This work was supported by grants from the Fonds National de la Recherche Scientifique (FNRS-Cr&#233;dit de Recherche J.0022.20) and the Soci&#233;t&#233; Francophone du Diab&#232;te (SFD-Roche Diabetes Care).C.M.S. is the recipient of a Ph.D. fellowship from the FRIA (FNRS). M.A.D.-L.d.C. received a fellowship from the Wallonie-Bruxelles International Excellence Program.
The authors thank Alice Monnier for her contribution to the development of this protocol and Caroline Bouzin for her expertise and technical help in the image acquisition process. We also thank the 2IP-IREC imaging platform for access to the cryostat and the microscopes (2IP-IREC Imaging Platform, Institute of Experimental and Clinical Research, Universit&#233; Catholique de Louvain, 1200 Brussels, Belgium). Finally, the authors would like to thank Nicolas Dubuisson, Romain Versele, and Michel Abou-Samra for constructive criticism of the manuscript. Some of the figures of these article were created with BioRender.com.

The authors have no conflicts of interest to declare.

The protocol detailed here describes an efficient method to quantify LDs tagged with Bodipy on a fiber-type- and subcellular-specific basis. In recent years, classical lipid dyes, such as ORO or Sudan Black B, have been substituted with a new array of cell-permeable, lipophilic, fluorescent dyes that bind to neutral lipids (e.g., Bodipy). Available as different conjugates, Bodipy has been proven very effective at tagging LDs to study their morphology, dynamics, and interaction with other organelles, not only in different...

Skeletal muscle lipid infiltration, known as myosteatosis, increases with obesity and ageing. Myosteatosis is negatively correlated with muscle mass and strength and with insulin sensitivity1. Moreover, recent studies indicate that the degree of myosteatosis could be used as a prognostic factor for other conditions such as cardiovascular disease2, non-alcoholic fatty liver disease3, or cancer4. Lipids can accumulate in skeletal muscle between muscle fibers as extramyocellular lipids or within the fibers, as intramyocellular lipids (IMCLs). IMCLs are predominantly stored as triglycerides in lipid droplets (LDs) that are used as metabolic fuel during physical exercise5,6. However, when lipid supply exceeds demand, or when mitochondria become dysfunctional, IMCLs will be implicated in muscle insulin resistance, as seen in metabolically unhealthy, obese individuals and in type 2 diabetes patients7. Intriguingly, endurance athletes have similar, if not higher, levels of IMCLs to those found in obese patients with type 2 diabetes mellitus, while maintaining high insulin sensitivity. This phenomenon is described as the &#34;athlete&#39;s paradox&#34;8,9, and is explained by a more nuanced appraisal of muscle LDs, related to their size, density, localization, dynamics, and lipid species composition.
First, LD size is inversely correlated to insulin sensitivity and physical fitness10,11. In fact, smaller LDs exhibit a relatively greater surface area for lipase action and, thus, potentially have a greater capacity to mobilize lipids12. Second, LD density (number/surface) plays a controversial role in insulin action8,10; yet, it seems to be increased in athletes. Third, the subcellular localization of LDs is important, since LDs located just below the surface membrane (subsarcolemmal or peripheral) exert a more deleterious effect on insulin sensitivity than central ones8,9,13. The latter provide fuel to central mitochondria, which have a greater respiratory activity and are more specialized to meet the high energy demand required for contraction14. By contrast, peripheral LDs supply subsarcolemmal mitochondria, which are involved in membrane-related processes8. Finally, beyond triglycerides, specific complex lipids within the muscle may be more deleterious than others. For instance, diacylglycerol, long-chain acyl-CoA, and ceramides may accumulate in muscle when the triglyceride turnover rate is low, thereby impairing insulin signaling9,15. Returning to the &#34;athlete&#39;s paradox,&#34; endurance athletes have a high number of smaller central LDs with elevated turnover rates in type I (oxidative) fibers, while obese and diabetic patients have larger peripheral LDs with low turnover rates in type II (glycolytic) fibers8,15,16. In addition to their role in energy storage and release, LDs via derived fatty acids (FA) and a coat protein (perilipin 5) could also function as critical players involved in the transcriptional regulation of FA oxidation and mitochondrial biogenesis8. Because of their crucial implications in physiology and pathophysiology, in-depth studies on LDs dynamics and functions are warranted.
Although there are several techniques to study IMCLs, they are not all suitable to accurately quantify LD size, density, and distribution in a fiber-specific manner. For example, the assessment of IMCLs by magnetic resonance spectroscopy, while being non-invasive, offers a level of resolution that is not sufficient to study the size and precise location of LDs within the fiber, and it is not fiber-type specific17,18. Likewise, biochemical techniques performed on whole-muscle homogenates19 cannot assess the location and size of lipids. Consequently, the most adequate method to analyze LD morphology and location is quantitative transmission electronic microscopy13, but this technique is expensive and time-consuming. Therefore, confocal fluorescence imaging on preparations with dyes such as Oil Red O (ORO)20,21, monodansylpentane (MDH)22, or Bodipy23,24,25, has emerged as the best tool for these studies.
Here, a complete protocol is described, including tissue sampling and processing, Bodipy staining, and confocal image acquisition and analysis to quantify LD size, number, and localization in mouse muscle cryosections. Since IMCLs are not evenly distributed among oxidative and glycolytic fibers, and each fiber type regulates LD dynamics differently, the study of IMCLs must be fiber-type specific16,25,26,27. Therefore, this protocol uses immunofluorescence on serial sections to identify myosin heavy chain (MyHC) isoform(s) expressed by each fiber. Another advantage of this protocol is the simultaneous processing of a glycolytic (extensor digitorum longus, EDL) and an oxidative (soleus) muscle placed side-by-side before freezing (Figure 1). This simultaneous processing not only saves time but also avoids variability due to separate processing of the samples.
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/63718/63718fig01.jpg" /> 
Figure 1: Schematic overview of the procedure. After muscle dissection (1), similar-size selected muscles are prepared and frozen together (2). Serial transverse sections of 10 &#181;m are obtained using a cryostat and directly mounted on adhesion slides (3). From two serial slides, the first (4A) is immunolabeled for laminin and stained with Bodipy to recognize LDs and the second (4B) is immunostained with antibodies against MyHCs for the recognition of muscle fiber types. Images are acquired using a confocal microscope for Bodipy (5A) and an epifluorescence microscope for muscle fiber types (5B). Images are analyzed in Fiji by applying a threshold and quantifying particles (6A) to obtain the number, average size, density, and percentage of the total area occupied by LDs (7) or counting cells (6B) to obtain the percentage of fibers of each type in the section (7). Abbreviations: LDs = lipid droplets; EDL = extensor digitorum longus; MyHCs = myosin heavy chain isoforms. <a href="https://jove.unicartagenaproxy.elogim.com/files/ftp_upload/63718/63718fig01large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

<table><tbody><tr><td>&lt;strong&gt;Equipment&lt;/strong&gt;</td><td></td><td></td><td></td></tr><tr><td>AxioCam 506 mono 6 Mpix camera</td><td>Zeiss</td><td></td><td></td></tr><tr><td>AxioCam MRm 1.4MPix CCD camera&amp;nbsp;</td><td>Zeiss</td><td></td><td></td></tr><tr><td>Chemical hood</td><td>Potteau Labo</td><td>EN-14175</td><td></td></tr><tr><td>Confocal microscope</td><td>Zeiss</td><td>LSM800</td><td></td></tr><tr><td>Cork discs (&amp;oslash; 20 mm, 3 mm thick)&amp;nbsp;</td><td>Electron Microscopy Sciences</td><td>63305</td><td></td></tr><tr><td>Cryo-Gloves</td><td>Tempshield</td><td>16072252</td><td></td></tr><tr><td>Cryostat&amp;nbsp;</td><td>Thermo Scientific&amp;nbsp;</td><td>Microm Cryo Star HM 560</td><td></td></tr><tr><td>Dissecting Stereo Microscope SMZ745</td><td>Nikon</td><td></td><td></td></tr><tr><td>Dry Ice</td><td></td><td></td><td></td></tr><tr><td>Dumont Forceps</td><td>F.S.T</td><td>11295-10</td><td></td></tr><tr><td>Epifluorescence microscope</td><td>Zeiss</td><td>AxioImage-Apotome Z1</td><td></td></tr><tr><td>Extra Fine Bonn Scissors</td><td>F.S.T</td><td>14084-08</td><td></td></tr><tr><td>FisherBrand Disposable Base Molds (0.7 x 0.7 cm)</td><td>ThermoFisher</td><td>22-363-552</td><td>Used to cut a piece to hold the muscle on the cork for freezing</td></tr><tr><td>Glass petri dish (H 25&amp;nbsp;mm, &amp;oslash; 150&amp;nbsp;mm)</td><td>BRAND Petri dish, MERK</td><td>BR455751</td><td>Used to place the muscles on ice during dissection</td></tr><tr><td>ImmEdge Hydrophobic barrier PAP Pen</td><td>Vector Labs</td><td>H-4000</td><td>Used to create an hidrophobic barrier around the muscle sections</td></tr><tr><td>Incubator</td><td>MMM Medcenter</td><td>Incucell 707&amp;nbsp;</td><td></td></tr><tr><td>Microscope Cover Glasses (24x50 mm)</td><td>Assistent&amp;nbsp;</td><td>40990151</td><td></td></tr><tr><td>Microscope Slide Boxes&amp;nbsp;</td><td>Kartell</td><td>278</td><td>Used as humid chambers for immunohistochemistry</td></tr><tr><td>Neck holder</td><td>Linie zwo</td><td>SB-035X-02</td><td>Used as strap to hold the stainless steel tumbler</td></tr><tr><td>No 15 Sterile Carbon Steel Scalpel Blade</td><td>Swann-Morton</td><td>0205</td><td></td></tr><tr><td>Paint brushes</td><td>Van Bleiswijck</td><td>Amazon B07W7KJQ2X</td><td>Used to handle cryosections</td></tr><tr><td>Permanent Marker Pen Black</td><td>Klinipath/VWR</td><td>98307-R</td><td>Used to label slides</td></tr><tr><td>Pierce Fixation Forceps</td><td>F.S.T</td><td>18155-13</td><td></td></tr><tr><td>Polystyrene Box&amp;nbsp;</td><td></td><td></td><td>H 12 cm x L 25 cm x W 18 cm, used as a liquid nitrogen container and to transport the samples to the cryostat</td></tr><tr><td>Scalpel Handle, 125 mm (5&quot;), No. 3</td><td>Aesculap</td><td>BB073R</td><td></td></tr><tr><td>Stainless Steel Cup 10oz&amp;nbsp;</td><td>Eboxer</td><td>B07GFCBPFH</td><td>Tumbler to fill with isopentene for muscle freezing</td></tr><tr><td>Superfrost Ultra Plus slides</td><td>ThermoFisher</td><td>J1800AMNZ</td><td></td></tr><tr><td>Surgical tweezers 1/2 teeth</td><td>Medische Vakhandel</td><td>1303152</td><td>Also called &quot;Rat teeth tweezers&quot;</td></tr><tr><td>Vannas Spring Scissors - 3 mm Cutting Edge</td><td>F.S.T</td><td>15000-00</td><td></td></tr><tr><td>Weighing boats</td><td>VWR international</td><td>611-2249</td><td></td></tr><tr><td>Whole-Slide Scanner for Fluorescence</td><td>Zeiss</td><td>Axio Scan.Z1</td><td></td></tr><tr><td>&lt;strong&gt;Reagents&lt;/strong&gt;</td><td></td><td></td><td></td></tr><tr><td>Alexa Fluor 405 Goat Anti-Mouse IgG2b</td><td>Sigma-Aldrich</td><td>SAB4600477</td><td>Used at a final concentration of 1:500</td></tr><tr><td>Alexa Fluor 488 Goat Anti-Mouse IgG1</td><td>ThermoFisher</td><td>A-21121</td><td>Used at a final concentration of 1:500</td></tr><tr><td>Alexa Fluor 568 Goat Anti-Mouse IgM</td><td>Abcam</td><td>ab175702</td><td>Used at a final concentration of 1:1,000</td></tr><tr><td>Alexa Fluor 647 goat anti rat-IgG (H+L) secondary antibody</td><td>ThermoFisher</td><td>A-21247</td><td>Used at a final concentration of 1:500</td></tr><tr><td>BODIPY-493/503 (4,4-difluoro-1,3,5,7,8-pentametil-4-bora-3a,4a-diaza-s-indaceno)</td><td>ThermoFisher</td><td>D3922</td><td>Used at a final concentration of 1 &amp;micro;g/mL</td></tr><tr><td>BODIPY-558/568 C12 (4,4-Difluoro-5-(2-Thienyl)-4-Bora-3a,4a-Diaza-s-Indacene-3-Dodecanoic Acid)</td><td>ThermoFisher</td><td>D3835</td><td>Used at a final concentration of 1 &amp;micro;g/mL</td></tr><tr><td>DAPI (4&#039;,6-diamidino-2-phenylindole)</td><td>ThermoFisher</td><td>D1306</td><td>Used at a final concentration of 0.5 &amp;micro;g/mL</td></tr><tr><td>Dimethyl Sulfoxide (DMSO)</td><td>Sigma-Aldrich</td><td>D-8418</td><td>Used to solve Bodipy for the 1 mg/mL stock solution. &lt;strong&gt;CAUTION&lt;/strong&gt;: Toxic and flammable. Vapors may cause irritation. Manipulate in a fume hood. Avoid direct contact with skin. Wear rubber gloves, protective eye goggles.</td></tr><tr><td>Formaldehyde solution 4%, buffered, pH 6.9</td><td>Sigma-Aldrich</td><td>1004969011</td><td>&lt;strong&gt;CAUTION&lt;/strong&gt;: May cause an allergic skin reaction. Suspected of causing genetic defects. May cause cancer. Manipulate in a fume hood. Avoid direct contact with skin. Wear rubber gloves, protective eye goggles.</td></tr><tr><td>Isopentane GPR RectaPur</td><td>VWR international</td><td>24872.298</td><td>&lt;strong&gt;CAUTION&lt;/strong&gt;: Extremely flammable liquid and vapor. May be fatal if swallowed and enters airways. May cause drowsiness or dizziness. Repeated exposure may cause skin dryness or cracking. Wear protective gloves/protective clothing/eye protection/face protection.</td></tr><tr><td>Liquid Nitrogen</td><td></td><td></td><td>&lt;strong&gt;CAUTION&lt;/strong&gt;:&amp;nbsp; Extremely cold. Wear gloves. Handle slowly to minimize boiling and splashing and in well ventilated areas. Use containers designed for low-temperature liquids.</td></tr><tr><td>Mouse on mouse Blocking Reagent&amp;nbsp;</td><td>Vector Labs</td><td>MKB-2213-1</td><td>Used at concentration of 1:30</td></tr><tr><td>Myosin heavy chain Type I (BA-D5-s Primary Antibody) Gene: MYH7, monoclonal bovine anti mouse IgG2b</td><td>DSHB University of Iowa</td><td>BA-D5-supernatant</td><td>Used at a final concentration of 1:10</td></tr><tr><td>Myosin heavy chain Type IIA (SC-71-s Primary Antibody) Gene:&amp;nbsp; MYH2, Monoclonal bovine anti mouse IgG1</td><td>DSHB University of Iowa</td><td>SC-71-supernatant</td><td>Used at a final concentration of 1:10</td></tr><tr><td>Myosin heavy chain Type IIX (6H1-s Primary Antibody), Gene:&amp;nbsp; MYH1, Monoclonal rabbit anti mouse IgM</td><td>Developmental Studies Hybridoma Bank, University of Iowa</td><td>6H1-supernatant</td><td>Used at a final concentration of 1:5</td></tr><tr><td>Normal Goat Serum (NGS)</td><td>Vector Labs</td><td>S-1000</td><td></td></tr><tr><td>PBS 0.1 M</td><td></td><td></td><td>Commonly used on histology laboratories</td></tr><tr><td>ProLong Gold Antifade Mountant</td><td>Invitrogen&amp;nbsp;</td><td>P36930</td><td></td></tr><tr><td>Rat anti-Laminin-2 (&amp;alpha;-2 Chain) primary antibody (monoclonal)</td><td>Sigma-Aldrich</td><td>L0663</td><td>Used at a final concentration of 1:1,000</td></tr><tr><td>Tissue-Tek O.C.T compound</td><td>Sakura&amp;nbsp;</td><td>4583</td><td></td></tr><tr><td>&lt;strong&gt;Software&lt;/strong&gt;</td><td></td><td></td><td></td></tr><tr><td>Adobe Illustrator CC</td><td>Adobe Inc.</td><td></td><td>Used to design the figures</td></tr><tr><td>Adobe Photoshop</td><td>Adobe Inc.</td><td></td><td>Confocal software</td></tr><tr><td>BioRender</td><td>https://biorender.com/</td><td></td><td>Used to design the figures</td></tr><tr><td>Fiji/ImageJ</td><td>https://imagej.net/software/fiji/</td><td></td><td>Used to analyse the acquired images</td></tr><tr><td>Microsoft PowerPoint</td><td>Microsoft</td><td></td><td>Used to reconstruct the histology of the whole muscle after scanning the fiber types</td></tr><tr><td>Zen Blue 2.6</td><td>Zeiss</td><td></td><td>Used to reconstruct the histology of the whole muscle after scanning the fiber types</td></tr></tbody></table>

fiber type and subcellular-specific analysis of lipid droplet content in skeletal muscle

Skeletal muscle lipid infiltration, known as myosteatosis, increases with obesity and ageing. Myosteatosis has also recently been discovered as a negative prognostic factor for several other disorders such as cardiovascular disease and cancer. Excessive lipid infiltration decreases muscle mass and strength. It also results in lipotoxicity and insulin resistance depending on total intramyocellular lipid content, lipid droplet (LD) morphology, and subcellular distribution. Fiber type (oxidative vs glycolytic) is also important, since oxidative fibers have a greater capacity to utilize lipids. Because of their crucial implications in pathophysiology, in-depth studies on LD dynamics and function in a fiber type-specific manner are warranted.
Herein, a complete protocol is presented for the quantification of intramyocellular lipid content and analysis of LD morphology and subcellular distribution in a fiber type-specific manner. To this end, serial muscle cryosections were stained with the fluorescent dye Bodipy and antibodies against myosin heavy chain isoforms. This protocol enables the simultaneous processing of different muscles, saving time and avoiding possible artifacts and, thanks to a personalized macro created in Fiji, the automatization of LD analysis is also possible.

The protocol described herein provides an efficient method to easily quantify LDs in a fiber type and subcellular-specific manner. It shows how, by freezing together two muscles of similar size, such as the EDL and the soleus, the time and resources spent on the following steps are reduced by half.
A complete protocol is provided for immunostaining, image acquisition, and analysis of the different MyHC isoforms expressed in adult mouse muscles. This protocol is based on the one first designed ...

Watch this Scientific Journal Video about Fiber Type and Subcellular-Specific Analysis of Lipid Droplet Content in Skeletal Muscle at JoVE.com

Fiber Type and Subcellular-Specific Analysis of Lipid Droplet Content in Skeletal Muscle

Increasing evidence indicates that excessive infiltration of lipids inside skeletal muscle results in lipotoxicity and diabetes. Here, we present a complete protocol, including tissue processing, staining with Bodipy, image acquisition, and analysis, to quantify the size, density, and subcellular distribution of lipid droplets in a fiber-type specific manner.

Skeletal muscle lipid infiltration, known as myosteatosis, increases with obesity and ageing. Myosteatosis has also recently been discovered as a negative ...

fiber-type-subcellular-specific-analysis-lipid-droplet-content

Universidad de Cartagena UDEC

Research

JoVE Journal

Medicine

3.9K Views.  Université Catholique de Louvain. Increasing evidence indicates that excessive infiltration of lipids inside skeletal muscle results in lipotoxicity and diabetes. Here, we present a complete protocol, including tissue processing, staining with Bodipy, image acquisition, and analysis, to quantify the size, density, and subcellular distribution of lipid droplets in a fiber-type specific manner.

Video: Fiber Type and Subcellular-Specific Analysis of Lipid Droplet Content in Skeletal Muscle

Lipid Droplet Isolation for Quantitative Mass Spectrometry Analysis

Lipid droplets are vital to the replication of a variety of different pathogens, most prominently the Hepatitis C Virus (HCV), as the putative site of virion morphogenesis. Quantitative lipid droplet proteome analysis can be used to identify proteins that localize to or are displaced from lipid droplets under conditions such as virus infections. Here, we describe a protocol that has been successfully used to characterize the changes in the lipid droplet proteome following infection with HCV. We use Stable Isotope Labeling with Amino Acids in Cell Culture (SILAC) and thus label the complete proteome of one population of cells with &#34;heavy&#34; amino acids to quantitate the proteins by mass spectrometry. For lipid droplet isolation, the two cell populations (i.e.&#160;HCV-infected/&#34;light&#34; amino acids and uninfected control/&#34;heavy&#34; amino acids) are mixed 1:1 and lysed mechanically in hypotonic buffer. After removing the nuclei and cell debris by low speed centrifugation, lipid droplet-associated proteins are enriched by two subsequent ultracentrifugation steps followed by three washing steps in isotonic buffer. The purity of the lipid droplet fractions is analyzed by western blotting with antibodies recognizing different subcellular compartments. Lipid droplet-associated proteins are then separated by SDS-polyacrylamide gel electrophoresis (SDS-PAGE) followed by Coomassie staining. After tryptic digest, the peptides are quantified by liquid chromatography-electrospray ionization-tandem mass spectrometry (LC-ESI-MS/MS). Using this method, we identified proteins recruited to lipid droplets upon HCV infection that might represent pro- or antiviral host factors. Our method can be applied to a variety of different cells and culture conditions, such as infection with pathogens, environmental stress, or drug treatment.

Lipid droplets are important organelles for the replication of several pathogens, including the Hepatitis C Virus (HCV). We describe a method to isolate lipid droplets for quantitative mass spectrometry of associated proteins; it can be used under a variety of conditions, such as virus infection, environmental stress, or drug treatment.

Lipid droplets are vital to the replication of a variety of different pathogens, most prominently the Hepatitis C Virus (HCV), as the putative site of ...

Biochemistry

Isolation of Cellular Lipid Droplets: Two Purification Techniques Starting from Yeast Cells and Human Placentas

Lipid droplets are dynamic organelles that can be found in most eukaryotic and certain prokaryotic cells. Structurally, the droplets consist of a core of neutral lipids surrounded by a phospholipid monolayer. One of the most useful techniques in determining the cellular roles of droplets has been proteomic identification of bound proteins, which can be isolated along with the droplets. Here, two methods are described to isolate lipid droplets and their bound proteins from two wide-ranging eukaryotes: fission yeast and human placental villous cells. Although both techniques have differences, the main method - density gradient centrifugation -&#160;is shared by both preparations. This shows the wide applicability of the presented droplet isolation techniques.

In the first protocol, yeast cells are converted into spheroplasts by enzymatic digestion of their cell walls. The resulting spheroplasts are then gently lysed in a loose-fitting homogenizer. Ficoll is added to the lysate to provide a density gradient, and the mixture is centrifuged three times. After the first spin, the lipid droplets are localized to the white-colored floating layer of the centrifuge tubes along with the endoplasmic reticulum (ER), the plasma membrane, and vacuoles. Two subsequent spins are used to remove these other three organelles. The result is a layer that has only droplets and bound proteins.

In the second protocol, placental villous cells are isolated from human term placentas by enzymatic digestion with trypsin and DNase I. The cells are homogenized in a loose-fitting homogenizer. Low-speed and medium-speed centrifugation steps are used to remove unbroken cells, cellular debris, nuclei, and mitochondria. Sucrose is added to the homogenate to provide a density gradient and the mixture is centrifuged to separate the lipid droplets from the other cellular fractions.

The purity of the lipid droplets in both protocols is confirmed by Western Blot analysis. The droplet fractions from both preps are suitable for subsequent proteomic and lipidomic analysis.

Two techniques for isolating cellular lipid droplets from 1) yeast cells and 2) human placentas are presented. The centerpiece of both procedures is density gradient centrifugation, where the resulting floating layer containing the droplets can be readily visualized by eye, extracted, and quantified by Western Blot analysis for purity.

Lipid droplets are dynamic organelles that can be found in most eukaryotic and certain prokaryotic cells. Structurally, the droplets consist of a core of ...

Bioengineering

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.

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 ...

Analysis of Lipid Signaling in Drosophila Photoreceptors using Mass Spectrometry

The activation of phospholipase C&#946; (PLC&#946;) is an essential step during sensory transduction in Drosophila photoreceptors. PLC&#946; activity results in the hydrolysis of the membrane lipid phosphatidylinositol 4,5 bisphosphate [PI(4,5)P2] leading ultimately to the activation of transient receptor potential (TRP) and TRP like (TRPL) channels. The activity of PLC&#946; also leads subsequently to the generation of many lipid species several of which have been proposed to play a role in TRP and TRPL activation. In addition, several classes of lipids have been proposed to play key roles in organizing the cell biology of photoreceptors to optimize signaling reactions for optimal sensory transduction. Historically, these discoveries have been driven by the ability to isolate Drosophila mutants for enzymes that control the levels of specific lipids and perform analysis of photoreceptor physiology in these mutants. More recently, powerful mass spectrometry methods for isolation and quantitative analysis of lipids with high sensitivity and specificity have been developed. These are particularly suited for use in Drosophila where lipid analysis is now possible from photoreceptors without the need for radionuclide labeling. In this article, the conceptual and practical considerations in the use of lipid mass spectrometry for the robust, sensitive, and accurate quantitative assessment of various signaling lipids in Drosophila photoreceptors are covered. Along with existing methods in molecular genetics and physiological analysis such lipid is likely to enhance the power of photoreceptors as a model system for discoveries in biology.

Analysis of Lipid Signaling in Drosophila Photoreceptors using Mass Spectrometry

The manuscript presents versatile, robust, and sensitive mass spectrometry protocols to identify and quantify several classes of lipids from Drosophila photoreceptors.

The activation of phospholipase C&#946; (PLC&#946;) is an essential step during sensory transduction in Drosophila photoreceptors. PLC&#946; activity ...

Shotgun Lipidomics of Rodent Tissues

Lipids play a vital role as essential components of all prokaryotic and eukaryotic cells. Constant technological improvements in mass spectrometry have made lipidomics a powerful analytical tool for monitoring tissue lipidome compositions in homeostatic as well as disease states. This paper presents a step-by-step protocol for a shotgun lipid analysis method that supports the simultaneous detection and quantification of a few hundred molecular lipid species in different tissue and biofluid samples at high throughput. This method leverages automated nano-flow direct injection of a total lipid extract spiked with labeled internal standards without chromatographic separation into a high-resolution mass spectrometry instrument. Starting from sub-microgram amounts of rodent tissue, the MS analysis takes 10 min per sample and covers up to 400 lipids from 14 lipid classes in mouse lung tissue. The method presented here is well suited for studying disease mechanisms and identifying and quantifying biomarkers that indicate early toxicity or beneficial effects within rodent tissues.

Shotgun mass spectrometry-based lipidomics delivers a sensitive quantitative snapshot of a broad spectrum of lipid classes simultaneously in a single measurement from various rodent tissues.

Lipids play a vital role as essential components of all prokaryotic and eukaryotic cells. Constant technological improvements in mass spectrometry have ...

Assessing Hepatic Steatosis with 3D Fluorescence Lipid Droplet Reconstruction

Analysis of Fluorescent-Stained Lipid Droplets with 3D Reconstruction for Hepatic Steatosis Assessment

Lipid droplets (LDs) are specialized organelles that mediate lipid storage and play a very important role in suppressing lipotoxicity and preventing dysfunction caused by free fatty acids (FAs). The liver, given its critical role in the body's fat metabolism, is persistently threatened by the intracellular accumulation of LDs in the form of both microvesicular and macrovesicular hepatic steatosis. The histologic characterization of LDs is typically based on lipid-soluble diazo dyes, such as Oil Red O (ORO) staining, but a number of disadvantages consistently hamper the use of this analysis with liver specimens. More recently, lipophilic fluorophores 493/503 have become popular for visualizing and locating LDs due to their rapid uptake and accumulation into the neutral lipid droplet core. Even though most applications are well-described in cell cultures, there is less evidence demonstrating the reliable use of lipophilic fluorophore probes as an LD imaging tool in tissue samples. Herein, we propose an optimized boron dipyrromethene (BODIPY) 493/503-based protocol for the evaluation of LDs in liver specimens from an animal model of high-fat diet (HFD)-induced hepatic steatosis. This protocol covers liver sample preparation, tissue sectioning, BODIPY 493/503 staining, image acquisition, and data analysis. We demonstrate an increased number, intensity, area ratio, and diameter of hepatic LDs upon HFD feeding. Using orthogonal projections and 3D reconstructions, it was possible to observe the full content of neutral lipids in the LD core, which appeared as nearly spherical droplets. Moreover, with the fluorophore BODIPY 493/503, we were able to distinguish microvesicles (1 &#181;m &lt; d &#8804; 3 &#181;m), intermediate vesicles (3 &#181;m &lt; d &#8804; 9 &#181;m), and macrovesicles (d &gt; 9 &#181;m), allowing the successful discrimination of microvesicular and macrovesicular steatosis. Overall, this BODIPY 493/503 fluorescence-based protocol is a reliable and simple tool for hepatic LD characterization and may represent a complementary approach to the classical histological protocols.

Author Spotlight: Analysis of Fluorescent-Stained Lipid Droplets with 3D Reconstruction for Hepatic Steatosis Assessment  

Herein, we demonstrate an optimized BODIPY 493/503 fluorescence-based protocol for lipid droplet characterization in liver tissue. Through the use of orthogonal projections and 3D reconstructions, the fluorophore allows for successful discrimination between microvesicular and macrovesicular steatosis and may represent a complementary approach to the classical histological protocols for hepatic steatosis assessment.

Lipid droplets (LDs) are specialized organelles that mediate lipid storage and play a very important role in suppressing lipotoxicity and preventing ...

A Guide to Examining Intramuscular Fat Formation and its Cellular Origin in Skeletal Muscle

Fibro-adipogenic progenitors (FAPs) are mesenchymal stromal cells that play a crucial role during skeletal muscle homeostasis and regeneration. FAPs build and maintain the extracellular matrix that acts as a molecular myofiber scaffold. In addition, FAPs are indispensable for myofiber regeneration as they secrete a multitude of beneficial factors sensed by the muscle stem cells (MuSCs). In diseased states, however, FAPs are the cellular origin of intramuscular fat and fibrotic scar tissue. This fatty fibrosis is a hallmark of sarcopenia and neuromuscular diseases, such as Duchenne Muscular Dystrophy. One significant barrier in determining why and how FAPs differentiate into intramuscular fat is effective preservation and subsequent visualization of adipocytes, especially in frozen tissue sections. Conventional methods of skeletal muscle tissue processing, such as snap-freezing, do not properly preserve the morphology of individual adipocytes, thereby preventing accurate visualization and quantification. To overcome this hurdle, a rigorous protocol was developed that preserves adipocyte morphology in skeletal muscle sections allowing visualization, imaging, and quantification of intramuscular fat. The protocol also outlines how to process a portion of muscle tissue for RT-qPCR, enabling users to confirm observed changes in fat formation by viewing differences in the expression of adipogenic genes. Additionally, it can be adapted to visualize adipocytes by whole-mount immunofluorescence of muscle samples. Finally, this protocol outlines how to perform genetic lineage tracing of Pdgfr&#945;-expressing FAPs to study the adipogenic conversion of FAPs. This protocol consistently yields high-resolution and morphologically accurate immunofluorescent images of adipocytes, along with confirmation by RT-qPCR, allowing for robust, rigorous, and reproducible visualization and quantification of intramuscular fat. Together, the analysis pipeline described here is the first step to improving our understanding of how FAPs differentiate into intramuscular fat, and provides a framework to validate novel interventions to prevent fat formation.

The replacement of healthy muscle tissue with intramuscular fat is a prominent feature of human diseases and conditions. This protocol outlines how to visualize, image, and quantify intramuscular fat, allowing the rigorous study of the mechanisms underlying intramuscular fat formation.

Fibro-adipogenic progenitors (FAPs) are mesenchymal stromal cells that play a crucial role during skeletal muscle homeostasis and regeneration. FAPs build ...

Developmental Biology

Exploring the Regulation of Lipid Droplet Catabolism through Lipophagy

Macroautophagy, commonly referred to as autophagy, is a highly conserved cellular process responsible for the degradation of cellular components. This process is particularly prominent under conditions such as fasting, cellular stress, organelle damage, cellular damage, or aging of cellular components. During autophagy, a segment of the cytoplasm is enclosed within double-membrane vesicles known as autophagosomes, which then fuse with lysosomes. Following this fusion, the contents of autophagosomes undergo non-selective bulk degradation facilitated by lysosomes. However, autophagy also exhibits selective functionality, targeting specific organelles, including mitochondria, peroxisomes, lysosomes, nuclei, and lipid droplets (LDs). Lipid droplets are enclosed by a phospholipid monolayer that isolates neutral lipids from the cytoplasm, protecting cells from the harmful effects of excess sterols and free fatty acids (FFAs). Autophagy is implicated in various conditions, including neurodegenerative diseases, metabolic disorders, and cancer. Specifically, lipophagy -- the autophagy-dependent degradation of lipid droplets -- plays a crucial role in regulating intracellular FFA levels across different metabolic states. This regulation supports essential processes such as membrane synthesis, signaling molecule formation, and energy balance. Consequently, impaired lipophagy increases cellular vulnerability to death stimuli and contributes to the development of diseases such as cancer. Despite its significance, the precise mechanisms governing lipid droplet metabolism regulated by lipophagy in cancer cells remain poorly understood. This article aims to describe confocal imaging acquisition and quantitative imaging analysis protocols that enable the investigation of lipophagy associated with metabolic changes in cancer cells. The results obtained through these protocols may shed light on the intricate interplay between autophagy, lipid metabolism, and cancer progression. By elucidating these mechanisms, novel therapeutic targets may emerge for combating cancer and other metabolic-related diseases.

Lipophagy is a selective form of autophagy that involves the degradation of lipid droplets. Dysfunctions in this process are associated with cancer development. However, the precise mechanisms are not yet fully understood. This protocol describes quantitative imaging approaches to better understand the interplay between autophagy, lipid metabolism, and cancer progression.

Macroautophagy, commonly referred to as autophagy, is a highly conserved cellular process responsible for the degradation of cellular components. This ...

Cancer Research

Analysis of Lipid Droplet Content in Fission and Budding Yeasts using Automated Image Processing

Lipid metabolism and its regulation are of interest to both basic and applied life sciences and biotechnology. In this regard, various yeast species are used as models in lipid metabolic research or for industrial lipid production. Lipid droplets are highly dynamic storage bodies and their cellular content represents a convenient readout of the lipid metabolic state. Fluorescence microscopy is a method of choice for quantitative analysis of cellular lipid droplets, as it relies on widely available equipment and allows analysis of individual lipid droplets. Furthermore, microscopic image analysis can be automated, greatly increasing overall analysis throughput. Here, we describe an experimental and analytical workflow for automated detection and quantitative description of individual lipid droplets in three different model yeast species: the fission yeasts Schizosaccharomyces pombe and Schizosaccharomyces japonicus, and the budding yeast Saccharomyces cerevisiae. Lipid droplets are visualized with BODIPY 493/503, and cell-impermeable fluorescent dextran is added to the culture media to help identify cell boundaries. Cells are subjected to 3D epifluorescence microscopy in green and blue channels and the resulting z-stack images are processed automatically by a MATLAB pipeline. The procedure outputs rich quantitative data on cellular lipid droplet content and individual lipid droplet characteristics in a tabular format suitable for downstream analyses in major spreadsheet or statistical packages. We provide example analyses of lipid droplet content under various conditions that affect cellular lipid metabolism.

Here, we present a MATLAB implementation of automated detection and quantitative description of lipid droplets in fluorescence microscopy images of fission and budding yeast cells.

Lipid metabolism and its regulation are of interest to both basic and applied life sciences and biotechnology. In this regard, various yeast species are ...

Immunology and Infection

Quantification of Skeletal Muscle Fatty Infiltration via Decellularization

Decellularization-Based Quantification of Skeletal Muscle Fatty Infiltration

Fatty infiltration is the accumulation of adipocytes between myofibers in skeletal muscle and is a prominent feature of many myopathies, metabolic disorders, and dystrophies. Clinically in human populations, fatty infiltration is assessed using noninvasive methods, including computed tomography (CT), magnetic resonance imaging (MRI), and ultrasound (US). Although some studies have used CT or MRI to quantify fatty infiltration in mouse muscle, costs and insufficient spatial resolution remain challenging. Other small animal methods utilize histology to visualize individual adipocytes; however, this methodology suffers from sampling bias in heterogeneous pathology. This protocol describes the methodology to qualitatively view and quantitatively measure fatty infiltration comprehensively throughout intact mouse muscle and at the level of individual adipocytes using decellularization. The protocol is not limited to specific muscles or specific species and can be extended to human biopsy. Additionally, gross qualitative and quantitative assessments can be made with standard laboratory equipment for little cost, making this procedure more accessible across research laboratories.

Author Spotlight: Decellularization-Based Quantification of Skeletal Muscle Fatty Infiltration  

The present study describes decellularization-based methodologies for visualizing and quantifying intramuscular adipose tissue (IMAT) deposition through intact muscle volume, as well as quantifying metrics of individual adipocytes that comprise IMAT.

Fiber Type and Subcellular-Specific Analysis of Lipid Droplet Content in Skeletal Muscle

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A Guide to Examining Intramuscular Fat Formation and its Cellular Origin in Skeletal Muscle

Exploring the Regulation of Lipid Droplet Catabolism through Lipophagy

Analysis of Lipid Droplet Content in Fission and Budding Yeasts using Automated Image Processing

Decellularization-Based Quantification of Skeletal Muscle Fatty Infiltration