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Are spinal nerves myelinated and unmyelinated at the same time?


I was trying to answer this question when I remembered that the somatic axon is myelinated, while both sympathetic and parasympathetic preganglionic axons are also myelinated. Are they only myelinated or are they both myelinated and unmyelinated? Thanks

Edit:

This is a question from an old lecture quiz (previous year) and unfortunately I do not have the answers for the questions. The wording of the question was

Spinal Nerves are

a.) myelinated

b.) unmyelinated

c.) answers a and b

d) none of the above"


Spinal nerves are mixed nerves containing afferent and efferent neurons of various types. Anatomically, they protrude from the spinal column bilaterally at each vertebral level. They contain both myelinated fibers (e.g., A fibers) and unmyelinated fibers (e.g., C fibers).

The answer is (c): both myelinated and unmyelinated.

Please note that spinal nerves are NOT located in the spinal cord, despite the fact that neurons in each spinal nerve will either originate or terminate there. A spinal nerve is a peripheral structure. It starts at the point where the dorsal and ventral root converge (See April's Clinical Anatomy, Ch. 1). If you have the opportunity to observe back surgery or dissect a human cadaver, you can see and touch a spinal nerve. They are cable like structures containing bundles of many many many axons, but no neuron in its entirety. A nerve is not a neuron. This is a common misconception.

Just to add, the details here may seem like trivia, but they are not. Similar questions (and questions that require you to understand the distinction between a nerve and a neuron) are often asked on US medical licensing exams. This is because they are relevant to understanding and interpreting neurological symptoms and physical exam findings.


Yes, there is grey and white matter in the spinal chord and the parasympathetic nervous system, so they are both myelinated and unmyelinated:

Here is an image of parasympathetic nerves:

Primary afferent fibers of the dorsal roots are either myelinated or unmyelinated. In the rat, cat, and monkey, thin unmyelinated and thick myelinated fibers are located in the lateral and medial parts of the dorsal roots, respectively, in the dorsal root entry zone (Light and Perl, 1979a).


Distinguish between: (a) afferent neurons and efferent neurons (b) impulse conduction in a myelinated nerve fibre and unmyelinated nerve fibre (c) aqueous humor and vitreous humor (d) blind spot and yellow spot (f) cranial nerves and spinal nerves.

NCERT Solutions for Class 11 Biology Chapter 21 Neural Control and Coordination
Important Board Questions
Neural Control and Coordination
NCERT Books for Session 2020-2021
CBSE Board and UP Board
Question : 12

1 Answer

Aashvi Patel

(a) Afferent neurons and Efferent neurons
Afferent Neurons:- Afferent neuron conducts nerve impulses toward the brain or the spinal cord.
Efferent Neurons:- Efferent neuron conducts nerve impulses from the brain or spinal cord to the effector organs such as muscles or glands.

(b) Impulse conduction in a myelinated nerve fibre and unmyelinated nerve fibre
Impulse Conduction in a Myelinated Nerve Fibre:-
1. Myelinated nerve fibre, the action potential is conducted from one node to another.
2. Conduction of impulses is faster.
Impulse Conduction in an Unmyelinated Nerve Fibre:-
1. Unmyelinated nerve fibre, the action potential is not conducted from node to node. It is carried along the whole length of the nerve fibre.
2. Conduction of impulses is slower.

(c) Aqueous humor and vitreous humor:-
Aqueous Humour:- Thin, watery fluid present between the cornea and the lens.
Vitreous Humour:- Transparent gel present between the lens and the retina.

(c) Blind spot and yellow spot:-
Blind Spot:-
1. Blind spot is a spot on the retina present at the point of origin of the optic nerve.
2. Photoreceptor cells are absent from this region.
3. Insensitive to light as both rods and cones are absent.
Yellow Spot:-
1. Yellow spot is a small area on the retina present at the posterior pole of the eye, lateral to the blind spot.
2. Only cones are present in this region.
3. Sensitive to bright light as cones are present.

(f) Cranial nerves and spinal nerves.
Cranial Nerves:-
1. Cranial nerves arise from the brain.
2. 12 pairs of cranial nerves.
Spinal Nerves:-
1. Spinal nerves arise from the spinal cord.
2. 31 pairs of spinal nerves


What is a Nerve

A nerve is a bundle of axons of a large number of neurons in the peripheral nervous system. A nerve is a wire-like structure which transmits nerve impulses in the form of chemical and electrical signals between central nervous system and sensory or effector organs. Each axon in a nerve is wrapped with a connective tissue layer called endoneurium. The axon bundle of the nerve is wrapped by a connective tissue layer called epineurium. A fascicle is a group of neurons. A cross section of a nerve is shown in figure 1.

Figure 1: A Cross-section of a Nerve

Three types of nerves are found based on the direction of transmitting signals within the nervous system. They are sensory nerves, motor nerves, and mixed nerves. Sensory nerves are also known as afferent nerves, and carry information from the sensory receptors into the central nervous system. Motor nerves carry nerve impulses from the central nervous system to the effector organs. Mixed nerves contain both sensory and motor neurons within the same nerve.

Based on the way they are connected to the central nervous system, two types of nerves can be identified as cranial nerves and spinal nerves. The parts of the head are innervated by cranial nerves, which are connected with the brain. Twelve cranial nerves are found in humans. Most of the body parts are innervated by spinal nerves, which are connected to the spinal cord. Nerve fibers can be divided based on their diameter, velocity of the signal conduction, and the myelinated state of the axons as well. The group A nerves have a large diameter. They are myelinated and have a high signal conduction velocity. The group B nerves have a small diameter. They are also myelinated, but their conduction velocity is low. The group C nerves are unmyelinated, with a small diameter, and low conduction velocity. The nerves in the left upper limb of humans are shown in figure 2.

Figure 2: Nerves in the Left Upper Limb


Myelin

Myelin is a whitish, fatty substance that forms a sheath around many vertebrate nerve fibers. Myelin insulates the nerves and permits the rapid transmission of nerve impulses. The white matter of the brain is composed of nerve fibers covered in myelin. It is essential for the proper functioning of the nervous system. It is an outgrowth of a type of glial cell.

In humans, myelination begins early in the 3rd trimester, although little myelin exists in the brain at the time of birth. During infancy, myelination occurs quickly, leading to a child’s fast development, including crawling and walking in the first year. Myelination continues through the adolescent stage of life.

The nerve cell is made up of a cell body and a long, projecting nerve fibre called the axon that is responsible for the transmission of electrical impulses from the cell body to receiving neurons, glands and muscles. Myelin is considered a defining feature of the vertebrates or animals with a spinal column, although similar sheaths have also evolved in some invertebrates. The myelin sheath of nerve fibres was first discovered and described by Rudolf Virchow in 1854.

Structure and Functions of Myelin

Myelin is made by two different types of support cells. In the central nervous system (CNS) the brain and spinal cord cells called oligodendrocytes wrap their branch-like extensions around axons to create a myelin sheath. In the nerves outside of the spinal cord, Schwann cells produce myelin. Regardless of where it is in the nervous system, all myelin performs the same function, enabling efficient transmission of electrical signals.

Myelin is composed of about 40% water and the dry mass is composed of about 80% lipids and 20% protein. The mainly lipid composition of the myelin gives it a white hue, hence the reference to the brain’s “white matter.” The main lipid found in myelin is a glycolipid called galactocerebroside. Other major myelin constituents include myelin basic protein (MBP), proteolipid protein (PLP) and myelin oligodendrocyte glycoprotein (MOG). Within the myelin, there are cross linked hydrocarbon chains composed of sphingomyelin which strengthens the myelin sheath.

The main purpose of a myelin sheath is to increase the speed at which impulses propagate along the myelinated fiber. Along unmyelinated fibers, impulses continuously move as waves, but, in myelinated fibers, they “hop” or propagate by saltatory conduction. Myelin decreases capacitance and increases electrical resistance across the cell membrane (the axolemma).

In the 1870s, French physician Louis-Antoine Ranvier noted that the myelin sheath is discontinuous, covering most of the nerve fiber but with gaps at regular intervals along the axon. Scientists later learned that charged particles called ions can cross the axon only at these myelin gaps, which became known as the “nodes of Ranvier.”

In the 1930s and 1940s, scientists found that this passage of ions helps maintain the electrical signal, allowing it to travel quickly down an axon. The signal appears to “jump” from one node to the next in a process called saltatory conduction.

Myelin Disorders

Loss of myelin is a problem for many CNS disorders, including stroke, spinal cord injury, and, most notably, multiple sclerosis (MS). MS is a chronic, disabling disease of the CNS that affects more than 2.3 million people worldwide. MS results from the accumulation of damage to myelin and the underlying nerve fibers it insulates and protects.

The majority of patients (about 80%) develop relapse-remitting MS, where neurological symptoms occur in episodes, with no deterioration seen between those episodes. Around 10 years after initial onset of the condition, about half of these patients go on to develop progressive neurological deterioration, referred to as secondary progressive MS.

The remaining 20% of MS patients experience a continuous neurological deterioration with no periods where symptoms seem to improve. This is referred to as primary progressive MS.


Feature: My Human Body

Would you like your brain to make new neurons that could help you become a better learner? When it comes to learning new things, what college student wouldn’t want a little more brain power? If research about rats applies to humans, then sustained aerobic exercise (such as running) can increase neurogenesis in the adult brain, and specifically in the hippocampus, a brain structure important for learning temporally and/or spatially complex tasks, as well as memory. Although the research is still at the beginning stages, it suggests that exercise may actually lead to a “smarter” brain. Even if the research results are not ultimately confirmed for humans, though, it can’t hurt to get more aerobic exercise. It is certainly beneficial for your body, if not your brain!


Neuromuscular Junction

Muscle cells innervated by motor neurons that arise from spinal cord. Activation of motor neuron triggers contraction of muscle cell. The axon of motor neurons form synapse on the surface of skeletal muscle cells at the neuromuscular junction.

Motor neurons innervate skeletal muscle cells at neuromuscular junctions.

Neuromusclular junctions are rarely seen in histological samples and require special preparation of muscle tissue. This image shows a single axon from a motor neuron splitting to form synapses on multiple skeletal muscle cells. An axon and the muscle cells it innervates form a motor unit. Some motor neurons innervate one or a few muscle cells whereas other motor neurons can innervate hundreds of muscle cells. Motor axons terminate in a neuromuscular junction on the surface of skeletal muscle cells. The neuromuscular junction occurs at the center of the muscle cell so that action potentials being in the middle of the cell and propagate towards the end of the muscle cell.

Motor neurons form synapses on muscle cells at the neuromuscular junction.

Neuromuscular Junction Synapse

The neuromuscular junction is composed of a pre-synaptic axon terminus and a post-synaptic muscle cell. Upon depolarization of the axon, synaptic vesicles containing the neurotransmitter acetylcholine fuse with the membrane, releasing acetylcholine into the cleft that separates the axon from the skeletal muscle cell. Acetylcholine binds to receptors on the post-synaptic membrane. The acetylcholine receptor is a ligand-gated ion channel that opens after binding acetylcholine. The open channel depolarizes the membrane which can trigger opening of voltage-gated sodium channels. The opening of the sodium channels initiates an action potential that propagates along the cell membrane and into the T-tubules.

The neuromuscular junction contains a synapse between the axon terminus and skeletal muscle cell.

Note the basal lamina that surrounds the muscle cell. The basal lamina is similar to the basement membrane of epithelia. The basal lamina of skeletal muscle cells contains acetylcholinesterase which is an enzyme that digests acetylcholine. Acetylcholinesterase helps limit the duration of each contractile event.


MATERIALS AND METHODS

Transgenic mice overexpressing NGF in the skin under the control of the K14 keratin promoter (Albers et al., 1994) were used. Mice were screened for the presence of the K14-NGF transgene using slot-blot analysis of DNA isolated from tail samples. The background strain (Bl6/C3H) in which NGF-OE mice were generated was used as the control strain.

Nerve fiber counts. Mice were anesthetized and perfused transcardially with 4.0% paraformaldehyde and 2.0% glutaraldehyde in 0.1 m phosphate buffer (PB) (NGF-OE,n = 3 control, n = 3). Segments of the saphenous nerve from the midthigh were post-fixed, washed in PB, dehydrated in graded alcohols, embedded in Spurr’s resin (EM Corp., Chestnut Hill, MA), and cut at 0.7–0.8 nm on an ultramicrotome. Ultrathin sections were stained with lead citrate and uranyl acetate, photographed on an electron microscope (EM) (H7000 Hitachi Ltd., Tokyo, Japan), and the images were assembled into montages. Myelinated and unmyelinated nerve fibers were analyzed with a computer using NIH Image 1.4 software. The entire nerve was analyzed for axon counts of myelinated and unmyelinated fibers, and 150 axons were measured in each group for the axon diameters. NGF is also known to increase the survival of sympathetic neurons (Johnson et al., 1980 Ruit et al., 1990 Davis et al., 1996), and some of these sympathetic neurons become myelinated in NGF-OE mice. Therefore, to determine the contribution of sympathetic fibers to the number of myelinated axons, we surgically removed the upper three lumbar sympathetic ganglia from adult NGF-OE (n = 2) of one side under brief anesthesia of Avertin. The sympathetic contribution to the saphenous nerve has not been reported for the mouse however, in rat, the upper three lumbar sympathetic ganglia contain the sympathetic neurons that project into the saphenous nerve (Baron et al., 1988). In the experiments reported here, there was no visible fusion between the ganglia of both sides. Five days after sympathectomy, mice were reanesthetized and perfused, and sections of the ipsilateral and contralateral saphenous nerve were removed, fixed, cut, and analyzed as described above.

Neurophysiological recordings. An in vitroskin–nerve preparation was used to record from single, functionally identified cutaneous sensory neurons in NGF-OE (n = 8) and control (n = 8) mice (Koltzenburg et al., 1997). The saphenous nerve with its cutaneous innervation territory was removed from the hindlimb, placed corium side up in a tissue bath, and superfused (15 ml/min) with oxygen-saturated, synthetic interstitial fluid containing (in m m ): 123 NaCl, 3.5 KCl, 0.7 MgSO4, 1.7 NaH2PO4, 2.0 CaCl2, 9.5 sodium gluconate, 5.5 glucose, 7.5 sucrose, and 10 HEPES, pH 7.45 ± 0.05, at 32° ±0.5°C. The nerve was desheathed, and extracellular recordings were made from functionally isolated afferent fibers using gold wire electrodes. Action potentials were acquired using a low-noise differential amplifier, stored on a personal computer, and later analyzed with a template-matching program (Forster and Handwerker, 1990).

Fibers were first identified by probing the skin with a glass rod, a stimulus that is known to activate all mechanically sensitive units (Kress et al., 1992 Koltzenburg et al., 1997). The conduction velocity of each mechanosensitive fiber was determined by electrically stimulating the receptive field with supramaximal square wave pulses of 0.1–1.0 msec duration using a Teflon-coated steel electrode (1–5 MΩ impedance, shaft diameter of 300 μm, uninsulated tip diameter of 5–10 μm). Units conducting faster than 10 m/sec were classified as large myelinated Aβ fibers, units conducting between 1.2 and 10 m/sec were classified as thin myelinated Aδ fibers, and units conducting slower than 1.2 m/sec were unmyelinated C fibers (Koltzenburg et al., 1997).

The mechanical threshold was determined using calibrated von Frey filaments (self-constructed from nylon thread of various diameters, tip diameter of 0.8 mm, range of force of 1–362 mN). Mechanical responsiveness was then systematically characterized by using a computer-driven, feedback-controlled stimulator that applied a standard series of ramp and sustained force stimuli (200 msec rise time, 10 sec duration, plateau force of 5–300 mN, interstimulus interval of 60–120 sec). Data were analyzed for 11 sec beginning with the onset of force.

Based on their response pattern to mechanical stimuli, myelinated fibers could be clearly classified in both strains of mice as described previously into four subpopulations (Koltzenburg et al., 1997 Stucky and Koltzenburg, 1997). Because large myelinated fibers tend to be sampled more frequently than thin myelinated or unmyelinated fibers, the proportion of each subtype of fiber encountered within each category (Aβ, Aδ, C) was determined. Large myelinated (Aβ) fibers had low mechanical thresholds and were classified as either slowly adapting (SA) if they responded tonically to sustained force or rapidly adapting (RA) if they responded only at the onset or offset of the force. Thin myelinated (Aδ) fibers were classified as either A fiber mechanonociceptors (AM nociceptors) if they responded tonically to high-intensity force or D-hair receptors if they were activated by very low mechanical force (<1 mN) and responded with high frequency to the onset and offset of force. Unmyelinated C fibers and AM nociceptors were tested first for their response properties to mechanical stimuli and then further classified by their response properties to noxious thermal stimuli.

Heat sensitivity was determined by applying a linear heat ramp (32–47°C in 15 sec) with a feedback-controlled lamp focused through the translucent bottom of the tissue bath onto the epidermal side of the skin. The resulting temperature was measured at the corium side (top) of the skin by a thermocouple inserted into the skin. The temperature at the corium side of the skin corresponds to an ∼5 C° higher temperature on the epidermal surface (Reeh, 1986). Therefore, the maximal heat stimulus given to each nociceptor was 52°C, a temperature that has been shown to maximally activate heat-sensitive AM and C fiber nociceptors in rodents without damaging the skin preparation (Reeh, 1986 Koltzenburg et al., 1997). Cold stimuli were given by isolating the receptive field with a metal ring and applying a bolus of ice-cold synthetic interstitial fluid for 10 sec, resulting in a minimum skin temperature of 4–8°C. Care was taken to avoid mechanical stimulation during the thermal stimuli. A fiber was considered to be heat- or cold-sensitive if three or more action potentials were evoked during the stimulation.

Statistical analysis. All values are given as mean ± SEM or as median and interquartile range of the 25th and 75th percentile. Parametric and nonparametric statistical tests were performed as appropriate after fulfillment of all necessary prerequisites using the Statistica software package (StatSoft, Tulsa, OK).


Discussion

Although various treatments have been shown to promote axonal regeneration in the spinal cord, relatively little information is available on the precision with which regenerating axons reestablish projections to their original target areas. The DR crush model is useful for such studies, because specific classes of sensory fibers can be labeled in peripheral nerves, allowing an anatomical determination of their spinal projections. Using this model, we found that two treatments that promote robust anatomical and functional regeneration differed dramatically in terms of the specificity of this regeneration. Different classes of sensory axons projected diffusely through the dorsal columns and dorsal horn when regeneration was promoted by the soluble Nogo receptor peptide, sNgR. In contrast, a 2-wk regimen of systemic artemin treatment led to the reestablishment of projections of three different classes of sensory axons to their appropriate topographic locations within the cord.

One successful approach to stimulating regeneration of axons in the CNS has been to block the inhibition to growth caused by oligodendrocytes. Many previous studies have used a contusion model of SCI, blocking the inhibition caused by CNS myelin with antibodies or peptides directed against Nogo ligands or the Nogo receptor NgR1. (See ref. 18 for a discussion of these studies.) Several groups have reported significant improvements in spinal cord function through blockage of this inhibition, but determining the degree to which normal spinal circuitry is reestablished has proven difficult. We found that interfering with Nogo receptor signaling with sNgR also promoted robust regeneration of myelinated sensory axons into the cord, with restoration of synaptic function and behavioral recovery of the affected limb (8).

In the present study, we assessed the degree to which regeneration promoted by sNgR is topographically specific. Although myelinated axons supplying both muscle and skin regenerated through the inhibitory barrier of the DREZ, both classes of axons projected throughout the dorsal white matter and superficial laminae of the dorsal horn. This projection pattern is distinctly different from normal and also from the undamaged afferents on the contralateral side of the cord, where these afferents project only within laminae III–VI of the dorsal horn. This indicates that reduction of myelin inhibition with sNgR treatment allows robust regeneration but with an inappropriate anatomical distribution, suggesting that some inhibition might be required to direct growth to the appropriate regions of the spinal cord.

Regeneration of sensory axons after DR crush also has been reported using various neurotrophic factors, including nerve growth factor (NGF), neurotrophin 3 (NT3), and glial-derived neurotrophic factor (GDNF). Expression of NGF or fibroblast growth factor by viral transfection of cells in the dorsal horn was found to promote massive regeneration of unmyelinated CGRP + axons, restoring nociceptive sensation in the affected limb (19). These projections are not limited to the most dorsal laminae of the gray matter, however rather, they occupy a large fraction of the entire dorsal horn. Intrathecal infusions of NT3 or GDNF promote the regeneration of large-diameter sensory afferents through the DREZ and into the dorsal horn gray matter, restoring functional synaptic transmission with spinal neurons and leading to significant behavioral improvement (15). However, these regenerated afferents occupy an aberrant position in lamina IIo and grow along the pial surface, abnormal locations for this class of sensory axon (16, 17).

Similar to the effects of intrathecal GDNF or NT3 treatment, systemic injections of ART promote regeneration of sensory axons through the DREZ and into the gray matter of the spinal cord despite the presence of myelin (9). This finding might be explained by a dynamic inhibitory influence of myelin. Intact myelin and myelin debris produced by damaged axons have different inhibitory properties (18). Inhibition mediated by myelin produced by oligodendrocytes increases during the first week after DR crush injury as myelin degenerates (5). Myelin debris might expose an increased number of epitopes that bind receptors mediating growth cone collapse. Indeed, in the absence of DR injury, transplanted primary afferent neurons from mice into rat DRG grow past the DREZ and into the spinal cord within an environment of intact myelin. Growth is inhibited when DRs are crushed at the time of transplantation (5). Ramer et al. (16) postulated that with the support of certain growth factors, such as NT3, regenerating sensory axons might enter the spinal cord before inhibition mediated by myelin debris is increased. Our data support this hypothesis 2 wk after injury, regenerated axons were found in the gray matter in more ventral locations with ART treatment than with sNgR treatment. Thus, rapid growth promoted by growth factors such as NT3, GDNF, and ART might promote growth of sensory axons through the DREZ before inhibitory mechanisms are up-regulated.

Unlike the effects of intrathecal NGF or NT3 treatment, however, systemic administration of ART after DR crush promotes regeneration of both myelinated and unmyelinated sensory neurons. This positive result is confounded by the expression patterns of GFRα3 in neuronal subpopulations in the DRG. Based on immunohistochemical staining, GFRα3 is expressed largely on unmyelinated neurons. Variable expression on myelinated neurons, ranging from 0 to 14% of the total number of myelinated neurons in the DRG, has been reported (9, 12, 13). Behavioral recovery attributed to regeneration of unmyelinated axons is more robust than that by myelinated axons, consistent with a larger fraction of unmyelinated neurons expressing GFRα3 (9). Nonetheless, substantial regeneration of myelinated DRG neurons is observed with administration of ART. It is possible that these neurons express a low level of GFRα3 that cannot be detected with standard immunohistochemical techniques.

In contrast to the results with other treatments that promote regeneration, we report here that regeneration of sensory axons with systemic ART is topographically specific. We characterized the projection pattern of regenerated myelinated muscle and cutaneous afferents and found that these projections are localized to the appropriate regions of the spinal cord, avoiding areas that are normally occupied by unmyelinated nociceptive afferents. Furthermore, regenerated CGRP-expressing nociceptive axons project to the dorsal laminae of the dorsal horn, in a distribution similar to that of undamaged nociceptive axons. Because the effects of myelin proteins are unlikely to be interrupted with ART treatment, we hypothesize that myelin proteins might contribute to the guidance of growing axons toward appropriate regions of the gray matter. Data from other experiments are consistent with this hypothesis. For example, the rostrocaudal projections of regenerating axons with NT3 and GDNF treatment avoid the white matter, instead projecting rostrocaudally in the gray matter, where the influence of these proteins is less significant (15, 16). Thus, the presence of intact myelin at early time points after DR injury might act in concert with other guidance cues in the cord to allow a short window of opportunity for axons to regenerate before inhibition is increased by myelin debris and a glial scar develops.

The question arises as to why sensory regeneration promoted by systemic treatment with ART is more specific than that promoted by other neurotrophic factors. One possibility is that the other factors have been tested either by infusion into the cerebrospinal fluid (15–17) or by viral infection directly in the spinal cord (19, 20). Direct neurotropic effects of these neurotrophic factors within the spinal cord may overpower molecular cues that could otherwise guide growing axons back to their original target areas in the cord. In contrast, we found that systemic application of ART, whether injected intradermally, subcutaneously, or into peripheral nerves, results in increased ART levels within the DRG, but not within the spinal cord itself. ART contains a heparin-binding site that could possibly mediate its binding to the extracellular matrix within the DRG. Heparin binding is a requirement for the optimal activation of GFRα3 (21). Thus, stable positioning of ART near these receptors might maximize interactions, especially on the larger neurons that do not express high levels of GFRα3. Therefore, systemic ART would be capable of up-regulating growth programs directly in sensory neurons within the DRG without interfering with guidance cues in the cord. If this hypothesis were correct, it could motivate a change in strategies for promoting regeneration in the CNS. Rather than attempting to overcome the inhibitory environment within the area of axon regeneration, effective treatments should be developed to stimulate growth programs in neuronal cell bodies. Recent experiments demonstrating robust axon growth via stimulation of the mTOR pathway in retinal ganglion cells and of Arg1 in sensory neurons suggest that these approaches can be successful (22, 23), although the specificity of the regeneration that they promote remains unknown.

To the best of our knowledge, no other agent or program of treatment has been shown to promote topographically specific regeneration of axons in the adult mammalian CNS after DR injury. Our findings suggest that molecular cues capable of directing the growth of regenerating sensory axons to their targets are present in the adult mammalian spinal cord. Treatment with artemin may thus enable functional restoration of specific sensory input to the spinal cord after brachial plexus injury. A more general implication of our findings is that other guidance cues within the gray matter also may be available to guide the regeneration of other classes of spinal axons, such as those damaged in contusion injuries. Although artemin is unlikely to promote regeneration of these axons, which do not express known artemin receptors, nevertheless it might be possible to develop generalized strategies for promoting specific regeneration of these axons while keeping intact the inhibitory influences in the spinal cord that may guide appropriate targeting of regenerated axons.


Methods and Materials

This study protocol was approved by the Stanford University Administrative Panel on Laboratory Animal Care.

Dorsal Root Isolation

Adult male Sprague-Dawley rats (weight 200–350 g, n = 23) were anesthetized using 1–2% enflurane and 70% N 2 O in oxygen. After induction, the trachea was exposed, cannulated, and connected to a semiclosed circle anesthesia circuit for continued maintenance of anesthesia. End-tidal carbon dioxide and respiratory rates were continuously monitored. With a surgical microscope, a laminectomy extending from the T12 thoracic vertebra to the L6 lumbar vertebra followed by a long dural incision was used to expose the dorsal surface of the spinal cord and the dorsal roots. Individual lumbar dorsal roots were dissected from adjacent roots and cut proximally near their point of entry into the spinal cord and distally near their exit from the spinal canal. Each isolated root was transferred immediately to an artificial cerebrospinal fluid (aCSF) solution with the following composition (mM): NaCL 123, KCl 5, CaCl 2 2, MgSO 4 1.3, NaHCO 3 26, NaH 2 PO 4 1.2, and glucose 10, bubbled continuously with 95% O 2 /5% CO sub 2. As measured in the perfusion chamber at 37 degrees C +/-0.3 degree C, aCSF pH was in the range of 7.35–7.40 and P CO 2 was 35–40 mmHg.

Stimulation and Recording

Each root was placed in a Teflon perfusion chamber (internal volume 1.5 ml) for stimulation and recording Figure 1. The ends of each root were trimmed, and the perineurium was left intact. The proximal end of the root (2–3 mm) was led through a slotted partition into a recording compartment, while a suction-type stimulating electrode was attached to the distal end within the perfusion chamber. The slot in the partition was sealed with silicone-based vacuum grease, and the aCSF in that compartment was covered with mineral oil. Using standard single-fiber microdissection techniques, the proximal end of the root was progressively divided into small fascicles, each typically containing one to three electrophysiologically distinguishable axons. Action potentials were recorded using chlorided silver wire electrodes.

Figure 1. The recording and perfusion chamber arrangement. ACSF in = artificial cerebrospinal fluid inlet AMP = preamplifier and amplifier/signal conditioning circuits connected to an oscilloscope and computer data acquisition system Stim. = isolated constant voltage nerve stimulator Temp. control = automated temperature controller and monitor.

Figure 1. The recording and perfusion chamber arrangement. ACSF in = artificial cerebrospinal fluid inlet AMP = preamplifier and amplifier/signal conditioning circuits connected to an oscilloscope and computer data acquisition system Stim. = isolated constant voltage nerve stimulator Temp. control = automated temperature controller and monitor.

Supramaximal (1.5 x threshold) constant voltage stimuli (0.2 ms in duration at 0.3 Hz) were delivered to the intact distal end of the isolated root while single-fiber action potentials in a proximal fascicle were amplified, displayed on a digital storage oscilloscope, and recorded for computer analysis (Figure 2). The stimulus parameters were selected to minimize activity-dependent changes in axon properties. The length of the root from the tip of the stimulating electrode to the recording electrode was measured to calculate conduction velocity from single axon conduction latency measurements. Latencies were measured with an adjusted resolution ranging from 0.1 micro second for the shortest latencies to 5 micro second for the longest. The length of the root exposed to the perfusate was measured between the tip of the stimulating electrode and the partition separating the recording and perfusion compartments (average length+/-SD 19+/-3.8 mm).

Figure 2. An example of single-fiber action potentials recorded simultaneously in two unmyelinated dorsal root axons (conduction velocities 0.72 and 0.37 m/s). The stimulus artifact followed by a compound action potential can be seen at the beginning (left side) of the trace. Time and voltage calibrations are shown in the figure.

Figure 2. An example of single-fiber action potentials recorded simultaneously in two unmyelinated dorsal root axons (conduction velocities 0.72 and 0.37 m/s). The stimulus artifact followed by a compound action potential can be seen at the beginning (left side) of the trace. Time and voltage calibrations are shown in the figure.

The exposed portion of the root was continuously superfused with aCSF at 37 degrees+/-0.3 degree C and a flow rate of approximately 8 ml/min. In pilot experiments, single axon recordings in this preparation were stable (< 5% change in conduction velocity or action potential amplitude) for as long as 6 h.

Drug Exposure Protocol

Each single fiber preparation was tested for stability under control conditions for 15–30 min. After control measurements, the aCSF perfusate was switched to one containing a low concentration of lidocaine (150 or 260 micro Meter Astra, Westboro, MA), selected from pilot study data to be at or below the EC 50 level. Latency was measured at 1-min intervals until the unit failed or for at least 15 min to ensure steady-state conditions. Maximum conduction velocity slowing typically occurred within 5–10 min of lidocaine exposure. Axons unblocked at the lowest concentration of lidocaine were tested at higher lidocaine concentrations (e.g., 260 then 540 micro Meter) for additional 15-min intervals. Each dorsal root was exposed to this sequence of lidocaine concentrations only once. Data were accepted from those axons demonstrating complete recovery (latency and amplitude within 5% of control values) after return to control aCSF.

Vagus Nerve Experiments

In a separate series of experiments, cervical portions of vagus nerves were removed from 14 adult male rats using the anesthetic technique previously described (see methods, dorsal root isolation). Similarly, in vitro perfusion, stimulation, single-fiber recording, and drug exposure were accomplished in the manner previously described for the dorsal root experiments.

The vagus nerve was selected because of the availability of comparable data in other species and because it has been used in most previous single fiber and whole nerve studies.

Data Analysis and Statistics

Fisher's exact test was used to determine the statistical significance of differences in the incidence of conduction block between dorsal root and vagal axons, between myelinated (CV > 3 m/s) and unmyelinated (CV < 1.4 m/s) axons, and between long (> 20 mm) and short (< 15 mm) exposure lengths. Multiple comparisons were evaluated using analysis of variance and Bonferroni's t test. Estimates of EC 50 were calculated from a least-squares fit to a sigmoid function bounded by 0% and 100%. EC 50 95% confidence intervals were estimated from this model.


PATHOBIOLOGY OF NONMYELINATING SCHWANN CELLS

Schwann Cell Response to Wallerian Demyelination

Transection of a nerve fiber initiates Wallerian degeneration of the distal stump (see Jessen and Mirsky, this issue). The mechanisms of axonal degeneration are intrinsic to the axon, and not dependent on other cell types (Glass et al., 1993 ). However, axotomy sets in motion a series of responses in the Schwann cells of the injured fibers as well as in some of the Schwann cells of neighboring fibers. These responses differ between myelinating and nonmyelinating fibers both are discussed here because the loss of the axon converts each previously myelinating Schwann cells into an NMSC at least until regeneration of a “myelinatable” axon into that Schwann cell occurs and new myelination supervenes.

Responses of Myelinating Schwann Cells

After axonal transection, the axons in the distal stump pass through a latent period during which they appear relatively normal and conduct effectively, when locally stimulated. This latent period is abruptly terminated by conversion of the axoplasm to granular amorphous debris, probably as a consequence of increased calcium entry and release from intra-axonal stores. As the axon disappears, the myelin sheath undergoes longitudinal segmentation into the characteristic ovoids. Viewed in transverse section, the sheath begins to collapse and fold inward. With time the individual myelin ovoids shorten.

By the second day after transection (in the rat), the process of Schwann cell proliferation commences (Bradley and Asbury, 1970 Clemence et al., 1989 Oaklander et al., 1987 Pellegrino et al., 1986 Weinberg and Spencer, 1975 ). The nucleus of the now-denervated myelin-bearing Schwann cell comes to lie in approximate center of the fiber, as viewed both longitudinally and transversely. This reflects the fact that the myelin sheath is first interrupted in this central region of the old internode, forming two ovoids, one on either side of the Schwann cell nucleus. The first Schwann cell division results in two daughter cells one peripheral to the other (Stoll et al., 1989 ). Over the next few days, these daughter Schwann cells continue to divide. Schwann cells in Schwann cells peaks on Days 3–4 in the rat and the mouse. The total number of divisions, and thus the number of daughter Schwann cells, appears to be influenced by the original internodal distance, so that longer internodes have more daughters. The daughter Schwann cells remain within the basal lamina of the original fiber (Stoll et al., 1989 ) this contrasts with their behavior following demyelination, as described below. The resulting longitudinal chain of Schwann cells within the basal lamina forms the classical Bungner band. This arrangement appears particularly favorable to the support of regenerating sprouts (Taniuchi et al., 1988 ).

Responses of Schwann Cells of Motor Nerve Terminals

Terminal SCs, like their counterparts within nerves, undergo marked changes in response to damage to their axons. As within the peripheral nerves, there is a latent period following nerve section. In the case of NMJs, this latency is directly dependent upon the distance between the nerve lesion and the motor endplate (Miledi and Slater, 1970 ). In most cases, transmission across the synapse ceases with 24 h (Miledi and Slater, 1968 ), after which the nerve terminal begins to show frank signs of Wallerian degeneration, including mitochondrial changes. Electron microscopy shows that SC processes begin to invade the synaptic cleft, separating the nerve terminal from the muscle fiber the SCs at the junction participate in the removal of the axon debris by phagocytosis (Miledi and Slater, 1970 ). In contrast to the reports for NMSCs along the nerve, proliferation does not seem to be an early feature of the response of tSCs, and the cell number does not begin to change appreciably until the return of the nerve (Love and Thompson, 1998 ), implying that the major impetus for mitosis in these cells comes from an axonal mitogen, namely NRG. However, Hess et al. ( 2007 ) suggest a role for muscle NT3 in determining tSC number. Schwann cells move to occupy the terminal gutters of the endplate and appose the acetylcholine receptors there (Letinsky et al., 1976 Miledi and Slater, 1970 ). However, with increasing time after denervation, the occupancy of the synaptic site by these SC processes declines. In the frog, the tSCs at denervated endplates come to release quanta of acetylcholine that produce potential fluctuations in the muscle fibers that resemble the miniature endplate potentials (MEPPs) in innervated muscle (Birks et al., 1960b ). Katz and colleagues found this phenomenon when they denervated muscles in attempt to show that MEPPs were generated by acetylcholine released from the nerve terminal. They found that the MEPPs did disappear, but then reappeared days later as the SCs began to release quanta of acetylcholine. A similar phenomenon is seen only rarely at NMJs in mammals, apparently because, in contrast to the frog, the SCs remain in “synaptic” apposition to the denervated endplate for only a short period of time (Miledi and Slater, 1968 ). It is interesting to note that mammalian SCs exposed to muscle-conditioned medium come to express choline acetyltransferase, a synthetic enzyme for acetylcholine (Brockes et al., 1979 ).

Surprisingly, we still have meager information about the changes in gene expression that occur in tSCs following denervation. One would like to know to what extent the tSCs of the nerve resemble those of the NMSCs of the peripheral nerves both before and after denervation. One of the most robust immunological markers of the “reactive” state of tSCs is the intermediate filament protein nestin (Kang et al., 2007 ), which is upregulated within 1–2 days of denervation, even as the most commonly used immunological marker for SCs, S100B, declines in expression (Perez and Moore, 1968 ). Reactive tSCs upregulate the low affinity nerve growth factor receptor (p75) (Reynolds and Woolf, 1992 ). Most likely, if the cells are like those along the nerve, they also upregulate expression of NRG and NRG receptors (Carroll et al., 1997 ). Indeed, aspects of the reactive state of tSCs are induced in these cells by extrinsic application of NRG or by induction with tSCs of transgenic expression of a constitutively active NRG receptor (Hayworth et al., 2006 Heuser and Reese, 1973 Trachtenberg and Thompson, 1997 ), suggesting that autocrine signaling induced by loss of nerve contact may play a role in the responses of these cells.

One of the most remarkable responses of tSCs to denervation, a response first described by Reynolds and Woolf ( 1992 ), although alluded to in earlier reports (Letinsky et al., 1976 ), is the profusion of processes of these cells from the denervated endplate. These processes grow steadily with time and are quite dynamic (Kang et al., 2007 ) some of these processes can reach lengths of up to 800 μm. This process growth occurs in concert with the expression by the cells of the protein GAP-43 (Mehta et al., 1993 ). These processes are then retracted upon reinnervation of endplates by their motor axons (Reynolds and Woolf, 1992 Son and Thompson, 1995a ). Actually, this growth response is probably not fundamentally different from those of many SCs in the peripheral nerves: these SCs in the nerve extend processes from the cut ends of nerves (Reynolds and Woolf, 1992 Son and Thompson, 1995a ), and most likely extend processes up and down the endoneurial sheaths within the nerves (Cook and Bishop, 2004 ). These processes have been shown to act as a substrate for the extension of regenerating and of sprouting axons (Son and Thompson, 1995a , b ). Gutmann and Young ( 1944 ) extended earlier studies of Cajal and his students and furnished a rather complete study of the reinnervation of muscle fibers in mammals. They showed that regenerating axons commonly overshoot the sites of their former synaptic contacts, often extending axons (they called them “escaped fibers”) to innervate adjacent muscle fibers. This phenomenon of escaped fibers is explained by the pathways laid down by the growth of the tSCs during the period of denervation. Since the tSCs processes often grow to link adjacent endplates in the denervated muscle (or form these links during the early stages of reinnervation, see Love et al., 2003 ), a regenerating axon returning to one endplate has a preformed pathway to regenerate and reinnervate adjacent endplates. This kind of growth explains why motor axons change the distribution of the fibers they innervate upon regeneration (see Karpati and Engel, 1968 ). Following simple nerve damage, such as a nerve crush, axons commonly regenerate by growing quickly down their endoneurial tubes and return to the endplates to which these tubes guide them (see Nguyen et al., 2002 ). However, following more severe damage, such as a nerve cut, the axons likely cross the lesion site asynchronously, grow down these tubes, and arrive within muscle regions which have few other axons. In such cases, growth of escaped fibers provides each successful axon a means of reinnervating as many muscle fibers as possible. A similar phenomenon also occurs after damage to only a portion of the nerve supply to a muscle. Such “partial denervation” leads to a mixture of denervated and innervated fibers in the muscle. The response to this is “sprouting” of the remaining axons (Brown et al., 1981 ). This sprouting is also elicited by growth of processes of tSCs that occurs from the denervated endplates (Son and Thompson, 1995a ) (see Fig. 3). Thus, this “reactive” property of tSCs provides for restoring as much innervation as possible to denervated muscles as quickly as possible. A similar reinnervation-promoting role of the extension of processes occurs at sites of nerve damage (Son and Thompson, 1995b ).

Processes extended by terminal SCs guide the growth of nerve sprouts between endplates on adjacent muscle fibers. The rat soleus muscle shown here was partially denervated 3 days before these images were made. Two immunostains were applied. Panel A: The stain for neurofilament. Panel B: The stain for nestin, an intermediate filament protein that is not expressed in tSCs at innervated junctions but is expressed by them upon denervation, that is, when they become “reactive” and extend processes. Panel C is a merged image of A and B. Two endplates are present and are labeled 1 and 2 in each panel. Panel A shows that endplate 1 is innervated by an axon (arrow) that enters from below. There is no nestin label associated with this axon or with most of the junction. However, the SCs above endplate 2 are labeled by nestin, suggesting it was denervated the absence of an axon stained with neurofilament in the SCs in the old endoneurial sheath (arrowhead in panel B) also reactive and labeled with nestin confirms this denervation. However, endplate 2 has been innervated by a sprout (double arrowhead in A) that has grown from endplate 1 to endplate 2. This sprout is associated with a process that itself is nestin immunoreactive and therefore derived from the tSCs on endplate 2. Thus, the sprout appears to have grown from endplate 1 to endplate 2 by following SC processes that have grown from endplate 2. Modified from Son and Thompson ( 1995a ). Scale Bar 10 μm.

Are SCs actually required to guide this kind of nerve growth? Two sets of observations suggest that they may be. In neonatal animals, tSCs frequently undergo apoptosis in response to denervation (see above and Trachtenberg and Thompson, 1996 ). As this would be expected to remove the source of the SC processes in the muscle, one would expect a reduction in the ability of neurons in these animals to sprout, and such is in fact reported (Lubischer and Thompson, 1999 ). Also interesting is the observation that the SCs within the endoneurial tubes distal to the site of nerve lesion begin to disappear if reinnervation is delayed and such delays in reinnervation lead to much poorer regeneration (Sulaiman and Gordon, 2000 ).

tSCs occupy a precarious position at the nerve terminal. They apparently adhere to the nerve terminal very tightly, yet unlike their NMSCs in the nerve, they do not completely wrap the nerve terminals: the synaptic side of the terminal is apposed to the muscle fiber rather than to SCs. If tSCs penetrated into the space between the nerve and the muscle fibers and wrapped the terminals as NMSCs do axons in the nerve, then the result would be failure of neuromuscular transmission. Somehow, the tSCs must be excluded from the synaptic cleft. One obvious way to accomplish this would be for the nerve terminal to have a greater adhesion for the muscle fiber than the SC. Another means would be to repel the SCs from the synaptic cleft. Sanes and his collaborators have found evidence for the latter mechanism. Laminin β-2 is a specialized component of the so-called “synaptic basal lamina”—the basal lamina between the muscle fiber and nerve terminal (Sanes et al., 1990 ). This laminin-β-2 is absent from the basal lamina that surrounds the SCs yet connects to the synaptic basal lamina at the edges of the NMJ. Genetic deletion of laminin β-2 results in animals that have a progressively weaker neuromuscular synaptic transmission (Noakes et al., 1995 ). This weakness is correlated with a progressive intrusion of SCs into the synaptic cleft (Miner et al., 2006 Patton et al., 1998 ). Tissue culture substrates coated with laminin-β-2 are nonadherent for SCs and their processes (Patton et al., 1998 ). This presents a rather compelling picture of how the distribution of SC processes is regulated at the junction. However, the role of laminin-β-2 is likely to be more complicated as it has recently been shown to be important in the alignment of presynaptic active zones with the tips of the postsynaptic folds and their high density of acetylcholine receptors (Nishimune et al., 2004 ), and the failure of this alignment may generate neuromuscular weakness that then indirectly leads to SC intrusion.

Although tSCs are reported to penetrate the synaptic cleft after denervation (see above) and after damage to muscle fibers (Jirmanova, 1975 ), they may also play a role in neuromuscular pathology. Human patients with myasthenia gravis can be treated with anticholinesterases in attempt to increase the transmission at damaged NMJs by slowing the breakdown of acetylcholine released from the nerve terminal. Such patients often present muscle biopsies that show evidence of Schwann cell intrusion between nerve terminals and muscle fibers. Similar intrusion of SC processes is seen in experimental animals treated with anticholinesterases (Engel et al., 1973 ). Interestingly, Feng et al. ( 1999 ) have shown that a genetic deletion of a collagen isoform in mouse results in the lack of acetylcholinesterase at their junctions. Although the early response to this deletion is damage to the muscle fiber at synapses, presumably from the excess of neurotransmitter, with time these animals recover at least partially and the recovery is characterized by the extension of tSC processes into the synaptic clefts of the NMJs. This intrusion appears to compensate for the loss of cholinesterase by reducing the area of synaptic apposition between nerve terminal and muscle fibers. These observations suggest that SCs may monitor synaptic transmission between the nerve terminal and muscle fiber and, at least in extreme circumstances, make adjustments in the area of contact.

Responses of Remak Schwann Cells

Remak Schwann cells respond to denervation by entering the cell cycle in a fashion analogous to that of myelinated fibers. Intriguingly, they also proliferate in response to degeneration of neighboring myelinated fibers (Murinson and Griffin, 2004 ). This was shown by an experiment in which L5 ventral root, which contains very few unmyelinated fibers, was transected. As a result, the axons that underwent Wallerian degeneration in the sciatic nerve were restricted to a group of myelinated motor fibers. By Day 3 after the ventral root transection, the Schwann cells of the degenerating motor fibers had entered the cell cycle, as expected from the foregoing. However, the number of their Schwann cells incorporating 3 H-thymidine was greatly exceeded by the number of Remak Schwann cells entering the cell cycle. Because almost no unmyelinated axons were injured yet their Schwann cells proliferated abundantly, this observation raises the possibility that a diffusible Schwann cell mitogen is released into the endoneurial space when myelinated fibers degenerate. In contrast to Remak Schwann cells, the myelin-maintaining Schwann cells ensheathing uninjured axons never entered the cell cycle in this model (Murinson and Griffin, 2004 ).

Growth Factor Production by Denervated Schwann Cells

In the early stages after axotomy, the denervated Schwann cells produce increased amounts of growth factors. Recent data have shown that different Schwann cell populations produce different growth factors (Murinson et al., 2005a ). For example, after transection of the ventral root the myelinated motor fibers produce pleotrophin and GDNF, but not members of the neurotrophin family, such as NGF or BDNF. In contrast, denervated dorsal root fibers produce NGF, BDNF, NT3, and GDNF but no pleotrophin. Surprisingly, these patterns were at least partly maintained in transplantation studies. In these models, reinnervation of Schwann cells from sensory nerves were reinnervated by motor axons, and the converse. For example, sensory nerve grafts were placed into the transected femoral motor nerve and the motor axons allowed to regenerate through the graft. The motor branch was then transected above the graft. The Schwann cells degenerating within the graft—Schwann cells which had originally ensheathed sensory fibers but more recently had ensheathed motor axons—expressed a predominantly “sensory pattern” of growth factors when denervated. When the converse experiment, transplanting ventral roots into the femoral sensory system, was done, transection of the sensory nerve resulted in a predominantly “motor” pattern of growth factor expression within the graft (Hoke et al., 2006 ).

Death of Denervated Schwann Cells

Denervated Schwann cells develop heterochromatic chromatin and atrophic cell processes, undergo apoptosis, and die, leaving only a serpentine basal lamina as an indicator of their earlier presence. After a few months, this basal lamina fragments and disappears. This sequence has been extensively documented in neonatal rodent nerves, in which Schwann cells undergo prompt apoptotic death in the nerve distal to axotomy (Grinspan et al., 1996 ). Some reports suggest that no comparable death occurs in older animals (Grinspan et al., 1996 ). Other reports document extensive Schwann cell death and loss after prolonged denervation in the adult animals. The factors responsible for the differences in these data sets and the resulting conclusions include differences in timing and the expression of autocrine survival factors (Meier et al., 1999 ). In neonatal animals denervated Schwann cells can die within 3 days of denervation, and they are largely gone within 1 month (Ebenezer et al., 2007 Grinspan et al., 1996 Trachtenberg and Thompson, 1996 ). In adults Schwann cell death occurs much later, usually after several months. Studies of adult animals that followed Schwann cells for only a few after axotomy found little death, whereas studies in rodents and in man that extended for several months of denervation (Ebenezer et al., 2007 Pellegrino et al., 1986 Sulaiman et al., 2002 ) identified extensive Schwann cell loss.

Schwann cell expression of p75 is upregulated in denervated Schwann cells and falls with time after axotomy. Signaling through p75 has been shown to mediate Schwann cell death in cultured Schwann cells, and presumptive evidence suggests that NGF might trigger death of denervated Schwann cells (Ferri and Bisby, 1999 Petratos et al., 2003 Syroid et al., 2000 ). If p75-dependent mechanism is important in the death of denervated Schwann cells, then other p75 ligands must be capable of initiating the death process in Schwann cells. This conclusion is based on the fact that Schwann cells in transected ventral roots and motor fibers, in which NGF production is scanty, are lost as rapidly as those in transected dorsal roots. Survival of noninnervated Schwann cells can be prolonged by neurotrophin-3 and insulin-like growth factor (Meier et al., 1999 ).

Schwann Cell Responses in Demyelination

The causes of demyelination include immune-mediated, inflammatory heritable, and toxic etiologies, but many of the Schwann cell responses are shared (see Scherer and Wrabetz, this issue). A fundamental response is proliferation of Schwann cells. The Schwann cells of demyelinating fibers divide to form redundant nonmyelinating daughter cells along the demyelinated internode. The perikayal region of the parent Schwann cell can enter S phase at a time when the myelin looks relatively undamaged (Stoll et al., 1989 ), but entering S phase appears to represent a commitment to subsequent demyelination. This conclusion came from experiments that use local application to rat sciatic nerves of lysolecithin, to produce focal demyelination of the underlying internodes. Pulsed 3 H-thymidine was taken up by the Schwann cell nuclei of myelinated internodes in the first days after application, occasionally at a stage when only mild myelin vesiculation was present (Griffin et al., 1990 ). However, 2 weeks after administration of the marker there were no labeled nuclei within myelinated internodes. Rather, by Day 5–7 all of the labeled myelin-forming Schwann cells “amputated” their myelin sheaths. The anucleate Schwann cell fragments and myelin remnants shed by this amputation were taken up by macrophages. Interestingly, application of the Schwann cell mitogen neuregulin I Type III to myelinating cultures can produce demyelination (Harrisingh et al., 2004 Zanazzi et al., 2001 ). Taken together, these data suggest that a myelinating Schwann cell that is “driven” to enter the cell cycle is obligated to amputate its myelin sheath and demyelinate.

Demyelination of long internodes triggers several cycles of Schwann cell division. Some of the daughter Schwann cells leave the basal lamina formed by the original Schwann cell but remain overlying the demyelinated segment, often forming a circlet of Schwann cells around the fiber. Some of these daughter Schwann cells successfully ensheath segments of the demyelinated internode and form new short remyelinated internodes. With time, some of these new internodes can lengthen and “evict” neighboring short internodes. A pathological hallmark of remyelinated internodes is the presence of multiple short, thin myelin sheaths of irregular length. Demyelination also triggers proliferation of neighboring Remak Schwann cells (Griffin et al., 1987 ), in a fashion similar to that seen in Wallerian degneration (Murinson et al., 2005a ). The morphological sequence that some of these daughters of Remak Schwann cells can remyelinate nearby demyelinated internodes.

In disorders in which demyelination occurs once or twice and successful remyelination supervenes, the circlet of supernumerary Schwann cells around the old demyelinated segment die and disappear. In other disorders, particularly models characterized by recurrent demyelination and remyelination, the supernumerary Schwann cells remain and can become extensive, forming rings of Schwann cells that are termed onion bulbs. In short-term experimental models of demyelination, the Schwann cells forming onion bulbs are not innervated, but in heritable demyelinating neuropathies and occasionally in inflammatory nerve diseases they are found to ensheath unmyelinated axons. The origin of these axons is not resolved. They have been hypothesized to represent collateral sprouts arising during the period of demyelination. Alternatively, they could represent pre-existing Remak fibers in which the Schwann cells have been “attracted” by an unknown signal to migrate and overlie demyelinated segments.

This latter mechanism may sound implausible, but it has been suggested by observations on a model of paranodal demyelination produced by the neurotoxin 3, 3″-iminodipropionitrile (IDPN). Systemic administration of IDPN results in impaired neurofilament transport and the formation of large neurofilamentous axonal swellings beginning just beyond the initial segment and occupying the first several internodes (Griffin et al., 1987 ) (see Fig. 4). In rodents, large myelinated axons can double their perimeter within 3 days. The number of lamellae in the middle of the internode remains unchanged, but the myelin sheath attachment sites “slip” into the old internodes, so that the length of the internode is reduced. As a consequence the paranodes are demyelinated (see Fig. 4), but true segmental demyelination—demyelination of a whole internode—does not occur, and the myelin-forming internodal Schwann cells never enter the cell cycle. However, within the first few days the demyelinated paranodes come to be ensheathed by new Schwann cells. These cells go on to form new short intercalated internodes in the old paranode (Griffin et al., 1987 ). One source of these new Schwann cells appears to be neighboring unmyelinated fibers. These Remak Schwann cells enter the cell cycle as the paranodal demyelination develops (see Fig. 4). In addition, the perikayal region of the nearby Remak Schwann cells appear to migrate to the demyelinated paranode, “dragging” the axonal segments of the Remak fiber along with it. The paranodal remyelination in the IDPN model could reflect asymmetrical division of the Remak Schwann cell.

Effects on Schwann cells of the neurofilamentous axonal swelling produced by IDPN. This spinal motor neuron is undergoing paranodal demyelination as a consequence of the axonal enlargement (A) Nearby Remak Schwann cells enter S phase. The EM autoradiogram in (B) demonstrates silver grains, produced by 3 H-thymidine incorporation, over the nucleus of this Remak Schwann cell, in the ventral root of an IDPN-treated rat. In (C) a daughter Schwann cell (SC) has migrated to and ensheathed the old paranodal region of a swollen motor axon. The arrows identify the outermost myelin terminal loops of the adjacent internodes. B and C are modified from Griffin et al. ( 1987 ).

IDPN-induced paranodal demyelination also leads to short intercalated internodes in a nerve that has only myelinated motor fibers, the L5 ventral root of the rat (Griffin et al., 1987 ). This observation indicates that there must be source of new Schwann cells for repair in addition to Remak Schwann cells. The possibility of a population of Schwann cell precursors in adult nerves is largely unexplored.