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Immunolocalization of dystrophin and neurofilament protein in muscle spindles of normal, mdx-dystrophic,… Nahirney, Patrick Charles 1993

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IMMUNOLOCALIZATION OF DYSTROPHIN AND NEUROFILAMENT PROTEIN INMUSCLE SPINDLES OF NORMAL, MDX-DYSTROPHIC, AND DENERVATED MICEByPATRICK CHARLES NAHIRNEYB.Sc. (Biology), Washington State University, 1990A THESIS SUBMITTED IN PARTIAL FULFILLMENT OFTHE REQUIREMENTS FOR THE DEGREE OFMASTER OF SCIENCEinTHE FACULTY OF GRADUATE STUDIES(Department of Anatomy)We accept this thesis as conformingto the required standardTHE UNIVERSITY OF BRITISH COLUMBIAApril 1993©PATRICK CHARLES NAHIRNEY, 1993In presenting this thesis in partial fulfilment of the requirements for an advanceddegree at the University of British Columbia, I agree that the Library shall make itfreely available for reference and study. I further agree that permission for extensivecopying of this thesis for scholarly purposes may be granted by the head of mydepartment or by his or her representatives. It is understood that copying orpublication of this thesis for financial gain shall not be allowed without my writtenpermission.(Signature)Department of ^AV ,4 7-0t-1The University of British ColumbiaVancouver, CanadaDate^tw/i__ ci 3DE-6 (2/88)THESIS ABSTRACTDystrophin is a high molecular weight protein localized under the sarcolemma of normalextrafusal muscle fibers but absent in skeletal muscles of Duchenne muscular dystrophypatients and in the mdx mouse. Muscle spindles in the soleus of 32-week-old normal and age-matched mdx mice were examined by immunocytochemical methods to determine thelocalization of dystrophin in polar and equatorial regions of the intrafusal fibers. Spindles wereserially-sectioned in transverse and longitudinal planes, and they were double-labelled with anantibody to dystrophin and with a 200 kD neurofilament protein antibody which revealed theirsensory innervation. By fluorescence microscopy, intrafusal fibers in the soleus of mdx micewere deficient in dystrophin throughout their lengths, whereas their sensory nerve-terminalsstained intensely with the nerve-specific antibody and appeared unaltered in dystrophy. Inthe normal soleus, polar regions of bag and chain fibers exhibited a peripheral rim ofsarcolemmal staining equivalent to that seen in the neighboring extrafusal fibers. Dystrophinlabelling in equatorial regions of normal intrafusal fibers, however, showed dystrophin-deficientsegments alternating in a spiral fashion with positive-staining domains along the sarcolemma.Double-labelling for dystrophin and neurofilament protein showed that these dystrophin-deficient sites were subjacent to the annulospiral sensory-nerve wrappings terminating on theintrafusal fibers. Additionally, it was found that chronic denervation of muscle spindles innormal mice did not affect the expression of dystrophin either at these sites or at the non-sensory regions of the sarcolemma. The results of this study suggest that dystrophin is notan integral part of the subsynaptic sensory-membrane in equatorial regions of normalintrafusal fibers, and, that the neurotrophic effect of sensory innervation is not the principlecause of this unique arrangement of dystrophin in equatorial regions. In dystrophy, intrafusalfibers display the same primary defect in muscular dystrophy as seen in the extrafusal fibers.However, because of their small-diameters, capsular investment, and relatively low tensionoutputs, dystrophic intrafusal fibers may be less prone to the sarcolemmal membranedisruption that is characteristic of extrafusal fibers in this disorder.TABLE OF CONTENTSTHESIS ABSTRACT ^  i iLIST OF FIGURES  ivACKNOWLEDGMENTS ^  vINTRODUCTIONA. Muscular dystrophy ^  1Discovery of the Duchenne Muscular Dystrophy Gene ^ 2Proposed Structure and Function of Dystrophin  3Synthesis of an Antibody to Dystrophin ^  4B. The Mammalian Muscle Spindle ^  6Capsule ^  7Intrafusal Fibers ^  8Innervation  11Pathology ^  15C. Immunofluorescence Microscopy ^  17D. Thesis Objectives ^  19METHODSHistochemistry ^  21Immunofluorescence Microscopy ^  22Denervation ^  23OBSERVATIONSGeneral Morphology ^  25Histochemistry  26Immunofluorescence Microscopy ^  27Denervation ^  30FIGURES ^  32DISCUSSIONHistochemistry ^  60Immunolocalization of Dystrophin in Muscle Spindles ^ 62Intrafusal Fibers in Dystrophic Mice ^  65Denervation ^  67CONCLUSIONS AND FUTURE DIRECTIONS ^  68LITERATURE CITED ^  70UST OF FIGURESFIGURE 1^Light Micrograph of the Normal Mouse Soleus Stained with H & E ^ 33FIGURE 2^Light Micrograph of the Mdx-Dystrophic Soleus Stained with H & E ^ 33FIGURE 3^Serial Sections of a Muscle Spindle in the Normal Soleus ^ 35FIGURE 4^Serial Sections of a Muscle Spindle in the Mdx-Dystrophic Soleus ^ 35FIGURE 5^Myosin ATPase Staining of the Normal Mouse Soleus ^ 37FIGURE 6^Myosin ATPase Staining of the Mdx-Dystrophic Soleus  37FIGURE 7^Myosin ATPase Staining of Intrafusal Fibers in the Normal Soleus ^ 39FIGURE 8^Myosin ATPase Staining of Intrafusal Fibers in the Mdx-DystrophicSoleus ^  39FIGURE 9^Immunofluorescence Micrographs of Anti-Dystrophin and Anti-Neurofilament Protein Staining in the Normal Soleus ^ 41FIGURE 10^Immunofluorescence Micrographs of Anti-Dystrophin and Anti-Neurofilament Protein Staining in the Mdx-Dystrophic Soleus ^ 43FIGURE 11^Immunofluorescence Micrographs of Polar and Equatorial Regionsof a Normal Muscle Spindle ^  45FIGURE 12^Light and Immunofluorescence Micrographs of an Mdx-DystrophicMuscle Spindle ^  45FIGURE 13^Serial Sections and Three-Dimensional Reconstruction of a NormalMuscle Spindle Stained for Dystrophin and Myosin ATPase ^ 47FIGURE 14^Immunofluorescence and Light Micrographs of a Longitudinal Sectionof a Normal Muscle Spindle ^  49FIGURE 15^Immunofluorescence and Phase-Contrast Micrographs of a NuclearBag Fiber ^  51FIGURE 16^Immunofluorescence and Phase-Contrast Micrographs of a NuclearChain and Nuclear Bag Fiber ^  53FIGURE 17^Transverse sections of 21-day and 42-day Denervated Normal SoleusMuscles ^  55FIGURE 18^Denervated Normal Muscle Spindle at 21 Days ^  57FIGURE 19^Denervated Normal Muscle Spindle at 42 Days  59ivACKNOWLEDGMENTSI am deeply grateful to my advisor Dr. William Ovalle who provided me withencouragement and guidance throughout my program. It was an enjoyable and an invaluableeducational experience to be working with him.I also extend my sincere thanks to Dr Pierre Dow who not only opened my mind toresearch but also gave me support and encouragement along the way. My regards also to theother members of my committee, Dr. Bernard Bressler and Dr. Wayne Vogl for their interestand advice in my education.My sincere gratitude is extended to the Department of Anatomy for creating acomfortable environment in which to seek an overall understanding of science and research. Aspecial thanks goes to Roseanne Mclndoe for her assistance in the laboratory and inmaintaining our animal colony.Finally, I would like to express my thanks to my family for their patience and support. Iam greatly indebted to them for all their efforts and encouragements throughout the pastfew years.This work was supported by grants to Dr. W.K. Ovalle from the Muscular DystrophyAssociation of Canada and by a University Graduate Fellowship and British Columbia MedicalResearch Services Scholarship to P.C. Nahirney.INTRODUCTIONA. Muscular DystrophyMuscular dystrophy is a term used to describe the heterogeneous group ofgenetically-linked disorders that cause progressive weakness and wasting of the skeletalmuscles. The most common and devastating of these disorders, Duchenne musculardystrophy, was first described in the mid-1800's by Meryon (1852) and later by Duchenne(1868). The incidence of this disease in North America has been estimated to be one in 3500boys, and in approximately one-third of cases the disease is caused by a mutation in a genelocated on the X chromosome. In the remaining two-thirds of cases the defective gene isinherited on the X chromosome from a carrier mother (Worton, 1992).This genetic disorder exhibits no obvious clinical manifestation until the age of three tofive years, when proximal muscle weakness is first observed. The ensuing progressive loss ofmuscle strength usually leaves affected individuals wheelchair-bound by the age of 11, andresults in early death due to respiratory failure. To date there is no cure and no effectivetreatment although prednisone, a catabolic steroid, has, paradoxically, been shown tostabilize muscle strength for a period of up to three years (Brooke, 1989).The Becker type of muscular dystrophy (Becker and Kiener, 1955) is a milder form ofthe disease with an incidence rate of about one-tenth of that of the severe Duchenne form(Becker, 1964). It is characterized by a much later onset with loss of ambulation after theage of 16 years and a relatively normal lifespan (Becker, 1965). Although thought for yearsto be a distinct disease, it is now recognized that it is caused by mutations in the same gene,and therefore a milder version of the same disease (Baumbach et al., 1989; Koenig et al.,1989). Boys who lose ambulation between the ages of 12 and 16 years are said to have anintermediate phenotype.1Discovery of the Duchenne Muscular Dystrophy GeneThe basis of this disorder stemmed from molecular genetic studies using a strategybased on identification of sequences that mapped into a region of the X chromosome. Thisregion had previously been shown by Kunkel and colleagues (1985) to be deleted in a boy withseveral X-linked disorders, including Duchenne muscular dystrophy. The identification of aclone from this part of the X chromosome that detected deletions in 6-7% of Duchennepatients suggested that this was the site of the Duchenne gene. Conserved sequences fromthis region of the X chromosome were used to identify cDNA (complementary DNA) clonesmade by reverse-transcribing mRNA (messenger RNA). The sequences encoded in the cDNAwere shown to be deleted in a number of boys with Duchenne muscular dystrophy. Thisprovided evidence that the isolated cDNA clones were indeed from the gene that is mutatedin muscular dystrophy. The gene, known as the Duchenne muscular dystrophy gene, wasshown to encode a protein that Hoffman and Kunkel's laboratory (Hoffman et al., 1987)named dystrophin, and as expected, this protein was missing from the skeletal muscle tissueof boys with Duchenne muscular dystrophy (Monaco et al., 1988).The mdx mutant of the C57B1110SnJ strain of genetically dystrophic mouse hasrecently been introduced as an animal model applicable to the study of X-linked humanmuscular dystrophies (Bulfield et al., 1984). In several of the earliest pathological studies ofthe mdx mouse, it was recognized that some of the histopathological features resembledquite closely those seen in Duchenne muscular dystrophy (Bulfield et al., 1984; Anderson etal., 1987, 1988; Carnwath and Shotton, 1987; Torres et al., 1987; Coulton et al., 1988)while others did not fit at all well. More importantly, however, was the fact that the samebiochemical defect was shown to be present in mdx skeletal muscle, in that it lacked theprotein dystrophin (Hoffman et al., 1987). Analysis of the dystrophin gene in the mdx mouserevealed that the genetic defect was a point mutation involving a single base change within anexon (Sicinski et al., 1989). This feature of the mdx mouse made it a reliable model for2characterizing the primary pathological processes linking the absence of dystrophin tonecrosis in skeletal muscle fibers.Proposed Structure and Function of DystrophinOnce the cDNA had been isolated and sequenced, the deduced amino acid sequence ofdystrophin was compared to those sequences already present in protein databases. Bycomparison with such known protein sequences it was found that certain domains of theprotein were very similar to cytoskeletal proteins such as alpha-actinin and spectrin,suggesting that dystrophin is itself a cytoskeletal protein (Koenig et al., 1988). The aminoacid sequence predicted a rod-shape protein approximately 150 nm in length that could bedivided into four different domains: an N-terminal "actin-binding" domain (Hammond, 1987), amiddle domain formed by 25 triple helical segments, a cystein-rich domain, and a lesscharacterized C-terminal domain (Koenig et al., 1988). The first three domains showsignificant similarity to the three domains of alpha-actinin whereas amino acid sequences ofspectrin are similar to the succession of 25 triple-helical segments in the second domain. TheC-terminal of dystrophin, however, does not show any similarities to other known proteins butis thought to be the domain that mediates the attachment to an integral membraneglycoprotein in the sarcolemma (Campbell and Kahl, 1989).Dystrophin has been shown to be associated with a complex of sarcolemmalglycoproteins that are believed to provide a linkage to the extracellular matrix protein, laminin(Campbell and Kahl, 1989; Ervasti et al., 1990; Ohlendieck and Campbell, 1991; Ibraghimov-Beskrovnaya et al., 1992). The absence of dystrophin leads to a dramatic reduction in thesedystrophin-associated glycoproteins in the sarcolemma of patients with Duchenne musculardystrophy and mdx mice (Ervasti et al., 1990; Ohlendieck and Campbell, 1991). Morerecently, it has been demonstrated that a dystrophin-related protein named utrophin3(Matsumura et al., 1992; Tinsley et al., 1992), an autosomal homologue of dystrophin, isassociated with an identical or antigenically similar complex of sarcolemmal proteins in skeletaland cardiac muscle of both normal and mdx mice.Synthesis of an Antibody to DystrophinAdditional information about dystrophin came from the synthesis of fusion proteins inbacteria from chimeric genes in which a portion of the dystrophin cDNA was joined with abacterial gene (Hoffman et al., 1987). The resulting fusion protein was injected into rabbitsor sheep to make an antibody against the dystrophin part of the protein. Alternatively,synthetic peptides of approximately 13 amino acids were created utilizing the sequenceinformation, and these peptides were conjugated to larger molecules and injected intoanimals, again with the purpose of generating antibodies. The antibodies directed againstdystrophin then became important molecular tools for the analysis of dystrophin structureand function. In particular, Western blot analysis of muscle and other tissues demonstratedthat the antibodies recognized a protein of 427 kiloDalton (kD) molecular weight. The proteinwas present in skeletal muscle, cardiac muscle, smooth muscle, and brain of normal patientsand absent in those tissues of patients with Duchenne muscular dystrophy (Hoffman et al.,1987).Using these antibodies, immunocytochemical studies of dystrophin on tissue sectionsrevealed that, in skeletal muscle, dystrophin is localized at the sarcolemmal membrane ofextrafusal muscle fibers (Hoffman et al., 1987; Zubrzycka-Gaarn et al., 1988). Transmissionelectron microscopic studies coupled with immunogold labelling techniques have further shownthat this protein is situated on the cytoplasmic side of the sarcolemma (Watkins et al., 1988;Carpenter et al., 1990; Cullen et al., 1990; Squarzoni et al., 1992). Observations from these4studies suggested that the dystrophin molecules are linked in a form of a lattice immediatelybeneath the plasma membrane.Immunocytochemical studies have demonstrated elevated levels of dystrophin at themotor end-plate regions of extrafusal fibers in normal muscle (Miike et al., 1989; Shimuzu etal., 1989; Byers et al., 1991; Huard et al., 1991, 1992). In the study by Shimuzu andcoworkers (1989), the accumulation of dystrophin at the neuromuscular junction wasconfirmed by revealing acetylcholinesterase activity by light microscopy, whereas Huard'sgroup (1991) used alpha-bungarotoxin coupled to cascade blue to identify cholinergicreceptors in skeletal muscle. Their observations emphasize a relationship between dystrophinexpression and the site of innervation of skeletal muscle fibers. Additionally, studies of thecentral nervous system (Lidov et al., 1990) have revealed detectable amounts of dystrophinin postsynaptic regions of cerebral cortical neurons and cerebellar Purkinje cells, suggesting apossible role for dystrophin in synaptic transmission.Recent work on dystrophin localization in smooth muscle has provided evidence thatdystrophin is associated with specific regions of the smooth muscle sarcolemma (North et al.,1993). By immunofluorescence and immunoelectron microscopy, it was shown that twostructurally distinct smooth muscle sarcolemmal domains exist in an alternating patternparallel to the long axis of the smooth muscle cell. Dense plaque regions, known to be involvedin actin anchorage and characterized by a specific set of junctional proteins (Geiger andGinsborg, 1991), typically expressed a marked diminution of dystrophin whereas theintervening caveolae-rich areas of the sarcolemma showed relatively normal levels ofsarcolemmal labelling. This mutually exclusive colocalization of specific membrane-associatedproteins in smooth muscle suggests that dystrophin complements other cytoskeletal proteinsthat function in maintaining the integrity of the smooth muscle cell and its sarcolemma.Since the discovery of the DMD gene, the literature on dystrophin localization in theextrafusal fibers of skeletal muscle has escalated at a rapid rate. In spite of this, however,5very little has been published on the localization of dystrophin in the specialized intrafusalmuscle fibers of muscle spindles. The following section will review some of the characteristicfeatures of the mammalian muscle spindle with the aim of providing an appreciation for itshighly complex nature.B. The Mammalian Muscle SpindleMuscle spindles are specialized sensory stretch-receptors embedded within mostvertebrate skeletal muscles. They were first described by Hassall (1849) and later by KUhne(1864) who gave them the name Muskelspindeln in describing their spindle shape. It was Cajal(1888) who recognized this structure as a sense organ and also suggested the existence ofa specific motor innervation. Its sensory nature was elegantly confirmed by Sherrington(1894), and the first detailed studies of the anatomy of mammalian muscle spindles by Ruffini(1898) have provided the basis of our present understanding of this receptor.The mammalian muscle spindle consists of discrete bundles of encapsulated skeletalmuscle fibers, termed intrafusal fibers, that stand out by the comparative smallness of theirsize to the surrounding extrafusal fibers. These small-diameter muscle fibers are associatedwith sensory and motor nerve endings that terminate on specific regions along their lengths.Lying in close proximity, nerve axon bundles, extracapsular capillaries, and mast cells are alsooften encountered.The microscopic appearance of these receptors can be described in terms of the levelof section through each receptor (Sherrington, 1894). In a series of cross-sections, threeregions or zones can be identified according to the overall diameter of the receptor andextent of capsular investment. The equatorial zone represents the central or widest portionof the receptor. In this region, both an inner and an outer spindle capsule are present,separated by a large fluid-filled periaxial space. The so-called polar regions represent the two6tapered ends of the spindle. In these regions, only the outer capsule persists and in somecases the intrafusal fibers extend beyond the capsular limits and either terminate in theendomysium of neighboring extrafusal fibers or insert into tendon. Midway between thesetwo regions, the so-called juxtaequatorial zone is present.Muscle spindles can be described as consisting of four basic elements: A capsule,intrafusal fibers, and their sensory and motor nerves. Each of these will be addressed in thefollowing paragraphs.CapsuleThe muscle spindle capsule is composed of a multilayered outer capsule and a thin innercapsule that differ from each other both in structure and location (Merrillees, 1960; Landon,1966; Corvaja et al., 1969; Ovalle and Dow, 1983). The outer capsule consists of severalconcentric layers of flattened epithelial-like cells and connective tissue elements. It iscontinuous with both the extrafusal and the intrafusal endomysium as well as with theperineurium which surrounds the small bundles of nerve fibers supplying the receptor organs.A dilatation of the capsule is generally found at the central, equatorial region of the spindlegiving the receptor its characteristic fusiform shape (KUhne, 1864).The inner capsule or axial sheath (Barker, 1974) is most prominent in the equatorialregion where it may form an elaborate sheath around individual intrafusal fibers, or groups offibers and their sensory nerve endings. It consists of flattened cells that resemble theendoneurial cells of peripheral nerve and unlike the outer capsule cells, cells of the innercapsule lack an external lamina although collagen and elastic fibrils are known to occupy theparacellular regions (Cooper and Gladden, 1974).7In the equatorial region, a space is present between the outer and inner capsules.This so-called periaxial space contains a jelly-like material rich in acid mucopolysaccharides andhyaluronic acid (Brzezinski, 1961a, b; James, 1971; Fukami, 1982, 1986; Ovalle and Dow,1988). This substance bears some resemblance to vitreous humor and it is thought toinsulate as well as mechanically protect and lubricate the internal components of thereceptor in equatorial regions (Barker and Banks, 1986).It should also be mentioned that the outer capsule presents a significant barrier tothe passage of the exogenous protein tracer, horseradish peroxidase, and bears someresemblance to the permeability barrier formed by the perineurium of peripheral nerves (Dowet al., 1980). The horseradish peroxidase tracer was found to freely enter the spindle at itspolar regions, but in the equatorial zone, the outer capsule cells were effective in preventingthe indiscriminate penetration of this protein tracer into the periaxial space. Thus, a blood-muscle spindle barrier appears to exist in central equatorial zones, but not in more distalpolar regions.Intrafusal FibersThe specialized skeletal muscle fibers in a muscle spindle are unique in that they varymorphologically not only from the surrounding extrafusal fibers but also show variationsamongst themselves. Two distinctly different intrafusal fiber types can be identified on thebasis of their appearance and according to the arrangement of nuclei in their centralequatorial regions (Barker, 1948; Boyd, 1962; Cooper and Daniel, 1963). The nuclear bagfibers derive their name from the dilatation produced by the numerous tightly-packed,vesicular nuclei in the equatorial region. The nuclei are surrounded by a thin peripheral layerof sarcoplasm containing few myofibrils. At either end of this central portion there are twojuxtaequatorial regions in which the nuclei continue as a single central row surrounded by8sarcoplasm and a peripheral layer of contractile material. These fibers have the widestdiameter of the two, and are also the longest, often extending beyond the capsularinvestment of the spindle. Nuclear chain (chain) fibers, in contrast, contain in their equatorialregion a more conspicuous amount of contractile material which surrounds a single, centralrow of elongated nuclei. They are smaller in diameter and shorter in length, often endingwithin the capsule. There are, however, some that extend beyond the limits of the capsule.These are termed long chain fibers and were first described by Harker and coworkers(1977). In polar regions of both types of fibers the nuclei assume a peripheral locationunderneath the sarcolemma and resemble more closely the architecture of extrafusal musclefibers.It is now known that the nuclear bag fibers are themselves of two types, which differin their histochemistry, ultrastructure, and mechanical properties. They are designated asnuclear bag1 (bags) and nuclear bag2 (bag2) fibers (Ovalle and Smith, 1972) according totheir histochemical staining properties. Another important difference between bags and bag2fibers is that the latter are surrounded by prominent elastic fibers in their polar regionswhereas the former are not (Gladden, 1976). In terms of their differing mechanicalproperties, the nuclear bag fibers have been described as either dynamic nuclear bag (bags)or static nuclear bag fibers (bag2) (Boyd et al., 1975).The usual methods by which the three mammalian intrafusal fiber types are identifiedin light microscopy are the histochemical reactions for actomyosin adenosine triphosphatase(mATPase) and nicotinamide adenine dinucleotide-tetrazolamine reductase (NADH-TR). Thedifferential staining observed with these reactions in bag1, bag2, and chain fibers suggeststhe presence in each of a different myosin profile and energy requirements. The bags fiberexhibits low mATPase activity following alkaline preincubation and has a relatively low glycogencontent. In contrast, the bag2 fiber exhibits medium to high ATPase activity and has a medium9glycogen content. The chain fibers exhibit high ATPase activity and have a high glycogencontent (Banks et al., 1977).Recent immunofluorescence work with newly developed monoclonal antibodies hasproved to be a more elaborate and comprehensive technique in understanding the myosinprofiles (isoforms) that constitute each intrafusal fiber type. Studies of spindle fibers incapsular regions have revealed that antibodies specific for fast fiber myosins labelpredominantly the chain fibers and those specific for slow fiber myosins show a strongreaction in bag fibers (Rowlerson et al., 1985). Tonic myosin antibodies were shown to labelbag I fibers, and when used at higher concentrations exhibited a weak reaction with chainfibers. In some occasions it was noted that this antibody reacted with bag2 fibers. Antibodiesraised against embryonic and neonatal myosins (Sartore et al., 1982) do not bind to any adultextrafusal fibers and have little reaction with bags fibers. However, this antibody reacts veryintensely with chain fibers and moderately with bag2 fibers. Therefore, chain fibers appear tocontain a mixture of either embryonic, neonatal and fast fiber myosins, or a myosin isoformthat is recognized by both types of antibodies.Kucera and Walro (1989) have demonstrated that regional differences occur in themyosin heavy chain expression in the intrafusal fibers. In capsular regions of intrafusal fibers,myosin heavy chains are significantly decreased whereas extracapsular regions exhibit astrong reaction similar to that of the type I (slow) extrafusal fibers. Moreover, specificregional differences were noted between the three intrafusal fiber types. The highcorrelation between the various antibody-specific myosin isoforms and the contractile andelectrophysiological properties of intrafusal fibers have demonstrated thatimmunocytochemical techniques are reliable in the typing of intrafusal fibers.In other immunocytochemical studies, the expression of certain structural muscle fiberproteins was shown to be distributed in a non-uniform arrangement in the different regions ofthe intrafusal fibers. Maier and Zak (1990) showed that actin, tropomyosin, desmin, and10myosin heavy chains take on a striated appearance in polar regions, while the equatorappears non-striated. These observed immunofluorescent patterns suggest that, at theequator, these sarcoplasmic proteins are assembled into looser arrays than in thesarcomeres of the pole. This supports the concept that the equatorial zones arestructurally more flexible, an appropriate substrate for distorting the affixed sensory endingsduring an applied stretch.InnervationSeveral histological methods are commonly used to study motor and sensoryinnervation of mammalian muscle spindles (Barker and Ip, 1963; Gladden, 1970; Namba et al.,1967; Nahirney and Ovalle, 1992a, b, 1993). Some of these procedures are normallyperformed on muscle pieces by block impregnation with silver salt solutions followed by teasingapart the impregnated tissue, while others are perfomed on histological sections.The most commonly used procedure to study innervation patterns is the silverimpregnation technique (Barker and Ip, 1963) which enables the innervation pattern of entiremuscle spindles to be visualized more or less completely.There are, however, other techniques that allow for the identification of nerve fibersin skeletal muscle. Roden and coworkers (1991) have shown by immunocytochemistry thatthe 200 kD neurofilament protein, a component of the intermediate filaments found inneuronal tissue, is expressed at high concentrations in both the axons and terminal endings(motor end-plates) of nerves. This immunocytochemical staining method may be useful instudies of muscle innervation since it can be used in conjunction with other antibody labellingprocedures (Nahirney and Ovalle, 1992a, b, 1993).1 1The intrafusal fibers of muscle spindles are separately innervated by motor andsensory nerves which terminate on different regions along these fibers (Ruffini, 1898). Whilelarge myelinated sensory nerve fibers pass through the capsular sleeve to enter theequatorial space, the motor supply may enter the spindle either with afferent nerve fibers orindependently through the polar regions.The sensory innervation of mammalian muscle spindles comprises both primary andsecondary nerve endings, the latter being less abundant or in some cases absent. Theprimary sensory endings consist of spiral or annular terminations, each of which encircles thedensely nucleated equatorial regions of the intrafusal fibers (Ruffini, 1898). These endingscan be found on each of the three intrafusal fiber types and all are connected to the samegroup la afferent nerve fiber leaving the spindle. The group la afferent nerve fibers can rangein diameter from 7 jim to 15 gm. In the cat, the primary endings cover 33-37 % of the bag1fiber surface, 25% of the bag2 fiber surface, and 5-12 % of the surface of individual chainfibers (Banks et al., 1982; Banks, 1986). Boyd (1962) has shown that the region of theintrafusal fibers surrounded by the primary endings in cat muscle spindles extends for about300 um in the equatorial zone.Secondary endings can be found adjacent to the primary endings in thejuxtaequatorial regions of the muscle spindle. They consist of spiral terminations around eachnuclear chain fiber and less extensive flower-spray terminations on the nuclear bag fibers(Boyd and Smith, 1984). The spiral and spray terminations are connected to the same groupII afferent axon, averaging 8 j.tm in diameter. Some spindles have been shown to have nosecondary endings at all, whereas the most complex can have four or five such endings (Boydand Smith, 1984). The most common arrangement is for a spindle to have one primary endingand one secondary ending (Boyd and Smith, 1984). In the cat tenuissimus, the total terminalcontact area of secondary endings has been estimated to be 16-22% on individual chainfibers, 17% on bag2 fibers, and 8% on bag1 fibers (Banks et al., 1982).12The ultrastructure of both types of sensory endings is essentially identical (Adal,1969; Landon, 1972). Both contain numerous mitochondria, neurofilaments, microtubules,and axoplasmic vesicles. The endings lie in close apposition to the sarcolemma of the intrafusalfiber and, in the case of the primary endings, sit in troughs indented on the surface of theintrafusal fiber (Landon, 1966). In both types of ending, the sensory terminals are separatedfrom the sarcolemma of the adjacent muscle fiber only by the plasma membrane of the nerveterminal itself. That is, the external lamina of intrafusal fibers does not extend through thegap between the two plasma membranes of nerve and muscle, but continues over the surfaceof the sensory endings (Merrillees, 1960; Landon, 1966; Corvaja et al., 1969; Hennig, 1969).An intercellular gap of about 15-20 nm separates the adjacent cell membranes of theintrafusal fiber and sensory ending (Corvaja et al., 1969) and, in some primary endings, thepresence of finger-like prolongations which penetrate from the internal surface of theprimary sensory ending into the depth of the intrafusal fiber have been observed (Corvaja etal., 1969). Additionally, Schwann cells are not usually present on the sensory terminals(Merrillees, 1960), a feature that distinguishes sensory endings from motor terminals.The complex nature of motor innervation to muscle spindles has been a topic ofinterest and one of controversy since it was first described by Cajal in 1888. Since a singlespindle receives from eight to twenty-five branches of sensory or motor axons it has takenmany years for the patterns of innervation to be elucidated. To date, many detailed reviewsof the motor innervation to spindles have been published (Boyd, 1981; Boyd and Smith, 1984;Barker and Banks, 1986).The motor endings are distributed in the juxtaequatorial and polar regions ofmammalian intrafusal fibers. In general, two forms of motor innervation to intrafusal fibershave been described. The first type arises from gamma efferents, a group of axons withsmaller diameters and slower conduction velocities than those of alpha efferents which supplythe extrafusal fibers (Leksell, 1945). These nerve axons innervate all types of intrafusal13fibers and are sometimes referred to as fusimotor axons (Barker, 1974). They have beenestimated to account for 30% of the nerve fibers in the ventral roots (Boyd and Smith,1984). The second type are termed beta efferents or skeleto-fusimotor axons (Barker,1974). They commonly innervate both extrafusal and intrafusal fibers by way of collateralbranches. These nerve fibers are generally low in number and, when seen, are located in thedistal polar regions of intrafusal fibers (Barker, 1974).Both the gamma and beta efferents have been subdivided into static and dynamicsubgroups based upon the intrafusal fibers they innervate and how they influence theresponse of the primary and secondary endings (Matthews, 1962; Boyd et al., 1975; Boyd,1981). Generally speaking, gamma-static axons can innervate bag2 fibers, chain fibers, orboth. Motor axons supplying bags fibers are often of the gamma-dynamic and beta-dynamicvariety. The beta-static axons are usually restricted to the extracapsular regions of longchain fibers (Kucera, 1980).The gamma axons exhibit two types of terminal endings. The gamma-static axons thatterminate on either bag2 or chain fibers or both are called "trail endings" (Barker et al.,1970). They are typically found in the juxtaequatorial regions of these fibers and almost alllie superficially on the muscle fiber with little or no postsynaptic folding. The other terminalendings that are usually found in the midpolar zone correspond to the gamma-dynamic axonsand are termed "plate2 endings" or "P2 plates" (Barker et al., 1970). These axon terminalsare nearly all clearly indented into the surface of the muscle fiber, however, little or no foldingof the postsynaptic membrane is evident. "Plate1 endings" or "P1 plates" are thought to bethe terminations of beta axons (Barker et al., 1970). They are smaller than P2 plates, and, infact, resemble the plates of alpha motor axons on extrafusal fibers, however, show a lesserdegree of postsynaptic membrane folding. Beta-dynamic and beta-static axon endings havebeen shown to be found on bags and long chain fibers respectively, the majority of which arepresent in the extracapsular regions (Kucera, 1980).14Thus, it is evident that the complex innervation, both sensory and motor, of intrafusalfibers is in contrast to the innervation of extrafusal fibers which, as a rule, are innervated byonly one motoneuron, have a single motor end plate, and are not innervated by sensorynerves.PathologyDespite considerable progress in the basic knowledge of muscle spindles there is still nodisease that can be directly attributed to defective or abnormal muscle spindle function. Inmost diseases, pathological changes in spindles appear to be nonspecific and do not parallelthose in the surrounding extrafusal fibers (Lapresle and Milhaud, 1964; Patel et al., 1968).Nevertheless, numerous pathological changes have been recorded and, for the most part,have been consequences of either denervation experiments or neuropathies.Studies of spindle morphology after denervation have been undertaken by Onanoff(1890), Tower (1932, 1939), Boyd (1962), Barker et al. (1970), and SchrOder (1974a, b).Tower's studies (1932, 1939) revealed an increased thickness of the spindle capsules and ofthe fibrous tissue in the periaxial space after sensorimotor denervation and after ventralroot section of prolonged duration, but not after sensory root section or dorsalganglionectomy. In man, sensorimotor denervation, whether due to mononeuropathy,diabetes mellitus, or drug-induced neuropathy, results in striking capsular thickeningconsisting of increased numbers of lamellae of perineurial capsular cells and increasedamounts of collagen (De Reuck, 1974; Swash and Fox, 1974). In sensory denervation due totabes dorsalis, or to carcinomatous sensory neuropathy (Croft et al., 1965) deposition ofcollagen in the periaxial space is prominent only in the equatorial and juxta-equatorial regions,whereas in motor denervation, capsular thickening is marked only in polar regions (Swash andFox, 1974).15In both sensory and motor denervation, it has been reported that the nuclear chainfibers undergo earlier and more severe degenerative changes than the nuclear bag fibers(Swash and Fox, 1974) and, after a year or more, all the intrafusal muscle fibers showdegenerative changes, becoming clumped together in a poorly defined mass in the center ofthe periaxial space. After prolonged motor denervation, however, polar regions show similaratrophy of the intrafusal fibers, whereas equatorial regions show normal primary andsecondary sensory endings on nuclear bag fibers of only slightly reduced diameter (Swash andFox, 1974).Morphological studies of muscle spindles in patients with Duchenne muscular dystrophy(Lapresle and Milhaud, 1964; Cazzato and Walton, 1968) have shown swelling of periaxialspaces of spindle capsules, and degeneration of intrafusal fibers. In a quantitative study of230 muscle spindles by Swash and Fox (1976), the capsular thickness found in Duchennedystrophy was not as great as that found in denervated spindles and only a slight decrease indiameter and in number of intrafusal muscle fibers was observed. However, they noted thatin some spindles the intrafusal fibers consisted of a degenerate mass of centrally placedamorphous material, especially in areas of marked extrafusal fiber degeneration and fibrotictissue accumulation whereas in regions of relatively well-preserved extrafusal fibers, themuscle spindles were usually less abnormal. In silver-impregnated longitudinal sections, thepattern of motor and sensory innervation appeared normal (Swash and Fox, 1976). Theseabnormalities described in spindles in Duchenne muscular dystrophy have been proposed tocontribute to the relatively early loss of tendon reflexes often found in this condition (Swashand Fox, 1976).16C. Immunofluorescence MicroscopyImmunofluorescence is a special field of fluorescence microscopy that involves thedetection of antigens by way of immune reactions. The original technique was developed byCoons and Kaplan (1950), and modifications of this technique are currently the most widelyused microscopic fluorescence method in research. Originally restricted to fluoresceinisothiocyanate (FITC), immunolocalization techniques now utilize other, newly developedfluorochromes such as rhodamine B 200, tetramethyl-rhodamine-isothiocyanate (TRITC), andTexas Red.The basic principle of immunofluorescence is based on the reactions of specificantigens to specific antibodies. Antibodies are submicroscopic protein structures thatattach themselves to specific recognition sites on the surface of antigens. They are proteinsof the globulin group (immunoglobulins) that appear in plasma and tissue fluids after antigeninjection to the host organism. The immunoglobulins that are produced by the host reactspecifically and bind strongly to the antigen.Immunofluorescence is the coupling of immunoglobulins to substances that fluoresce(fluorochromes) rendering them visible in the microscope without causing loss of theantibody's biologic activity. Two basic methods are used for antigen localization inimmunofluorescence (Coons and Kaplan, 1950). The direct method is a simple detectionsystem that uses a solution containing a prelabelled antibody (i.e. conjugated to afluorochrome) on sections of tissue. The excess antibody is washed off, and the tissue isobserved under the fluorescence microscope. The second method is known as the indirect or"sandwich" method. To avoid the preparation and storing of specifically labelled antibodies foreach antigen, the indirect method utilizes anti-antibodies that are produced in anintermediate host of a different species. These anti-antibodies, or so-called secondaryantibodies, will react with every original (primary) antibody complex as if it were an antigen.The secondary antibody can be visualized by directly attaching a fluorochrome, or by17conjugating it to biotin which is then linked to a fluorochrome via the bacterial proteinstreptavidin. The biotin-streptavidin system offers the advantage of an amplification step andis therefore useful in the detection of antigens at low concentrations in tissues (Hsu et al.,1981a, b).Until recently the use of transmitted-light fluorescence microscopy was the onlymethod of observing fluorochromes. The few available exciter filters with their widetransmittance ranges did not allow the use of specific fluorescence techniques and was not avery reliable method for routine diagnosis. However, the situation has changed with theincreasing impact of the fluorochrome labelling techniques. Epifluorescence microscopy, asystem that uses incident-light excitation, has essentially replaced transmitted-light darkfieldillumination. In this type of microscope, illumination and observation can be made from thesame direction.In summary, the use of immunofluorescence can reveal microscopic details of specificantigen localization not obtainable by other histochemical methodologies. It is a powerfultechnique that can provide a better understanding of the organization of specificcomponents such as proteins within tissues. Furthermore, simultaneous localization of twospecific antigens (double-labelling) using two monospecific antibodies and two differentcolored fluorochromes is possible and is an effective way to study the relationship of oneantigen to another (Vandesande, 1983). In this respect, it would be advantageous to applythis method to the study of the localization of specific proteins in skeletal muscle and musclespindles.18D. Thesis ObjectivesAlthough there is extensive literature on dystrophin in normal and dystrophicextrafusal muscle fibers (Bonilla et al., 1988; Hoffman et al., 1988; Carpenter et al., 1990;Cullen et al., 1990; Byers et al., 1991), detailed information concerning the localization ofdystrophin in either normal or diseased intrafusal muscle fibers of muscle spindles is lacking.To more clearly understand the role this protein plays in skeletal muscle it is necessary todetermine its precise localization in the different types of muscle fibers that constitute askeletal muscle. The intrafusal fibers of muscle spindles present a unique array ofcharacteristics that differentiate them in both structure and function from the larger andmore numerous extrafusal fibers.Therefore, the present investigation was undertaken with the following aims:(1) To determine the localization of dystrophin along the lengths ofintrafusal fibers in normal and mdx-dystrophic skeletal muscle.(2) In view of the elevated levels of dystrophin reported to occur atextrafusal fiber motor-end-plates of normal skeletal muscle, it was ofinterest to determine the relationship between sensory nerveinnervation and dystrophin expression in the intrafusal fibers.(3)^To determine the effects of chronic denervation on dystrophinexpression in skeletal muscles and muscle spindles of normal mice.The following hypotheses were tested in this study. First of all, that intrafusal fibersof mdx-dystrophic mice exhibit a similar deficiency of dystrophin as that seen in theextrafusal fiber population. Secondly, in normal skeletal muscle, the sensory innervation ofintrafusal fibers plays a role in modulating the subsarcolemmal expression of dystrophin. Andthirdly, chronic denervation of normal skeletal muscle does not affect the expression of19dystrophin at synaptic and non-synaptic sarcolemmal membranes of extrafusal and intrafusalmuscle fibers.This study will attempt to bring together the current knowledge of dystrophinlocalization in skeletal muscle and will also provide new information about its distribution in theintrafusal fibers of muscle spindles in both normal and pathological states.20Eighteen normal (C57BI/10SnJ) and 8 dystrophic (C57BI/10mdx) male mice wereused in this study. Breeding pairs were originally obtained from Jackson Laboratories (BarHarbor, ME) and were raised and maintained in our animal facility.The soleus muscles of both normal and mdx mice at 32 weeks of age were used. Atotal of 24 muscles from 12 normal mice and 16 muscles from 8 dystrophic mice wereexamined in this study. Animals were first killed with an overdose of halothane. The soleusmuscles from both hindlimbs were rapidly excised and pinned at resting length on small woodenblocks between two thinly sliced pieces of liver (for transverse sectioning), or adhered with asmall drop of O.C.T. embedding medium (Miles Laboratories, Naperville, Ill.) to the wooden blockfor longitudinal sectioning. Specimens were quickly frozen by plunge immersion in isopentanecooled to -196° C in liquid nitrogen for 30 seconds and then transferred to a Forma ScientificBio Freezer maintained at -63° C for storage until sectioning and subsequent staining.Before sectioning, specimens were mounted on metal chucks in either transverse orlongitudinal orientations to the long axis of the muscle fibers and allowed to equilibrate to-20° C in a Bright Instruments 5030 series cryostat microtome (Huntington, England). Serialsections of 5-gm thickness were cut with a metal knife and collected on polylysine-coatedglass slides or coverslips. The slides were stored in a -20° C freezer for up to 2 weeks untilstaining procedures were performed.HistochemistrySections were stained with either hematoxylin and eosin (H & E) or with a modifiedprocedure to detect myosin adenosine-triphosphatase (mATPase) at acid (pH 4.6)preincubation (Johnson and Ovalle, 1986). This ATPase method was adopted because it has21the advantages of being less time consuming than previously published protocols, is relativelysafe since the ammonium sulfide step is eliminated, and can detect all three extrafusal fibertypes simultaneously using the same pH (Johnson and Ovalle, 1986).The following steps in this procedure are outlined as follows. Cryostat sections (5-gmthickness) were collected on coverslips and allowed to air dry at room temperature. Thecoverslips were preincubated for 3 min in barbitol acetate buffer (pH 4.6), washed twice for60 sec in basic medium (pH 9.4), and then incubated in an adenosine triphosphate (ATP) basicmedium (pH 9.4) at 37°C for 20 min. Following incubation, the sections were stained with a1% aqueous toluidine blue solution for 8 sec and then rinsed well with distilled water. Thecoverslips were then processed through an ascending series of ethanols (50, 70, 90, 100%),cleared in xylene, and mounted with Histoclad (Clay Adams, Parsippany, NJ).Intrafusal fibers were typed according to the bag1, bag2, and chain classification ofOvalle and Smith (1972). The histochemical profiles along the lengths of individual intrafusalfibers were determined by examining serial sections of the receptor at intervals of 70 gm.Selected transverse profiles of spindles were photographed on T-Max 100 Kodak film using aLeitz Orthoplan photomicroscope equipped with brightfield optics and a tungsten light source.lmmunofluorescenceTransverse and longitudinal sections were immunostained at room temperature in ahumidified chamber with the original polyclonal antibody (made in sheep) directed toward the60 kD fusion protein of dystrophin (kindly donated by Dr. Eric Hoffman from the University ofPittsburgh) diluted 1:1,000 in phosphate-buffered saline with bovine serum albumin (PBS BSA)at pH 7.3 for 2 hr. Control sections were incubated with pooled normal sheep serum. Afterrepeated washes (3 x 15 min) with PBS BSA, a secondary biotinylated anti-sheep IgG(Amersham, Oakville, ONT) diluted 1:200 was applied for 1 hr, washed (3 x 15 min) and22decorated with a streptavidin-Texas red conjugate (Amersham, Oakville, ONT) (45-minincubation). Sections were washed in PBS BSA (3 x 15 min), coverslips were appliedtemporarily, and dystrophin immunolabelling was examined with a Zeiss axiophot epi-fluorescence photomicroscope using a G 365 exciter filter. Muscle spindles were located inboth transverse and longitudinal planes and photographed on T-Max 400 Kodak (35 mm) blackand white print film pushed to 1600 ASA.Innervation patterns of extrafusal and intrafusal fibers were revealed byconcommittant use of a nerve-specific antibody (Roden et al., 1991) that recognizes a 200kD neurofilament protein. The coverslips of previously dystrophin-labelled slides were removedand sections were incubated for 1 hr with the anti-neurofilament (200 kD) antibody (Sigma,St. Louis, MO) diluted at 1:32, washed in PBS BSA (3 x 15 min), and then immunoreacted withan anti-rabbit-FITC conjugate (Sigma, St. Louis, MO) diluted 1:32 in PBS BSA for 30 min. Aftera final rinse in PBS BSA, coverslips were mounted with a 1:1 PBS BSA/glycerol medium.Previously photographed dystrophin-stained spindles were relocated and their innervationpatterns were then photographed using epifluorescence and a BP 450-490 exciter filter.DenervationSix male C57BI/10SnJ mice were used to study the effects of denervation bysectioning the sciatic nerve on dystrophin and neurofilament expression in normal skeletalmuscle. Denervation was performed on mature (32 wk) mice under halothane anesthesia. Anincision was made in the region just below the greater trochanter of the right femur and thesciatic nerve was located and teased apart from its surrounding connective tissue. A largesection (5 mm) of the nerve was removed to prevent the possibility of reinnervation. Thecutaneous incision was then closed and sutured. Upon regaining consciousness, thedenervated leg of the mouse was flaccid and lack of motor control in the leg was evident by23the failure to maintain a flexed or extended position. At 21 and 42 days post-denervationintervals, the animals were killed with an overdose of halothane. The soleus muscle from thedenervated limb was rapidly excised and placed between two thinly-sliced pieces of liver.Unoperated muscles from age-matched normal mice were placed alongside the denervatedmuscles to serve as controls for immunofluorescent and morphological evaluation.To determine if the animal was chronically denervated, the site of the nerve lesion wasrelocated. In all cases the two cut ends of the sciatic nerve had not reunited. Additionally,immunostaining with the neurofilament protein antibody was utilized to confirm whetherreinnervation had occurred. An absence of staining in axons and motor and sensory nerveterminals indicated non-reinnervation.Serial transverse cryostat sections were immunolabeled with the dystrophin antibody(60 kD) using the previously described labelling procedure and were subsequently observedunder epifluorescence. Denervated intrafusal fibers were examined throughout the differentregions of the receptors for dystrophin labelling. Sections of muscle spindles were also post-stained with H & E to provide morphological details.24OBSERVATIONSGeneral MorphologyThe mouse soleus muscle has been studied extensively by morphologists, histochemists,and physiologists (Jasch et al., 1982; Bressler et al., 1983; Ovalle et al., 1983; Johnson andOvalle, 1986; Ovalle and Dow, 1986) due to its small size, ease of accessibility, unique fibertype population and functional properties, and the relative abundance of muscle spindles. It isclassified as a slow-contracting muscle and, consequently, exhibits a proportionally highnumber of slow-twitch extrafusal fibers to other muscles in the leg. Located in the posteriorcompartment of the hindlimb just anterior to the large gastrocnemius, the soleus serves animportant role in the maintenance of posture (Barker, 1974).When viewed in transverse-section by light microscopy, the normal soleus musclecontained profiles of polygonally-shaped extrafusal fibers arranged in discrete pleomorphicfascicles (Fig. 1). Intramuscular nerve fascicles and blood vessels were also observed andcoursed through the perimysial component of the muscle. The extrafusal fibers werecharacterized by peripherally located nuclei and exhibited overall diameters ranging from 35to 70 gm. Very little intervening endomysial tissue was present between the muscle fibers.In H & E stained sections of the mdx soleus, the extrafusal fibers exhibited obvioussigns of pathological change including central nucleation, degeneration, and extreme variationin size and shape (Fig. 2). Extrafusal muscle fibers ranging from 20 gm to 90 gm in diameterwere encountered and evidence of fiber splitting was seen in a relatively high number of theextrafusal fibers. Additionally, the endomysial and perimysial connective-tissue showed highlevels of mononuclear cell infiltration and an overall increase in paracellular elements (Fig. 2).Muscle spindles in the normal soleus typically contained 4-5 intrafusal fibers, twolarge-diameter bag fibers and two or three small-diameter chain fibers (Fig. 3a, b, c). Bag25fibers usually extended well beyond the limits of the capsular sleeve, while the chain fibersterminated within capsular regions or shortly thereafter. In equatorial regions, the intrafusalfibers were encased by a delicate inner capsule that separated them from a large periaxialspace (Fig. 3a). The outer capsule formed a prominent layer around the receptor and wasoften flanked by nerves and blood vessels within the surrounding perimysial tissue.In contrast to the extrafusal fibers, intrafusal fibers in the mdx soleus displayed arelatively minor degree of histological change in dystrophy. In sections stained with H & E (Fig.4a, b, c), the total number and overall appearance of the intrafusal fibers was similar to thosein the age-matched normal soleus. The number of intrafusal fibers ranged from 3 to 6 andthe lengths of the fibers did not show a significant difference from that of their normalcounterparts. Capsular thickening, however, was particularly evident in equatorial regions ofmdx spindles (Fig. 4c). It has been suggested that this may either be due to the relativeincrease in overall endomysial connective tissue within the muscle or a morphologicaladaptation of the receptor to protect itself from the disease process (Ovalle and Dow,1986).HistochemistryThe histochemical profiles of extrafusal fibers from sections of the normal (Fig. 5) andthe mdx (Fig. 6) soleus muscle are illustrated. Acid preincubation (pH 4.6) of the sectionsrevealed two distinct extrafusal fiber types based on their staining properties. At this pH,darkly staining fibers were presumed to be of the type I (slow-oxidative) variety and pale-staining fibers were presumed to be of the type IIA (fast-glycolytic) variety. Very fewintermediate staining fibers were observed. In this study, quantitative measurements of fibertype distribution were not determined, however, it was noted that the mdx soleus appeared26to contain a proportionally high number of type I fibers in comparison to its normalcounterpart.Three intrafusal fiber types were clearly distinguished in polar regions of all spindlesusing the mATPase reaction at an acid (pH 4.6) preincubation (Johnson and Ovalle, 1986). Inequatorial regions, however, all intrafusal fibers of both normal and mdx spindles were poorlystained with this technique (Figs. 7a, 8a). Outside of this region, bag2 fibers were darklystained along their remaining lengths whereas chain fibers exhibited little reaction and werepale staining (Figs. 7b, 8b). The staining pattern of bags fibers was variable. Within thejuxtaequatorial zone, bag1 fibers were observed to be lightly stained while in the more distalpolar regions these fibers characteristically stained darkly (Figs. 7c, 8c). These stainingprofiles were noted in both the normal and dystrophic intrafusal fibers with little variation ineither the intensity or distribution of mATPase staining between corresponding intrafusalfiber types.lmmunofluorescence microscopyFrozen sections of the normal mouse soleus immunolabelled with the dystrophinantibody consistently exhibited a strong reaction at the sarcolemmal domains of theextrafusal fibers (Fig. 9a). These muscle fibers were homogeneous in size and appearanceand were typically polygonal in shape with little intervening endomysial connective tissue.Additionally, a more intense staining reaction for dystrophin was particularly evident at themotor end-plate regions of these fibers (Fig. 9a) which were confirmed as such bycolocalization in double-labelled sections with the 200 kD neurofilament-protein antibody (Fig.9b). Presynaptic terminals of these neuromuscular junctions closely abutted the extrafusalfibers and were typically crescent-shaped in transverse section (Fig. 9b). Intramuscularnerve fascicles and sensory terminals in muscle spindles also displayed a high reactivity withthe neurofilament-protein antibody (Fig. 9b).27Extrafusal fibers of the mdx soleus, in contrast, displayed little or no immunostainingfor dystrophin indicating the specificity of the antibody (Fig. 10a). There were, however, afew extrafusal fibers in the mdx muscle (about 0.01 %) that exhibited either partial orcomplete dystrophin staining at their sarcolemmal membranes. These dystrophin-positiveextrafusal fibres, previously reported by Karpati et al. (1990) in mdx mice, may reflect asomatic back-mutation in the extrafusal fiber nuclei.Concommitant immunolabelling with the neurofilament-protein antibody revealedstrong reactions in intramuscular nerve fascicles, individual axons, and sensory terminals ofmuscle spindles in the mdx soleus (Fig. 10b). Terminal arborizations contacting extrafusalfibers were distributed throughout the muscle and were assumed to be motor-nerveterminals (Fig. 10b). No dystrophin reactivity, however, was seen in the sarcolemmalmembranes in contact with these terminals.In muscle spindles of the normal soleus, the polar regions of intrafusal fibers displayed athin homogeneous peripheral rim of dystrophin immunolabelling similar to that observed at thenon-synaptic sarcolemmal membranes of neighbouring extrafusal fibers (Fig. 11a). Equatorialand juxtaequatorial regions of intrafusal fibers, in contrast, displayed an inconsistentdystrophin staining. Portions of the sarcolemma exhibited either decreased or deficientdystrophin expression, whereas other areas retained a relatively normal labelling intensity(Fig. 11 b). Subsequent labelling of the same sections with the neurofilament-protein antibodyrevealed sensory nerve-terminals of the annulospiral type encircling the perimeters of theintrafusal fibers (Fig. 11c). Those areas deficient in dystrophin (Fig. 11b) consistenlycorresponded to the contact sites of the sensory nerve terminals (Fig. 11c).Serial cross-sections (70 gm intervals) of spindles in the normal soleus were cut todetermine the extent to which these dystrophin-deficient sites are found along the lengths ofthe intrafusal fibers (Figs. 13a-f). Alternate serial-sections, in addition, were treated todetect mATPase (Figs. 13a'-f') making it possible to histochemically differentiate the28intrafusal fibers into bag1 (pale), bag2 (dark staining), and chain (pale) fibers (Ovalle and Smith,1972). These sections were used, subsequently, to reconstruct a muscle spindle along itsentire length (Fig. 13). Deficiencies of dystrophin at the intrafusal fiber sarcolemma wereobserved in all three fiber types in the dilated intracapsular zones (equatorial andjuxtaequatorial regions) of the muscle spindle, occupying a region approximately 280-320 gmin length. The centralized portion of equatorial regions exhibited a relatively high frequencyof dystrophin-staining deficiencies (Figs. 13c, d) whereas more distal juxtaequatorial regionsdisplayed fewer focal deficiencies (Figs. 13b, e). Outside of this region and towards each pole,the sarcolemmal dystrophin expression of the intrafusal fibers was homogeneous andresembled that of the surrounding extrafusal fibers (Figs. 13a, f). No differences indystrophin labelling between nuclear bag and chain fibers or between bags and bag2 fiberswere evident in polar regions.In longitudinal sections of normal intrafusal fibers (Fig. 14a) a more clearly demarcatedstaining pattern for dystrophin was observed. At low magnification, a distinct transition froma continuous sarcolemmal labelling in polar regions to a discontinuous pattern in the sensoryequatorial regions was evident (Fig. 14a). The sarcolemmal membranes of both nuclear bagand nuclear chain fibers were characterized by alternating dystrophin-positive anddystrophin-deficient segments (Figs. 15a, 16a). These dystrophin-negative sarcolemmalsegments measured 5 - 7 gm in length and were found in the highly-nucleated areas of themuscle fibers (Fig. 15b). In double-labelled longitudinal sections, an intensely-labelled networkof nerves arranged in an annulospiral fashion (Fig. 16b) were revealed immunofluorescently bythe 200 kD neurofilament-protein antibody. The areas of sarcolemma subjacent to thesenerve-terminal wrappings (primary sensory endings) corresponded to the dystrophin-deficient segments seen previously. Secondary endings (flower-spray endings) were difficultto identify with certainty due to the limitations of the neurofilament staining method and,subsequently, could not be resolved as distinct terminations on the intrafusal fibers.29When immunolabelled with the dystrophin antibody (Fig. 12b) and counterstained withH & E (Fig. 12a), fluorescence microscopy showed a complete absence of dystrophin labellingin the intrafusal fibers of muscle spindles in the mdx soleus. This finding is in agreement withother immunocytochemical studies of skeletal muscle (Miike et al., 1989; Samitt and Bonilla,1990; Tanaka et al., 1990) where dystrophin staining was briefly noted in muscle spindles. Inthe sections labelled with the nerve-specific antibody, the sensory nerve-terminals in mdxmuscle spindles appeared unaltered morphologically and terminated in a similar fashion to thatobserved in the age-matched normal soleus (Fig. 12c).DenervationFollowing sciatic denervation of normal muscle, obvious morphological alterations in themuscle architecture of the soleus were observed. The extrafusal fibers exhibited aprogressive decrease in size and an increase in the number of centrally located nuclei. Nonormal nerves were observed in either of the two denervation periods and the absence ofneurofilament protein labelling in both nerves and nerve terminals confirmed that the musclewas not reinnervated. Although a decrease in extrafusal fiber size was observed, no changesin either the dystrophin labelling pattern or its intensity was seen in either the 21-day (Fig.17) or 42-day (Fig. 18) denervations. Additionally, elevated levels of dystrophin seen atmotor end-plates in normal muscle were also apparent in denervated muscle (Figs. 17, 18),however, morphological confirmation of the the motor endings was not possible sinceneurofilament protein labelling was absent.In muscle spindles, several morphological characteristics were altered as a result ofdenervation. Capsular thickening in equatorial regions was evident in both groups (Figs. 19a,20a) with a marked increase seen in the 42-day denervated spindles (Fig. 20a). An overallincrease in the cellular components of the spindle capsule and the surrounding endomysial andperimysial connective tissues was also noted (Figs. 19a, 20a). Intrafusal fibers only showed aslight decrease in cross-sectional size and did not appear to parallel the dramatic decrease in30size of the extrafusal fibers. Dystrophin labelling was homogeneously distributed at thesarcolemma in polar regions, whereas equatorial regions displayed a heterogeneousexpression of dystrophin at their sarcolemmal membranes. Deficient zones of dystrophinlabelling were presumed to be sites of degenerating or necrotic sensory nerve terminals (Figs.19b, 20b) and were consistent with those seen in intrafusal fibers of muscle spindles in thenormal soleus (Figs. 12b, 13b-e). This was evident in all intrafusal fiber types examined in thisstudy. In addition, neurofilament protein expression was consistently absent in sensory-terminal regions of the muscle spindles in the denervated soleus at both the 21-day and 42-day denervation periods (Fig. 19c, 20c).31Fig. 1. Transverse frozen section of a normal 32-week-old mouse soleus muscle stainedwith H & E. Regularly-sized extrafusal fibers predominate in the field and appear polygonal inshape with relatively little intervening connective tissue. A muscle spindle is indicated (curvedarrow). x325. Bar = 50 gm.Fig. 2. Comparative view of the soleus from a 32-week-old mdx-dystrophic mousestained with H & E. Large variations in extrafusal fiber size and central nucleation areprominent features. A marked increase in connective tissue can be seen in the interstitium. Amuscle spindle is indicated in the center of the field (curved arrow). x325. Bar = 50 pm.32•^N.r ^I•Is■•.^•^-v411011r^;•P•ir -lb^•^111 _ z.P v^ i  •- ' - 4 &MIA"' . • %t^. ,...^,1^el^t.,,, 1%,e •-•••4!,• .0,•-0 •I33Fig. 3. Higher magnification views of a muscle spindle in the normal soleus stained with H& E. The same muscle spindle is seen sectioned through its equatorial (3a), proximal polar (3b),and distal polar (3c) regions. x820. Bar = 101.1m.Fig. 4. Serial sections of a muscle spindle in the mdx-dystrophic soleus stained with H &E. The spindle has been sectioned through its equatorial (4a), proximal polar (4b), and distalpolar (4c) regions. x820. Bar = 10gm.34_^• .^„ ..ki• .t1 111,• e. - q41•4435Fig. 5. Low magnification view of a portion of the normal soleus stained for myosinATPase at pH 4.6. Note the mosaic-like appearance of the slow-twitch (dark staining) andfast-twitch (pale staining) extrafusal fiber types. x325. Bar = 50gm.Fig. 6. Comparative view of a portion of the mdx-dystrophic soleus stained for myosinATPase at pH 4.6. Groups of slow-twitch (dark staining) extrafusal fibers separated byrelatively low numbers of fast-twitch (pale staining) extrafusal fibers are seen. x325. Bar =50p.m.36•••1•••••••• .11•• ft••1114i. 4re-• giA^41• „ .W\r--"F.^0‘,^,^•37Fig. 7. High magnification views of a muscle spindle in the normal soleus stained formyosin ATPAse at pH 4.6. Sections have been taken from equatorial (7a), proximal polar (7b),and distal polar (7c) regions. Note the change in staining properties along the lengths of thenuclear bag fibers. x820. Bar = 101.1m.Fig. 8. Serial sections of a muscle spindle in the mdx-dystrophic soleus muscle stainedfor myosin ATPase at pH 4.6. The spindle has been sectioned through equatorial (8a), proximalpolar (8b), and distal polar (8c) regions. Note the change in staining properties along thelengths of the nuclear bag fibers. x820. Bar = 10µm.383 9Fig. 9. Pair of fluorescence micrographs of the same transverse section of the normalsoleus double-labelled with anti-dystrophin (a) and anti-neurofilament protein (b). The equatorof a spindle with four intrafusal fibers (curved arrows), a neuromuscular junction on anextrafusal fiber (straight arrows), and a nerve fascicle (asterisks) are indicated. x300. Bar =30 um.406a..AApt‘K.m11441Fig. 10. Micrographs of the same section of the mdx soleus labelled with anti-dystrophin (a), anti-neurofilament protein (b), and post-stained with H & E (c). The equator ofa spindle (curved arrows), a neuromuscular junction on an extrafusal fiber (straight arrows),and a nerve fascicle (asterisks) are indicated. Note the absence of dystrophin in both theextrafusal and the intrafusal fibers (a). x300. Bar = 301.1m.424 3Fig. 11. Polar (a) and equatorial (b) regions of a normal spindle stained with anti-dystrophin show homogeneous staining at the periphery of polar intrafusal fibers (IF) anddystrophin-deficient areas (arrows) in the equator (b). Double-labelling of the equator withanti-neurofilament protein (c) shows sensory nerve terminals. x900. Bar = 10p.m.Fig. 12. Transverse section of an mdx spindle stained with H & E (a), anti-dystrophin(b), and anti-neurofilament protein (c). Intrafusal fibers (IF) show no dystrophin labelling attheir periphery (b), whereas sensory terminals (curved arrows) appear normal (c). x900. Bar= 101.tm.44ire?cr]45Fig. 13. Transverse serial sections (70grn intervals) of a normal spindle immunolabelledfor dystrophin (a-f). Alternate sections are stained for myosin ATPase (a'-f'). Dystrophin inintrafusal fibers is reduced periodically in the equator (arrowheads). The three-dimensionalreconstruction on the right shows polar (Pole) and equatorial (Eq) regions of the musclespindle. x360. Bar = 3011m.46Fig. 14. Low magnification view of a longitudinal section of a muscle spindle in thenormal soleus immunolabelled for dystrophin (a) and post-stained with H & E (b). Note thechange in intrafusal fiber sarcolemmal staining from continuous in the polar region (Pole) todiscontinuous in the equator (Eq). x425. Bar = 30gm.484 9R 1'to,4 .Eq••+4PoleFig. 15. Longitudinal section a a normal nuclear-bag fiber in the equatorial regionviewed with fluorescence (a) and phase-contrast (b) microscopy. Note the periodic pattern ofdystrophin expression along the sarcolemma (thin arrows) of the muscle fiber. Interveningregions (white arrowheads) are deficient in dystrophin. The phase-contrast image revealspunctate elevations (black arrowheads) corresponding to sensory nerve-terminals wheredystrophin is absent. x1,630. Bar = 10gm.5051Fig. 16. Longitudinal views of two intrafusal fibers in the normal soleus double-labelledfor dystrophin (a) and neurofilament protein (b). Dystrophin-deficient regions of thesarcolemma (arrows in a ) are consistent with those areas in contact with the annulospiralsensory endings (arrows in b). The phase contrast image (c) reveals a nuclear bag (nb) and anuclear chain (nc) fiber. x1330. Bar = 10gm.5253Fig. 17. Transverse sections of 21-day (a) and 42-day (b) denervated normal soleusmuscles immunolabelled for dystrophin. Note the persistence of dystrophin in all the extrafusalfibers. Elevated levels of dystrophin are seen at the sarcolemmal membranes of some of theextrafusal fibers (arrows). Small portions of age-matched control soleus muscles (left) areincluded for comparison. x300. Bar = 50um.5455Fig. 18. Equatorial region of a 21-day denervated muscle spindle stained with H & E (a),anti-dystrophin (b), and anti-neurofilament protein (c). Dystrophin is deficient on portions ofthe intrafusal fiber sarcolemma (arrowhead). Note the complete absence of neurofilamentprotein labelling (c) in the sensory regions of the muscle spindle. x625. Bar = 2011m.56o5 7Fig. 19. Equatorial region of a 42-day denervated muscle spindle stained with H & E (a),anti-dystrophin (b), and anti-neurofilament protein (c). Several focal deficiencies in dystrophinlabelling can be seen at the sarcolemmal membranes of the intrafusal fibers (arrowheads).Note the overall increase in paracellular and cellular elements in the connective-tissue capsule(a). x625. Bar = 201.tm.58coft■41059DISCUSSIONThis study has shown for the first time how dystrophin is localized along the lengths ofintrafusal fibers in both normal and mdx-dystrophic skeletal muscle, and these observationshave recently been published (Nahirney and Ovalle, 1993). Although there have been manystudies over the past five years on dystrophin and its localization in skeletal muscle, only a fewhave briefly alluded to the dystrophin staining characteristics of these fibers in musclespindles (Miike at al., 1989; Samitt and Bonilla, 1990; Tanaka et al., 1990; Zhao et al., 1992).Use of the present methodologies has provided a more detailed description of the musclespindle in normal and dystrophic states and, in addition, has revealed important featuresabout dystrophin that directly contribute to our understanding of the functional role thisprotein plays in normal skeletal muscle fibers.HistochemistryThree intrafusal fiber types in muscle spindles of the normal and mdx-dystrophic soleuscould be distinguished on the basis of mATPase staining characteristics. Moreover, regionalvariations in staining intensity were observed, similar to those noted in the study by Johnsonand Ovalle (1986), and also to those seen in rat (Soukup, 1976; Banks et al., 1977; Khan andSoukup, 1979), cat (Banks et al., 1977; Bakker and Richmond, 1981; Kucera, 1981), monkey(Ovalle and Smith, 1972), and human (Kucera and Dorivini-Zis, 1979).At acid preincubation (pH 4.6), equatorial regions of all intrafusal fiber types showedlittle or no histochemical reactivity. In juxtaequatorial regions, the bag1 fibers were acid-labile whereas in distal polar (extracapsular) regions they were acid-stable. Thus, a stainingpattern that gradually changed in a sequential fashion from dark to light in the bags fiber wasobserved as serial sections of a spindle were followed from the polar end toward the equator.60Bag2 fibers, on the other hand, showed a high acid reactivity in juxtaequatorial and polarregions, whereas chain fibers consistently exhibited low acid reactivity throughout theirlengths.These histochemical staining properties were characteristic of both normal anddystrophic intrafusal fibers. In contrast to the histochemical changes seen in extrafusal fibertypes in dystrophy, as noted in this study and in other murine dystrophies (Ovalle et al.,1983; Wirtz et al., 1983), the mATPase reactivity of intrafusal fibers appeared to beunaffected by the disease process. In mdx-dystrophic mice, bag1, bag2, and chain fibers couldbe clearly differentiated based on reactivity in an acid preincubation. Moreover, bag1 fibersshowed the same regional variability as that seen in its normal counterpart.It has been postulated that the variations in mATPase staining of the intrafusal fibersare a consequence of their specific motor innervation (Yellin, 1969; Milburn, 1973). Thisfusimotor influence can be disputed since Zelena and Soukup (1974) have demonstrated thatselectively de-efferented spindles at birth eventually differentiate and mature to obtain theirnormal histoenzymatic characteristics. Additionally, Kucera (1981) has shown no correlationbetween regional ATPase activity and the location, number, or type of motor endingsdetermined by cholinesterase staining. The histochemical profiles of these fibers has,therefore, been suggested to be controlled by different neurotrophic factors than those ofthe extrafusal fibers. Intrafusal fiber integrity may instead be influenced by beta innervationin polar regions, or by sensory innervation in equatorial regions.It has also been shown that the three types of intrafusal fibers exhibit differentisoforms of myosin in capsular (Pierobon-Bormioli et al., 1980; to Kronnie et al., 1981;Rowlerson et al., 1985; Maier et al., 1988) and extracapsular polar (Kucera and Walro,1989) regions. Furthermore, it has been reported that, like mATPase staining, theexpression of myosin heavy chains varies along the lengths of intrafusal fibers, whereas inextrafusal fibers a uniform distribution of these isoforms is observed (Kucera and Walro,611989). Their observations revealed that intracapsular regions of intrafusal fibers containeda relatively high concentration of slow-tonic and neonatal myosin isoforms while extracapsularregions of these fibers displayed a strong reactivity to antibodies specific for slow-twitch andfast-twitch myosin isoforms. A relationship between mATPase staining properties and theexpression of myosin heavy chains has been described by Kucera and Walro (1989) in theextracapsular regions of intrafusal fibers, however, intracapsular regions were not shown tohave such correlations. Similarly, extrafusal fibers have been shown to have a relationshipbetween myosin composition and mATPase reactivity (Staron and Pette, 1986).lmmunolocalization of Dystrophin in Muscle SpindlesThe topographic localization of dystrophin in the intrafusal fibers of muscle spindlesfrom normal and dystrophic mice was determined in this study. The results from this worksupport the original findings of previous workers (Miike et al., 1989; Sammit and Bonilla, 1990;Tanaka et al., 1990; Zhao et al., 1992) that dystrophin is localized under the sarcolemma ofthe intrafusal fibers in normal muscle. However, in this study the consistency of dystrophinexpression was not homogeneous along the lengths of these fibers. By serial inspection oftransverse and longitudinal sections, all three intrafusal fiber types displayed a homogeneoussarcolemmal labelling for dystrophin along the regions of each fiber not in contact withsensory terminals. On the other hand, the sensory equatorial zone exhibited discontinuousimmunolabelling marked by short dystrophin-negative segments of sarcolemma interposed withpositive-staining regions. With the nerve-specific antibody, these dystrophin-deficient zonesin the intrafusal fibers were clearly identified as areas of the sarcolemma immediatelyunderneath sensory nerve-terminals.Electron microscopic studies of mammalian spindles have revealed characteristicsabout the sensory nerve terminals which suggest that a neurosecretory process may be62involved at these sites (Boyd and Smith, 1984). Small membrane-bound vesicles with contentsthought to be neurochemical in nature have been seen in both primary and secondary nerveendings. Additionally, this hypothesis is supported by deafferentation and developmentalstudies which have shown that sensory nerves to spindles are required for differentiation(Zelenã, 1957; Milburn, 1973) and continued maintenance (Tower, 1932; SchrOder, 1974a,b; Schrtider et al., 1978) of the intrafusal fibers. It is possible that sensory nerve innervationplays a trophic role which promotes a modification in the structure of the subsynapticmembrane (i.e. one that inhibits dystrophin expression at this site). Alternatively, either ahomologue of dystrophin (Pons et al., 1991) or a dystrophin-related protein (Ohlendieck etal., 1991 b), both of which have been recently described at neuromuscular junctions ofextrafusal fibers, may, in fact, assume the role of dystrophin at these locations.Maier and Mayne (1993) have recently reported regional differences in theorganization of some of the cytoskeletal components at the equator of chicken intrafusalfibers. In their study, the immunocytochemical expression of the cytoskeletal proteins, alpha-actinin and filamin, was primarily limited to the region subjacent to the myosensory junctionsin the equatorial zones. Diametrically apposing nonsensory regions, on the other hand, didnot exhibit significant labelling for these proteins. However, they did express a sharplydelineated and narrow intrafiber crescent of vinculin colocalized with a crescent of talin. Anasymmetric distribution of beta-1-integrin molecules in the membrane was also noted andcoincided with the expression of vinculin and talin. They have proposed that this selectivearrangement is related to the mooring of the individual components of the myosensoryjunction. Towards the end of the equator and in juxtaequatorial and polar regions, vinculinand the other proteins gradually became distributed equally around the intrafusal fibers, achange that paralleled the decreasing number of contacts made by sensory terminals.63In the present study, dystrophin exhibited a similar variation in distribution at thesarcolemma of normal murine intrafusal fibers. The absence of dystrophin at the myosensoryjunction suggests that similar sarcolemmal attachment modifications occur at these sites.In several ultrastructural studies of muscle spindles, specialized junctions between themembranes of primary sensory terminals and adjacent intrafusal fibers have been described(During and Andres, 1969; Smith and Ovalle, 1972; Kennedy et al., 1975). Their structuralresemblance to fascia adherentes in cardiac muscle (Kennedy et al., 1975) suggests thatthese membrane specializations in spindles may serve as mechanical anchoring sites, therebyinhibiting displacement of the apposed nerve terminal and muscle fiber membranes.In a scanning electron microscopic study (Patten and Ovalle, 1991), the three-dimensional morphology of sensory endings has shown that these focal adhesion points appearas punctate bands that link the closely apposed membranes at irregularly spaced intervals.Kennedy and coworkers (1975, 1980) have proposed that these junctions transmitlongitudinally applied shear forces directly to the nerve cell membrane. This distortion of themembrane results in an increase in Na+ conductance which leads to generation of receptorpotentials (Hunt et al., 1987). These junctions may also provide an alternate method ofsarcolemmal adhesion to underlying myofibrillar structures since it is known that a peripheralrim of subsarcolemmal myofilaments characterizes the equatorial region of intrafusal fibers(Ovalle, 1972). Although dystrophin is thought to be a critical link between the sarcolemmaand the underlying contractile elements of a muscle fiber (Hoffman et al., 1987), its absenceunder sensory nerve-terminals of spindles may be suggestive of a mechanical adaptation ofthe intrafusal fiber for its role as a sensory transducer.Another ultrastructural feature of sensory nerve-terminals on intrafusal fibers is theabsence of an intervening basal (external) lamina in the nerve-muscle cleft (Merrillees, 1960;Landon, 1966; Corvaja et al, 1967; Hennig, 1969). This is in contrast to motor nerve-64endings where an external lamina intervenes between the presynaptic terminal and thesarcolemma of both intrafusal (Landon, 1966) and extrafusal (Sanes and Chiu, 1983) fibers.It has been shown that dystrophin maintains a tight association with integralmembrane glycoproteins of 156 and 50 kD in skeletal muscle fibers (Ohlendieck et al., 1991a)and that dystrophin-associated glycoproteins link dystophin to the extracellular matrix(lbraghimov-Beskrovnaya et al., 1992). The lack of an association of the subsynapticsarcolemma at the sensory nerve-terminal with an external lamina might somehow be relatedto the absence of dystrophin at these sites. If incorporation of the glycoproteins isdependent on the presence of an external lamina, then the synaptic sarcolemma would bevoid of attachment points which may function as 'dystrophin receptors'. The functional roleof the dystrophin-glycoprotein complex is not well established but it has been postulated toeither stabilize the plasma membrane, maintain a non-uniform distribution of a membraneglycoprotein such as a cell receptor or ion channel, or it could be a link between dystrophinand the extracellular matrix (Campbell and Kahl, 1989).Intrafusal fibers in Dystrophic MiceThe complete absence of dystrophin in spindles of the mdx mouse indicates that theintrafusal fibers exhibit the same primary defect in muscular dystrophy as seen in thesurrounding extrafusal muscle fibers. From a physiological perspective, however, the absenceof dystrophin in mdx intrafusal fibers may not be as detrimental as it is for the extrafusalfiber population. Muscle spindles in the dystrophic soleus observed in this study did not exhibitdrastic morphological changes from their normal counterparts, and no abnormally-sized ornecrotic intrafusal fibers were encountered. It is possible that their small diameters,substantially lower tension-outputs, and capsular investment may decrease the chance of65focal sarcolemmal tearing and subsequent fiber necrosis, the process that is thought tooccur primarily in the larger diameter extrafusal fibers (Karpati and Carpenter, 1986).Light and electron microscopic studies of muscle spindles in other murine dystrophieshave revealed similar results. Failure to detect abnormalities in hindlimb spindles of either thedy (Meier, 1969; Yellin, 1974) or the dy 2J (James and Meek, 1979; Ovalle and Dow, 1986;Johnson and Ovalle, 1986) mouse, both autosomal recessive in nature, may be indicative ofthe lack of involvement of the receptors in the disease process. There has been, however,descriptions of atrophy in polar regions of intrafusal fibers and also capsular thickening inequatorial regions of spindles in the 1-year-old dy 2J mouse (Ovalle and Dow, 1986). Thethickening of the capsule in equatorial regions has been postulated to be an adaptiveresponse of the receptor to sequester the delicate sensory regions of the intrafusal fibers(Ovalle and Dow, 1986).Sensory signal-transduction in spindles of the mdx mouse at the level of the sensorynerve/intrafusal fiber junctions is not likely to be affected in this animal model for dystrophyfor two reasons. First, sensory-endings revealed immunofluorescently in double-labelledsections with the nerve-specific antibody appeared to be unaltered in dystrophy; theyterminated in a normal annulospiral fashion similar to that observed in the age-matchedcontrol soleus. Secondly, even though dystrophin is absent in dystrophic intrafusal fibers, it ispossible that this feature has little bearing on sensory transduction since, in normal muscle,dystrophin is also either significantly reduced or absent in the sarcolemma immediatelyunderlying sensory terminals.66DenervationThe speculation that trophic factors emanating from the nervous system play a role indetermining the expression of dystrophin was tested in the denervation experiments ofnormal mature mice. It was predicted that if trophic factors from sensory nerve terminalswere absent, the expression of dystrophin at these sites would increase to a level equivalentto that seen in nonsynaptic regions of the sarcolemma. The results of this experimentrevealed that at both 21-day and 42-day post-denervation time periods, absent or very lowlevels of dystrophin were still persistent in the equatorial region of the intrafusal fibers. Fromthese observations, it was apparent that a neurotrophic influence was not the primary causeof the dystrophin deficiency and that alternate factors were involved.There is the possibility that post-denervation times may not have been long enough fortermination of this neurosecretory process. This, however, is unlikely since it has previouslybeen shown by Barker et al. (1970) that all traces of motor and sensory nerve endings oncat intrafusal fibers disappear 96 hours after muscle nerve-section. In the present study,several morphological changes within the muscle had already occurred which were indicativeof denervation. The atrophy of extrafusal fibers and, to a lesser degree, the intrafusal fibers,along with the absence of neurofilament protein expression in nerves and nerve terminalswere obvious signs of a significant deprivation of nerve influences. Additionally, capsularthickening in equatorial regions and an overall increase in the cellular components of theendomysial and perimysial tissue were evident in both time periods with a significant increaseseen in the 42-day denervated spindles. The changes observed in the overall morphology ofmuscle spindles emphasize the importance of innervation in the maintenance of spindlestructure.The unaltered immunoreactivity of the dystrophin antibody in both the extrafusal andintrafusal fibers of denervated normal muscle indicates that this protein is not affected bythe absence of neural stimulation or neurotrophic factors. Moreover, the extrafusal fibers67continued to show elevated levels of dystrophin at distinct regions of their sarcolemma.These regions closely resembled the intensely-stained motor end-plate regions seen on theextrafusal fibers in the normal soleus and were presumed to be areas of degenerated motorneuromuscular junctions. These observations suggest that innervation does not play a crucialrole in the adult in determining the expression of dystrophin in the subsarcolemmal lattice.CONCLUSIONS AND FUTURE DIRECTIONSThe overall purpose of this study was to investigate more closely the localization ofdystrophin in mouse muscle spindles in both normal and diseased states and, then to use thisdata to help elucidate the possible functions of dystrophin. The results can be summarized asfollows:1. In normal mouse skeletal muscle, dystrophin was found in all regions of thesarcolemma of intrafusal fibers except in those areas in contact with sensorynerve terminals.2. In mdx-dystrophic muscle, the intrafusal fibers were deficient in dystrophinthroughout their lengths, even though sensory innervation patterns appearednormal.3.^Chronic denervation of the normal mouse soleus did not significantly affect theexpression or distribution of dystrophin in the intrafusal fibers of musclespindles.From these results several conclusions can be drawn regarding the function ofdystrophin. First, the sensory innervation to an intrafusal fiber specifically affects thesubsarcolemmal distribution of dystrophin. The exact reason why this occurs remainsunclear, however, evidence from the chronically denervated spindles in this study indicates68that a neuromodulatory role in the form of a hormone or a chemical transmitter is unlikely. Amore plausible explanation for this occurrence would be the unique ultrastructure of thesensory nerve/muscle junction itself. In contrast to motor endings where a basal laminaexists in the nerve-muscle cleft, sensory endings lack an intervening basal lamina. At thesesites, the basal lamina of the intrafusal fiber is continuous with the external surface of thesensory nerve terminal. This feature of the sensory nerve/muscle junction suggests thatdystrophin expression is dependent on the presence of a basal lamina. It is possible thatcomponents of the basal lamina are required for the mechanical linking of dystrophin via theintegral membrane proteins to the extracellular matrix. The functional role for dystrophinwould, therefore, predict one that is mechanical in nature.Secondly, it can also be concluded that the sensory synaptic domains of thesarcolemma in equatorial regions of intrafusal fibers are structurally similar with respect todystrophin expression in both normal and mdx muscle tissue, and, that the presence ofdystrophin in these zones is not required for normal sensory nerve/muscle fiber interaction.This, however, does not eliminate the possibility of dystrophin-like proteins assuming the roleof dystrophin at these sites. Further studies, involving high resolution microscopic techniquescoupled with immunogold labelling may provide a better understanding of the exactlocalization of dystrophin at sensory junctions in muscle spindles. In addition,immunolocalization of other cytoskeletal components, such as cell adhesion proteins anddystrophin-associated and dystrophin-related proteins, would be beneficial for comparing andcontrasting differences that exist between synaptic and nonsynaptic membranes in skeletalmuscle.69LITERATURE CITEDAdal, M.N. 1969 The fine structure of the sensory region of cat muscle spindles.Ultrastruct. Res., 26: 332-354.Anderson, J.E., W.K. Ovalle, and B.H. Bressler^1987^Electron microscopic andautoradiographic characterization of hindlimb muscle regeneration in the mdx mouse.Anat. Rec., 219: 243-257.Anderson, J.E., B.H. Bressler, and W.K. Ovalle 1988 Functional regeneration in the hindlimbskeletal muscle of the mdx mouse. J. Mus. Res. Cell Mot., 9: 499-515.Bakker, G.J., and F.J.R. Richmond 1981 Two types of muscle spindles in cat neck muscles: ahistochemical study of intrafusal fiber composition. J. Neurophysiol., 45: 973-986.Banks, R.W. 1986 Observations on the primary sensory ending of tenuissimus muscle spindlesin the cat. Cell Tissue Res., 246: 309-319.Banks, R.W., D. Barker, and M.J. Stacey 1982 Form and distribution of sensory terminals incat hindlimb muscle spindles. Philos. Trans. R. Soc. Lond. (Biol.), 299: 329-364.Banks, R.W., D.W. Harker, and M.J. Stacey 1977 A study of mammalian intrafusal musclefibres using a combined histochemical and ultrastructural technique. J. Anat., 123: 783-796Barker, D. 1948 The innervation of the muscle spindle. Q. J. Micr. Sc., 89: 143-186.Barker, D. 1974 The morphology of muscle receptors. In: Muscle Receptors. Handbook ofSensory Physiology. C.C. Hunt, ed. Springer-Verlag, Berlin, Vol. 3, Pt. 2, pp. 1-190.Barker, D., and R.W. Banks 1986 The muscle spindle. In: Myology. A.G. Engel and B.O.Banker, eds. McGraw-Hill, New York, Vol. 1, pp. 309-341.Barker, D., and M.C. Ip 1963 A silver method for demonstrating the innervation of mammalianmuscle in teased preparations. J. Physiol., 169: 73-82.Barker, D., M.J. Stacey, and M.N. Adal 1970 Fusimotor innervation in the cat. Philos. Trans.R. Soc. Lond. (Biol.), 258: 315-346.Baumbach, L.L., J.S. Chamberlain, P.A. Ward, N.J. Farwell, C.T. Caskey 1989 Molecular andclinical correlations of deletions leading to Duchenne and Becker muscular dystrophies.Neurology, 39: 465-474.Becker, P.E. 1964 Myopathien. In: Becker, P.E. (ed.) Humangenetik. Ein kurzes Handbuch,vol. 3. Georg Thieme, Stuttgart.Becker, P.E. and F. Kiener 1955 Eine neue X-chromosomale Muskeldystrophie. Archiv furPsychiatrie and Nervenkrankheiten, 193: 427-48.70Bonilla, E., C.E. Samitt, A.F. Miranda, A.P. Hays, G. Salviati, S. DiMauro, L.M. Kunkel, E.P.Hoffman, and L.P. Rowland 1988 Duchenne muscular dystrophy: deficiency ofdystrophin at the muscle cell surface. Cell, 54: 447-452.Boyd, I.A. 1962 The structure and innervation of the nuclear bag muscle fibre system andthe nuclear chain muscle fibre system in mammalian muscle spindles. Phil. Trans. B., 245:81-136.Boyd, I.A. 1981 The muscle spindle controversy. Sci. Prog. Oxf., 67: 205-221.Boyd, I.A., M.H. Gladden, P.N. McWilliam, and J. Ward 1975 "Static" and "dynamic" nuclearbag fibres in isolated cat muscle spindles. J. Physiol., 250: 11-53.Boyd, I.A., and R.S. Smith 1984 The muscle spindle. In: Peripheral Neuropathy. P.J. Dyck, P.K.Thomas, E.H. Lambert, and R. Bunge, eds. W.B. Saunders, Philadelphia, Vol. 1, pp. 171-202.Bressler, B.H., L.G. Jasch, W.K. Ovalle, and C.E. Slonecker 1983 Changes in isometriccontractile properties of fast-twitch and slow-twitch skeletal muscle of C57BL/6Jdy2J/dy2J dystrophic mice during postnatal development. Exp. Neurol., 80: 457-480.Brooke, M.H., G.M. Fenichel, and R.C. Griggs 1989 Duchenne muscular dystrophy: patternsof clinical progression and effects of supportive therapy. Neurology, 39: 475-481.Brzezinski, D.K. von 1961a Untersuchungen zur Histochemie der Muskelspindeln. I.Mitteilung: Topochemie der Polysaccharide. Acta Histochem., 12: 75-79.Brzezinski, D.K. von 1961b Untersuchungen zur Histochemie der Muskelspindeln. II.Mitteilung: Zur Topochemie und Function des Spindelraulmes und der Spindelkapsel. ActaHistochem., 12: 277-288.Bulfield, B., B.G. Siller, P.A. Wight, and K.J. Moore 1984 X chromosome-linked musculardystrophy (mdx) in the mouse. Proc. Natl. Acad. Sci. USA, 81: 1189-1192.Burghes, A.H., C. Logan, X. Hu, B. Belfall, R.G. Worton, and P.N. Ray 1987 A cDNA clone fromthe Duchenne/Becker muscular dystrophy gene. Nature, 328: 434-437.Byers, T.J., L.M. Kunkel, and S.D. Watkins 1991 The subcellular distribution of dystrophin inmouse skeletal, cardiac, and smooth muscle. J. Cell Biol., 115: 411-421.Cajal, S.R. 1888 Terminaciones nerviosas en los husos musculares de la rana. Riv. Trim. Histol.Norm. y Patol., 1.Campbell, K.P., and S.D. Kahl 1989 Association of dystrophin and an integral membraneglycoprotein. Nature (Lond.), 338: 259-262.Carnwath, J.W., and D.M. Shotton 1987 Muscular dystrophy in the mdx mouse:histopathology of the soleus and extensor digitorum longus muscles. J. Neurol. Sci., 80:39-54.Carpenter, S., G. Karpati, E. Zubrzycka-Gaarn, D.E. Bulman, P.N. ray, and R.G. Worton 1990Dystrophin is localized to the plasma membrane of human skeletal muscle fibers byelectron-microscopic cytochemical study. Muscle Nerve, 13: 376-380.71Cazzato, G., and J.N. Walton 1968 The pathology of the muscle spindle: a study of biopsiedmaterial in various muscular and neuromuscular diseases. J. Neurol. Sci., 7: 15-70.Coons. A.H., and M.H. Kaplan 1950 Localization of antigens in tissue cells. II. Improvements ina method for the detection of antigen by means of fluorescent antibody. J. Exp. Med., 91:1-1 3 .Cooper, S., and P.M. Daniel 1963 Muscle spindles in man: their morphology in the lumbricalsand deep muscles of the neck. Brain, 86: 563-586.Cooper, S., and M.H. Gladden 1974 Elastic fibres and reticulin of mammalian muscle spindlesand their functional significance. Q. J. Exp. Physiol., 59: 367-385.Corvaja, N., V. Marinozzi, and 0. Pompeiano 1969 Muscle spindles in the lumbrical muscles ofthe adult cat. Electron microscopic observation and functional considerations. Arch. Ital.Biol. 107: 365-543.Coulton, G.R., J.E. Morgan, T.A. Partridge, and J.C. Sloper 1988 The mdx mouse skeletalmuscle myopathy: 1. A histological, morphometric and biochemical investigation.Neuropath. Appl. Neurobiol., 14: 53-70.Croft, P.B., R.A. Henson, H. Urich, and M. Wilkinson 1965 Sensory neuropathy with bronchialcarcinoma: a study of four cases showing serological abnormalities. Brain, 88: 501-522.Cullen, M.J., J. Walsh, L.V.B. Nicholson, and J.B. Harris 1990 Ultrastructural localization ofdystrophin in human muscle by using gold immunolabelling. Proc. R. Soc. Lond. B., 240:197-210.DeReuck, J. 1974 The pathology of the human muscle spindle: a light microscopic, biometricand histochemical study. Acta Neuropath. (Berl.), 30: 43-55.Dow, P.R., S.L. Shinn, and W.K. Ovalle 1980 Ultrastructural study of a blood-muscle spindlebarrier after sytemic administration of horseradish peroxidase. Am. J. Anat., 157: 375-3 8 8.Duchenne, G.B.A. 1868 Recherches sur la paralysie musculaire pseudohypertrophique ouparalysie myo-sclerosique. Arch. Gen. Med., 11: 5-25; 179-209; 305-321; 421-443;552-588.During, M., and K.H. Andres 1969 Zur feinstruktur der muskelspindel von mammilia. Anat.Anz., 124: 566-573.Ervasti, L.M., K. Ohlendieck, S.D. Kahl, M.G. Gayer, and K.P. Campbell 1990 Deficiency of aglycoprotein component of the dystrophin complex in dystrophic muscle. Nature, 345:315-319.Fukami, Y. 1982 Further morphological and electrophysiological studies on snake musclespindles. J. Neurophysiol., 47: 810-826.Fukami, Y. 1986 Studies of capsule and capsular space of cat muscle spindles. J. Physiol.(Lond.), 376: 281-297.Geiger, B., and D. Ginsberg 1991 The cytoplasmic domain of adherens-type junctions. CellMotil. Cytoskeleton, 20: 1-6.72Gladden, M.H. 1970 A modified pyridine-silver stain for teased preparation of motor andsensory nerve endings in skeletal muscle. Stain Technol., 45: 161-164.Gladden, M.H. 1976 Structural features relative to the function of intrafusal muscle fibers inthe cat. Progr. Brain Res., 44: 51-67.Hammond, R.G. 1987 Protein sequence of DMD gene is related to actin-binding domain ofalpha-actinin. Cell, 51: 1-9.Harker, D.W., L. Jami, Y. Laporte, and J. Petit 1977 Fast conducting skeletofusimotor axonssupplying intrafusal chain fibers in the cat peroneus-tertius muscle. J. Neurophysiol., 40:791-799.Hassell, A.H. 1849 The Microscopic Anatomy of the Human Body. Vol. I. Samuel Highley,London.Heilig, R., C. Lemaire, and J.L. Mandel 1987 A 230 kb cosmid walk in the Duchenne musculardystrophy gene: detection of a conserved sequence and of a possible deletion proneregion. Nucl. Acids Res., 15: 9129-9142.Hennig, G. 1969 Die Nervenendigungen der rattenspindel im elektronenmikroskopischen bild.Z. Zellforsch, 96: 275-294.Hoffman, E.P., R.H. Brown, Jr., and L.M. Kunkel 1987 Dystrophin: the protein product of theDuchenne muscular dystrophy locus. Cell, 51: 919-928.Hoffman, E.P., S.C. Watkins, H.S. Slayter, and L.M. Kunkel 1989 Detection of a specificisoform of alpha-actinin with antisera directed against dystrophin. J. Cell Biol., 108: 503-51 0 .Hsu, S.M., L. Raine, and H. Fanger 1981a A comparative study of peroxidase-antiperoxidasemethod and an avidin-biotin-complex method for studying polypeptide hormones withradioimmunoassay antibodies. Am. J. Clin. Pathol., 74: 32-40.Hsu, S.M., L. Raine, and H. Fanger 1981b The use of avidin-biotin-peroxidase complex (ABC)in immunoperoxidase techniques. A comparison between ABC and unlabelled antibody PAPprocedures. J. Histochem. Cytochem., 29: 577-580.Huard, J., L.P. Fortier, C. Labrecque, G. Dansereau, and J.P. Tremblay 1991 Is dystrophinpresent in the nerve terminal at the neuromuscular junction? An immunohistochemicalstudy of the heterozygote dystrophic (mdx) mouse. Synapse, 7: 135-140.Huard, J., L.P. Fortier, G. Dansereau, C. Labrecque, and J.P. Tremblay 1992 A light andelectron microscopic study of dystrophin localization at the mouse neuromuscularjunction. Synapse, 10: 83-93.Hunt, C.C. 1974 The physiology of muscle receptors. In: Muscle Receptors. Handbook ofSensory Physiology. C.C. Hunt, ed. Springer-Verlag, Berlin, Vol. 3, Pt. 2, pp. 191-234.Hunt, C.C., R.S. Wilkinson, and Y. Fukami 1978 Ionic basis of the receptor potential in primaryendings of mammalian muscle spindles. J. Gen. Physiol., 71: 683-698.73lbraghimov-Beskrovnaya, 0., J.M. Ervasti, C.J. Leveille, C.A. Slaughter, S.W. Sernett, and K.P.Campbell 1992 Primary structure of dystrophin-associated glycoproteins linkingdystrophin to the extracellular matrix. Nature (Lond.), 355: 696-702.Jasch, L.G., B.H. Bressler, W.K. Ovalle, and C.E. Slonecker 1982 Abnormal distribution ofproteins in the soleus and extensor digitorum longus of dystrophic mice. Muscle Nerve, 5:462-470.James, N.T. 1971 The histochemical demonstration of mucopolysaccharide in the lymphspace of muscle spindles. J. Anat., 110: 163.James, N.T., and G.A. Meek 1979 Ultrastructure of muscle spindles in C57BI/6J dy2J/dy2 Jdystrophic mice. Experentia, 35: 108-109.Johnson, M.I., and W.K. Ovalle 1986 A comparative study of muscle spindles in slow and fastneonatal muscles of normal and dystrophic mice. Am. J. Anat., 175: 413-427.Karpati, G., and S. Carpenter 1988 Small-caliber skeletal muscle fibers do not sufferdeleterious consequences of dystrophic gene expression. Am. J. Med. Gen., 25: 653-658.Karpati, G., E. Zubrzycka, S. Carpenter, D.E. Bulman, P.N. Ray, and R.G. Worton 1990 Agerelated conversion of dystrophin negative to positive fiber segments of skeletal but notcardiac muscle fibers in heterozygote mdx mice. J. Neuropathol. Exp. Neurol., 49: 96-105Kennedy, W.R., J. deF. Webster, and K.S. Yoon 1975 Human muscle spindles: fine structure ofthe primary sensory ending. J. Neurocytol., 4: 675-695.Kennedy, W.R., R.E. Poppele, and D.C. Quick 1980 Mammalian muscle spindles. In: ThePhysiology of Peripheral Nerve Diseases. A.J. Sumner, ed. Saunders, Philadelphia, pp. 74-1 3 3Khan, M.A., and T. Soukup 1979 Histoenzymatic study of rat intrafusal muscle fibers.Histochemistry, 62: 179-189.Koenig, M., E.P. Hoffman, C.J. Bertelson, A.P. Monaco, C. Feener, and L.M. Kunkel 1987Complete cloning of the Duchenne muscular dystrophy (DMD) cDNA and preliminarygenomic organization of the DMD gene in normal and affected individuals. Cell, 50: 509-51 7.Koenig, M., A.P. Monaco, and L.M. Kunkel 1988 The complete sequence of dystrophinpredicts a rod-shaped cytoskeletal protein. Cell, 53: 219-228.Koenig, M., A.H. Beggs, and L. Moyer 1989 The molecular basis for Duchenne versus Beckermuscular dystrophy: correlation of severity with type of deletion. Am. J. Hum. Genet., 45:498-506.to Kronnie, G., Y. Donselaar, T. Soukup, and W. van Raamsdonk 1981 Immunohistochemicaldifferences in myosin composition among intrafusal muscle fibers. Histochemistry, 73: 65-74.Kucera, J. 1980 Histochemical study of long nuclear chain fibers in the cat muscle spindle.Anat. Rec., 198: 567-580.74Kucera, J. 1981 Histochemical profiles of cat intrafusal muscle fibers and their motorinnervation. Histochemistry, 73: 397-418.Kucera, J., and K. Dorovini-Zis 1979 Types of human intrafusal muscle fibers. Muscle Nerve,2: 437-451.Kucera, J., and J.M. Walro 1989 Nonuniform expression of myosin heavy chain isoforms alongthe length of cat intrafusal muscle fibers. Histochemistry, 92: 291-299.KUhne, W. 1864 Uber die Endigung der Nerven in den NervenhOgeln der Muskeln. VirchowsArch. Path. Anat., 30: 187-220.Kunkel, L.M., A.P. Monaco, W. Middlesworth, H.D. Ochs, and S.A. Latt 1985 Specific cloning ofDNA fragments absent from the DNA of a male patient with an X chromosome deletion.Proc. Natl. Acad. Sci. (USA), 82: 4778-82.Landon, D.N. 1966 Electron microscopy of muscle spindles. In: Control and Innervation ofSkeletal Muscle. B.L. Andrew, ed. Livingstone, Edinburgh, pp. 96-111.Landon, D.N. 1972 The fine structure of developing muscle spindles in the rat. J. Anat., 111:512-513.Lapresle, J., and M. Milhaud 1964 Pathologie du fuseau neuromusculaire. Rev. Neurol., 110:9 7-1 2 2.Leksell, L. 1945 The action potential and excitatory effects of the small ventral root fibersto skeletal muscle. Acta Physiol. Scand. (Suppl.), 10: 1-84.Lidov, H.G.W., T.J. Byers, S.C. Watkins, and L.M. Kunkel 1990 Localization of dystrophin topostsynaptic regions of central nervous system cortical neurons. Nature, 348: 725-727.Maier, A., B. Gambke, and D. Pette 1988 Immunohistochemical demonstration of embryonicmyosin heavy chains in adult mammalian intrafusal fibers. Histochemistry, 88: 267-271.Maier, A., and R. Zak 1990 Arrangement of cytoskeletal filaments at the equator of chickenintrafusal muscle fibres. Histochemistry, 93: 423-428.Maier, A, and R. Mayne 1993 Regional differences in organization of the extracellular matrixand cytoskeleton at the equator of chicken intrafusal muscle fibers. J. Mus. Res. Cell Mot.,14: 35-46.Matthews, P.B.C. 1962 The differentiation of two types of fusimotor fibre by their effectson the dynamic response of muscle spindle primary endings. Q. J. Exp. Physiol., 47: 324-3 3 3.Matsumara, K., J.M. Ervasti, K. Ohlendieck, S.D. Kahl, and K.P. Campbell 1992 Association ofdystrophin-related protein with dystrophin-associated proteins in mdx mouse muscle.Nature, 360: 588-591.Merrillees, N.C.R. 1960 The fine structure of muscle spindles in the lumbrical muscles of therat. J. Biophys. Biochem. Cytol., 7: 725-742.Meryon, E. 1852 On granular and fatty degeneration of the voluntary muscles. Med. Chir.Trans., 35: 73-84.75Miike, T., M. Miyatake, J. Zhao, K. Yoshioka, and M. Uchino 1989 Immunohistochemicaldystrophin reaction in synaptic regions. Brain Dev., 11: 344-366.Milburn, A. 1973 The early development of muscle spindles in the rat. J. Cell Sci., 12: 175-1 9 5 .Monaco, A.P., R.L. Neve, C. Colletti-Feener, C.J. Bertelson, D.M. Kurnit, andL.M. Kunkel 1986Isolation of candidate cDNAs for portions of the Duchenne muscular dystrophy gene.Nature, 323: 646-650.Nahirney, P.C., and W.K. Ovalle 1992a Immunocytochemical localization of dystrophin andneurofilament protein in muscle spindles of normal and dystrophic mice. Anat. Rec., 232:64A-65A.Nahirney, P.C., and W.K. Ovalle 1992b The expression of dystrophin in intrafusal musclefibers of normal and mdx mice. Proc. Can. Fed. Biol. Soc., 35: 54A.Nahirney, P.C., and W.K. Ovalle 1993 Distribution of dystrophin and neurofilament protein inmuscle spindles of normal and mdx-dystrophic mice: an immunocytochemical study. Anat.Rec., 235: 501-510.Namba, T., T. Nakamura, and D. Grob 1967 Staining for nerve fiber and acetylcholinesteraseactivity in fresh frozen sections. Am. J. Clin. Path., 47: 74-77.North, A.J., B. Galazkiewicz, T.J. Byers, J.R. Glenney, Jr., and J.V. Small 1993 Complementarydistributions of vinculin and dystrophin define two distinct sarcolemma domains in smoothmuscle. J. Cell Biol., 120: 1159-1167.Ohlendieck, K., and K.P. Campbell 1991 Dystrophin-associated proteins are greatly reducedin skeletal muscles from mdx mice. J. Cell Biol., 115: 1685-1694.Ohlendieck, K., J.M. Ervasti, J.B. Snook, and K.P. Campbell 1991a Dystrophin-glycoproteincomplex is highly enriched in isolated skeletal muscle sarcolemma. J. Cell Biol., 112: 135-1 4 8 .Ohlendieck, K., J.M. Ervasti, K. Matsumura, S.D. Kahl, C.J. Leveille, and K.P. Campbell 1991bDystrophin-related protein is localized to neuromuscular junctions of adult skeletal muscle.Neuron, 7: 499-506.Onanoff, M.I. 1890 Sur la nature des faisceux neuromusculaires. C.R. Seanc. Soc. Biol., 42:432-433.Ovalle, W.K. 1972 Fine structure of rat intrafusal muscle fibers. The equatorial region. J.Cell Biol., 52: 382-396.Ovalle, W.K., B.H. Bressler, L.G. Jasch, and C.E. Slonecker 1983 Abnormal distribution of fibertypes in the slow-twitch soleus muscle of the C57BI/dy 2J/dy2 -1 dystrophic mouse duringpostnatal development. Am. J. Anat., 168: 291-304.Ovalle, W.K., and P.R. Dow 1983 Comparative ultrastructure of the inner capsule of themuscle spindle and the tendon organ. Am. J. Anat., 166:343-357.Ovalle, W.K., and P.R. Dow 1986 Alterations in muscle spindle morphology in advanced stagesof murine muscular dystrophy. Anat. Rec., 216: 111-126.76Ovalle, W.K., and P.R. Dow 1988 The capsular sleeve of muscle spindles in mouse and man withspecial reference to the cytoskeleton. In: Mechanoreceptors. Development, Structureand Function. P. Hnik, T. Soukup, R. Vejsada, and J. Zelena, eds. Plenum, New York, pp.255-261.Ovalle, W.K., and R.S. Smith 1972 Histochemical identification of three types of intrafusalmuscle fibers in the cat and monkey based on the myosin ATPase reaction. Can. J. Physiol.Pharm., 50: 195-202.Patel, A.N., V.S. Lalitha, and D.K. Dastur 1968 The spindle in normal and pathological muscle;an assessment of the histological changes. Brain, 91: 737-750.Patten, R.M., W.K. Ovalle 1991 Muscle spindle ultrastructure revealed by conventional andhigh-resolution scanning electron microscopy. Anat. Rec., 230: 183-198.Pierobon Bormioli, S., S. Sartore, M. Vitadello, and S. Schiaffino 1980 Slow myosins invertebrate skeletal muscle. An immunofluorescence study. J. Cell Biol., 85: 672-681.Pons, F., N. Augier, J.O.C. Leger, A. Robert, F.M.S. Tome, M. Fardeau, T Voit, L.V.B. Nicholson,D. Mornet, and J.J. Leger 1991 A homologue of dystrophin is expressed at theneuromuscular junctions of normal individuals and DMD patients, and of normal and mdxmice. FEBS (Fed. Eur. Biochem. Soc.) Lett., 282: 161-165.Roden, R.L., S.P. Donahue, G.A. Schwartz, J.G. Wood, and A.W. English 1991 200 kDneurofilament protein and synapse elimination in the rat soleus muscle. Synapse, 9: 239-243 .Rowlerson, A., L. Gorza, and S. Schiaffino 1985 Immunohistochemical identification of spindlefibre types in mammalian muscle using type-specific antibodies to isoforms of myosin. In:The Muscle Spindle. I.A. Boyd and M.H. Gladden, eds. Macmillan, London, pp. 29-34.Ruffini, A. 1898 On the minute anatomy of the neuromuscular spindles of the cat, and ontheir physiological significance. J. Physiol. (Lond.), 23: 190-208.Samitt, C.E., and E. Bonilla 1990 lmmunocytochemical study of dystrophin at themyotendinous junction. Muscle Nerve, 13: 493-500.Sanes, J.R., and A.Y. Chiu 1983 The basal lamina of the neuromuscular junction. Cold SpringHarbor Symp. Quant. Biol., 48: 667-678.Sartore, S., L. Gorza, and S. Schiaffino 1982 Fetal myosin heavy chains in regeneratingmuscle. Nature, 298: 294-296.SchrOder, J.M. 1974a The fine structure of de- and reinnervated muscle spindles. I. Theincrease, atrophy, and hypertrophy of intrafusal muscle fibers. Acta Neuropathol. (Bed.),30: 109-128.SchrOder, J.M. 1974b The fine structure of de- and reinnervated muscle spindles. II.Regenerated sensory and motor nerve terminals. Acta Neuropathol. (Berl.), 30: 129-144 .SchrOder, J.M., P.T. Kemme, and L. Scholz 1979 The fine structure of denervated andreinnervated muscle spindles: Morphometric study of intrafusal muscle fibers. ActaNeuropathol. (Berl.), 46: 95-106.77Sherrington, C.S. 1894 On the anatomical constitution of nerves of skeletal muscles; withremarks on recurrent fibres in the ventral spinal nerve-root. J. Physiol. (Lond.), 17: 211-2 5 8.Shimuzu, T., K. Matsumura, Y. Sunada, and T. Mannen 1989 Dense immunostainings on bothneuromuscular and myotendon junctions with an anti-dystrophin monoclonal antibody.Biomed. Res., 10: 405-409.Sicinski, P., Y. Geng, A.S. Ryder-Cook, E.A. Barnard, M.G. Darlison, and P.J. Barnard 1989 Themolecular basis of muscular dystrophy in the mdx mouse: a point mutation. Science, 244:1 5 78-1 5 8 O.Smith, R.S., and W.K. Ovalle 1972 The structure and function of intrafusal muscle fibers. In:Progress in Muscle Biology. R.G. Cassens, ed. Marcel Dekker, New York, Vol. 1, pp. 147-2 2 7.Soukup, T. 1976 Intrafusal fiber types in rat limb muscle spindles. Morphological andhistochemical characteristics. Histochemistry, 47: 43-57.Squarzoni, S., P. Sabatelli, M.G. Maltarello, A. Cataldi, R. DiPrimio, and N.M. Maraldi 1992Localization of dystrophin COOH-terminal domain by the fracture-label technique. J. CellBiol., 118: 1401-1410.Staron, R.S., and D. Pette 1986 Correlation between myofibrillar ATPase activity and myosinheavy chain composition in rabbit muscle fibers. Histochemistry, 86: 19-23.Sugita, H., K. Arahata, T. Ishiguro, Y. Suhara, T. Tsukahara, S. Ishiura, C.H. Eguchi, I. Nonaka,and E. Ozawa 1988 Negative immunostaining of Duchenne muscle dystrophy (DMD) andmdx muscle surface membrane with antibody against synthetic peptide fragmentpredicted from DMD cDNA. Proc. Jpn. Acad., 69: 37-39.Swash, M., and K.P. Fox 1974 The pathology of the muscle spindle; effect of denervation. J.Neurol. Sci., 22: 1-24.Swash, M., and K.P. Fox 1976 The pathology of the muscle spindle in Duchenne musculardystrophy. J. Neurol. Sci., 29: 17-32.Tanaka, H., K. Ikeya, and E. Ozawa 1990 Difference in the expression pattern of dystrophinon the surface membrane between the skeletal and cardiac muscles of mdx carrier mice.Histochemistry, 93: 447-452.Tinsley, J.M., D.J. Blake, A. Roche, U. Fairbrother, J. Riss, B.C. Byth, A.E. Knight, J. Kendrick-Jones, G.K. Suthers, D.R. Love, Y.H. Edwards, and K.E. Davies 1992 Primary structure ofdystrophin-related protein. Nature, 360: 591-593.Torres, L.B.F. and L.W. Duchen 1987 The mutant mdx: inherited myopathy in the mouse.Morphological studies of nerves, muscles and endplates. Brain, 110: 269-299.Tower, S. 1932 Atrophy and degeneration in the muscle spindle. Brain, 55: 77-90.Tower, S. 1939 The reaction of muscle to denervation. Physiol. Rev., 19: 1-48.Vandesande, G.^1983^Immunohistochemical double staining techniques. In:Immunohistochemistry. A.C. Cuello, ed. Wiley & Sons, Chichester, pp. 257-272.78Watkins, S.C., E.P. Hoffman, H.S. Slayter, and L.M. Kunkel 1988 Immunoelectron microscopiclocalization of dystrophin in myofibres. Nature (Lond.), 333: 863-866.Wirtz, P., H.M. Loermans, P.G. Peer, and A.G. Reintjes 1983 Postnatal growth anddifferentiation of muscle fibers in the mouse: II. A histochemical and morphometricalinvestigation of dystrophic muscle. J. Anat., 137: 127-142.Worton, R.G. 1992 Duchenne muscular dystrophy: gene and gene product; mechanism ofmutation in the gene. J. lnher. Metab. Dis., 15: 539-550.Yellin, H. 1969 A histochemical study of muscle spindles and their relationship to extrafusalfiber types in the rat. Am. J. Anat., 125: 31-46.Zelend, J. 1957 The morphogenetic influence of innervation on the ontogenetic developmentof muscle spindles. J. Embryol. Exp. Morphol., 5: 183-292.Zhao, J., K Yoshioka, M. Miyatake, and T. Miike 1992 Dystrophin and a dystrophin-relatedprotein in intrafusal muscle fibers, and neuromuscular and myotendinous junctions. ActaNeuropathol., 84: 141-146.Zubrzycka-Gaarn, E.E., D.E. Bulman, G. Karpati, A.H.M. Burghes, B. Belfall, H.J. Klamut, J.Talbot, R.S. Hodges, P.N. Ray, and R.G. Worton 1988 The Duchenne muscular dystrophygene product is localized in sarcolemma of human skeletal muscle. Nature (Lond.), 333:466-496.79

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