1

Differentiation of Axonal Membrane Systems, the Axolemma, and the Axoplasmic Matrix

Mark H. Ellisman, James D. Lindsey, Clayton Wiley-Livingston, and S. Rock Levinson

In this chapter we will discuss the interactions between axonal membrane systems and the "microtrabecular lattice" (MTL). In order to do this a brief review of some relatively new but fundamental observations on the membrane systems within myelinated is needed.

I. MEMBRANE SYSTEMS OF THE

The coexistance of several different intra-axonal membrane systems is a concept which has only recently gained general acceptance. Most likely this is due to the complexity and intimacy of these various systems. The first system to be described in depth was the axo• plasmic reticulum (Droz et al., 1975; Tsukita and Ishikawa, 1976). This reticulum consists of an anastomotic network of fine tubules which course down the axon (diagramed in Fig. 1). Although well known individually for some time, multivesicular bodies have only recently been grouped with dense lumened and clear lumened cisternae to make up the second major intra-axonal membrane system (Weldon, 1975; Bunge, 1977; Broadwell and Brightman, 1979). This second system characteristically appears able to sequester tracer molecules such as thorium dioxide or horseradish peroxidase from the extracellular environment. A third intra-axonal membrane system has been described but has not yet received the level of general acceptance of the first two. This last group is best seen as distinct from the axoplasmic

Mark H. Ellisman and James D. Lindsey • Department of Neurosciences, University of California, San Diego, School of Medicine, La Jolla, California 92093. Clayton Wiley-Livingston • Department of Pa• thology, University of California, San Diego, School of Medicine, La Jolla, California 92093. S. Rock Levinson. Department of Physiology, University of Colorado Health Sciences Center School of Medicine, Denver, Colorado 80262.

D. C. Chang et al. (eds.), Structure and Function in Excitable Cells 3 © Plenum Press, New York 1983 4 Mark H. Ellisman et al.

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Figures 1 and 3. An artist's representation of two of the membrane systems of the axon drawn approximately to scale in the same axoplasmic space. Fig. 1. The axoplasmic reticulum, an interconnected membrane system. Fig. 3. Orthograde transport compartments or vectors, discrete vesicular and vesiculotubular entities of the axon. Figure 2. Diagram illustrating the design of the cold block system. The cylinder represents axons ofthe saphenous while the squares represent cold and warm aluminum blocks. Figure 4. Diagram illustrating the design of double cold block experiments. The cylinder represents the axons within the saphenous nerve while the squares represent the selectively warmed and chilled blocks of aluminum applied to the skin just above the nerve. Beneath the two 4°C blocks, fast axonal transport is arrested whereas under the 3rC blocks it continues in a normal fashion. After 3-4 hr, transported material moving faster than 25 mm/day in either direction will have cleared from the region under the central warm block and no new material will have entered. Axonal Membranes and the Cytoskeleton 5 reticulum in thick sections and primarily contains clear lumened elongated cisternae and vesicles. The functional physiology of these various membrane systems remains obscure. It is now generally accepted that rapidly transported macromolecules move in association with membranous structures of some sort. This concept has recently received further support by the finding that all rapidly transported macromolecules pass through the Golgi apparatus in the cell body enroute to destinations along the axolemma or beyond (Hammerschlag et at., 1982). At least one of these macromolecules, the Na + + K + ATPase is known to be rapidly transported (Baitinger and Willard, 1981) and is interesting in the context of this volume since it is known to be involved in contributing to properties of excitable membranes. The axoplasmic reticulum first became suspect as a possible vector for the fast transport of such macromolecules because of its shape and extent (for reviews see Schwartz, 1979; Grafstein and Forman, 1980). Newly obtained evidence, however, is decreasing the attractiveness of this notion. This new evidence comes from experiments designed to block fast transport focally in a minimally disruptive manner. In a recent electron microscopic study, Tsukita and Ishikawa (1980) showed that blocking axoplasmic transport by gluing a cooled aluminum block over mouse saphenous nerve (Fig. 2), results in an accumulation of membranous material moving in the orthograde direction on the proximal side of the cold zone, while retrogradely moving components on this same proximal side continue to move centrally away from this site. Conversely, retrogradely moving materials distal to the chilled zone accumulate on the distal side of the cold block. With this single system it is possible to examine the morphology of orthograde and retrograde compartments following a suitable period oftime for accumulation. A study using similar logic was recently conducted by Smith (1980). Smith's paradigm makes use of a mechanical block on single teased fibers that are observed at both light and electron microscopic levels. We have been conducting similar in vivo experiments in our own laboratory, to examine the mobility of the axoplasmic reticulum and also to determine the relationship of motile elements to the cytoskeleton (Ellisman and Lindsey, 1981, 1983). Micrographs taken of tissue from the proximal side of the blockade, in all studies, demonstrate an accumulation of vesicular or vesiculotubular structures (Tsukita and Ishikawa, 1980; Smith, 1980; Ellisman and Lindsey, 1981, 1983). These types of discrete membrane bounded compartments accumulate where the logic of these experiments would predict the orthogradely moving components to selectively accumulate. Figure 3 depicts the form of these orthograde transport compartments or "vectors" (as we will refer to them) summarized from electron micrographs. The form of these compartments does not resemble that of the axoplasmic reticulum (depicted in Fig. 1), and we do not find axoplasmic reticulum accu• mulating against either distal or proximal sides of the cold block. Due to these two incon• sistencies, we became curious about the relationship of the axoplasmic reticulum to the motile vectors and the actual mobility of the former. A double cold block apparatus was designed (Fig. 4) and used to examine the mobility of the axoplasmic reticulum. This apparatus maintains a small length of nerve at 37°C between two areas that are cold blocked. After 3-4 hr, any axoplasmic elements moving faster than 25 mmJday should have moved to either the proximal or distal edge of the warm

Fig u re 5. In the middle of the warm region between the two cold blocks, fast transported elements have presumably cleared from the . Here in these stereopair micrographs of a I II-m thick section, viewed with the high voltage electron microscope (HVEM), the only type membranous elements remaining in the axoplasm are those morphologically and histochemically resembling the axoplasmic reticulum (the 3D effect is best when viewed with the aid of a 2 x stereoviewer). x 19,000, 100 tilt. 6 Mark H. Ellisman et at.

Figure 6. Axoplasm adjacent to proximal side of a cold blocked region illustrating the axoplasmic reticulum elements (arrows) that are clearly separate from discrete orthograde vector type elements (arrowheads). Continuities between these membrane systems were not observed in the large number of micrographs examined. x 73,000. Axonal Membranes and the Cytoskeleton 7 zone. In the center of the wann zone all rapidly moving organelles should be absent leaving components which would not have been subject to rapid transport (Fig. 5). Using this procedure, we found that numerous clearly defined elements of the axoplasmic reticulum remain in such central regions while discrete elements virtually disappear without apparent fusions (Fig. 6). These observations support the view that the axoplasmic reticulum is a distinct membrane system, separate from the discrete motile membranous compartments (vectors) which carry rapidly transported macromolecules. The functional significance of this axoplasmic reticulum is not established. Whether a complicated system of membranous tubules within the axon, such as this, would contribute significantly to the electrical properties of the axon is unknown. Serial section analysis of guinea pig ear (Reiter, 1966) and tracer experiments using cultured neurites have fueled speculation that this system has occasional continuity with the axolemma, not unlike the transverse tubule system of muscle (Bunge, 1977; Weldon, 1975). The axoplasmic reticulum may have some properties not unlike the sarcoplasmic reticulum of muscle in that it appears capable of sequestering calcium (Duce and Keen, 1978; Henkart et al., 1978). Thus, this membrane system may be important in regulating the ionic milieu of the axon as well as contribute capacitance to its static electrical properties.

II. THE MICRO TRABECULAR LA TTICE OF THE AXON AND ITS INTERACTIONS WITH MEMBRANOUS COMPONENTS OF THE AXON

Now that the three different membrane systems in the axon have been illustrated we will describe how the entire axonal cytoskeleton including the MTL specializes upon inter• action with the discrete membranous vectors of the axoplasm. and microtubules are crosslinked in a periodic manner by MTL cross• linking components in both chemically fixed and rapidly frozen and etched preparations (Ellisman and Porter, 1980; Ishikawa and Tsukita, 1982). In our high voltage electron microscopic (HVEM) studies we found that discrete cisternae often appeared asymmetrically connected by these crosslinkages to microtubules or neurofilaments as may be seen in the example of a small vesicle in Fig. 7. Such asymmetry was commonly observed, with one end and the sides of the vector connected to fibrous components and the other not. The nonconnected end exhibiting an absence of crosslinkages was often associated with a void in the axoplasmic space without neurofilaments, microtubules, or crossbridges. The im• mediately obvious question here is: does this asymmetry systematically reflect the direction of motion of a vector? More specifically, one might ask, are the voids systematically on the leading or trailing ends of vectors? By using the single cold block method (illustrated in Fig. 2) to focally stop transport, we were able to look at vectors fixed while moving in a predictable direction. This was accomplished by looking at sections, 4-6 mm proximal to the area of vector accumulation. Here orthograde vectors would be moving toward the dam while retrograde vectors will

Fig u re 7. High magnification HVEM stereopair micrographs of peripheral nerve axoplasm. Many small vesicular cisternae (arrow) are larger in diameter than the average distance between adjacent neurofilaments and microtubules. The crossbridging connections are often disconnected on one end of such cisternae. x 68,000. Figure 8. An orthograde vector predicted to be moving from left to right. Crossbridges are seen on the presumptive leading end (arrows) and a clear area behind. x 75,000. - -- -

Figure 9. An artist's representation of the asymmetry found for orthograde and retrograde vectors. The cell body lies towards the left in each diagram. (a) Orthograde vectors are crosslinked at leading ends and disconnected on trailing ends. (b) Retrograde vectors are also crosslinked to the microtubules and neurofilaments on their leading ends and disconnected on their trailing ends. Figu re 10. HVEM stereopair micrographs of a longitudinal thick section through a myelinated peripheral nerve axon. Linkages to the plasma membrane (solid arrows) and discrete membranous cisternae of axonal smooth endoplasmic reticulum (SER) (open arrows) are visible. x 45,000. Axonal Membranes and the Cytoskeleton 9 have had sufficient opportunity to clear the region between the analyzed zone and the cold block as well as the analyzed region itself. Likewise a converse argument can be made for the presence of moving retrograde vectors and absence of orthograde vectors in a zone 4-6 mm distal to the distal edge of the cold block. At ligature lesions, evidence for a delayed turnaround of orthogradely moving material has been found (Bisby, 1977). Whether such a turnaround occurs at a cold block is still unknown. Suggestive evidence against this possibility arises from the dramatic depletion of multivesicular and multilamellar bodies which are known retrograde vectors (Tsukita and Ishikawa, 1980; LaVail et al., 1980; Ellisman and Lindsey, 1983), from the proximal analyzed region. Although further work is still needed, it is thus likely that the discrete compartments within the proximal zone analyzed are predominantly orthograde vectors. By maintaining the proximodistal orientation of the axon throughout the processing and to the electron micrographs, we were able to determine that the asymmetry of crossbridges, the voids, and the organization of microtubules and neurofilaments, all related to both orthograde and retrograde vector direction. We found that there is a predominance of cross• bridges on the leading ends of vectors and an absence of crossbridges (with significant voids creating enlarged spaces between microtubules or neurofilaments) on the trailing ends of both types of vectors. We suspect that this asymmetry has something to do with the actual movement of vectors during rapid axonal transport. Figure 8 is an orthograde vector pre• sumably moving from left to right down this axon. The drawings presented in Figs. 9a and b summarize our observations on the asymmetry of crossbridges on both the orthograde and retrograde vectors.

III. CYTOSKELETAL SPECIALIZATION OF THE AXOLEMMA

A. Internodal Zone In addition to MTL crossbridges attaching to transport vectors, one also finds cross• bridges between the microtubules and neurofilaments (Ellisman and Porter, 1980) and, most importantly for what we will consider next, from microtubules and neurofilaments to the axolemma. Figure 10 is an example of an axolemmal-cytoskeletal connection in an internodal region of a myelinated axon. Here, the subaxolemmal (cortical) specialization of the plasma membrane is relatively simple. A relatively thin layer of cortical material from which wispy elements project deeper into the axoplasm, connect to "core cytoskeletal components," the so-called fibrous proteins, microtubules, and neurofilaments.

B. Nodal Zone A more complex cortical specialization or axoplasmic matrix specialization is found where the axolemma is highly specialized at the or in initial segment regions (Peters, 1966). Figure 11 is an HVEM stereopair of a thick section illustrating what is generally referred to as the subaxolemmal densification at the node of Ranvier. Upon close examination of this micrograph, however, one notices that this densification is composed

Figure 11. HVEM stereo pair micrographs of a central node of Ranvier. Note that the special• ization of the axolemma at the nodal membrane contains a band of fine filaments seen here in cross section (arrows). Also visible in this micrograph is an example of the axoplasmic reticulum (arrowheads). x 13,000. 10 Mark H. Ellisman et al.

Figure 12. An HVEM stereopair capturing virfually an entire node within the thick section. The nodal membrane is seen enjace (arrowhead) and here an array of fine filaments like those seen in Fig. 11 are visible (small arrows). x 30,000. Axonal Membranes and the Cytoskeleton 11 of a series of cross-sectioned filaments, here appearing as a series of 40 A diameter dots. In fortunately oriented ultrathin sections the periodicity of the "dense layer" beneath the nodal membrane is also noticeable. The organization of the cortical filaments of the subnodal zone is best appreciated in high voltage electron micrographs of thick sections in which a portion of the nodal membrane is tangentially included within the section (Fig. 12). The nodal membrane thus viewed en face appears as a translucency with the filamentous components of the cortical densification just beneath the axolemma. From this perspective the fine filaments are seen to be arranged in parallel, forming a wide band beneath the nodal membrane. This precise arrangement of very fine filaments appears to be restricted to the nodal zone of the axolemma. Noteworthy and very evident in Fig. 11 is a system of subsurface cisternae characteristic of the paranodal zone of axoplasm (Ellisman, 1977). Several other types of membrane organelles are commonly subjacent to the nodal and paranodal regions of the axolemma. For instance, notice the multivesicular body in Fig. 12 just below the membrane of the nodal zone. Multivesicular bodies are observed frequently at nodes of Ranvier. These organelles are part of the neuronal lysosomal system and we suspect they are involved in the turnover of nodal membrane proteins by endocytocis followed by their retrograde transport to the cell body. Evidence for endo- and/or exocytotic activity at the node of Ranvier is especially evident during development (Wiley-Livingston and Ellisman, 1980). For example, Fig. 13 is a micrograph of a node of Ranvier from a 9-day-old rat exhibiting a coated vesicle and many smooth vesicles intimately associated with the nodal axolemma. Evidence for continuity between axoplasmic vesicular components and the axolemma in the nodal zone has been obtained in freeze-fracture replicas (Fig. 14). Here, dimples may represent either endo- or exocytosis. Occasionally, in such replicas we find a fracture through a region of the nodal zone, where such depressions occur, that also fracture into the underlying axoplasm (Fig. 15). From these micrographs it is evident that dimples in the axolemma are directly connected to vesicular elements within the axoplasm. Thus, in the nodal area there are several types of cisternae and there is evidence for membrane exchange between the axolemma and membrane bounded components of the axoplasm. The filaments of the subnodal densification are illustrated (Fig. 16). Also illustrated in this diagram are some of the specializations characteristic of the paranodal zone including the subaxolemmal cisternae and variations in particle size and density exposed in freeze-fracture replicas (Ellisman, 1976, 1979; Rosenbluth, 1976; Wiley and Ellisman, 1980).

C. Paranodal Zone In the paranodal zone where the glial loops of the myelinating cells attach, connections between the core cytoskeletal structure of the axoplasm and small tufts of wispy material

Figure 13. A spinal root from a 7-day-old rat. During the early stages of myelination, evidence of endocytotic and exocytotic events at the nodal membrane includes invaginations and evaginations of the axolemma, the appearance of dense core and coated vesicles (arrow), as well as the presence of multilarnmelar and multivesicular bodies. x 24,000. Figure 14. An electron micrograph of a freeze-fracture replica from rat peripheral nerve. Evidence of endo- or exocytotic events are seen here as depressions in the nodal membrane (arrows). x 11,000. Figure 15. A stereopair of freeze-fractured node of Ranvier, nodal membrane, partially "crossfractured" into axoplasm. Many vesicles (arrows) in the axoplasm show apparent continuity with the nodal membrane axonal P face (AX PF), suggesting that active membrane, and/or membrane protein turnover occurs here. x 21 ,000. 12 Mark H. Ellisman et al.

16

Figure 16. This diagram illustrates some of the structural features of the node of Ranvier observed either in sections (transmission electron microscopy (TEM) and HVEM), on left HVEM or in freeze-fracture replicas, on right (see text and Figs. 11, 12, 21-25). Figure 17. Cytoskeletal connections at the paranodal zone viewed here in stereo. Filamentous connections from the neurofilaments to the glial-axonal junction (G-A J) are seen in the axon (arrows) while cross-sectioned 70 A microfilaments are visible within each glial loop (small arrows). x 50,000. Axonal Membranes and the Cytoskeleton 13 of the axolemma are also seen. Figure 17 is a stereopair of a thin section from the paranodal region illustrating crossbridges between axoplasmic neurofilaments and the wispy material of the axolemma immediately subjacent to a paranodal glial loop. Neurofilaments are found here in parallel arrays, very closely applied to the axolemma, and with a regularity of spacing. These features are detailed in Figs. 17-19, which represent sections through this region of different orientation. Perhaps most notable in Fig. 17 are the cross-sectioned arrays of 70 A micro filaments distributed just above the glial membrane in the paranodal glial loop . Several of these microfilaments appear to be arranged in the cortex of each glial loop adjacent to the glial-axonal junction (GAl) as it encircles the axon. The 70 A glial filaments can be seen best when this area of glial and axonal membrane apposition is captured tangentially or en face in thin sections (Figs. 18 and 19). Figures 18 and 19 are serial sections oriented such that they capture the cytoplasm of the series of glial tenninalloops just before (Fig. 18) and just after (Fig. 19) they are applied to the axon at the paranodal junction. The glial filaments seen in cross section in Fig. 17 are longitudinally exposed in Figs. 18 and 19, as part of a helical circle within the glial loop around the axon. Deeper into the axon, Fig. 19 includes the paranodal junction and some of the axoplasm. Neurofilaments just below the axolemma in the axoplasm are clearly orthogonally arranged with respect to the 70 A filaments of the glial loops. Also notable in this series of figures (Figs. 17-19) are more subaxolemmal cisternae of the paranode, probably a specialization of the axoplasmic reticulum (mentioned above, see also Figs. 11 and 12). The entire area and the superpositioning of the glial and axonal filaments may be appreciated in stereopairs of thick sections viewed with the aid of HVEM (Fig. 20). In this thick section one views the axon from a point outside the paranodal zone. Here, one can pick out filaments just above the glial membrane before this membrane is involved in fonning the glial axonal junction of the paranode. Slender tubular elements (subsurface cisternae of the ) appear in several of these paranodalloops. This is most often seen in young nodes and to best advantage with freeze-fracture as illustrated in Fig. 21. The viewing perspective of Fig. 21 is from the axoplasm looking outward toward parts of the helical glial loops enshrouding the paranodal zone of the axolemma. The tubular cisternae of the glial loop seen in cross section in Fig. 17 are clearly visible in this replica. Thus, the cytoskeleton of the axolemma and the cytoskeleton of the glial loop as well as subsurface cisternal systems of both are highly specialized in the paranodal zone.

IV. SPECIALIZA TION OF THE MAMMALIAN PARA NODE AND NODE EXPOSED BY FREEZE-FRACTURE

A. Paranodal Zone The preceding two sections consider the cortical specialization of the nodal and par• anodal zones including the characteristic linkages of these regions with the core cytoskeleton. While stereo HVEM techniques reveal the three-dimensional aspects of the cortical zone

Figure 18. The 70 A microfilaments of the glial loops seen in cross section in Fig. 17 are visible in longitudinal orientation in Fig. 18 which is an en face section through the G-A J. The filaments contained in the glial loop are notable in the cortical cytoplasm where the plasma membrane is involved in forming the G-A J (arrows). x 22,000. Figure 19. This is the next serial section after that shown in Fig. 18, here cutting deeper into the axon, the axoplasmic contribution to the junctional complex is included. Microfilaments of the glial loop are still visible (arrows) and the parallel arrays of neurofilaments in the cortical axoplasm (open arrows) are orthogonally oriented to the glial microfilaments. Also note the fragment of the axoplasmic reticulum (arrowhead) very often observed in this cortical axoplasm of the paranodal zone involved in the junctional complex. x 25,000. 14 Mark H. EI/isman et al.

Figure 20. Many of the structural features detailed in the serial sections (Figs. 18 and 19) are visible when thick sections of this paranodal region are viewed with the HVEM . Here, view is from the glial side of the junction into the axoplasm. Orthogonal superpositioning of glial microfilaments and axonal neurofilaments may be observed in the area inscribed by the box. Also notable in such images are elements of a tubular cisternal system contained within the glial loops (arrows). x 26,000.

Figure 21. Freeze-fracture of paranode exposing axonal E face (EF) and glial P face (PF). In this micrograph the tubular cisternae of the glial loops noted above in the stereo pair (Fig. 20), are cross-fractured (arrows). x41,OOO. Axonal Membranes and the Cytoskeleton 15 and subsurface cisternae of this region to best advantage, freeze-fracture replicas reveal the membrane molecular architecture of these zones most effectively. Where the paranodal junction is formed, the membranes of both glia and axon exposed in freeze-fracture replicas contain many particles and associated pits in highly ordered arrays (Wiley and Ellisman, 1980). The protoplasmic fracture face (PF face) of the paranodal axolemma contains rows of slightly elongated particles forming "dimers." We have examined the structural relation• ship between intramembranous specializations contributed by the myelinating cell (glial PF face and EF face) and elements contributed by the axon (axonal PF face and EF face) in forming the glia-axonal junctional complex with high resolution freeze-fracture techniques. Some of the results of our analysis are shown in Figs. 22-25 and are summarized in a composite diagram (Fig. 26). The junction appears as a highly ordered aggregate of both axonal and glial membrane components. We have speculated that some of these components may be involved in exchanges of ions and/or metabolites between axon and myelinating cell.

B. Nodal Zone The PF face and EF face of the nodal zone exposed in freeze-fracture replicas contains particles of a much more heterogenous size distribution (Ellisman, 1976, 1979). There are only about 2000 particles/,.,..m2 exposed in both leaflets of the nodal zone in aldehyde fixed preparations. The types of particles and associated pits exposed in these membranes are illustrated in Figs. 22-25 and detailed in the figure legends. They range in size from 75-200 A. The most frequent sized particle is 90-100 A on PF faces while both the PF face and EF face reveal a high density of elongated "rod-shaped" particles. The relationship between Na + + K + ATPase of the node or the K + or Na + channels and specific particles is presently unknown. It is known, however, that the Na + + K + ATPase is a 90-100 A freeze-fracture particle (Deguchi et al., 1977). In order to define the particle candidates one would like to know whether the distribution of the excitable membrane pumps and channels is limited to the nodal zone which includes the paranode, and/or extends over the entire axolemma (under the sheath as well).

V. IMMUNOCYTOCHEMICAL EVIDENCE FOR THE NODAL MOSAIC

A. Electric Organ The distribution of at least two of these important components of the excitable mem• branes within the axolemma, Na + channels and the Na + + K + ATPase, have been ex• amined with immunoelectron microscopic techniques (Wood et al., 1977; Ellisman and Levinson, 1982; Ellisman et al., 1982b). To determine the actual distribution of the Na+ channel we have raised antibodies to the tetrodotoxin binding component (TTXR), purified from the electric organ of the South American eel Electrophorus electricus (Miller et al., 1982; Ellisman and Levinson, 1982). These antibodies were first used to localize the Na+ channel in light and electron micrographs of eel electric organ and then along myelinated axons of the eel spinal cord (Ellisman and Levinson, 1982). The distribution of anti-TTXR antibodies is demonstrated by peroxidase reaction product in the light micrograph of the thick epoxy section shown in Fig. 27. The resultant dark staining is limited to the innervated and less convoluted faces of the electroplax (solid arrows) while the highly invaginated noninnervated faces are comparatively unstained (open arrows). 16 Mark H. Ellisman et al.

Figure 22. A freeze-fractured E face of a glial loop (GL EF) displaying rows of 160 A particles. The rows (arrows) are spaced 360 A apart while the particles within the rows are separated by 200 A. The underlying axonal P face (AX PF) exhibits rows of dirneric particles. A star indicates the location of a tight junction between adjacent glial loops. 10-day-old rat. X 93,000. Axonal Membranes and the Cytoskeleton 17

Figure 26. In this illustration both P face (dirneric particle distribution) and E face morphology are demonstrated. The paranodal region shows glial loops abutting the scalloped axolemma. Circumferential rows of dimeric particles are shown in the axonal P face. Particles within the glial membranes are shown in cross section as they are positioned above the axolemmal particles. The outermost glial loops interdigitate above the nodal axolemma. The nodal region bulges between the paranodal regions; this region is shown as it would appear in the axonal E face. Large and small particles are closely packed in a uniform annulus around the axon. The basement membrane surrounding the fiber is smooth and closely applied to the membrane.

Figure 23. A P face of a terminal glial loop (GL PF) displaying rows of 160 A particles (filled arrows). The rows are spaced 360 A apart, while the particles within the rows are separated by 200 A. Rows of 75 A particles (open arrows) are centered between the rows of 160 A particles. IO-day-old rat. x 93,000. Figure 24. In a series of transparent overlays, micrographs such as that shown here have been used to project the position of axonal specializations with respect to glial specializations. The rows (arrows) of dimeric particles in the axonal P face (AX PF) are seen to be positioned between the rows of 160 A particles in the glial E face (GL EF) (arrowheads). The 160 A glial particles fall on a line of co linearity between dimeric particles of adjacent rows (see text and Fig. 26). 12-day-old rat. x 117,000. Figure 25. Using transparent overlays as in Fig. 24, it is possible to project the position of specializations on the axonal E face (AX EF) with respect to specializations on the glial P face (GL PF). Rows of large glial particles (closed arrows) alternate with rows of small glial particles (open arrows). The rows of small glial particles are superimposed over the obscure pits of the axonal E face (indicated by small arrows), while the row of large glial particles are superimposed over the ridges of the axonal E face. A star indicates the position of a tight junction between adjacent glial loops. I4-day-old rat. x 117,000. 18 Mark H. Ellisman et al.

Figures 27 and 28. Light micrographs of thick epoxy sections from electrocytes in the main electric organ. Fig. 27. The peroxidase reaction product loca~izing the tetrodotoxin binding protein (Na + channel) stains the innervated face (solid arrow) while the noninnervated face is unstained (open arrows). Fig. 28. A preadsorbed control is presented for comparison. x 200. Axonal Membranes and the Cytoskeleton 19

An example of the control employing antibodies against ITXR which have preabsorbed with TIXR is presented in Fig. 28. No significant enhancement of staining in innervated over noninnervated faces was evident in either the pre-adsorbed control or in preparations first incubated in normal rabbit serum. When viewed at higher magnification with the electron microscope, the microinvagin• ations (caveole) of both the innervated and noninnervated surfaces may be observed (Fig. 29). Although caveole of both surfaces are approximately the same dimensions, those of the innervated face (in continuity with the plasma membrane of the innervated face) exhibited positive staining for ITXR, while those of the noninnervated face did not. Thus, only the innervated surface, including caveole, appears to contain the antigenic determinant for the anti-TIXR antibodies.

B. Myelinated Nerve Having established the distribution of sodium channels (ITXR) on the electroplax, and finding this to be in agreement with physiological evidence for their distribution, we applied similar procedures to examine the distribution along myelinated axons. Vibratome sections of dorsal spinal column of electrophorus, exposed to the immunochemical reagents revealed focal sites of very dense staining when examined in thin sections. The focal staining was restricted to the nodal zone of the axolemma in axons retaining paranodal junctions and an intact myelin sheath (Fig. 30). In order to determine whether this focal staining was a result of limited diffusion of immunochemical reagents into the paranodal zone some preparations were exposed to much more vigorous vibratome sectioning. This often resulted in desheathing ofaxons after primary fixation but prior to immunocytochemical reactions. The staining of axons with the axolemma made accessible in this manner is nonetheless restricted to nodal foci (Fig. 31). No staining was observed in either the preabsorbed IgG control preparation or tissues incubated first in normal rabbit serum instead of anti-ITXR antibodies. Thus, the distribution of antigenic determinants for the anti-TIXR antibodies (sodium channels) ap• pears to be restricted to the nodal zone of the node of Ranvier. We have also used antibodies raised against rat Na + + K + ATPase to examine the distribution of this protein on myelinated mammalian axons. The distribution at the node is the same as that shown in detail above for the Na+ channel. Wood and co-workers (1977) used antisera against eel electric organ Na + + K + ATPase to examine its distribution in knife fish. Our experiments have required examining the distribution of the Na + + K + ATPase in rat and mouse nerves. The antibodies raised against the eel protein do not cross react significantly with rat Na+ + K+ ATPase, so we have purified the rat Na+ + K+ ATPase and raised antibodies to it (Ariyasu et al., 1982; Schenk et al., 1982). These antibodies raised against the purified rat kidney Na + + K + ATPase cross react with rat

Figure 29. Electron micrograph of the immunoreacted electrocytes innervated (solid arrows) and noninnervated surfaces (open arrows). Two apposed surfaces are presented, note that the caveolae of the innervated face stain while those of the noninnervated do not. x 3,800. Figure 30. Node of Ranvier from the dorsal columns of the eel spinal cord. An experimental node, partially demyelinated by the action of the vibratome is presented here. The arrows indicate the nodal zone reaction product localizing the Na+ channels. x 13,000. Figure 31. This node was completely demyelinated prior to the immunocytochemical reaction and only the nodal zone (arrows) is focally stained for Na + channels. x 16,000. 20 Mark H. Ellisman et al. neuronal and glial Na + + K + ATPases and also with the enzyme in the mouse nervous system. Thus, both Na + + K + ATPase and sodium channels appear to be located in the nodal zone of the node of Ranvier. Correlation of the Na + channel with a specific group of particles exposed by freeze-fracture electron microscopy at the node of Ranvier would enable direct measurement of the number of channels per ,...m2 • This may soon be feasible as the TTXR has an apparent molecular weight of 250,000 dalton (Levinson and Ellory, 1973; Agnewet al., 1978; Miller et al., 1982) and on the basis of its solubility and other physical charac• teristics is likely to be an integral membrane protein (Miller et al., 1982; Ellisman et al., 1982a,b). Thus, by analogy with the acetylcholine receptor complex or rhodopsin (Darszon et al., 1980) one would expect to visualize a correlated freeze-fracture particle of approx• imately 100 A. Quantitative freeze-fracture examination of nodal, paranodal, and internodal zones of myelinated axons has revealed approximately 2000 appropriately sized particles per square micron on both protoplasmic (PF face) and external (EF face) fracture faces of the nodal membrane (Ellisman, 1979; Wiley and Ellisman, 1980; Kristol et al., 1977, 1978) where the TTXRs and Na + + K + ATPases are located. Some of the particles found on the nodal PF face undoubtedly represent the N a + + K + ATPase since this protein is known to be exposed by freeze-fracturing (Deguchi et al., 1977). A correlation of sodium channels with freeze-fracture particles is further complicated by differences between numbers of particles found at nodes and the current predicted values for the number of channels per square micron of nodal membrane. Estimates of the number of sodium channels/,...m2 have been made using physiological (Nonner et al., 1975) and pharmacological (Ritchie and Rogart, 1977) techniques. These estimated density values are approximately 50ool,...m2 and 1O,0001,...m2 respectively. Direct determination of whether the sodium channel is exposed by freeze-fracture and if so, what size and shape it is, including knowledge of which fracture face(s) it partitions to, will undoubtedly help reconcile the differences between density estimates obtained using different techniques. It is noteworthy in this regard that the data of Kristol and co-workers (1977) on excitable and inexcitable nodes of Stenarchus implicate EF face particles as correlates for Na + channels since far fewer of these are found in the membranes of inexcitable nodes. Restriction of Na + channels to the nodal zone of the axolemma is one aspect of the regional specialization of the axolemma. We do not know what mechanisms are used by the axon and/or myelinating cell to promote or retain such localized membrane specialization. It appears however to be independent of myelination or the "trapping" of nodal components by the paranodal junctions (Ellisman, 1976, 1979; Wiley-Livingston and Ellisman, 1981). Another possible way of restricting the lateral mobility of Na + channels at nodes might be through linkages with the extracellular matrix. Perhaps a more attractive proposal would be to suggest that the lateral mobility of membrane components at the node is restricted by specific elements of the axoplasmic cytoskeleton. Dense granular material just beneath the nodal membrane which appears to be composed of fine filaments in high voltage electron micrographs (as detailed above) could be the morphological correlate of such a cytoskeletal anchoring system. Understanding the relation of this cortical material to the Na + channel or Na+ + K+ ATPase of the node will depend upon further study. The localization of TTX binding protein and Na + + K + ATPase to the nodal zone eliminates the two activities of these proteins as functional correlates for the paranodal zone of the axon. Recent voltage clamping work on demyelinated mammalian preparations (Ritchie and Chui, 1981) has suggested that the paranodal zone may be involved in delayed recti• fication and thereby one may speculate that the paranodal junction is involved in this activity. Axonal Membranes and the Cytoskeleton 21

Further work in progress is aimed to define the locations of specific macromolecules involved in nodal excitability and discover how they are anchored or aggregate in their specific locations so as to form a functional mosaic. Axonal transport may play an important role in modulating the turnover of proteins at the node such as the Na + + K + ATPase or the sodium channel. Thus, the cytoskeletal membrane interactions that occur both at the level of the node of Ranvier and at sites where transport vectors interact with microtubules or neurofilaments may be functionally linked by molecular sorting codes which specify the location where axonally transported membrane proteins are to be added to the axolemma (Ellisman, 1982).

ACKNOWLEDGMENTS. The authors wish to acknowledge the assistance of Thomas Deerinck, Derek Leong, Judith Nichol, and Dolores Taitano. This was supported by grants from the NIH to M.H.E. (#NS14718) and S.R.L. (#NS15879), as well as grants from the Muscular Dystrophy Association and the National Multiple Sclerosis Society to M.H.E. M.H.E. is an Alfred P. Sloan Research Fellow. S.R.L. is a Research Career Development awardee. C.W.-L. was supported by the National Institute of General Medical Sciences Training Grant (#GM07198) and the National Multiple Sclerosis Society. J.D.L. was a NSF Predoctoral Fellow (#SP179-22285).

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