Thus, this paper outlines how to use isolated and fully functional mitochondria from skeletal muscle to measure protein import capacity in order to promote a greater understanding of the methods involved and an appreciation of the importance of the pathway for organelle turnover in exercise, health, and disease.

Mitochondria are key metabolic organelles that exhibit a high level of phenotypic plasticity in skeletal muscle. The import of proteins from the cytosol is a critical pathway for organelle biogenesis, essential for the expansion of the reticulum and the maintenance of mitochondrial function. Therefore, protein import serves as a barometer of cellular health.

Mitochondria are key metabolic and regulatory organelles that determine the energy supply as well as the overall health of the cell. In skeletal muscle, ...

Determination of Glucan Chain Length Distribution of Glycogen Using the Fluorophore-Assisted Carbohydrate Electrophoresis (FACE) Method

Glycogen particles are branched polysaccharides composed of linear chains of glucosyl units linked by &#945;-1,4 glucoside bonds. The latter are attached to each other by &#945;-1,6 glucoside linkages, referred to as branch points. Among the different forms of carbon storage (i.e., starch, &#946;-glucan), glycogen is probably one of the oldest and most successful storage polysaccharides found across the living world. Glucan chains are organized so that a large amount of glucose can quickly be stored or fueled in a cell when needed. Numerous complementary techniques have been developed over the last decades to solve the fine structure of glycogen particles. This article describes Fluorophore-Assisted Carbohydrate Electrophoresis (FACE). This method quantifies the population of glucan chains that compose a glycogen particle. Also known as chain length distribution (CLD), this parameter mirrors the particle size and the percentage of branching. It is also an essential requirement for the mathematical modeling of glycogen biosynthesis.

Determination of Glucan Chain Length Distribution of Glycogen Using the Fluorophore-Assisted Carbohydrate Electrophoresis FACE Method

In the present protocol, the Fluorophore-Assisted Carbohydrate Electrophoresis (FACE) technique is used to determine the chain length distribution (CLD) and the average chain length (ACL) of glycogen.

Glycogen particles are branched polysaccharides composed of linear chains of glucosyl units linked by &#945;-1,4 glucoside bonds. The latter are attached ...

Routine Collection of High-Resolution cryo-EM Datasets Using 200 KV Transmission Electron Microscope

Cryo-electron microscopy (cryo-EM) has been established as a routine method for protein structure determination during the past decade, taking an ever-increasing share of published structural data. Recent advances in TEM technology and automation have boosted both the speed of data collection and quality of acquired images while simultaneously decreasing the required level of expertise for obtaining cryo-EM maps at sub-3 &#197; resolutions. While most of such high-resolution structures have been obtained using state-of-the-art 300 kV cryo-TEM systems, high-resolution structures can be also obtained with 200 kV cryo-TEM systems, especially when equipped with an energy filter. Additionally, automation of microscope alignments and data collection with real-time image quality assessment reduces system complexity and assures optimal microscope settings, resulting in increased yield of high-quality images and overall throughput of data collection. This protocol demonstrates the implementation of recent technological advances and automation features on a 200 kV cryo-transmission electron microscope and shows how to collect data for the reconstruction of 3D maps that are sufficient for de novo atomic model building. We focus on best practices, critical variables, and common issues that must be considered to enable the routine collection of such high-resolution cryo-EM datasets. Particularly the following essential topics are reviewed in detail: i) automation of microscope alignments, ii) selection of suitable areas for data acquisition, iii) optimal optical parameters for high-quality, high-throughput data collection, iv) energy filter tuning for zero-loss imaging, and v) data management and quality assessment. Application of the best practices and improvement of achievable resolution using an energy filter will be demonstrated on the example of apo-ferritin that was reconstructed to 1.6 &#197;, and Thermoplasma acidophilum 20S proteasome reconstructed to 2.1-&#197; resolution using a 200 kV TEM equipped with an energy filter and a direct electron detector.

High-resolution cryo-EM maps of macromolecules can be also achieved by using 200 kV TEM microscopes. This protocol shows the best practices for setting accurate optics alignments, data acquisition schemes, and selection of imaging areas that are all essential for the successful collection of high-resolution datasets using a 200 kV TEM.

Cryo-electron microscopy (cryo-EM) has been established as a routine method for protein structure determination during the past decade, taking an ...

High-Throughput Quantification of the Synthesis and Distribution of mtDNA

High-Throughput Image-Based Quantification of Mitochondrial DNA Synthesis and Distribution

The vast majority of cellular processes require a continuous supply of energy, the most common carrier of which is the ATP molecule. Eukaryotic cells produce most of their ATP in the mitochondria by oxidative phosphorylation. Mitochondria are unique organelles because they have their own genome that is replicated and passed on to the next generation of cells. In contrast to the nuclear genome, there are multiple copies of the mitochondrial genome in the cell. The detailed study of the mechanisms responsible for the replication, repair, and maintenance of the mitochondrial genome is essential for understanding the proper functioning of mitochondria and whole cells under both normal and disease conditions. Here, a method that allows the high-throughput quantification of the synthesis and distribution of mitochondrial DNA (mtDNA) in human cells cultured in vitro is presented. This approach is based on the immunofluorescence detection of actively synthesized DNA molecules labeled by 5-bromo-2'-deoxyuridine (BrdU) incorporation and the concurrent detection of all the mtDNA molecules with anti-DNA antibodies. Additionally, the mitochondria are visualized with specific dyes or antibodies. The culturing of cells in a multi-well format and the utilization of an automated fluorescence microscope make it easier to study the dynamics of mtDNA and the morphology of mitochondria under a variety of experimental conditions in a relatively short time.

Author Spotlight: High-Throughput Image-Based Quantification of Mitochondrial DNA Synthesis and Distribution  

A procedure for studying the dynamics of mitochondrial DNA (mtDNA) metabolism in cells using a multi-well plate format and automated immunofluorescence imaging to detect and quantify mtDNA synthesis and distribution is described. This can be further used to investigate the effects of various inhibitors, cellular stresses, and gene silencing on mtDNA metabolism.

The vast majority of cellular processes require a continuous supply of energy, the most common carrier of which is the ATP molecule. Eukaryotic cells ...

MyDoBID- A Novel Technique for Identifying Muscle Fiber Types Efficiently

Fiber Type Identification of Human Skeletal Muscle

The technique described here can be used to identify specific myosin heavy chain (MHC) isoforms in segments of individual muscle fibers using dot blotting, hereafter referred to as Myosin heavy chain detection by Dot Blotting for IDentification of muscle fiber type (MyDoBID). This protocol describes the process of freeze-drying human skeletal muscle and isolating segments of single muscle fibers. Using MyDoBID, type I and II fibers are classified with MHCI- and IIa-specific antibodies, respectively. Classified fibers are then combined into fiber type-specific samples for each biopsy.
The total protein in each sample is determined by Sodium Dodecyl-Sulfate Polyacrylamide Gel Electrophoresis (SDS-PAGE) and UV-activated gel technology. The fiber type of samples is validated using western blotting. The importance of performing protein loading normalization to enhance target protein detection across multiple western blots is also described. The benefits of consolidating classified fibers into fiber type-specific samples compared to single-fiber western blots, include sample versatility, increased sample throughput, shorter time investment, and cost-saving measures, all while retaining valuable fiber type-specific information that is frequently overlooked using homogenized muscle samples. The purpose of the protocol is to achieve accurate and efficient identification of type I and type II fibers isolated from freeze-dried human skeletal muscle samples.
These individual fibers are subsequently combined to create type I and type II fiber type-specific samples. Furthermore, the protocol is extended to include the identification of type IIx fibers, using Actin as a marker for fibers that were negative for MHCI and MHCIIa, which are confirmed as IIx fibers by western blotting. Each fiber type-specific sample is then used to quantify the expression of various target proteins using western blotting techniques.

Author Spotlight: Deciphering the Mysteries of Skeletal Muscle Fiber Types Using the MyDoBID Technique 

This protocol demonstrates single-fiber isolation from freeze-dried human skeletal muscle and fiber-type classification according to Myosin heavy chain (MHC) isoform using the dot blotting technique. Identified MHC I and II fiber samples can then be further analyzed for fiber type-specific differences in protein expression using western blotting.

Quantification of Subcellular Glycogen Distribution in Skeletal Muscle Fibers using Transmission Electron Microscopy

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