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Year : 2022  |  Volume : 65  |  Issue : 5  |  Page : 291-299
Electron microscopy in the diagnosis of skeletal muscle disorders: Its utility and limitations

Department of Neuropathology, National Institute of Mental Health and Neurosciences, Bengaluru, Karnataka, India

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Date of Submission16-Nov-2021
Date of Decision31-Jan-2022
Date of Acceptance06-Feb-2022
Date of Web Publication11-May-2022


Electron microscopy (EM) has a substantial role in the diagnosis of skeletal muscle disorders. The ultrastructural changes can be observed in muscle fibers and other components of the muscle tissue. EM serves as a confirmatory tool where the diagnosis is already established by enzyme histochemistry staining. Although it is indispensable in the diagnosis of rare forms of congenital myopathies not appreciated by light microscope, such as cylindrical spiral myopathy, zebra body myopathy, fingerprint body myopathy, and intranuclear rod myopathy, in cases not subjected to histochemical staining, it is required for definitive diagnosis in certain groups of muscle disorders, which includes congenital myopathies, metabolic myopathies in particular mitochondrial myopathies and glycogenosis, and in vacuolar myopathies. It does not have diagnostic implications in muscular dystrophies and neurogenic disorders. In the recent past, despite the availability of advanced diagnostic techniques, electron microscopy continues to play a vital role in the diagnosis of skeletal muscle disorders. This review gives an account of ultrastructural features of skeletal muscle disorders, the role of EM in the diagnosis, and its limitations.

Keywords: Electron microscopy, inclusions, muscle fibers, neuromuscular disorders, ultrastructure

How to cite this article:
Santhoshkumar R, Narayanappa G. Electron microscopy in the diagnosis of skeletal muscle disorders: Its utility and limitations. Indian J Pathol Microbiol 2022;65, Suppl S1:291-9

How to cite this URL:
Santhoshkumar R, Narayanappa G. Electron microscopy in the diagnosis of skeletal muscle disorders: Its utility and limitations. Indian J Pathol Microbiol [serial online] 2022 [cited 2022 May 24];65, Suppl S1:291-9. Available from: https://www.ijpmonline.org/text.asp?2022/65/5/291/345045

   Introduction Top

Electron microscopy (EM) plays a vital role in unraveling the ultrastructural features of normal and pathological alterations of various subcellular structures in cells/tissue in response to changes in the microenvironment. Muscle constitutes 40%–50% of the total body weight.[1] Skeletal muscle endures several variations in response to disease and trauma. In this contemporary era of molecular diagnostic testing, the use of minimally invasive genetic testing and noninvasive biomarkers approach is chosen over invasive skeletal muscle biopsy. Nevertheless, skeletal muscle biopsy remains an important investigative tool in the diagnosis of diverse groups of muscle disorders. EM of muscle offers insight into the pathophysiological mechanisms and in some instances can guide molecular genetic analysis.

Skeletal muscle biopsy subjected primarily to routine histological, enzyme histochemical stains, immunohistochemical markers, and biochemical investigations contributed not only to the understanding of the muscle physiology and pathology but also in identifying several new disease entities.

EM serves as a confirmatory tool where the diagnosis is already established by enzyme histochemistry staining. It is indicated when the diagnosis is not forthcoming by conventional methods and in some rare forms of congenital myopathies such as cylindrical spiral myopathy, zebra body myopathy, fingerprint body myopathy, and intranuclear rod myopathy not discernible by light microscope. In cases not subjected to histochemical staining, it aids in offering definitive diagnosis in certain groups of muscle disorders, which includes congenital myopathies, metabolic myopathies (in particular, mitochondrial myopathies and glycogenosis), and vacuolar myopathies. It does not have diagnostic implications in muscular dystrophies and neurogenic disorders.

EM is not a routine diagnostic procedure. It is following pathologists' observation on light microscopy that the biopsy is subjected to EM studies. Nevertheless, as a routine practice, a tiny bit of skeletal muscle fixed in 2.5% buffered glutaraldehyde soon after biopsy proves useful. This article details ultrastructural pathological changes in all components of the muscle fibers, the utility of EM in diagnosis, and its limitations.

   Tissue Processing Top

The biopsied tissue following overnight fixation in 2.5% buffered glutaraldehyde is postfixed in 1% buffered osmium tetroxide for 1 h, gradually dehydrated in ethanol, cleared in propylene oxide, embedded in epoxy resin (Araldite Cy212), and allowed to polymerize undisturbed in an oven maintained at 60°C for 48 h. The semithin sections (1-μm thickness) stained with 1% toluidine blue are viewed under a light microscope to select an area of interest for further ultrathin (60–70-nm thickness) sectioning. Ultrathin sections collected on copper/nickel grids are double-stained with saturated uranyl acetate in methanol and 0.2% aqueous lead citrate. Stained grids are scanned and representative areas are captured using a transmission electron microscope (TEM). The resultant images are termed electron micrographs.

   Interpretation of the tissue Top

EM helps in differentiating normal and abnormal fibers. It is used to evaluate structural components such as sarcolemma, the myofilaments and Z-band abnormalities, sarcoplasmic inclusions, alterations of the sarcotubular system, the nuclear changes, mitochondrial aberrations, glycogen aggregation, the surrounding connective tissue (endomysium and perimysium), the blood vessels, nerve twigs, and the neuromuscular junctions.[2]

  • Extracellular matrix and sarcolemma: Extracellular matrix (ECM) in skeletal muscle comprises epi, peri, and endomysium and plays a very important role in muscle development, adaptation, and its physiology. The bulk of ECM consists of collagen VI synthesized by fibroblasts.[3] Endomysium, a thin layer of connective tissue, surrounds each muscle fiber which continues as perimysium to group fibers into a fascicle, and groups of fascicles are collected into muscle bellies surrounded by epimysium. Several ultrastructural changes in endomysium have been defined. Gubbay et al.[4] described abnormal proliferation of collagen and its fusion to the sarcolemma in a case of congenital muscular dystrophy (CMD). Fidzianska et al.[5] described the presence of atypical fibroblasts similar to myofibroblasts with bundles of filaments in the cytoplasm and abnormal collagen fibrils in the extracellular space in CMD patients. In the severe form of Ullrich muscular dystrophy, failure of abnormal collagen VI to anchor the basal lamina was detected.[6]

Muscle cells are protected and connected to the extracellular matrix by sarcolemma consisting of an outer basement membrane/basal lamina (BL) and inner plasmalemma [Figure 1]a.[2],[7] While BL is vital for maintaining the integrity of the muscle fiber,[8] plasmalemma forms a protective covering of the muscle cells and has a dynamic role in muscle physiology. Numerous proteins, ion channels, and receptors are present in the sarcolemma. Sarcolemmal-specific proteins and dystrophin-associated complexes link to the extracellular matrix.[7] The pathological alterations of sarcolemma comprise absence of plasmalemma [Figure 1]b in the necrotic fibers, segmental loss of plasma membrane at the region of fusion of autophagic vacuoles in X-linked myopathy with excessive autophagy (XMEA) [Figure 1]c, discontinuous sarcolemma (muscular dystrophies) [Figure 1]d, papillary projections (muscular dystrophies, inflammatory myopathies) [Figure 1]e, separation of plasmalemma and basal lamina, and accumulation of glycogen between plasmalemma and basal lamina (glycogen storage disorders, GSD). Atrophic fibers display redundant/folded basal lamina due to the inability of the fiber to reach its normal size[7] [Figure 1]f. Numerous caveolae (flask-shaped infoldings) are seen on the inner surface of the plasmalemma which open into extracellular space in normal skeletal muscle and have an important role in signaling pathway.[9] Marked reduction in number and distribution of caveolae is noted in CAV3 mutations, a gene that encodes the caveolin-3 protein.[10] Thickening of the basal lamina occurs in a wide range of neuromuscular disorders. Indentations of the plasmalemma is seen at the site of satellite cells which are undifferentiated mononuclear stem cells with scanty cytoplasm and few organelles and shares common basal lamina with the parent fiber but has a distinct plasmalemma. It plays a crucial role in muscle repair and regeneration.
Figure 1: Ultrastructural features of skeletal muscle cell membrane- Normal and disease condition. (a) Normal myofiber with smooth outer basal lamina (arrow) and inner plasma membrane (arrowhead) ×6000; (b) Total absence of plasmalemma in a case of muscular dystrophy, ×4000; (c) Segmental loss of plasma membrane at the region of fusion of autophagic vacuoles (arrow) in XMEA, ×6000; (d) Focal discontinuous membrane (arrow) in MD, ×4000; (e) Papillary projections of the sarcolemma (arrow) and basement membrane merged with extracellular matrix (arrowhead) in MD, ×3000; (f) Redundant basal lamina (arrow) in an atrophic fiber, from a case of spinal muscular atrophy ×1500

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The sarcolemma show variation of the myofiber surface at specialized structures such as neuromuscular junctions (NMJs) and myotendinous junctions (MTJs). A NMJ is a specialized region of synapse between the motor neuron nerve terminal and its muscle fiber[11] where the plasmalemma is thrown into deep postsynaptic folds/clefts and the presynaptic nerve terminal that is covered by Schwann cell processes. Numerous synaptic vesicles and mitochondria are concentrated at the junction. The double layer of basal lamina extends into the cleft's anchors' junction-specific proteins (acetylcholine esterase, agrin, etc.). NMJs are responsible for converting the electrical signal received from the motor nerve terminal into electrical activity in the muscle fibers.[11] While alterations in the NMJ occur in a variety of disorders, NMJs are seldom seen in the biopsy unless specifically taken at the motor endplate. In myasthenia gravis, the folds of the junction are either reduced or absent and the debris accumulates as electron-dense material between the nerve and the muscle membrane. Another complex infoldings of the sarcolemma are MTJs, which are special structural regions connecting skeletal muscle to the tendon.[12] The plasmalemma at a MTJ appears as a thick layer of electron-dense material that merges with the Z line of the myofibrils.

  • Myonuclei: Muscle cells are multinucleated. Myonuclei are oval, heterochromatic [Figure 2]a and positioned beneath the sarcolemma and parallel to the myofibrils. One or two nucleoli are seen in each nucleus. Nuclear abnormalities in muscle diseases include abnormal positioning, that is, central nucleus (centronuclear myopathy, myotubular myopathy, and myotonic disorders) and internalized nuclei-single or multiple (various muscle diseases). The nuclear changes include irregularities in shape, pyknotic appearance, extrusion of nucleoplasm into the sarcoplasm, and changes in chromatin distribution which are seen in muscular dystrophy due to aberrant emerin[13] or lamin A/C proteins.[14] Peripheral chains of abnormal nuclei is reported in desmin deficiency.[15] The nuclei aggregate in atrophic fibers. Intranuclear rods, filamentous, and tubular inclusions have been observed in muscle diseases with different etiologies. Though rods are usually sarcoplasmic, Schröder et al.[16] described intranuclear rods with an ACTA1 mutation. Tubular filamentous inclusions [Figure 2]b, [Figure 2]c are a diagnostic feature of inclusion body myositis (IBM) and oculopharyngeal muscular dystrophy (OPMD).[17] In OPMD, tubular filamentous inclusions with a diameter of ~8.5 nm are restricted to nuclei, while in IBM, it ranges between ~15 and 21 nm and is seen both in the sarcoplasm and nucleus; 15–21-nm filaments are also seen in distal myopathies, myofibrillar myopathies, and in cases with MYH2 gene mutations.[18]
  • Myofibrils: Myofibrils are the major cytoskeletal component of muscle fibers. Each myofibril is composed of bundles of myofilaments that run parallel along the length of the myofiber and are aligned to form repeated structures called the sarcomeres, the functional unit of the muscle cell. Each sarcomere comprises an alternating arrangement of anisotropic (A band) and isotropic (I band) regions, giving a striated pattern. The A band is bisected by a dense M line with a pale H zone on either side, while the I band with a dense Z line. The sarcomere is the region between two adjacent Z lines [Figure 3]a. Diseased muscle undergoes alterations of varying degrees. Nevertheless, artefactual changes, the plane of section, and the extent of changes in normal muscle should be considered while observing the changes in unhealthy muscle.[7]
Figure 2: Ultrastructural features of normal and abnormal myonucleus. (a) Normal myonucleus (Nu), oval-shaped, with chromatin material within the nucleoplasm, ×11000; (b) Abnormal myonucleus filled with tubular filamentous inclusions from a case of IBM, ×3000 and (c) High magnification of tubular filamentous inclusions in longitudinal section (arrow) and transverse section (arrowhead), ×5000

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Figure 3: Ultrastructural alterations of myofibrils seen in various muscle diseases. (a) Normal-longitudinal section of myofibrils showing striated pattern consisting of A band, I band, Z band, H zone, and M line, ×4000; (b) Focal Z-band streaming (arrow), ×9300; (c) Nemaline rods (arrow) in NRM, ×2500; (d) Core area (asterisk), ×11000; (e) Multi mini-cores (asterisk), ×2900; (f) Electron-dense granular aggregates (asterisk), ×13000; (g) Selective disorganization of I and Z band leading to the formation of granulofilamentous aggregates (asterisk) in a case of myofibrillar myopathy, ×9300; (h)Subsarcolemmal accumulation of filamentous material (asterisk) in a case of myofibrillar myopathy, ×9300; (i) Nucleus (asterisk), perinuclear aggregates of granular and filamentous material forming reducing bodies (arrow) in a case of FHL1opathy, ×1500

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The most common abnormalities observed in all genetic and acquired skeletal muscle disorders are the loss and variable distortion of myofilament patterns. In hyalinized or necrotic fibers, the myofibrils appear as structureless dense mass retracted from the sarcolemma. Sarcolemmal tubes completely devoid of myofilaments with a few surviving mitochondria are seen in spinal muscular atrophies and muscular dystrophies. Focal loss with displaced triads and mitochondria and narrowing and splitting of myofibrils are noted in most diseases. In addition, selective loss of the I band and A band has been documented in some conditions. Varying degrees of disorganization, disorientation, and disruption of myofibrils occur.

The most common structural alteration associated with myofibrillar disorganization includes alterations in the Z band in the form of Z band streaming [Figure 3]b, Z band expansions, and aggregates of Z band material due to defects in the various sarcomeric and extra-sarcomeric proteins. Nemaline rods originate from the Z band and have similar Z band electron density [Figure 3]c and lattice structure and show positive labeling with Z band proteins such as α-actinin and α-actin. They are numerous in nemaline rod myopathy (NRM) caused due to mutations in various structural and/or regulatory thin filament proteins.[19] Cores either center or eccentric are regions demarcated from the rest of the myofiber with reduced oxidative enzyme activity on light microscopy.[20],[21] On EM, core regions have a disorganized sarcomeric structure with a paucity of mitochondria [Figure 3]d and involve several sarcomeres. The core disease is chiefly associated with RYR1 mutation. Disruption restricted to the focal area is seen in target fibers in neurogenic disorders and mini-core disease [Figure 3]e due to SEPN1 or RYR1 mutation.[22] The coexistence of multiple morphological abnormalities (central nucleus, nemaline rods, and core) has been reported.[23] Similarly, Malfatti et al.[24] described combined cap disease and nemaline myopathy with TPM3 mutation. “Caps” are subsarcolemmal regions with substantial loss of thick filaments and show the presence of thickened Z bands and small rods with emanating thin filaments. In contrast, “hyaline body,” though it appears similar to “caps,” is highlighted by ATPase stain and shows a finely demarcated region with accumulation of subsarcolemmal amorphous, granular, and vaguely filamentous structure with interspersed glycogen region highlighting the accumulation of degraded thick myosin filaments.[25],[26],[27]

In myofibrillar myopathies (MFM), the architectural changes of the myofibrils cause granular [Figure 3]f, granulofilamentous [Figure 3]g, or filamentous inclusions [Figure 3]h[28] due to alterations in several myofibrillar Z disc protein and/or cytoskeletal associated and/or chaperone protein.[29] FHL1-related myopathy with granular and granulofilamentous inclusions is reported.[30] Reducing bodies (RBs) [Figure 3]i are round to oval nonmembrane bound structures with closely packed electron dense fibrillar material interspersed with glycogen and seen in close proximity to the nucleus.[31],[32] These bodies are due to FHL1 mutation but are seen in other conditions such as glycogen storage disorder (GSD II).

Although light microscopy highlights morphological features, EM confirmation is highly reliable for classic forms of congenital myopathies and protein surplus myopathies where myofibrillar alterations are very prominent.

  • Mitochondria: In normal skeletal muscle fibers, mitochondria are round to oval, present in pairs, in parallel orientation, adjacent to I band as seen in the longitudinal section. The number, distribution, and size vary depending on the physiological state of the fiber. Type I fibers have higher numbers as compared to type II fibers. Subsarcolemmal aggregation seen in some diseases may be nonspecific and may be seen in normal muscle. Abnormalities include variation in size, shape, number, distribution, and alterations in cristae (fragmented, concentric lamellar pattern, swollen). In addition, linear inclusions in the matrix, paracrystalline inclusions between folds of inner mitochondrial membrane (type 1, parking lot type) and type 2 inclusions amid the outer and inner membrane space are seen. The presence of vacuoles, lipid, and/or glycogen is also noted. Some of the ultrastructural alterations are illustrated in [Figure 4].
Figure 4: Ultrastructural alterations of mitochondria from a case of mitochondrial myopathy. (a) Subsarcolemmal loss of myofilaments, aggregation of abnormal mitochondria with paracrystalline inclusions, ×13000; (b) High magnification of paracrystalline inclusions, rectangular consisting of parallel lines, ×50000; (c) Mitochondria with abnormal cristae pattern (arrow) and electron-dense particles in the mitochondrial matrix (asterisk), ×18500; (d) Mitochondria with concentric lamellar pattern (asterisk) and vacuoles (arrow), ×30000

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An increase in mitochondria and variation in size is seen in regions of MTJs, subsarcolemmal region of denervated fibers, lobulated fibers (wedge pattern), and in ragged red fibers (RRFs) appreciated on modified Gomori trichrome (MGT). It is to note that RRFs with prominent subsarcolemmal and sarcoplasmic aggregation of mitochondria are seen in normal orbicularis oculi and in skeletal muscles of age group above 40 years, in a range similar to those used in the diagnosis of mitochondrial disorders.[33] Ultrastructurally, RRFs show proliferation and collection of abnormal mitochondria and paracrystalline inclusions. Though the presence of aberrant mitochondria is a feature of primary mitochondrial diseases caused due to mitochondrial DNA (mtDNA) and/or nuclear DNA mutations,[34],[35] they can also be seen as secondary change in Duchenne muscular dystrophy (DMD),[36] Limb-girdle muscular dystrophy (LGMD), IBM,[37] OPMD,[38] MFMs,[39] and metabolic diseases.[40],[41]

  • Transverse tubule (T-tubule) and Sarcoplasmic reticulum (SR): T-tubule and SR form a network of membranous structures throughout the myofiber. T-tubules are invaginations of plasmalemma at regular intervals and inserted between myofibrils. The SR is a membrane-bound structure between and around the myofibril and forms terminal cisterns or lateral sacs. A T-tubule with two lateral sacs on either side constitutes the triad at the level of A/I junction across the length of the myofiber. The lateral sacs contain granular electron-dense material and are thus easily distinguished from the T-tubule. The triads are responsible for the mechanism of excitation-contraction coupling.[42] Abnormalities due to the disease process include dilation, proliferation, displacement, vesicle formations, and honeycomb structure. Excessive proliferation of SR and T-tubule results in tubular aggregates [Figure 5]a, which stain intensely by nicotinamide adenine dinucleotide-tetrazolium reductase (NADH-Tr), MGT, and myoadenylate deaminase. There are various types seen in the subsarcolemmal and/or intermyofibrillar region. They are derived from terminal cisterns[43]and appear as single-walled [Figure 5]b, double-walled with granular core [Figure 5]c, and may also have several microtubules within a dilated tubule.[7],[44] They are detected as a nonspecific feature in several neuromuscular conditions, including exercise-induced myalgia and cramps,[45] phosphoglycerate mutase deficiency (GSD X),[46] periodic paralysis,[47],[48] inflammatory myopathies,[48] alcohol and chronic drug-induced myopathy,[49] congenital myasthenic syndrome associated with STIM1 mutation[50] or ORAI1,[51],[52],[53],[54] congenital limb-girdle myasthenia,[55] congenital myopathies,[56] distal myopathy with mtDNA alterations,[57] and necrotizing autoimmune myopathy associated with 3-hydroxy-3-methyl-glutaryl-CoA reductase (HMGCR) antibody.[58] Dilation of T-tubule [Figure 5]d and SR are nonspecific features and require an appropriate distinction between fact and artifact. SR dilation is common in DMD muscle and is also seen in periodic paralysis.
Figure 5: Ultrastructural alterations of sarcoplasmic reticulum and T-system from a case of myopathy with tubular aggregates. (a)Subsarcolemmal tubular aggregates (asterisk) at low magnification, ×2500; (b) Single-walled tubular aggregates (asterisk) along with dilated SR containing electron-dense granular aggregates (arrow), ×12000; (c) Double-walled tubular aggregates forming honeycomb pattern (asterisk), ×15000; (d) Dilated tubules in the subsarcolemmal region (arrow), ×6000

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  • Storage material: Lipid and glycogen are normally present in the intermyofibrillar region and are the source of energy in muscle. Dysregulation in their metabolic pathway due to enzyme deficiency result in excess accumulation of lipid and glycogen. In lipid storage disorders (LSDs), lipid droplets appear as empty rounded spaces of uniform diameter between the myofibrils and in the subsarcolemmal region. Some vacuoles coalesce to form larger vacuoles [Figure 6]a. In glycogen storage disorders, glycogen aggregation is seen in the subsarcolemmal region [Figure 6]b, between the myofibrils, or within membrane vacuoles.
  • Blood vessels: Capillaries within the diseased muscle can show pathological changes. Thickening of the basal lamina (BL) is seen in diabetics and other neuromuscular conditions [Figure 6]c. Degenerative features of basal lamina such as the formation of loops, tubuloreticular inclusions [Figure 6]d in endothelial cells of capillaries and arterioles in different forms of myositis (dermatomyositis, IBM and viral myositis),[59],[60] and myelinic figures or myeloid bodies within membrane-bound structures called autophagic vacuoles (AV) [Figure 6]e or nonmembrane vacuoles are seen in IBM and distal myopathy.
  • Non-specific structures: Several structures are seen in muscle cells that do not have a specific role in the disease process but are found across various neuromuscular diseases, for example, concentric laminated bodies (CLBs) [Figure 6]f, filamentous bodies (FBs) [Figure 6]g, and cytoplasmic bodies (CBs) [Figure 6]h. CLBs are hollow cylindrical structures with glycogen particles in the central region. Some workers consider it to be myofibrillar origin while others are from mitochondria.[61],[62] FBs are a bundle of thin filaments beneath the sarcolemma. CBs have a granular electron-dense core and filaments emanating from the periphery. The presence of picornavirus-like crystals, which are aggregates of glycogen, has been described.[63],[64]
Figure 6: Ultrastructural changes in storage disorders, blood vessel and non-specific features. (a) Accumulation of lipid and mitochondria in the subsarcolemmal and perinuclear region (asterisk) from a case of LSD, ×1500; (b) Subsarcolemmal aggregation of glycogen in GSD (asterisk), ×3000; (c) Blood vessel with a thickened basement membrane from a case of myopathy with tubular aggregates, ×4000; (d) Tubuloreticular inclusions in endothelial cells from a case of dermatomyositis, ×4000; (e) Membrane-bound autophagic vacuole with electron-dense degraded material, from a case of MFM, ×6800; (f) Concentric laminated bodies, ×8000; (g) Filamentous body, ×4000 and (h) Cytoplasmic body, ×3000, from a case of CM, mitochondrial disease and LSD, respectively; (i) Membrane-bound granular osmiophilic deposits (GRODS, arrow) in the subsarcolemmal region from a case of NCL, infantile form, ×2000

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Role of skeletal muscle biopsy in neurometabolic and other disorders

Muscle biopsy is useful in the diagnosis of various neurometabolic disorders as storage material results in the formation of the inclusions that are seen not only in muscle fibers but also in satellite cells, blood vessels, endomysial fibroblasts, endomysial and perimysial nerve twigs, and motor endplates. Some of these disorders include neuronal ceroid lipofuscinosis (NCL) by the presence of granular osmiophilic deposits; GRODS [Figure 6]i; curvilinear, fingerprint profiles, and rectilinear inclusions within muscle fibers; endothelial and smooth muscle cells of vessels; and Schwann cell cytoplasm of myelinated and/or unmyelinated fibers. Mucopolysaccharidosis and oligosaccharidosis are characterized by the presence of membranous cytoplasmic bodies (MCBs) and lamellar inclusions.[65] In lipidosis, vacuoles, lamellar, trilaminar, MCB, and Zebra bodies (ZBs) are noted, whereas in Fabry disease, MCBs, lamellar inclusions, and ZBs are present in endothelial and smooth muscle cells of vessels; granular osmiophilic material (GOM) inclusions in vascular smooth muscles are noted in CADASIL/CARASIL. Endomysial nerve twigs and motor endplates show dilated axons with vesiculotubular profiles in infantile neuroaxonal dystrophy (INAD). In peroxisomal disorders (adrenoleukodystrophy (ADL)/adrenoleukomyeloneuropathy (ALMN)), the spicular clefts with glycogen and mitochondria are seen in endomysial nerve twigs.[65] As some features/inclusions such as MCBs and ZBs are common to most neurometabolic disorders such as mucopolysaccharidosis, oligosaccridosis, and Fabry disease, the diagnosis should be further ascertained by biochemical and molecular testing.

   Application in research Top

The potentialities of EM as a research tool for biochemical and physiological investigations by the use of light microscopic staining techniques such as enzyme histochemistry has enabled locating the sites of chemical constituents or enzymes within the cell and various organelles. Immunoelectron microscopy has provided specific information about the localization of dystrophin protein to the plasmalemma and the functional interrelationships of various myofibrillar proteins.

Freeze fracture, a unique electron microscopic technique, has aided in providing planar views of the internal organization of membranes and has helped in understanding the functional morphology of neuromuscular junctions.

Recently, cryo-electron tomography (cryo-ET), which enables the three-dimensional (3D) structural characterization of macromolecular complexes with nanometer-scale resolution in their native environment, has been extensively used both for research and diagnosis.

Further, to explore muscle biology and to understand the pathomechanism underlying the muscle disease, in vivo and in vitro model systems are extensively used. The most widely used are the mammalian (mice) and nonmammalian model organisms (Drosophila melanogaster, Caenorhabditis elegans, Danio rerio).[66]

   Limitations Top

Diagnosis of muscle biopsy by electron microscope has certain limitations. Most important factors include a small portion of tissue for investigation, and every phase of the processing of skeletal muscle from fixation to embedding and sectioning is critical as any deviation during these procedures induces artifacts resulting in distortion of the muscle sample. It is time-consuming and needs expertise for interpreting the tissue. The biopsy may appear normal if not chosen from the affected site. The ultrastructural abnormalities may not be disease-specific and hence must be correlated with clinical and light microscopic features.

   Conclusion Top

The role of electron microscope in the diagnosis of muscle disease though limited due to availability of high-throughput molecular techniques, it has enabled to define several new nosologically distinct entities such as central core disease, nemaline myopathy, and cylindrical spirals and to establish an accurate diagnosis and in classifying these conditions. It is helpful in the diagnosis of metabolic disorders, particularly, mitochondrial disorders, and in the elucidation of pathogenetic mechanisms. It gives insight into pathophysiologic mechanisms and can guide molecular genetic testing. Recently, advances in EM techniques such as the use of TEM and SEM (scanning EM) as complementary approaches with 3D reconstruction has identified structures not previously known by conventional EM, while the advent of cryo-EM has enabled 3D characterization of macromolecules in their native state and thus offers insights into conformational landscape and biological functions leading to new discoveries.


The authors acknowledge the support extended by all staff and faculty, Electron Microscopy Lab and Department of Neuropathology, NIMHANS, Bengaluru.

Declaration of patient consent

We certify that we have obtained all appropriate patient consent forms. The form is duly signed by the patient(s) with the content that he/she/they have given his/her/their consent for the use of his/her/their clinical and investigative information, images, and other findings to be reported in the journal. The patients have been explained and they understand that they are anonymized to conceal their identity.

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Conflicts of interest

There are no conflicts of interest.

   References Top

Tortora GJ, Derrickson BH. Principles of Anatomy & Physiology. 3rd ed. New Jersey: John Wiley & Sons, Inc; 2011.  Back to cited text no. 1
Dubowitz V, Sewry CA, Oldfors A. Muscle Biopsy: A Practical Approach. Saunders Ltd.; 2013.  Back to cited text no. 2
Purslow PP. The structure and role of intramuscular connective tissue in muscle function. Front Physiol 2020;11:495.  Back to cited text no. 3
Gubbay SS, Walton JN, Pearce GW. Clinical and pathological study of a case of congenital muscular dystrophy. J Neurol Neurosurg Psychiatry 1966;29:500-8.  Back to cited text no. 4
Fidzianska A, Goebel HH, Lenard HG, Heckmann C. Congenital muscular dystrophy (CMD)-A collagen formative disease? J Neurol Sci 1982;55:79-90.  Back to cited text no. 5
Ishikawa H, Sugie K, Murayama K, Awaya A, Suzuki Y, Noguchi S, et al. Ullrich disease due to deficiency of collagen VI in the sarcolemma. Neurology 2004;62:620-3.  Back to cited text no. 6
Sewry CA. Electron microscopy of human skeletal muscle: Role in diagnosis. Curr Diagn Pathol 2002;8:225-31.  Back to cited text no. 7
Sanes JR. The basement membrane/basal lamina of skeletal muscle. J Biol Chem 2003;278:12601-4.  Back to cited text no. 8
Gazzerro E, Sotgia F, Bruno C, Lisanti MP, Minetti C. Caveolinopathies: From the biology of caveolin-3 to human diseases. Eur J Hum Genet 2010;18:137-45.  Back to cited text no. 9
Minetti C, Bado M, Broda P, Sotgia F, Bruno C, Galbiati F, et al. Impairment of caveolae formation and T-system disorganization in human muscular dystrophy with caveolin-3 deficiency. Am J Pathol 2002;160:265-70.  Back to cited text no. 10
Rodríguez Cruz PM, Cossins J, Beeson D, Vincent A. The neuromuscular junction in health and disease: Molecular mechanisms governing synaptic formation and homeostasis. Front Mol Neurosci 2020;13:610964.  Back to cited text no. 11
Charvet B, Ruggiero F, Le Guellec D. The development of the myotendinous junction. A review. Muscles Ligaments Tendons J 2012;2:53-63.  Back to cited text no. 12
Gayathri N, Taly AB, Sinha S, Suresh TG, Gorai D. Emery dreifuss muscular dystrophy: A clinico-pathological study. Neurol India 2006;54:197-9.  Back to cited text no. 13
[PUBMED]  [Full text]  
Gotzmann J, Foisner RP. Lamins and Emerin in Muscular Dystrophy: The Nuclear Envelope Connection. In: Madame Curie Bioscience Database [Internet]. Austin (TX): Landes Bioscience; 2000-2013. Available from: https://www.ncbi.nlm.nih.gov/books/NBK6513/?report=classic.  Back to cited text no. 14
Santhoshkumar R, Preethish-Kumar V, Polavarapu K, Reghunathan D, Chaudhari S, Satyamoorthy K, et al. A novel L1 linker mutation in DES resulted in total absence of protein. J Mol Neurosci 2021;71:2468-73.  Back to cited text no. 15
Schröder JM, Durling H, Laing N. Actin myopathy with nemaline bodies, intranuclear rods, and a heterozygous mutation in ACTA1 (Asp154Asn). Acta Neuropathol 2004;108:250-6.  Back to cited text no. 16
Coquet M, Vital C, Julien J. Presence of inclusion body myositis-like filaments in oculopharyngeal muscular dystrophy. Ultrastructural study of 10 cases. Neuropathol Appl Neurobiol 1990;16:393-400.  Back to cited text no. 17
Tajsharghi H, Oldfors A. Myosinopathies: Pathology and mechanisms. Acta Neuropathol 2013;125:3-18.  Back to cited text no. 18
Laitila J, Wallgren-Pettersson C. Recent advances in nemaline myopathy. Neuromuscul Disord 2021;31:955-67.  Back to cited text no. 19
Goebel HH. Congenital myopathies. Semin Pediatr Neurol 1996;3:152-61.  Back to cited text no. 20
Hudgson P. The Diagnostic Value of Electron Microscopy: The Value of Electron Microscopy in Muscle Biopsies. Proceedings of the Royal Society of Medicine, UK, 1970;63:470-4.  Back to cited text no. 21
Nucci A, Queiroz LS, Zambelli HJ, Martins Filho J. Multi-minicore disease revisited. Arq Neuropsiquiatr 2004;62:935-9.  Back to cited text no. 22
Dhinakaran S, Kumar RS, Thakkar R, Narayanappa G. Coexistence of central nucleus, cores, and rods: Diagnostic relevance. Ann Indian Acad Neurol 2016;19:201-4.  Back to cited text no. 23
[PUBMED]  [Full text]  
Malfatti E, Schaeffer U, Chapon F, Yang Y, Eymard B, Xu R, et al. Combined cap disease and nemaline myopathy in the same patient caused by an autosomal dominant mutation in the TPM3 gene. Neuromuscul Disord 2013;23:992-7.  Back to cited text no. 24
Cancilla PA, Kalyanaraman K, Verity MA, Munsat T, Pearson CM. Familial myopathy with probable lysis of myofibrils in type I fibers. Neurology 1971;21:579-85.  Back to cited text no. 25
Shingde MV, Spring PJ, Maxwell A, Wills EJ, Harper CG, Dye DE, et al. Myosin storage (hyaline body) myopathy: A case report. Neuromuscul Disord 2006;16:882-6.  Back to cited text no. 26
Goebel HH, Laing NG. Actinopathies and myosinopathies. Brain Pathol 2009;19:516-22.  Back to cited text no. 27
Claeys KG, Fardeau M, Schröder R, Suominen T, Tolksdorf K, Behin A, et al. Electron microscopy in myofibrillar myopathies reveals clues to the mutated gene. Neuromuscul Disord 2008;18:656-66.  Back to cited text no. 28
Ferrer I, Olivé M. Molecular pathology of myofibrillar myopathies. Expert Rev Mol Med 2008;10:e25.  Back to cited text no. 29
Santhoshkumar R, Preethish-Kumar V, Mangalaparthi KK, Unni S, Padmanabhan B, T SK, et al. A dominant C150Y mutation in FHL1 induces structural alterations in LIM2 domain causing protein aggregation in human and Drosophila indirect flight muscles. J Mol Neurosci 2021;71:2324-35.  Back to cited text no. 30
Brooke MH, Neville HE. Reducing body myopathy. Neurology 1972;22:829-40.  Back to cited text no. 31
Goebel HH, Halbig LE, Goldfarb L, Schober R, Albani M, Neuen-Jacob E, et al. Reducing body myopathy with cytoplasmic bodies and rigid spine syndrome: A mixed congenital myopathy. Neuropediatrics 2001;32:196-205.  Back to cited text no. 32
McKelvie P, Satchi K, McNab AA, Kennedy P. Orbicularis oculi: Morphological changes mimicking mitochondrial cytopathy in a series of control normal muscles. Clin Exp Ophthalmol 2012;40:497-502.  Back to cited text no. 33
Challa S, Kanikannan MA, Murthy JM, Bhoompally VR, Surath M. Diagnosis of mitochondrial diseases: Clinical and histological study of sixty patients with ragged red fibers. Neurol India 2004;52:353-8.  Back to cited text no. 34
[PUBMED]  [Full text]  
Gayathri N, Deepha S, Sharma S. Diagnosis of primary mitochondrial disorders -Emphasis on myopathological aspects. Mitochondrion 2021;61:69-84.  Back to cited text no. 35
Moore TM, Lin AJ, Strumwasser AR, Cory K, Whitney K, Ho T, et al. Mitochondrial dysfunction is an early consequence of partial or complete dystrophin loss in mdx mice. Front Physiol 2020;11:690.  Back to cited text no. 36
Oldfors A, Moslemi AR, Jonasson L, Ohlsson M, Kollberg G, Lindberg C. Mitochondrial abnormalities in inclusion-body myositis. Neurology 2006;66 (2 Suppl 1):S49-55.  Back to cited text no. 37
Wong KT, Dick D, Anderson JR. Mitochondrial abnormalities in oculopharyngeal muscular dystrophy. Neuromuscul Disord 1996;6:163-6.  Back to cited text no. 38
Jackson S, Schaefer J, Meinhardt M, Reichmann H. Mitochondrial abnormalities in the myofibrillar myopathies. Eur J Neurol 2015;22:1429-35.  Back to cited text no. 39
Debashree B, Kumar M, Keshava Prasad TS, Natarajan A, Christopher R, Nalini A, et al. Mitochondrial dysfunction in human skeletal muscle biopsies of lipid storage disorder. J Neurochem 2018;145:323-41.  Back to cited text no. 40
Verity MA. Infantile Pompe's disease, lipid storage, and partial carnitine deficiency. Muscle Nerve 1991;14:435-40.  Back to cited text no. 41
Al-Qusairi L, Laporte J. T-tubule biogenesis and triad formation in skeletal muscle and implication in human diseases. Skeletal Muscle 2011;1:26.  Back to cited text no. 42
Salviati G, Pierobon-Bormioli S, Betto R, Damiani E, Angelini C, Ringel SP, et al. Tubular aggregates: Sarcoplasmic reticulum origin, calcium storage ability, and functional implications. Muscle Nerve 1985;8:299-306.  Back to cited text no. 43
Engel WK, Bishop DW, Cunningham GG. Tubular aggregates in type II muscle fibers: Ultrastructural and histochemical correlation. J Ultrastruct Res 1970;31:507-25.  Back to cited text no. 44
Brumback RA, Staton RD, Susag ME. Exercise-induced pain, stiffness, and tubular aggregation in skeletal muscle. J Neurol Neurosurg Psychiatry 1981;44:250-4.  Back to cited text no. 45
Salameh J, Goyal N, Choudry R, Camelo-Piragua S, Chong PS. Phosphoglycerate mutase deficiency with tubular aggregates in a patient from Panama. Muscle Nerve 2013;47:138-40.  Back to cited text no. 46
Yoshimura K, Morihata H, Takeda K, Sakoda S, Yuan JH, Nakano S. [A case of hyperkalemic periodic paralysis presenting progressive myopathy with tubular aggregates]. Rinsho Shinkeigaku 2018;58:663-7.  Back to cited text no. 47
Rosenberg NL, Neville HE, Ringel SP. Tubular aggregates: Their association with neuromuscular diseases, including the syndrome of myalgias/cramps. Arch Neurol 1985;42:973-6.  Back to cited text no. 48
Chui LA, Neustein H, Munsat TL. Tubular aggregates in subclinical alcoholic myopathy. Neurology 1975;25:405-12.  Back to cited text no. 49
Engel AG, Shen X-M, Selcen D, Sine SM. Congenital myasthenic syndromes: Pathogenesis, diagnosis, and treatment. Lancet Neurol 2015;14:420-34.  Back to cited text no. 50
Shahrizaila N, Lowe J, Wills A. Familial myopathy with tubular aggregates associated with abnormal pupils. Neurology 2004;63:1111-3.  Back to cited text no. 51
Nesin V, Wiley G, Kousi M, Ong EC, Lehmann T, Nicholl DJ, et al. Activating mutations in STIM1 and ORAI1 cause overlapping syndromes of tubular myopathy and congenital miosis. Proc Natl Acad Sci U S A 2014;111:4197-202.  Back to cited text no. 52
Garibaldi M, Fattori F, Riva B, Labasse C, Brochier G, Ottaviani P, et al. A novel gain-of-function mutation in ORAI1 causes late-onset tubular aggregate myopathy and congenital miosis. Clin Genet 2017;91:780-6.  Back to cited text no. 53
Endo Y, Noguchi S, Hara Y, Hayashi YK, Motomura K, Miyatake S, et al. Dominant mutations in ORAI1 cause tubular aggregate myopathy with hypocalcemia via constitutive activation of store-operated Ca2 314; channels. Hum Mol Genet 2015;24:637-48.  Back to cited text no. 54
Furui E, Fukushima K, Sakashita T, Sakato S, Matsubara S, Takamori M. Familial limb-girdle myasthenia with tubular aggregates. Muscle Nerve 1997;20:599-603.  Back to cited text no. 55
Narayanappa G, Nalini A, Thaha F. Congenital myopathy with tubular aggregates: Report on two siblings from India. J Child Neurol 2009;24:874-8.  Back to cited text no. 56
Garrard P, Blake J, Stinton V, Hanna MG, Reilly MM, Holton JL, et al. Distal myopathy with tubular aggregates: A new phenotype associated with multiple deletions in mitochondrial DNA? J Neurol Neurosurg Psychiatry 2002;73:207-8.  Back to cited text no. 57
Madigan NN, Liewluck T, Milone M, Naddaf E. Necrotizing autoimmune myopathy with tubular aggregates. Neurology 2019;93:313-4.  Back to cited text no. 58
Bronner IM, Hoogendijk JE, Veldman H, Ramkema M, van den Bergh Weerman MA, Rozemuller AJM, et al. Tubuloreticular structures in different types of myositis: Implications for pathogenesis. Ultrastruct Pathol 2008;32:123-6.  Back to cited text no. 59
Katzberg HD, Munoz DG. Tubuloreticular inclusions in inclusion body myositis. Clin Neuropathol 2010;29:262-6.  Back to cited text no. 60
Gambarelli D, Hassoun J, Pellissier JF, Berard M, Toga M. Concentric laminated bodies in muscle pathology. Pathol Eur 1974;9:289-96.  Back to cited text no. 61
Payne CM, Curless RG. Concentric laminated bodies. Ultrastructural demonstration of muscle fiber type specificity. J Neurol Sci 1976;29:311-22.  Back to cited text no. 62
Fukuyama Y, Ando T, Yokota J. Acute fulminant myoglobinuric polymyositis with picornavirus-like crystals. J Neurol Neurosurg Psychiatry 1977;40:775-81.  Back to cited text no. 63
Ben-Bassat M, Machtey I. Picornavirus-like structures in acute dermatomyositis. Am J Clin Pathol 1972;58:245-9.  Back to cited text no. 64
Ceuterick-de Groote C, Martin JJ. Extracerebral biopsy in lysosomal and peroxisomal disorders. Ultrastructural findings. Brain Pathol 1998;8:121-32.  Back to cited text no. 65
Sparrow J, Hughes SM, Segalat L. Other model organisms for sarcomeric muscle diseases. Adv Exp Med Biol 2008;642:192-206.  Back to cited text no. 66

Correspondence Address:
Gayathri Narayanappa
Retired Professor, Department of Neuropathology, National Institute of Mental Health and Neurosciences, Bengaluru, Karnataka- 560 029
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Source of Support: None, Conflict of Interest: None

DOI: 10.4103/ijpm.ijpm_1113_21

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