Evolution of Myelinated Nervous Systems

2.23 Evolution of Myelinated Nervous Systems B I Roots, University of Toronto, Toronto, ON, Canada R M Gould, University of Illinois at Chicago, Chicago, IL, USA ª 2007 Elsevier Inc. All rights reserved.

2.23.1 2.23.2 2.23.3 2.23.4 2.23.5

Introduction Myelin Sheaths in Invertebrates Morphological Features of Vertebrate Myelin Biochemical and Molecular Features of Vertebrate Myelin Sheaths Summary and Conclusions

Glossary desmosome myelin internode

myelin sheath

nodes of Ranvier

oligodendrocyte (OL) Schwann cell

Specialized adherens junction between neighboring cells. A myelin sheath that defines the region between two nodes of Ranvier or a node of Ranvier and the initial segment or distal region where myelin sheaths terminate. Found in both vertebrate and invertebrate nervous systems, they are multilayered membranes that surround large-caliber axons and facilitate saltatory conduction. Axonal membrane where sodium channels are concentrated. These are seen as segmented interruptions in the myelin sheaths. Central nervous system glial cell that produces myelin sheaths around one or more (upwards of 40) internodes. Derived from the neural crest, this cell either produces single myelinated nerves in the peripheral nervous system or establishes nonmyelinating relationships with one to multiple small-caliber peripheral nerve axons. The cells bordering the squid giant axon are often referred to as Schwann cells. As cells in the peripheral nervous system that associate with axons, this definition may apply.

2.23.1 Introduction Associations between glial cells and large-caliber axons are common features among invertebrate and vertebrate nervous systems examined (Schweigreiter et al., 2006). At one extreme are squid giant axons, which reach 1 mm or more in diameter and 10–20 cm in length in widely dispersed species. Tens of thousands of glial cells

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associate with each giant axon (Villegas and Villegas, 1984; Brown and Abbott, 1993). Another well-studied animal model, the sea lamprey, has many, if not all, of its larger-caliber axons surrounded by glial cells (Bertolini, 1964; Merrick et al., 1995). The reasons why glial cells first appeared in ancient nervous systems, why they began to associate selectively with larger axons, remain a mystery. Clearly, they not only contribute to efficiency of nerve impulse propagation by preventing/reducing cross-talk, they also provide structural support and nutrition for the axons and other larger neurites with which they associate. The largeness of squid giant axons allows investigators to separate axoplasm from the axolemmal membrane and surrounding glial cells by simple extrusion (Brown and Lasek, 1990). This feature led investigators to discover that glial cells transfer a subset of newly synthesized proteins to giant axons (Gainer et al., 1977; Lasek et al., 1977; Sheller et al., 1995). It seems reasonable that similar mechanisms exist in other large-caliber axons for, although protein transfer was not measured directly, a transfer of glial-derived proteins and other molecules is thought to underlie the longterm survival of these axons following their separation from neuronal soma (Hoy et al., 1967; Bittner, 1991). Although efforts to understand better the multifaceted nature of interactions between neurons and glial cells are ongoing (Kretzschmar and Pflugfelder, 2002; Oland and Tolbert, 2003; Edenfeld et al., 2005; Sattelle and Buckingham, 2006), our focus is on the specific interactions that result in the formation of myelinated axons. High-resistance myelin sheaths covering nearly the entire surface of large-caliber vertebrate axons are crucial to saltatory conduction, a process that brings both unprecedented efficiency and speed to axon signaling and allows the unmatched

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complexity of modern-day vertebrate nervous systems. Whereas many invertebrate and all vertebrate nervous systems (except cyclostomes) evolved the ability to form myelinated axons, both structural and biochemical evidence (see below) suggests that myelination originated independently several times in invertebrates and only once in vertebrates (Waehneldt, 1990). Furthermore, whereas the strategies used to make the vertebrate myelinated nervous system are being ever more clearly understood, strategies underlying invertebrate glial cell myelination are totally unknown and form the variety of structural features expressed (see below), probably quite varied (see Compensatory Innervation in Development and Evolution, Basic Nervous System Types: One or Many?, Origin and Evolution of the First Nervous System, Adult Neurogenesis and Neuronal Regeneration in the Teleost Fish Brain: Implications for the Evolution of a Primitive Vertebrate Trait).

2.23.2 Myelin Sheaths in Invertebrates Despite a long history, of well over a century, that includes many descriptions of multilayered myelinlike sheaths that surround invertebrate axons/neurites, current focus on invertebrate myelination is sparse (Bullock, 2004; Pan et al., 2006). In contrast to prominence of myelinated axons in vertebrate nervous systems, generally the numbers of axons covered with multi-layered membranes in annelid and crustacean nervous systems are small. In striking contrast to an invariant nature of vertebrate myelinated axons, the nature of invertebrate myelin sheaths is highly variable. Some myelin sheaths, as in the crayfish, Procambarus clarkii (Cardone and Roots, 1991, 1996), are loosely wound, whereas others, e.g., the eyestalk of the crab, Cancer irroratus (McAlear et al., 1958) and earthworm, Lumbricus terrestris (Gu¨nther, 1973, 1976; Roots and Lane, 1983) are highly compacted, more like vertebrate myelin. Among the annelids, myelin-like wrappings have been reported in members of three oligochaete families, i.e., two species of Lumbricidae, Eisenia foetida (Hama, 1959) and L. terrestris (Coggeshall, 1965; Gu¨nther, 1973, 1976; Roots and Lane, 1983), one species of Lumbriculidae, Lumbriculus variegatus (Drewes and Brinkhurst, 1990), and one Tubificid, Branchiura sowerbyi (Zoran et al., 1988). Three families of polychaetes, Capitellidae, Spionidae, and Maldanidae (Nicol, 1958), have large axons covered with multilayered sheaths. Among crustacea, Malacostracan crustacea, several shrimps, prawns, crayfish, and crabs (Holmes, 1942;

McAlear et al., 1958; Hama, 1966; Heuser and Doggenweiler, 1966; Kusano, 1966; Govind and Pearce, 1988; Cardone and Roots, 1991; Xu and Terakawa, 1999; Lenz et al., 2000; Weatherby et al., 2000) have axons covered with myelin-like sheaths. Resemblances between oligochaete and vertebrate sheaths include spiral winding and sheath thicknesses that are correlated with axon caliber. In E. foetida sheath thickness ranges from two to 30 lamellae, whereas in L. terrestris, sheaths contain 60–200 lamellae. Also, interlamellar spacing can be highly variable. The sheath of the L. terrestris median giant fiber, in particular, contains a mix of tightly compacted membranes that resemble vertebrate myelin, and redundant loops formed where the sheath buckles and retains substantial quantities of cytoplasm between membranes. Stacks of desmosome-like structures run serially in register across some sheaths, attaching lamellae to each other (Figure 1). Although these structures resemble vertebrate desmosomes in electron micrographs, they differ in both their intramembranous organization, as revealed by freeze-fracture (Roots and Lane, 1983), and in their protein composition (Pereyra and Roots, 1988). The sheaths found in crustaceans differ from those of annelids and vertebrates; they are concentric rather than spiral. Moreover, they come in two patterns. In the prawn Palaemonetes vulgaris (Heuser and Doggenweiler, 1966) and in shrimps, genus Penaeus (Xu and Terakawa, 1999), the concentric laminae join at a short seam reminiscent of vertebrate sheath mesaxons. The arrangement of seams is very regular with those of alternate laminae located on opposite sides of the axon. In the

*

*

* *

Figure 1 Myelin-like sheath in a longitudinal section of the median giant fiber of Lumbricus terrestris nerve cord. Note the desmosome-like structures (*) running in register across the sheath. Scale bar: 1 mm.

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copepods Undinula vulgaris, Neocalanus gracilis, and Euchaeta rimana, the lamellae form complete circles and lack seams (Weatherby et al., 2000). Another distinguishing feature of crustacean myelin is the location of glial cell nuclei, which are located in variable positions within the sheath (Heuser and Doggenweiler, 1966; Xu and Terakawa, 1999). The conditions whereby glial cell nuclei and perinuclear cytoplasm lie close to axons would tend to favor a potential metabolic transfer of glial proteins and other metabolites to axons. The myelin-like sheath of the crab, C. irroratus, in particular, bears a striking resemblance to vertebrate myelin. Glial cell nuclei lie outside the sheaths, compaction is similar to that of vertebrate myelin, and structures resembling Schmidt– Lanterman incisures and nodes of Ranvier are present (McAlear et al., 1958). At present it is unclear whether the sheath is spirally or concentrically wound. As in annelids and vertebrates, in crustaceans myelin thickness correlates with axon caliber. Lamellar numbers vary from 1 or 2 up to 50. In copepods, interlamellar spacing varies between 3 and 30 nm while the compact intralamellar (fused membranes) space remains constant, around 18 nm in thickness (Weatherby et al., 2000). In prawns, the interlamellar periodicity is more than 20 nm (Heuser and Doggenweiler, 1966), whereas in Penaeus shrimps it is 8 or 9 nm (Xu and Terakawa, 1999). Periodicity is species-dependent in both invertebrates and vertebrates (see following section; moreover, it is affected by tissue-processing methods (Kirschner and Blaurock, 1992; Roots, 1993). The periodicity of the fully compact myelin of Penaeus setiferus, determined by X-ray diffraction (probably the most accurate measurement), is 16 nm, a value similar to that in teleost peripheral myelin (Blaurock, 1986). Interruptions, functionally comparable to vertebrate nodes of Ranvier, occur in both annelid and crustacean sheaths. In many decapod crustaceans (prawns, shrimps, and crabs), the nodes are strikingly similar to vertebrate nodes, both in terms of general morphology (Retzius, 1890; Holmes et al., 1941; Holmes, 1942) and in the disposition of paranodal loops. As is the case in vertebrate loops (see below), these include structures that resemble septate desmosomes (McAlear et al., 1958; Heuser and Doggenweiler, 1966). Internodal distances are generally shorter than in vertebrates (Holmes et al., 1941). A more detailed comparison of crustacean and vertebrate nodes is available (Roots, 1984). Other shrimps, six species of the genus Penaeus, a number of copepods (Hsu and Terakawa, 1996; Xu

and Terakawa, 1999; Weatherby et al., 2000), and the earthworms E. foetida and L. terrestris (Hama, 1959; Gu¨nther, 1973, 1976) have completely different nodes. They have circular openings in the myelin sheath and are referred to as focal or fenestration nodes. In L. terrestris, there are two nodes of 10–15 mm diameter in each segment. In Penaeus shrimps, node diameter and internodal distance are both approximately proportional to fiber diameter. Node diameter varies between 5 and 50 mm and internodal distance from 3 to 12 mm (Xu and Terakawa, 1999). Another morphological feature, suggesting a novel mechanism of fast nerve conduction, occurs in shrimps of the genus Penaeus. A large gel-filled space is present between the axon and myelin sheath. This submyelinic space increases effective axon diameter and, as a consequence, conduction velocity. It is tightly sealed at the node regions, permitting saltatory conduction (Hsu and Terakawa, 1996; Xu and Terakawa, 1999). Conduction velocity has been measured in only a few invertebrate nerves. In the median giant fiber of the earthworm L. terrestris, which is 90 mm in diameter, it is 30 m s 1 (Gu¨nther, 1976). In the shrimp, Penaus japonicus, it is 90–190 m s 1 in fibers 120 mm in diameter (Kusano, 1966; Kusano and LaVail, 1971). For comparison, a rat fiber of 4.5 mm diameter with a sheath of about 50% of its total diameter conducts at 59 m s 1. Thus, vertebrate sheaths are far more effective in increasing conduction velocity. Reaction times for the escape responses of calanoid copepods have been measured (Lenz et al., 2000). The fastest responses were recorded in myelinated species, the escape response being initiated 2–5 times more rapidly than in the nonmyelinated species. Myelin is found only in the more recently evolved copepod superfamilies, which also live in more diverse habitats. They not only live in neritic and deep-water environments but also in regions of the ocean where faster reaction times are essential for avoiding predators (Hayward and McGowan, 1979; Parks, 1986; Hays et al., 1997; Lenz et al., 2000). Thus, during the course of copepod evolution, the development of myelin has allowed them to occupy new habitats. Very little is known about the distribution of sodium channels in invertebrate axons. Studies on the shrimp P. japonicus indicate that, as in vertebrates, sodium channels are concentrated at the nodes at a density of 530 channels mm 2 (Hsu and Terakawa, 1996; Xu and Terakawa, 1999). In the earthworm L. terrestris, sodium channels are also concentrated at the nodes (Gu¨nther, 1976; Roots, 1984, 1995b). Information on the chemical compositions of different invertebrate myelin sheaths is limited.

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Fortunately, techniques used to purify myelin membranes (based on size and density of the membrane) from vertebrates are adaptable to invertebrates (Pereyra and Roots, 1988; Waehneldt et al., 1989). Analysis of lipid and protein compositions in annelid and crustacean myelin shows marked differences from vertebrate myelin (see below) and from one another. Birefringence studies of earthworm myelin showed the sheath to be qualitatively similar to that of frog sciatic nerve, with protein contributing 30–40% of the total birefringence and the rest attributed to lipids (Taylor, 1942). In the shrimp, Penaus duorarum, a strikingly high proportion of lipids is found in isolated myelin; the lipid-to-protein ratio is 15 : 1 (Okamura et al., 1986b). Galactolipids, major constituents of vertebrate myelin sheaths (Morell and Quarles, 1999) are not present in annelid or crustacean myelin. Instead, glucocerebroside in amounts equivalent to galactocerebroside in vertebrate myelin is present in crustacean, though not in earthworm, myelin. Sphingomyelin is absent from earthworm nerve cord and, although it is found in crayfish (Cambarus clarki) nerves, it is structurally quite different from vertebrate sphingomyelin (Komai et al., 1973; Okamura et al., 1986a). Thus, there is an evolutionary trend in which glucocerebrosides in protostomes are replaced by galactolipids in deuterostomes. Although deuterostomes also synthesize glucocerebroside (Tamai et al., 1992), and do so under circumstances when galactolipid synthesis is blocked (Bosio et al., 1998), evolutionary inclusion of galactosphingolipids in myelin (Roots, 1995a) may have occurred to foster lipid–lipid and lipid– protein interactions needed for node/paranode stabilization (Popko, 2000) and/or raft-dependent signaling involved in myelin assembly (Boggs and Wang, 2001). The protein components of both annelid (earthworm L. terrestris) and crustacean (crayfish P. clarkii and pink shrimp P. duorarum) myelin are totally different from those of vertebrates. As in vertebrates, the protein pattern of earthworm myelin is relatively simple, with 80 and 42 kDa proteins predominating and 28–32 kDa proteins as minor components. The structure of these proteins is unknown and there is no cross-reactivity with antibodies to myelin proteins (see Section 2.23.4 for more information on these proteins), including myelin basic protein (MBP), proteolipid protein (PLP), myelin-associated glycoprotein (MAG), and 29,39-cyclic nucleotide 39-phosphodiesterase (CNP) (Pereyra and Roots, 1988; Cardone and Roots, 1990). In the pink shrimp, four major proteins, 21.5, 40, 78, and 85 kDa, and four minor proteins,

36, 41.5, 43, and 50 kDa, are found in purified sheath membranes. None of these proteins show cross-reactivity with antibodies that recognize mammalian MBP or PLP or trout MBP, 36 K or protein zero (P0) (Okamura et al., 1986b; Waehneldt et al., 1989). A monoclonal antibody generated to earthworm myelin-like membranes and showing cross-reactivity with 30–32 and 40 kDa proteins cross-reacts with 60–65, 42, and 40 kDa proteins in crayfish (P. clarkii) axonensheathing membranes (Cardone and Roots, 1996). Thus, earthworm and crayfish myelin membrane proteins have some antigenic epitopes in common. The presence of a myelin sheath confers several advantages in invertebrate survival. The startle reactions of earthworms, escape responses of crayfish, shrimp, and copepods, and retraction of eyestalks in crabs are behaviors in which speed is of paramount importance (Bullock, 1984). Alternatively, faster conduction may be achieved by simply increasing axon diameter, as occurs in squid giant axons (see above). However, this alternative is evolutionarily less favorable. A more efficient means of increasing conduction speed and, therefore, the rapidity with which escape mechanisms take place is the development of myelin sheaths. The nodes of invertebrate nerve fibers serve to allow saltatory conduction in a similar fashion to vertebrate nodes of Ranvier. It should be noted that the points of emergence of small collaterals serve as nodes in both vertebrates (Roots, 1984) and the shrimps Penaeus chinensis and P. japonicus (Xu and Terakawa, 1999).

2.23.3 Morphological Features of Vertebrate Myelin Myelination arose in the common gnathostome ancestor as cyclostomes; lamprey and hagfish totally lack myelin (Bullock et al., 1984) and all gnathostomes examined have myelinated nervous systems that are structurally rather invariant, as descriptions for mammalian myelin (Peters et al., 1991; Hildebrand and Mohseni, 2005) readily apply to nonmammalian myelin/myelination and vice versa (see below). Furthermore, the growing understanding of vertebrate myelination is nurtured by continued cross-fertilization, as studies with mammals and nonmammals are often interpreted interchangeably (Kagawa et al., 2001; Lobsiger et al., 2002; Jessen and Mirsky, 2004, 2005; Miller and Reynolds, 2004; Le et al., 2005; Richardson et al., 2006). Several reasons why nonmammals have been chosen for study are: (1) quail-chick

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chimeras allow ready oligodendrocyte (OL) lineage tracing (Cameron-Curry and Le Douarin, 1995; Pringle et al., 1998); (2) fishes and some amphibians have the ability to regenerate their central nervous system (CNS) (Sivron et al., 1990; Diekmann et al., 2005); (3) a general interest in comparative myelination (Jeserich and Rauen, 1990; Jeserich et al., 1990; Jeserich and Stratmann, 1992; Gould et al., 1995; Nguyen and Jeserich, 1998; Park et al., 2005); (4) a growing interest in comparative genomics (Aparicio et al., 2002; Gilchrist et al., 2004; Venkatesh and Yap, 2005); and (5) use of highthroughput genetic screens in zebra fish (Rossant and Hopkins, 1992; Sprague et al., 2003), that adapt to identifying proteins important for myelination (Lyons et al., 2005). The first electron microscopic studies showing the spiraling nature of peripheral nerve myelin formation were conducted with nonmammals, i.e., chick (Geren, 1954) and chameleon (Robertson, 1955) nerves. Subsequent electron microscope observations made on frog (Maturana, 1960) and toad (Stensaas and Stensaas, 1968) CNS sections show similar spiral wrapping of axons with elaborations of OL plasma membrane. More recently, antibody studies of PLP expression showed that frog OLs displayed identical structural appearances during development to mammalian OLs (Yoshida, 1997). We readily realize the morphological conservation of CNS and peripheral nervous system (PNS) myelination among gnathostomes with comparisons of images from developing and mature mammalian and nonmammalian species (Figures 2–4). Clearly, at both light and electron microscopic levels, multiple features, including selection of large-caliber axons, loose followed by tight wrapping, and development of nodal and paranodal specializations appear indistinguishable between these evolutionarily divergent species. At the time myelination begins in ventral spinal cord and medial longitudinal fascicle of elasmobranches (4–5 cm spiny dogfish fetuses) and mammals (newborn rats: Schwab and Schnell, 1989), one can clearly see that initial appearances of OLs and early myelin wrapping are quite similar (Figure 2). In both cases, early OLs hover around the edge of the tissue (Figures 2a and 2b, *). At the electron microscopic level, similar appearances of OL nuclei with chromatin clumped along the nuclear envelop and large regions of perinuclear cytoplasm filled with mitochondria, Golgi, rough and smooth endoplasmic reticulum, and numerous microtubules characteristic of myelinating mammalian OLs (Figure 2d) are also characteristic of myelinating spiny dogfish OLs. As development progresses in spiny dogfish ventral funiculi, the numbers of large

Figure 2 Transverse sections of developing ventral funiculi in spinal cords of spiny dogfish (left) and rat (right). a, From 4.5 cm spiny dogfish fetus showing the first appearances of OLs (small dark profiles lining the medial border) and initial ring-shaped myelin sheaths. b, From newborn rat pup showing similarly appearing early OLs (dark profiles) and beginning myelin sheaths. c, Electron micrograph from a 6 cm spiny dogfish fetus ventral spinal cord showing an OL (nucleus with clumped chromatin at upper left) and typical organelle-rich perinuclear cytoplasm with a few large-caliber axons in early stages of myelination. Many small unmyelinated axons also surround the OL. d, Electron micrograph from a P2 neonatal rat pup ventral spinal cord with a similar-appearing OL with surrounding large axons in early stages of myelination and clusters of many smallcaliber unmyelinated axons. Arrows in (a) and (b) point to early myelin sheaths and asterisks (*) mark the tissue border. Adapted from Schweigreiter, R., Roots, B. I., Bandtlow, C. E., and Gould, R. M. 2006. Understanding myelination through studying its evolution. Int. Rev. Neurobiol. 73, 219–273, Elsevier.

myelinated fibers increase dramatically. When the fetus has more than doubled its size, large numbers of myelinated fibers and few OLs occupy a cross section of ventral cord (Figure 3a). Even though the ventricle seems filled with large-caliber myelinated fibers, there are still many very small fibers that are totally disregarded. At the electron microscopic level, these fibers are still bundled tightly together (Figure 3b). That these axons are eventually myelinated can be seen when a comparable region of ventral spinal cord is imaged in the adult spiny dogfish (Figure 3c). By this time, i.e., after several years, the entire funiculus is composed of myelinated fibers of widely different caliber. An evolutionary important lesson is that OLs show an absolute selectivity for those axons that are enlarging while

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disregarding small-caliber axons. Evidence is emerging as to signaling molecules important for this recognition (see following section). As in the CNS, the Schwann cell-based PNS myelin is highly similar among vertebrates. Again, we will show examples from spiny dogfish to illustrate this point (Figure 4). We have followed the development of trigeminal nerve in spiny dogfish fetuses of 2–26 cm, the latter being the size at birth. In the smallest fetuses, bundles of axons are surrounded by an outer layer of immature Schwann cells, much like those seen in mice and rats (Figure 2; Jessen and Mirsky, 2005). When the fetuses reach 6 cm, nearly all the trigeminal axons are separated from all of their neighbors by Schwann cells (Figure 4A). A few have layers of noncompacted and compacted myelin (not shown). When the fetus reaches 13 cm in size, all of the axons are myelinated (Figure 4B). Myelin sheaths are related to Schwann cells in the same manner that has been described most frequently for the mammalian PNS (Peters et al., 1991).

Figure 3 a, Transverse section from the ventral funiculus in the spinal cord of a 13 cm spiny dogfish, myelination. Now there are many myelinated fibers and a few OL profiles (arrow) per cross section. b, What is not appreciated is the very large number of small nonmyelinated fibers that are still present in the ventral spinal cord and can only be seen when viewed through the electron microscope. c, In the same region of the adult spiny dogfish spinal cord, axons of wide-ranging sizes are all myelinated.

2.23.4 Biochemical and Molecular Features of Vertebrate Myelin Sheaths Based on structural differences (high lipid content and large expanses of compact membrane layers), methods that separated mammalian brain and peripheral nerve myelin from other membranes were developed in the 1960s (Albers, 1981; Quarles et al., 2005). Fortunately, these methods

Nu

Pn

(A)

(B)

(C)

Figure 4 Transverse section of developing trigeminal nerve in spiny dogfish fetuses that were either 4.5 cm (A) or 22 cm (B and C). A, During early development, axons (a) reach a critical size (all axons in the field have reached this size) when they become separated from all neighboring axons and are covered by single Schwann cells segmentally arranged. A large nucleus (Nu) of one Schwann cell indicates that the section is taken at the point where the Schwann cell soma is found. Collagen-filled (c) extracellular space separates each individually ensheathed axon. B, At later developmental stages all larger trigeminal axons are myelinated with sheaths of thicknesses that relate to axon caliber. As at earlier times, axons sectioned near the center of the myelinating cell display nuclei and/or perinuclear (Pn) Schwann cell cytoplasm. C, At high magnification enrichment of neurofilaments and microtubules in the axon along with a few motochondria and smooth membranes are visible. A thin rim of organelle-filled Schwann cell cytoplasm surrounds compact myelin and a basal lamina covers the surface.

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easily adapt to the isolation of myelin-like membranes from invertebrate neural tissues (see above) and from non-mammalian vertebrate neural tissues (Mehl and Halaris, 1970; Franz et al., 1981; Waehneldt et al., 1984). Analyses of myelin fractions from species representing each group of modern-day gnathostomes confirmed that, as in mammalian myelin preparations, they contain an abundance of lipids (70–80% of the isolated membrane fraction dry weight), including high portions of cholesterol, galactolipids, and ethanolamine plasmalogen (Bu¨rgisser et al., 1986; Kirschner and Blaurock, 1992), and a relatively simple protein profile (Waehneldt et al., 1986a). The major proteins in mammalian myelin, like those in invertebrate myelin, are of low molecular weight. In general antibodies recognizing mammalian myelin proteins recognize homologs in nonmammalian brain and nerve samples (Waehneldt et al., 1986a). These studies revealed that all vertebrate myelin sheaths contain a dominant protein, either P0 (one or two bands) or PLP (with a less abundant alternatively spliced variant, DM-20, absent only in amphibians; for review, see Waehneldt, 1990; Maisey, 1986). All peripheral nerve myelin, from elasmobranches to mammals, have P0-based myelin sheaths. The difference between fish and tetrapods lies in CNS myelin. Cartilaginous and bony fishes express P0, the same protein(s) they express in their PNS. In contrast, all tetrapod CNS myelin is PLP-based. This difference was first recognized through immunoblot comparisons of the proteins from fish and tetrapod myelin preparations (Franz et al., 1981; Waehneldt et al., 1985). This difference is apparent at the structural level as sheaths with P0 have wider intraperiod lines, double in appearance. The difference is seen by both electron microscopy and X-ray diffraction (Kirschner et al., 1984, 1989; Waehneldt et al., 1984; Kirschner and Blaurock, 1992). Recent evidence that further supports the correlation of P0 and wider intraperiod lines was obtained with transgenic mice with P0 substituted for PLP (Yin et al., 2006). A possible reason why the P0-based CNS myelin of teleost and cartilaginous fishes evolved to PLP-based CNS myelin in amphibians and other tetrapods was recently suggested (Schweitzer et al., 2006). Since the growth of the brain cases of tetrapods slows in adulthood, the tighter packing of a PLP-based CNS myelin would allow more myelin in a limited space. Although PLP/DM-20 (fish do not have exon 3b, the PLP-specific exon; see below) and P0 are co-expressed in amphibians (Takei and Uyemura, 1993; Yoshida and Colman, 1996) and teleost fishes

(Schweitzer et al., 2006), differences in intraperiod spacing (Kirschner and Blaurock, 1992) reflect the dominance of PLP in amphibian myelin and P0 in fish myelin. P0 is present at roughly 50% of total myelin protein, at least in mammalian PNS (Greenfield et al., 1973), and additional information on properties of P0 can be obtained in several reviews (Spiryda, 1998; Eichberg, 2002; Kirschner et al., 2004). It is type I (single transmembrane) glycoprotein, a smallest member, which has a single immunoglobulin domain, of the immunoglobulin superfamily, and has HNK-1-reactive antigens in many species (Quarles, 2002). Other posttranslational modifications include sulfation and fatty acylation. It is involved in adhesion at both the intraperiod and major dense lines (Kirschner et al., 1996; Shapiro et al., 1996). P0 homologues were not detected in ascidians (Gould et al., 2005), indicating that this protein likely evolved after the common gnathostome ancestor separated from the lineage leading to extant hemichordates. PLP and DM-20 are tetraspan proteins of an ancient family, termed lipophilins (Gow, 1997; Hudson, 2004; Gould et al., 2005; Schweitzer et al., 2006), with both N- and C-terminal regions in the cytoplasm. Although PLP is restricted to tetrapod CNS myelin, DM-20-like proteins are present in teleost fish (Tohyama et al., 1999, 2000; Brosamle and Halpern, 2002; Schweitzer et al., 2006), elasmobranch (Kitagawa et al., 1993; Sinoway et al., 1994), and amphibians (Yoshida and Colman, 1996). The difference between PLP and DM-20 is a 35-amino-acid sequence (exon 3b), present in PLP, that lies in the cytoplasmic loop between the second and third transmembrane domain. There are two different proteins that resemble DM-20, called M6A and M6B. The latter proteins are expressed in neurons and probably to much lesser extents in glial cells (Yan et al., 1996; Werner et al., 2001). Homologues of these DM-20/M6a/M6b proteins have been identified in insects (Stecca et al., 2000; Schweitzer et al., 2006) and basal chordates (Gould et al., 2005; Schweitzer et al., 2006) based on database screening. High conservation of their structure and genomic organization indicates a widespread evolutionary importance of this protein family. The high number of PLP/DM-20 sequences has been used as one base for determining phylogenetic relationships among vertebrate groups (Tohyama et al., 2000; Venkatesh et al., 2001). Another less abundant tetraspan protein of mammalian PNS myelin is PMP22, a protein studied in part because mutations are related to peripheral nerve diseases (Suter and Scherer, 2003; Young and Suter, 2003; Suter, 2004). It is found in zebra fish (Wulf et al., 1999)

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and a homologue was identified in ascidian (Gould et al., 2005), indicating that several tetraspan proteins, lipophilins, PMP22/claudins, and connexins may have been used for the original development of myelin sheaths. MBP, a family of alternatively spliced members, is present in all CNS and PNS myelin sheaths (Campagnoni, 1988; Martenson and Uyemura, 1992) and, because of its recognized importance, has been termed the executive myelin protein (Moscarello, 1997). MBP is a member of the intrinsically disordered, highly adapted proteins (Harauz et al., 2004). Perhaps with the openness of structure comes an attraction for post-translational modifications, which are numerous (Moscarello, 1997; Harauz et al., 2004). The only study of post-translational modification of MBP in a nonmammal was conducted with MBP isolated from spiny dogfish brain and spinal cord. Modifications include phosphorylation, deamidation, and likely deimidation and praline hydroxylation. Clearly studies of MBP post-translational modification in other nonmammalian species will help clarify our emerging understanding of the roles these proteins play in glial cell development and myelination. Classic MBPs, the series of alternatively spliced proteins present in myelin, are members of a larger family of genes of OL lineage (GOLLI) that include four additional proteins transcribed from two additional upstream promoters (Campagnoni and Campagnoni, 2004). MBPs are needed for major dense line (extrusion of intervening myelin), as mutant mice (shiverer and mld) and rats (Long Evans shaker) that lack MBP have little myelin and, when it occurs, it is poorly compacted and contains an abundance of trapped cytoplasm (Privat et al., 1979; Ganser and Kirschner, 1980; Kwiecien et al., 1998). Furthermore, mammalian MBP isoforms that contain exon-2 are expressed developmentally early (Barbarese et al., 1978), are enriched in radial component (Karthigasan et al., 1996), a structure unique to CNS myelin (see below) and are targeted to nuclei where they may function in transcriptional regulation (Pedraza et al., 1997). Interestingly, most likely the sequence for exon-2 does not occur in teleost fishes or amphibians based on analyses of the Xenopus, zebra fish and pufferfish databases (Gould, unpublished observations). Furthermore, only a single MBP homologue was identified in zebra fish (Brosamle and Halpern, 2002; Lyons et al., 2005). Two alternative splice variants have been reported in elasmobranches, though both lack exon-2 (Saavedra et al., 1989; Spivack et al., 1993). cDNA sequences in the pufferfish database suggest some

alternative splicing occurs there as well. Analysis of differences of MBP sequences can shed light on regions of the protein that might be used for signaling and for myelin compaction. A triproline region in mammalian and amphibian MBPs, thought to be structurally important in forming a hairpin loop, is absent in fish MBPs. No MBP homologues were detected in searches of the ascidian genome database (Gould et al., 2005), indicating that, like P0, MBP developed late in evolution. In addition to P0, MBP, and PLP/DM-20, there are a growing number of proteins associated with myelin sheaths proper and the process of myelination, based on comparison of earlier (Braun, 1984; Campagnoni, 1988) and later (Lazzarini et al., 2004) reviews. Evolutionary studies of myelin proteins occurred at the time when the major structural proteins in isolated myelin were the focus (Waehneldt et al., 1986b; Waehneldt, 1990; Jeserich and Waehneldt-Kreysing, 1992; Kirschner and Blaurock, 1992). Two minor-occurring proteins, CNP (Kurihara and Tsukada, 1967; Tsukada and Kurihara, 1992) and MAG (Everly et al., 1973), have long histories as myelin-related proteins. Immunocytochemical studies have demonstrated that they are located near, but not within, compact myelin (Trapp et al., 1988, 1989). CNP is present in bird and amphibian myelin (Kasama-Yoshida et al., 1997), though not in fish myelin (Kasama-Yoshida et al., 1997; Moll et al., 2003). A related protein, goldfish regeneration-induced CNPase homologue (gRICH) that lacks CNPase activity, has been identified in goldfish (Ballestero et al., 1995, 1997, 1999). A comprehensive review on CNP (Braun et al., 2004) can be consulted for additional information. Furthermore, a recent study of transgenic mice lacking CNP expression suggests a crucial role for this protein in axon–glial cell communication (Lappe-Siefke et al., 2003). Antibody-based evidence that MAG is ubiquitously expressed (Matthieu et al., 1986) had been disputed (Zand et al., 1991; Hammer et al., 1993) and the issue was not resolved until recent screening of pufferfish and zebra fish genomes identified several MAG, also called siglec 4 (sialic acid-binding protein), isoforms in these fish (Lehmann et al., 2004). Finding fish homologues of MAG, a protein associated with inhibition of myelin regeneration in the mammalian CNS (Filbin, 1995), will clearly impact efforts to understand the evolutionary loss of CNS regeneration. Another minor-occurring protein restricted to PNS myelin is PMP22, a protein involved in the process of myelin sheath formation (see above; Suter, 2004; Amici et al., 2006), present in amphibians (Xenopus genome project) and fish (Wulf

Evolution of Myelinated Nervous Systems 477

et al., 1999), and which has a homologue in the ascidian, Ciona intestinalis (Gould et al., 2005). Within compact CNS myelin are radial bands, structures seen with electron microscopy (Peters, 1961). Although not studied widely, they are also found in amphibian CNS myelin (Schnapp and Mugnaini, 1976; Tabira et al., 1978), though not any PNS myelin (Kosaras and Kirschner, 1990). Interestingly, the key structural proteins of radial bands, exon-2-containing MBP isoforms (Karthigasan et al., 1996) and myelin-associated oligodendrocytic basic proteins (MOBP) (Yamamoto et al., 1999), have not been identified in amphibians and other nonmammalian species. Nodes of Ranvier, regions between internodal myelin and bracketed by paranodal loops, contain high concentrations of sodium channels and other proteins that are delivered to these sites via axonal transport (Poliak and Peles, 2003; Salzer, 2003). In the PNS, they are covered by microvillar extensions of adjacent Schwann cells. In the CNS, they are covered by a totally different cell, which is related to CNS astrocytes (ffrench-Constant et al., 1986; Black et al., 1989; Butt et al., 1999) and expresses a proteoglycan, NG2 (Butt et al., 1999; Martin et al., 2001), that is well associated with the OL lineage (Dawson et al., 2000). Almost nothing is known about the structural properties of nonmammalian proteins located in nodes of Ranvier and surrounding paranodes and juxtaparanodes. From comparative studies of proteins present in compact myelin, it is likely that antibodies to mammalian nodal, paranodal, and juxtanodal proteins will identify nonmammalian homologues and it would be rather straightforward to determine whether proteins involved in node and paranode formation and function behave the same in nonmammalian settings. Interestingly, some of the key components of paranodal junctions, contactin, and neurexin family proteins are used in glial ensheathment in insects (Bhat, 2003; Banerjee et al., 2006). A totally unrelated area that has not been considered from an evolutionary viewpoint is the origin of myelin-forming Schwann cells and OLs. The few studies that have been done with chickens and frogs are enmeshed in studies of mammals without due recognition of evolutionary significance (Richardson et al., 2000, 2006; Qi et al., 2002; Miller and Reynolds, 2004; Noble et al., 2004; Rowitch, 2004; Jessen and Mirsky, 2005). A few studies have taken advantage of zebra fish with respect to understanding roles of olig family helix– loop–helix transcription factors (Park et al., 2002; Shin et al., 2003; Filippi et al., 2005). Hopefully, the future will include a re-assessment of these studies in the light of evolution.

In addition a number of screens have emerged, including ones that identify proteins: (1) synthesized from mRNAs transported along with MBP mRNA to OL processes (Gould et al., 2000); (2) essential for the development of myelin phenotypes in zebra fish, including transport of MBP mRNA to processes (Lyons et al., 2005) and clustering of sodium channels at nodes (Talbot, personal communication); and (3) based on the presence of a signal sequence that targets Schwann cell proteins to plasma membranes, where they may be involved in interactions with axons and/or the basal lamina. The first screen has been applied to spiny dogfish and shows that some OL process mRNAs, though surprisingly, not MBP mRNA, are common to both fish and mammals (Gould, unpublished observations). Studies of protein evolution are clearly impacting research on CNS myelin proteins with inhibitory influences on regeneration (Diekmann et al., 2005; Schweigreiter et al., 2006). Briefly, what these and others are doing is characterizing proteins, expressed in CNS myelin and/or OLs that inhibit regrowth of damaged CNS axons. It is well known that fish, some amphibians, chicks, and some mammals (Gaze, 1970; Stuermer et al., 1992; Ferretti et al., 2003) – the latter at premyelination stages only – are able to regenerate CNS axons following injury. Although it is not entirely clear why the ability to regenerate CNS axons is lost in birds and mammals following myelination, the presence of candidate proteins, including NOGO/RTN isoforms (Cadelli and Schwab, 1991; Chen et al., 2000; GrandPre´ et al., 2000; Prinjha et al., 2000), NOGO/RTN receptor (Fournier et al., 2001), MAG, and OL myelin glycoprotein (OMgp) (Spencer et al., 2003; Woolf, 2003; Raisman, 2004; Sandvig et al., 2004; Schwab, 2004; Domeniconi and Filbin, 2005; Schweigreiter et al., 2006) suggests that there was a need to inhibit axon growth/elongation in regions (myelin-rich) where these proteins are located. With the identification of some of these molecules in fish (see above references), it will be possible to compare structural features of the proteins and expression patterns as a means to understand how they might selectively inhibit mammalian CNS regeneration. Another important area that has not been considered from an evolutionary point of view is the axon– glial cell cross-talk involved in different stages of myelination. With rapid progress in mammalian systems, this is another area that should receive attention from evolutionary myelin biologists. Only when axons reach appropriate size, i.e., develop unique surface character, such as expressing sufficient amounts of neuregulin-1 type III on their surface, will OLs (Sherman and Brophy, 2005) and Schwann cells (Nave and Schwab, 2005; Sherman

478 Evolution of Myelinated Nervous Systems

and Brophy, 2005; Taveggia et al., 2005) myelinate them. Furthermore, myelin thickness, at least in peripheral nerves, is dependent on the same neuregulin-erbB receptor signaling (Michailov et al., 2004; Nave and Schwab, 2005). With knowledge of developmental time course of myelination in nonmammals, including birds (Ono et al., 1995), reptiles (Nadon et al., 1995), amphibians (Tabira et al., 1978), teleost (Jeserich and Rauen, 1990), and cartilaginous fishes (Gould et al., 1995), investigators will find ways of approaching evolutionary aspects of axon–glial cell interactions that are important for myelination.

2.23.5 Summary and Conclusions One might wonder why, with the rapid progress in mammalian cell biology and molecular biology, the availability of many mammalian mutants with defects in myelination, and the knockout and knockin strategies to influence myelin-related proteins of interest, one should devote and, probably more importantly, fund efforts to study myelination in nonmammalian species. From the information already presented, it is clear that there has been an evolutionary driving force for generating myelinated axons that includes not only all gnathostomes, but also many invertebrates. Particularly in invertebrates, a variety of strategies have been developed and, clearly, knowledge of these strategies will provide a broader base for understanding how interactions between neurons, their large-caliber axons, and the glial cells with which they interact were focused on myelination. Since none of the invertebrates with myelinated axons have been or (to our knowledge) are planned to be subjects of whole-genome studies, the best alternative to obtain sequence information on myelin proteins and proteins important for myelination would be through proteomic methods. It is our hope that investigators with available knowledge and facilities will consider earthworms, shrimps, and/or other annelids/crustaceans, obtain sequences of proteins present in myelin-like membranes, and use this knowledge as a starting point for developing understanding of how the proteins participate in neuron/ axon-generated myelin formation. For those interested in using nonmammalian vertebrate models, tremendous advances in genomic research involving zebra fish (Key and Devine, 2003), medaka (Shima and Mitani, 2004), two species of pufferfish (Aparicio et al., 2002; Jaillon et al., 2004), amphibians (Hirsch et al., 2002), chordates lacking myelin (amphioxus and sea lamprey) (Mulley and Holland, 2004), and urochordates

(Dehal et al., 2002; Meinertzhagen et al., 2004) have been made. This is an exciting time. Obtaining protein sequences in published databases is relatively easy and straightforward (Brosamle and Halpern, 2002; Lehmann et al., 2004). These groups identified zebra fish, and in some instances pufferfish, MBP, PLP-related DM-20, P0, and MAG. Protein sequence information can be used in many ways to advance myelin research. Because of evolutionary pressures, domains in protein sequences important for function are most highly conserved and sequence alignments can be used to determine these domains and searches can be performed to see if these domains have orthologues and/or paralogues. Overall this information should help in supporting and/or negating structural models of known myelin proteins (see Shapiro et al., 1996; Spiryda, 1998; Kirschner et al., 2004; for discussions on P0 structure and Martenson, 1992; for discussions on MBP structure). Clearly, the studies covered in this brief review are not comprehensive and we apologize to scientists whose research efforts and relevant studies were not cited.

Acknowledgments We would like to acknowledge the support of the National Science Foundation (grant IBN-0402188, 10 RMG) and grants from the NIH (NS 23868 and NS23320) and Natural Sciences and Engineering Research Council of Canada (grant A6052 to BIR), whose support helped allow us to write this article.

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Further Reading Lyons, D. A., Pogoda, H. M., Voas, M. G., et al. 2005. erbb3 and erbb2 are essential for Schwann cell migration and myelination in zebrafish. Curr. Biol. 15, 513–524. Roots, B. I. 1984. Evolutionary aspects of the structure and function of the nodes of Ranvier. In: The Node of Ranvier (eds. J. C. Zagoren and S. Fedoroff), pp. 1–29. Academic Press. Roots, B. I. 1995. The evolution of myelinating cells. In: NeuronGlia Interrelationships During Phylogeny. I: Phylogeny and Ontogeny of Glial Cells (eds. A. Vernadakis and B. I. Roots), pp. 223–248. Humana Press. Schweigreiter, R., Roots, B. I., Bandtlow, C. E., and Gould, R. M. 2006. Understanding myelination through studying its evolution. Int. Rev. Neurobiol. 73, 219–273.