Article |
Address correspondence to Don W. Cleveland, Ludwig Institute for Cancer Research, CMM-E/Room 3080, 9500 Gilman Drive, La Jolla, CA 92093-0670. Tel.: (858) 534-7811. Fax: (858) 534-7659. email: dcleveland{at}ucsd.edu
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Abstract |
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Key Words: myelin; neurofilaments; phosphorylation; axon caliber; radial growth
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Introduction |
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Despite these insights, the target(s) for such a kinase cascade has not been established. Analysis of neurofilament expression after axonal recovery from crush injury initially suggested a role for neurofilaments in establishing axonal diameter (Hoffman et al., 1987). Obligate heteropolymers of neurofilament light (NF-L), medium (NF-M), and heavy (NF-H) neurofilaments are the most abundant structural component of large myelinated axons. As with most neuronal proteins, they are synthesized in the neuronal soma and are subsequently transported into the axon via slow axonal transport. Once in the axon, neurofilaments are extremely long-lived proteins involved in establishing and maintaining the three-dimensional array of axoplasm.
Genetics in both mouse and quail unequivocally confirmed that neurofilaments are required for determining mature axonal diameter in myelinated regions. In mouse, loss of neurofilaments (Ohara et al., 1993; Eyer and Peterson, 1994; Elder et al., 1998; Jacomy et al., 1999; Zhu et al., 1997) markedly suppresses the many fold growth in axonal diameter (and volume) that initiates during myelination. Moreover, axonal diameter is sensitive to the subunit composition, as increased expression of any single neurofilament subunit inhibits radial growth (Monteiro et al., 1990; Cote et al., 1993; Collard et al., 1995; Tu et al., 1995; Wong et al., 1995; Marszalek et al., 1996; Xu et al., 1996), whereas simultaneous overexpression of NF-L and NF-M or NF-H increases overall axonal diameter (Xu et al., 1996). Expression of full-length neurofilament subunits and various truncation mutations in nonneuronal cells resulted in 10-nm fibers that formed COOH-terminal, phosphorylated cross-bridges that extended along the length of the filament (Nakagawa et al., 1995; Chen et al., 2000). In sciatic nerve, these COOH-terminal cross-bridges seem to span between adjacent neurofilaments and between neurofilaments and microtubules (Hirokawa et al., 1984; Rao et al., 2002). Thus, neurofilament-dependent structuring of axoplasm is likely to be mediated, at least in part, by these extended COOH-terminal tails of NF-M and NF-H.
Neurofilament-dependent radial growth is itself associated with phosphorylation of the COOH-terminal tail domains of both NF-M and NF-H, which in mice contain 7 and 51 KSP phosphorylation motifs, respectively. While neurofilaments are required for proper post-natal growth of axons, several lines of evidence suggest that neurofilament phosphorylation is essential for proper axonal diameter and that phosphorylation is regulated by myelinating cells (de Waegh et al., 1992; Yin et al., 1998). The KSP motifs of both NF-M and NF-H are nearly stoichiometrically phosphorylated (Julien and Mushynski, 1982) in the myelinated segments that undergo radial growth, whereas they are unphosphorylated in the narrow unmyelinated initial segment and the nodes of Ranvier (Hsieh et al., 1994). Further, axonal segments ensheathed by myelin-defective Schwann cells do not undergo post-natal growth, whereas segments of the same axons ensheathed by myelin-competent Schwann cells achieve large axonal diameters (de Waegh et al., 1992).
This has raised the possibility that the outside-in signal cascade originating from myelinating cells functions to activate axonal kinases (or inactivates phosphatases or both) that act directly on neurofilaments. We now use replacement of the NF-M and NF-H genes in mice to test this model of a myelination-dependent phosphorylation cascade targeting the tail domains of NF-M and NF-H as an essential feature of radial axonal growth.
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Results |
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NF-M tail and its phosphorylations are essential for radial growth of large myelinated motor axons
To determine how loss of the NF-M tail and its myelination-dependent phosphorylation sites affected radial axonal growth and survival, sizes of all axons from the fifth lumbar vertebra were determined at 2 mo, an age just after the initial burst of radial growth in mice with wild-type neurofilament content. Cross-sectional areas were measured for all axons, and equivalent diameters were calculated. Replacement of the NF-M or NF-H tails or both had no effect on the initial number of motor axons at 2 mo of age (Fig. 2 B) or on their survival to 6 mo (Fig. 2 B). While wild-type and NF-Mtail ventral root axons yielded bimodal distributions of axonal sizes, both the larger (>2 µm) and smaller (<2 µm) classes of myelinated fibers of NF-Mtail
axons failed to achieve diameters comparable to that seen in wild-type animals (Fig. 2 A).
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Axonal organization but not myelination-dependent radial growth is linked to cross-bridging between adjacent neurofilaments
To determine how the tail of NF-M and its myelination-dependent phosphorylation affects axonal organization of neurofilaments and microtubules and how this relates to establishment of axonal caliber, cross sections of motor axons were compared from wild-type, NF-Mtail, and NF-(M/H)tail
mice (Fig. 3, AC). Nearest neighbor spacing of neurofilaments was reduced (median values of 4539 nm at 2 and 6 mo were estimated from approximate Gaussian distributions of neurofilament spacing) by loss of the NF-M tail compared with wild type at both 2 and 6 mo of age (Fig. 3, D and E). Thus, NF-Mdependent cross-bridging between adjacent neurofilaments is one determinant of nearest neighbor spacing. In the presence of the NF-M tail, nearest neighbor spacing is not influenced by the much more heavily phosphorylated NF-H tail, as deletion of the entire NF-H tail does not alter nearest neighbor spacing (Rao et al., 2002). However, the NF-H tail partially compensates for the loss of NF-M tail; at both 2 and 6 mo, deletion of both tail domains reduced average spacing even further (from median values of 45 to 30 nm at 2 and 6 mo) (Fig. 3, D and E).
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In addition to altered nearest neighbor distances and neurofilament morphology, axoplasm was also less organized in the absence of both NF-Mtail and NF-(M/H)tail
mice. Axoplasm from both NF-Mtail
and NF-(M/H)tail
mice contained subregions in which the axoplasm frequently appeared "unoccupied" as compared with the more even distribution in wild-type axoplasm (Fig. 3, compare A and C). This effect was quantified by analyzing neurofilament clustering in both NF-Mtail
and NF-(M/H)tail
mice versus age-matched wild-type littermates (Fig. 4, A and B). Neurofilament clustering was defined as the ratio of average filament spacing to nearest neighbor distance, with higher ratios implying more clustered (less uniformly distributed) neurofilaments. Neurofilaments within each axon were redistributed into hexagonal arrays over a cross-sectional area equal to that of the axon (shown schematically in Fig. 4 A, left). Average neurofilament spacing was then defined to be the edge length of each hexagon. Using this measure, the array in wild-type mice diverged from the 1.0 value of a "perfectly ordered" array, yielding a clustering index of 1.8. After removal of either NF-M or both NF-M and NF-H tail domains (Fig. 4 B), neurofilaments in age-matched littermates were significantly more clustered, or less organized, with a clustering index of 2.5 in the NF-(M/H)tail
mouse.
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Slowed propagation of action potentials without NF-M taildependent expansion in axonal caliber
Motor axon conduction velocities were measured in the terminal segments of the sciatic nerves of 56-mo-old mice (five mice per genotype) from wild-type, NF-Htail, NF-Mtail
, and NF-(M/H)tail
mice (Fig. 5 A). Consistent with the loss in diameter along the distal half of the largest caliber myelinated fibers, conduction velocity of action potentials was reduced
30% in both NF-Mtail
and NF-(M/H)tail
mice (Fig. 5 A).
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Neurofilament synthesis and investment into proximal axons is suppressed after recovery from nerve crush (Hoffman et al., 1987), returning to normal levels only after recovery is complete. To determine if absence of major targets for the signaling cascade from myelinating cells to axons affects axonal recovery and regrowth after injury, sciatic nerves, at the level of the obturator tendon, were injured by crush, and the speed of recovery was assessed in NF-Htail, NF-Mtail
, NF-(M/H)tail
, and wild-type mice by measuring the spreading distance between the first and fifth digit of the ipsilateral foot as a percentage of the spreading distance before injury. Recovery profiles from all genotypes were indistinguishable throughout 3 wk after injury (Fig. 5 C).
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Discussion |
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The surprise in this mechanism underlying radial axonal growth is that it is interactions from the NF-M tail that are essential, rather than those of NF-H, which carries a much longer tail with many more sites (51 KSP sites in mice) that are nearly stoichiometrically phosphorylated. Truncation of NF-H by gene replacement in mice decreased the frequency of cross-bridges and increased the rate of radial growth, but did not alter the final caliber achieved (Rao et al., 2002). Thus, the myelin-derived signal results in an NF-H taildependent reduction in the rate, but not steady-state amount of axonal expansion.
As to how the NF-M tail mediates axonal growth in response to the myelin signal, it is clear that charge repulsion from neurofilament side arms does not play a key role. The much more heavily phosphorylated NF-H tail is neither necessary nor sufficient for neurofilament-dependent radial growth. Another simple model would be the assembly of a three-dimensional array of neurofilaments produced by nearest neighbor interactions that interlink the filaments into a space-filling, three-dimensional scaffold. This too cannot be completely correct because average interfilament spacing is altered in both NF-Mtail and NF-(M/H)tail
(Fig. 3, D and E) at both time points examined, yet at 6 mo, a time when NF-(M/H)tail
mice still have reduced interfilament distances relative to NF-Mtail
mice, the calibers of NF-Mtail
and NF-(M/H)tail
are indistinguishable (Fig. 2, C and D).
As the tail domains of NF-M and NF-H extend from the surface of the core filament, and site-specific phosphorylation apparently reduces the flexibility of this region (possibly due to local charge repulsion within the tail domain; Hisanaga and Hirokawa, 1988; Chen et al., 2000), the increased rigidity of phosphorylated neurofilament tail domains could serve two functions. First, semirigid neurofilament to neurofilament linkers, composed of NF-M and NF-H tail domains, could serve as hinge points allowing for a flexible cytoskeletal network that supports structure yet resists sudden alterations. Second, less flexible tail domains of either NF-M and/or NF-H mediate an increased spacing of adjacent neurofilaments. Increased interfilament distances may allow larger cytoskeletal linking proteins, including plectin (Wiche, 1998) and BPAG1n (Yang et al., 1996b), to associate efficiently with the neurofilament array. Increased rigidity or an increase in the number of nearest neighbor linkages stabilizes and straightens the core filament. Without these, as in the NF-(M/H)tail mice, the neurofilament array is bowed, yielding overall axoplasmic disorganization.
Further, the almost complete loss of projections from NF-(M/H)tail mice illustrates that nonneurofilament-derived cross-linking proteins, such as the plakin family of cytoskeletal linkers (Wiche, 1998), are relatively rare, or, more likely, efficient binding of these linkers to the neurofilament array requires that their binding sites be readily accessible through neurofilament tail domainmediated increase in interfilament distances or that their binding sites are affected by the NF-M and NF-H tail domains and their phosphorylation. The NF-M tail domain and the myelin-dependent phosphorylation may position cytoskeletal linking proteins in a manner that facilitates longer-range interactions between neurofilaments and either actin filaments, tubulin, or both.
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Materials and methods |
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Tissue preparation, morphological analysis, and nearest neighbor analysis
Mice were perfused intracardially with 4% paraformaldehyde in 0.1 M Sorenson's phosphate buffer, pH 7.2, and postfixed overnight in the same buffer. Samples were treated with 2% osmium tetroxide, washed, dehydrated, and embedded in Epon-Araldite resin. Thick sections (0.75 µm) for light microscopy were stained with toluidine blue. Cross sections of L5 motor and sensory axons were analyzed in five to six mice per genotype and each age group. Axonal diameters were measured using the Bioquant Software. Entire roots were imaged, imaging thresholds were selected individually, and the cross-sectional area of each axon was calculated and reported as a diameter of a circle of equivalent area. Axon diameters were grouped into 0.5-µm bins.
Mice were prepared for conventional EM analysis by intracardial perfusion with Ringers solution followed by 2% glutaraldehyde and 2% formaldehyde in 0.15 M sodium cacodylate buffer, pH 7.4, at 35°C for 5 min. L5 motor and sensory roots were dissected, fixed on ice for 12 h, washed five times for 5 min in cold 0.15 M cacodylate buffer, postfixed in 1% OsO4 in 0.15 M cacodylate buffer on ice for 1 h, and finally rinsed in cold double-distilled water five times for 5 min. After incubation with 1% uranyl acetate on ice for 16 h, roots were then rinsed in ice cold double distilled water three times for 3 min per rinse. Samples were dehydrated through an ethanol series, embedded by initially infiltrating with 50% ethanol and 50% durcupan resin for 1 h at room temperature, and then changed to 100% resin for 1 h and placed into fresh resin for an additional hour. The resin was cured in a 60°C vacuum oven for 2448 h. Samples were sectioned (6080 nm), collected on grids, stained for 10 min in 1% aqueous uranyl acetate followed by 2 min staining with lead salts, and then alyzed on a JEOL 1200EX electron microscope at 6080 kV.
Negatives were digitized with a 1024 x 1024 pixel CCD camera. Positions of individual neurofilaments were marked with an electronic pointer, and nearest neighbor distances were calculated using software written by Stephen Lamont (NCMIR; University of California, San Diego).
Methods for calculating neurofilament clustering
Average neurofilament spacing was determined by distributing identified neurofilaments in uniform arrays across the effective cross-sectional area of an axon. Cross-sectional area was estimated by tracing axoplasmic regions of the same digitized electron micrographs used to identify neurofilaments. Neurofilaments were organized in concentric hexagonal "rings" of equilateral triangles (Fig. 4 A), with average neurofilament spacing calculated as the side length of one triangle. Hexagons were selected due to their inherent ability to pack two-dimensional space optimally (Conway and Sloane, 1999). The number of counted neurofilaments (nNF) corresponded to ntri triangles, arranged into i hexagonal rings. The relationships between nNF, ntri, and I are given by the following recursive formulae:
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Visualization of neurofilament organization in the axon by quick-freeze, deep-etch analysis
Sciatic nerves of 45-mo-old NF-Mtail and their control littermate animals were dissected and incubated in oxygenated artificial cerebrospinal fluid containing (in mM, pH 7.3) 126 NaCl, 22 NaHCO3, 1 Na2HPO4, 2.8 KCl, 0.88 MgCl2, 1.45 CaCl2, and 3.5 glucose. Subsequently, nerves were sectioned with a razor blade, and the tissue was frozen by slamming against a liquid heliumcooled copper block (E7200; Polaron) as previously reported (Gotow et al., 1999). The frozen tissue was mounted onto the freeze fracture apparatus (BAF 400D; Balzers), fractured, and then deep etched and rotary replicated with platinum/carbon at an angle of 25°. The replicates were examined with a Hitachi H-300 electron microscope at 75 kV.
Nerve conduction velocity measurements
Nerve conduction velocities were measured in the sciatic nerve, interosseus muscle system of 5-mo-old mice (Calcutt et al., 1990). In brief, mice were anesthetized with halothane (4% in O2 for induction, 23% for maintenance), and rectal temperature was maintained at 37°C by a heating lamp and thermal pad connected to a temperature regulator and the rectal thermistor probe. The sciatic nerve was stimulated with single supramaximal square wave pulses (48 V and 0.05 ms duration) via fine needle electrodes placed at the sciatic notch and Achilles tendon. Evoked electromyograms were recorded from the interosseus muscles of the ipsilateral foot via two fine needle electrodes and displayed on a digital storage oscilloscope. The distance between the two sites of stimulation was measured using calipers, and conduction velocity was calculated as previously described (Calcutt et al., 1990). Measurements were made in triplicate from five animals per genotype, and the median was used as the measure of velocity. Statistical ANOVA was performed with subsequent Bonferroni multiple comparisons test post-hoc analysis using InStat.
Sciatic nerve regeneration measurements
Mice were placed under Metophane anesthesia, and the sciatic nerve was exposed via an incision in the flank followed by separation of underlying musculature by blunt dissection. The nerve is crushed using fine jewelers forceps at the level of the obturator tendon. To assess functional recovery of the injured limb, the mouse was induced to spread its toes by briefly lifting the hindlimbs off the bench. Distance from first to fifth digits was measured with a divider and expressed as a percentage of preinjury spread distance.
Activity wheels
Mice were placed in a single activity wheel chamber system (Lafayette Instruments) for 14 d. Activity was measured by the number of revolutions an animal would run during an 12-h period. Revolutions were counted using an optical sensor that detects wheel motion and were stored on an activity wheel counter (Lafayette Instruments). Revolutions were converted into kilometers based upon a 5-inch diameter activity wheel.
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Acknowledgments |
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This work has been supported by National Institutes of Health (NIH) grant NS 27036 to D.W. Cleveland and grant NS 38855 to N.A. Calcutt. Salary support for D.W. Cleveland is provided by the Ludwig Institute for Cancer Research. M.L. Garcia was supported, in part, by a postdoctoral fellowship from the NIH. Some of the imaging was conducted at the National Center for Microscopy and Imaging Research at San Diego, which is supported by the NIH through a National Center for Research Resources program grant (P41 RR04050) awarded to M. Ellisman.
Submitted: 28 August 2003
Accepted: 14 October 2003
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