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The Journal of Neuroscience, April 1, 2001, 21(7):2195-2205
Neurofilaments Consist of Distinct Populations That Can Be
Distinguished by C-Terminal Phosphorylation, Bundling, and Axonal
Transport Rate in Growing Axonal Neurites
Jason T.
Yabe1,
Teresa
Chylinski1,
Feng-Song
Wang2,
Aurea
Pimenta3,
Solomon D.
Kattar1,
Maria-Dawn
Linsley1,
Walter K-H
Chan1, and
Thomas B.
Shea1
1 Center for Cellular Neurobiology and
Neurodegeneration Research, Department of Biological Sciences,
University of Massachusetts-Lowell, Lowell, Massachusetts 01854, 2 Department of Biological Sciences, Purdue
University-Calumet, Hammond, Indiana 46323, and
3 Department of Neurobiology, University of Pittsburgh
School of Medicine, Pittsburgh, PA 15261
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ABSTRACT |
We examined the steady-state distribution and axonal transport of
neurofilament (NF) subunits within growing axonal neurites of NB2a/d1
cells. Ultrastructural analyses demonstrated a longitudinally oriented
"bundle" of closely apposed NFs that was surrounded by more widely
spaced individual NFs. NF bundles were recovered during fractionation
and could be isolated from individual NFs by sedimentation through
sucrose. Immunoreactivity toward the restrictive C-terminal phospho-dependent antibody RT97 was significantly more prominent on
bundled than on individual NFs. Microinjected biotinylated NF subunits,
GFP-tagged NF subunits expressed after transfection, and radiolabeled
endogenous subunits all associated with individual NFs before they
associated with bundled NFs. Biotinylated and GFP-tagged NF subunits
did not accumulate uniformly along bundled NFs; they initially appeared
within the proximal portion of the NF bundle and only subsequently were
observed along the entire length of bundled NFs. These findings
demonstrate that axonal NFs are not homogeneous but, rather, consist of
distinct populations. One of these is characterized by less extensive
C-terminal phosphorylation and a relative lack of NF-NF interactions.
The other is characterized by more extensive C-terminal NF
phosphorylation and increased NF-NF interactions and either undergoes
markedly slower axonal transport or does not transport and undergoes
turnover via subunit and/or filament exchange with individual NFs.
Inhibition of phosphatase activities increased NF-NF interactions
within living cells. These findings collectively suggest that
C-terminal phosphorylation and NF-NF interactions are responsible for
slowing NF axonal transport.
Key words:
neurofilaments; axonal transport; cytoskeleton; axon; phosphorylation; neuronal differentiation
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INTRODUCTION |
Despite decades of study,
controversy exists over several aspects of neurofilament (NF) axonal
transport. Models that have been advanced to describe the organization
of axonal NFs range from relentless translocation of a homogeneous
population of NFs (Lasek, 1986
; Lasek et al., 1992, 1993) to
translocation of only some NFs, whereas others incorporate into a
distinct cytoskeletal macrostructure that is essentially stationary
(Nixon and Logvinenko, 1986; Nixon, 1993, 1998). Contemporary
articulation of these models demonstrates them to display considerable
inherent similarity in that they both consider that NFs transport over
a broad range of rates and that these rates are derived from the
relative length of time the individual NFs are associated with their
transport vector (Lasek et al., 1993; Nixon, 1993, 1998). The essential difference among these models is whether axonal NFs constitute a single
population with a continuum of transport rates or whether the
differences in transport rates actually characterize distinct NF populations.
The form in which NF subunits undergo transport is also controversial.
Various studies provide evidence for the transport of NFs themselves
(Nixon and Lewis, 1986; Galbraith et al., 1999; Koehnle and Brown,
1999; Wang et al., 2000), transport of nonfilamentous punctate
assemblies of NF subunits (Yabe et al., 1999), and transport of
monomers or small oligomers (Takeda et al., 1994; Terada et al.,
1994; Hirokawa et al., 1997). Studies demonstrating the transport of
nonfilamentous punctate assemblies demonstrate that they can assume
filamentous form during transport and consider punctate structures only
as transitional forms (Yabe et al., 1999). Such studies, and those
indicating transport of filaments, are consistent with the possibility
that axonal NFs undergo turnover chiefly via replacement with new NF
polymer (for review, see Baas and Brown, 1997). Conversely, studies
supporting transport of monomers and/or small oligomers are interpreted
to indicate that the bulk of axonal NFs does not itself
translocate and instead undergoes turnover by
exchange/incorporation of transporting subunits (for review, see
Hirokawa et al., 1997).
Finally, controversy also exists regarding the nature of NF
interactions within axons and regarding how NFs exert their stabilizing influences on axonal cytoarchitecture. Certain studies indicate that
NFs undergo phosphorylation-dependent cross-linking and bundling (Leterrier and Eyer, 1987; Shaw and Hou, 1990; Nagakawa et al., 1995;
Leterrier et al., 1996). Resultant formation of an interconnected suprastructure could readily explain how NFs impart support to the
axon. However, the demonstration that NFs move apart on disruption of
axonal membrane integrity suggests that at least the majority of NFs
is not cross-linked physically (Brown and Lasek, 1993). NFs still can be envisioned to impart stability to axons in the absence
of physical cross-linking with the consideration that their extensive
phosphorylation fosters NF-NF repulsion and restricts their lateral motion.
In the present study we addressed the above controversies by examining
the organization and dynamics of NFs in growing axonal neurites in culture.
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MATERIALS AND METHODS |
Cell culture. NB2a/d1 cells were selected for these
analyses because (1) they express and phosphorylate all three NF
subunits (Shea et al., 1988, 1990), and (2) they accumulate a prominent bundle of Triton-insoluble axonal NFs that resembles NF bundles observed in axons in situ (Shaw and Hou, 1990). In addition,
although some "bundled" axonal NFs also have been reported in
certain cultured neurons (Brown, 1997), the ability to generate large
quantities of NB2a/d1 cells and to control their differentiation state
renders them particularly suitable for biochemical as well as
immunological and ultrastructural analyses.
Cells were cultured on poly-L-lysine-coated plates or
coverslips in DMEM (high glucose formulation) containing 10% horse
serum. Elaboration of axonal neurites was induced by the addition of 1 mM dibutyryl cyclic AMP (dbcAMP) for 3 d (Shea et al.,
1988). In some experiments the cells received 5 nM okadaic
acid (OA) for the final 24 hr of differentiation (Shea et al., 1993).
In some experiments at either 2 or 18 hr after microinjection, some cells were treated with 330 nM nocodazole for 2 hr.
Resultant perturbation of axonal microtubules by this treatment induces overall thinning of axonal caliber with periodic "beading" (Shea and Beermann, 1994).
Biotinylation and microinjection of NF subunits. Bovine
spinal cords were homogenized in 100 mM PIPES, pH 6.6, containing (in mM) 1 EGTA, 1 MgCl2, 1 PMSF, and 1 DTT plus 1 µg/ml leupeptin. NFs were recovered in the
void volume after Sepharose CL-4B gel chromatography (Takeda et al.,
1994). As described (Takeda et al., 1994), a single
assembly/disassembly cycle was sufficient to eliminate tubulin and
other contaminating proteins. Isolated NFs were disassembled in urea
buffer [100 mM phosphate buffer, pH 7.5, containing 6 M urea, 0.5 M PMSF, 1 µg/ml leupeptin, and (in mM) 1 EDTA, 1 EGTA, and 1 DTT] and then dialyzed
overnight at 4°C, followed by 1 hr at 37°C in assembly buffer
[containing (in mM) 20 PIPES, pH 6.6, 1 EGTA, 1 MgCl2, 1 EDTA, 1 DTT, 0.17 NaCl (Takeda et al.,
1994)]; they were recovered by centrifugation at 100,000 × g for 1 hr. The resultant once-cycled NFs were conjugated with 3-(N-maleimidylproprionyl) biocytin (25 ng/ml;
Molecular Probes, Eugene, OR) while in assembled form essentially as
described previously (Takeda et al., 1994). NF-H and NF-L subsequently
were purified by DE-52 chromatography in urea buffer; pure fractions (ascertained by SDS-gel electrophoresis and immunoblot analysis; see
Results) were combined and dialyzed overnight at 4°C against assembly
buffer to remove urea and then dialyzed for 6 hr against 10,000 volumes
of injection buffer [containing (in mM) 5 HEPES, 1 DTT, 1 EDTA, and 1 EGTA, pH 8.5], concentrated by centrifugation onto Centricon filters, and stored at 
80°C (Takeda et al., 1994). Just before injection, purified subunits were mixed 1:1 with 70 kDa
fluorescein-conjugated dextran (Molecular Probes) and clarified by
centrifugation for 10 min in a Beckman airfuge; the resulting supernatant (containing ~1 mg/ml NF protein) was microinjected as
described (Straube-West et al., 1996; Jung et al., 1998; Yabe et al.,
1999). Injected cells were located under fluorescein optics and
examined by phase-contrast microscopy; cells exhibiting any obvious
trauma resulting from microinjection were excluded from further
analyses. Notably, translocation of microinjected subunits into axonal
neurites of NB2a/d1 cells was prevented by administration of the
anti-microtubule drug nocodazole immediately after microinjection (Jung
et al., 1998; Yabe et al., 1999), indicating that microinjected subunits required an intact microtubule network to undergo active transport. These data confirm that no significant contribution to the
transport of biotinylated subunits into neurites is derived from
subunit diffusion and/or injection pressure.
Immunofluorescence and immunoelectron microscopy. Cells were
processed for endogenous and microinjected NF immunoreactivity at 2-18
hr after microinjection. To preserve axonal microtubules (MTs), we
rinsed the cells in warm Tris-buffered saline, pH 7.4 (TBS), and
fixed them under MT-stabilizing conditions [containing (in
mM) 60 PIPES, pH 6.9, 10 EGTA, and 2 MgCl2 plus 10 µM taxol (Shea,
1999)]; alternate cultures also received 1% saponin to extract
unassembled tubulin subunits. Alternatively, to deplete MTs and
associated proteins (some of which may cross-react with phospho-dependent anti-NF antibodies), we fixed most cells at 4°C in
the absence of taxol in 50 mM Tris, pH 6.8, containing 5 mM EDTA, 1 mM PMSF, and 50 µg/ml leupeptin,
with and without previous extraction for 15 min at 4°C with 1%
Triton X-100 (Shea et al., 1988). Triton extraction under these
conditions also depletes some NFs (Shea et al., 1990). Nonextracted
cells were incubated at 4°C in the same buffer without Triton X-100;
then all cultures were fixed for 15 min with 4% paraformaldehyde in
TBS and permeabilized by incubation with 0.1% Triton X-100 for 15 min
at room temperature. Because fixation without taxol at 4°C
effectively depleted most MTs and more clearly visualized the NFs (see
Fig. 1A-C), all cells except those in Figure 1,
A and B, were subjected to this latter Triton
X-100 extraction and fixation protocol. Some additional cultures were
processed by using a "splaying" technique (Brown, 1998) that
affords visualization of filamentous profiles.
Cultures were immunostained with 1:100 dilutions of monoclonal
antibodies directed toward phosphorylated (SMI-31) and
nonphosphor-ylated (SMI-32) NF epitopes (Sternberger Monoclonals,
Bethesda, MD), a monoclonal antibody (RT97) directed against a
developmentally delayed phospho-epitope of NF-H (Anderton et al.,
1982), a polyclonal antibody generated in this laboratory against
electrophoretically purified murine NF-L (L3), or a polyclonal
anti-biotin antibody (Jung et al., 1998), followed by 1:150 dilutions
of rhodamine- or fluorescein-conjugated anti-mouse or anti-rabbit IgG.
Incubations with primary antibodies were done either overnight at 4°C
or for 2 hr at 37°C. Incubations with secondary antibodies were for 2 hr at 37°C. For double-immunofluorescent analyses the primary antibodies were applied simultaneously and, after rinsing, secondary antibodies also were applied simultaneously.
For immunoelectron microscopy (EM), cells extracted with 1% Triton
X-100 at 4°C (see above) were fixed in 4% glutaraldehyde at 4°C;
rinsed; incubated (1 hr at room temperature) with 1:100 dilutions of
SMI-31, RT97, or anti-biotin; rinsed three times; and then incubated
with secondary antibodies conjugated to 5 or 10 nm colloidal gold
particles. Samples were dehydrated, embedded in resin, sectioned, and
stained with uranyl acetate by conventional methods and examined in a
Philips 300 EM (Shea et al., 1993). Positive identification of injected
cells required embedding within the culture dish and sectioning
parallel to the substrate; because of the difficulty of obtaining the
perikaryon and the entire axonal length (100 µm; Shea, 1999) within
individual sections, many analyses were performed on proximal or distal
regions of individual injected cells.
Construction of eGFP-NF-M and transfection. As described
previously (Yabe et al., 1999), a 2484 bp fragment encoding amino acids
1-824 of rat NF-M cDNA was isolated from the plasmid pRSVi-NF-M (generous gift of Dr. Ron Liem, Columbia University, New York, NY; Chin and Liem, 1989) by a double-restriction enzyme
digestion with HindIII and HincII (blunt end),
followed by agarose gel electrophoresis and gel extraction with JetSorb
(Genomed, Raleigh, NC). The purified fragment was subcloned into the
HindIII and SmaI (blunt end) restriction sites of
the eukaryotic expression vector peGFP-N3 (Clontech, Palo Alto,
CA) in frame with eGFP. After overnight ligation at 12°C the reaction
mixture was used to transform HB101 Escherichia coli.
Ampicillin-resistant colonies were isolated for mini-preparations of
plasmid, which were analyzed by restriction digestion for the NF-M cDNA
fused in frame with eGFP. Large-scale plasmid preparations for
transfections were performed with the Qiagen purification system
(Qiagen, Santa Clarita, CA) to generate in transfected cell lines NF-M
as a fusion protein with eGFP. All standard molecular biology
procedures were performed essentially as described by Sambrook et al.
(1989).
NB2a/d1 cells treated with dbcAMP for 2 d were transfected with
33.2 µg/ml eGFP-NF-M or eGFP without an NF-M insert in 238 µg/ml
Superfect (Qiagen) for 3 hr in the presence of 10% serum, after which
the medium was replaced (Yabe et al., 1999). Cells were incubated for
an additional 24 hr to allow for accumulation of eGFP-NF-M and for
continued elaboration of axonal neurites Stably transfected cells were
selected by the addition of 100 µg/ml GD14 (Life Technologies,
Grand Island, NY) to the medium and the isolation of surviving colonies
7-14 d later. Accumulation of full-length eGFP-conjugated NF-M was
confirmed by immunoblot analyses of stably transfected cultures (see below).
Densitometric analyses. Phase-contrast and epifluorescent
images of individual cells were captured via a Dage CCL-72 camera connected to a Scion LG-3 frame grabber housed in a Macintosh 7100AV
and operated by NIH Image software. Images were stored as TIFF or PICT files.
The relative distribution of NF immunoreactivity was determined via NIH
Image software. Only cells elaborating a single unbranching axonal
neurite were included in analyses. After automated background subtraction, areas of axonal neurites were encircled by using the
freehand tool of the program, and the resultant net densitometric value
was recorded. For determination of the relative distribution of NF
immunoreactivity within nocodazole-induced beads and thinned areas of
the shaft, representative beads were encircled, and the resulting
densitometric value was recorded. Then the identical circle was shifted
to an adjacent nonbeaded area of the axonal shaft. The ratio of
immunoreactivity within beads versus adjacent areas of the axonal shaft
was obtained by dividing the densitometric value of a given bead by
that of its adjacent shaft. Values that are presented represent the
mean and SEM derived from six to eight injected cells from two separate
experiments. Statistical comparisons were performed via Student's
t test.
To generate mean distribution profiles for multiple axons of
transfected cells, we measured the contour length of axonal neurites in
the digitized phase-contrast image by using the "neurite labeling macro" of the NIH program. The resulting length was divided into 10 equivalent segments. The start of segment one in these analyses was
defined as that region of the axon shaft that was distinguished clearly
from the perikaryon and any gradual thinning of the perikaryon. Accordingly, this first segment was entirely within the axonal shaft
and did not include the axonal hillock. Similarly, the end of segment
10 was defined as the final segment of the shaft just before the
swelling of the growth cone. Then the density of each segment in the
corresponding epifluorescent image was calculated by drawing a box
around each segment, shifting the box to an immediately adjacent
cell-free area, and recording individual background densities. The box
was shifted as little as possible for background recording, which
maintained the relative spatial localization within the microscopic
field of the selected background area relevant to the corresponding
selected axonal area; this eliminated the potential influence of
differential illumination across a given microscopic field. Resultant
data were exported to Excel spreadsheet software for subsequent calculations.
In additional analyses the relative distribution of colloidal gold
particles in immuno-EM analyses was quantified within proximal or
distal regions of axonal neurites of differentiated NB2a/d1 cells or
within bundled NFs versus adjacent regions of the same axons not
containing bundled NFs. In such analyses the percentage of gold
particles within the region of interest was calculated relative to the
total number of particles within the region of interest plus the
adjacent/opposing region.
Gel electrophoresis, autoradiography, and immunoblot analyses.
dbcAMP-treated cells on uncoated plates were pulse-radiolabeled for 15 min with [35S]methionine in
methionine-free DMEM without serum (500 µCi for each of 2 × 10 cm2 plates at ~80% confluency; Shea et
al., 1990). Some cultures were harvested immediately after labeling.
For additional cultures the medium was replaced with medium lacking
radiolabel but containing 10% horse serum and 20× the normal
concentration of methionine; incubation was continued for a total of 4 hr before harvesting. All cultures were incubated in methionine-free,
serum-free medium for 15 min before radiolabeling.
Radiolabeled cells were harvested according to established methods that
separate axonal neurites from perikarya (Shea et al., 1993). Briefly,
cultures were harvested by scraping with a rubber policeman in 50 mM Tris-HCl, pH 6.8, containing 5 mM EDTA, 1 mM PMSF, and 50 µg/ml leupeptin; they were homogenized
gently on ice (25 strokes) in a loose-fitting glass-Teflon homogenizer
and then centrifuged at 5000 × g for 5 min at 4°C.
This procedure sediments axonal fragments but leaves disrupted
perikarya in the supernatant (Shea et al., 1993). The axonal pellet was
resuspended in the same buffer containing 1% Triton X-100, and the
perikaryal supernatant was made 1% Triton X-100. Cytoskeletons
then were isolated separately from axonal- and perikaryal-enriched
fractions by homogenization (50 strokes in a tight-fitting glass-Teflon homogenizer) on ice, followed by centrifugation at 15,000 × g for 15 min at 4°C (Shea et al., 1988). Axonal
cytoskeletons were resuspended in the same buffer without Triton and
then sedimented for 15 min at 15,000 × g onto 1 ml of
1 M sucrose in the same buffer. Aliquots of
cytoskeletons from cell bodies and from the interface and pellet
generated after the above sucrose gradient centrifugation were
immunoprecipitated with a 1:150 dilution of a polyclonal antibody (R39)
that precipitates all NF subunits (Shea et al., 1997; Jung et al.,
1998). Gels were dried, and autoradiographs were generated. Aliquots of
immunoprecipitated material also were subjected to immunoblot analyses
with R39. Autoradiographs and immunoblots were digitized with a UMAX
flatbed scanner equipped with a "transparency adaptor" at 300 dpi
(Jung and Shea, 1999).
Immunoblots of Triton-insoluble cytoskeletons (100 µg) from
stably transfected cells were probed with a polyclonal anti-GFP antibody (Clontech) and a polyclonal antibody (M2) generated in this
laboratory against NF-M.
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RESULTS |
Axonal neurites contain a core of closely opposed NFs that exhibit
increased site-specific C-terminal phosphorylation and increased NF-NF
interactions
Consistent with previous studies (Shea et al., 1988),
intermediate-sized filaments were detected along the length of axonal neurites (Fig. 1A-C).
Reactivity with the relatively nonrestrictive (Shea et al., 1989)
phospho-dependent antibody SMI-31 indicated that these intermediate
filaments were NFs (Fig. 1C,D). Depletion of axonal MTs
highlighted that a significant portion of Triton-insoluble axonal NFs
was concentrated within the center of the axon with respect to the
longitudinal axis (Fig. 1C-E); these closely
apposed NFs will be referred to as "bundles" for the remainder of
this report. Bundled NFs were situated at a mean distance of
~17.5 ± 1.7 nm apart. The closest peripherally situated
individual NFs averaged 34.3 ± 5.3 nm apart and ranged to >60 nm
apart. Previous analyses (Gotow and Tanaka, 1994) have demonstrated
that bundled NFs could be distinguished from individual axonal NFs by
the increased prevalence of a developmentally delayed phospho-epitope
(RT97; Anderton et al., 1982). We therefore examined whether this was also the case in NB2a/d1 axonal neurites. Consistent with this possibility, SMI-31 immunoreactivity was distributed relatively evenly
throughout the axonal width, whereas RT97 was relatively more
concentrated within the central aspect of axonal neurites (Fig.
1D-F). Immuno-EM analyses confirmed that
bundled NFs displayed more than fourfold increased affinity
(p < 0.01) for RT97 than did individual NFs
(Fig. 1E), whereas individual and bundled NFs displayed equivalent SMI-31 immunoreactivity. Restriction of RT97 immunoreactivity within the central bundle was highlighted further, comparing the distribution of NF-L (as an index of total NFs) with that
of RT97 or SMI-31; SMI-31 codistributed throughout the axonal
diameter with NF-L, whereas RT97 was concentrated along the center of
axonal neurites (Fig. 1F).
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Figure 1.
NB2a/d1 axonal neurites contain a mixture of
closely opposed and relatively disbursed NFs that exhibit differential
reactivity toward phospho-dependent NF antibodies. A-C,
Regions of axonal neurites of differentiated NB2a/d1 cells.
A, B, Neurites fixed at 37°C under
MT-stabilizing conditions in the absence (A) and
presence (B) of 1% saponin (Shea, 1999). Note
prominent MT profiles along the neurite. C, An axonal
neurite extracted with 1% Triton X-100 at 4°C in the absence of
MT-stabilizing agents. Note the centrally located "bundle" of
closely apposed NFs (arrows) and the relative depletion
of MTs. On the basis of these findings, all cultures for the remainder
of this study were fixed at 4°C, with or without Triton X-100 and in
the absence of taxol, to deplete axonal MTs and to reveal more clearly
the Triton-insoluble NFs. A-C are presented at the same
magnification. D, Axonal neurites of cells processed for
immuno-EM with SMI-31 or RT97, as indicated, followed by colloidal
gold-conjugated secondary antibody. C presents an axonal
neurite, at smaller magnification, which also was processed for
immuno-EM with SMI-31. Note that SMI-31 immunoreactivity is distributed
among bundled and nonbundled individual NFs, whereas RT97
immunoreactivity is localized to bundled NFs. Arrows in
the neurite that was reacted with SMI-31 denote individual NFs
(peripheral to the bundle) that are decorated with colloidal gold,
whereas arrows in the neurite reacted with RT97 denote
individual NFs that are not decorated with colloidal gold.
E, Quantification of the distribution of SMI-31 and RT97
immunoreactivity on bundled and peripherally located individual NFs.
Note that the SMI-31 epitope is distributed relatively evenly between
bundled and individual NFs, whereas the RT97 epitope is markedly more
prevalent on bundled NFs. F, Double-immunofluorescent
analyses of the distribution of SMI-31 and NF-L and, in a second cell,
RT97 and NF-L. NF-L immunoreactivity is distributed evenly throughout
axonal neurites. Note that, like NF-L immunoreactivity, SMI-31
immunoreactivity is distributed relatively evenly throughout axonal
neurites. RT97 immunoreactivity, by contrast, is relatively
concentrated along the center of the axon with respect to its
longitudinal axis and does not codistribute with NF-L.
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Also consistent with previous studies (Shaw and Hou, 1990; Leterrier et
al., 1996), bundled NFs were recovered among individual NFs in
cytoskeletal preparations derived from NB2a/d1 axonal neurites; bundles
observed within such preparations also demonstrated markedly more RT97
immunoreactivity than did individual NFs (Fig.
2). The bulk of axonal NFs moves apart
when freed from the constraints of the axonal membrane, suggesting that
they are not cross-linked physically within axons (Brown and Lasek,
1993). However, recovery of some bundled NFs previously has been
interpreted to indicate that a population of NFs indeed is cross-linked
in some capacity in situ (Shaw and Hou, 1990). Bundled NFs
within NB2a/d1 axonal neurites therefore may represent a population of
NFs that have undergone more extensive NF-NF interactions than the
more peripherally situated individual NFs. In support of this notion
the NF bundles, but not individual NFs, sedimented through 1 M sucrose (Fig. 2).
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Figure 2.
Some NFs are recovered from axonal neurites in
bundled form. The panels present ultrastructural and immunological
analyses of cytoskeletons. Bundled NFs were detected among individual
NFs in sectioned cytoskeletal preparation (Isolated
NFs). Negative stain analyses demonstrated that bundles
sedimented via 1 M sucrose, whereas individual NFs did not
(Pelleted onto 1 M sucrose); the
accompanying immunoblots were processed for SMI-31 immunoreactivity,
and the 200 kDa region is presented. Immuno-EM analyses demonstrated
that RT97 immunoreactivity was associated selectively with bundled NFs
in sectioned cytoskeletal preparations.
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Because NF phosphorylation has been reported to mediate NF bundling in
cell-free analyses, we undertook to determine whether we could modulate
the extent of NF bundling within axonal neurites. To accomplish this,
we added the phosphatase inhibitor okadaic acid (OA) to cultures for
the final 24 hr of their dbcAMP treatment. OA previously has been
reported to increase the number of phospho-NFs within these cells (Shea
et al., 1993), to mediate increased C-terminal NF phosphorylation, and
to decrease NF axonal transport rate in situ (Veeranna et
al., 1995; Jung and Shea, 1999). We then quantified the number of
SMI-31-reactive filaments (as an index of total NFs) that were observed
within 20 nm of each other along their longitudinal length in
longitudinally oriented thin sections of axonal neurites from multiple
OA-treated and untreated cells; this distance was selected on the basis
of mean NF-NF distance within centrally oriented NF bundles (17 ± 1.7 nm; see above). We performed these analyses on peripheral areas
(i.e., close to the plasma membrane) rather than attempting to quantify
any OA-induced alterations in the size of the central NF bundle,
because the overall axonal caliber, as well as the size of central NF
bundles, varied from cell to cell. These analyses revealed a more than twofold increase (p < 0.05) in closely apposed
NFs (Fig. 3). This OA-mediated increase
in NF-NF association was unlikely to be derived artifactually solely
from an increase in total axonal phospho-NFs (Shea et al., 1993)
because, even after OA treatment, NFs occupied a very small percentage
of the axonal area within thin sections of peripheral axonal regions
(Fig. 3). These findings demonstrate a functional relationship between
NF phosphorylation and NF bundling, one interpretation of which is that
NF phosphorylation mediates NF bundling.
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Figure 3.
The phosphatase inhibitor okadaic acid
(OA) increases NF-NF associations within axonal
neurites. The panels present peripheral areas of axonal neurites from
OA-treated and untreated cells that were probed with SMI-31, followed
by colloidal gold-conjugated secondary antibody; the plasma membrane is
noted by arrows. The accompanying graph presents
quantification of the percentage of total SMI-31-immunoreactive
filaments observed within 20 nm of another SMI-31-immunoreactive
filament in longitudinally oriented sections. Note the more than
twofold increase in closely apposed NFs after OA treatment
(p < 0.05; Student's t
test).
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Transporting NF subunits exhibit a delay in association with
bundled NFs
Because more extensive C-terminal NF-H phosphorylation has been
associated with slower-moving NFs (Lewis and Nixon, 1988; Jung et al.,
2000), we hypothesized that individual and bundled NFs could be
distinguished by differential transport and/or subunit turnover. This
hypothesis was tested by microinjection of biotinylated NF subunits
(Fig. 4A,B),
transfection with a construct encoding GFP-tagged NF subunits (Fig.
4C,D), and pulse-chase metabolic radiolabeling of
endogenous NF subunits.
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Figure 4.
Biotinylated and GFP-tagged NF subunits and their
intracellular distribution after microinjection and transfection.
A, Coomassie blue staining after SDS-gel electrophoresis
of NFs isolated from bovine spinal cords (CBB) and
immunoblot analysis of this preparation (NFs) after
biotinylation and staining of chromatographically separated NF-H and
NF-L, as indicated. B, UV images of cells 2 hr after the
injection of biotinylated NF-H (Biotin) and
fluorescein-conjugated tracer (Tracer), as indicated.
C, Immunoblot analyses of Triton-insoluble cytoskeletons
from cells stably transfected with the eGFP-NF-M construct. In
addition to NF-M isoforms migrating between 97 and 145 kDa detected by
anti-NF-M antibodies (bracket), note the presence of an
NF-M-reactive isoform migrating at ~170 kDa that also is detected by
anti-GFP antibodies (arrow). Asterisks
indicate lower-molecular-weight C-terminal proteolytic products derived
from NF-M; note the labeling of these products by both GFP and NF-M
antibodies. Identity of these species as breakdown products was
confirmed by their increase at the expense of full-length eGFP-NF-M
after the incubation of additional samples at room temperature for
30-60 min (data not shown). D, Fluorescent and
corresponding phase-contrast image of an unextracted, transiently
transfected cell. E, The axonal neurite of a transiently
transfected cell processed under conditions that promote the splaying
of axonal NFs (Brown, 1998). The resultant loosening of bundled NFs
confirms the filamentous nature of axonal GFP fluorescence.
Inset presents the distal region, just before the growth
cone, of a second axonal neurite.
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As in previous immunofluorescent analyses (Jung et al., 1998; Yabe et
al., 1999), immuno-EM analyses revealed that biotinylated subunits were
detected along the entire axonal length by 2 hr after their injection
into perikarya. However, the resolution afforded by these
ultrastructural analyses revealed that biotinylated subunits were
distributed differentially among axonal NFs at early versus later times
after injection. Despite their distribution along the entire axonal
length within 2 hr, biotinylated subunits remained concentrated along
the periphery of the axon with respect to its longitudinal axis and
displayed relatively little localization within NF bundles (Fig.
5). This phenomenon was not derived by steric inhibition of antibody penetration, because, as shown above, SMI-31 immunoreactivity was distributed evenly among bundled and nonbundled axonal filaments at all of the times that were examined, and
RT97 immunoreactivity was even more prominent on bundled versus nonbundled NFs (see Fig. 1). Moreover, biotinylated subunits underwent progressive association with NFs in bundles along the axon and were
distributed relatively evenly among bundled and nonbundled NFs
throughout the axon by 18 hr after microinjection (Fig. 5).
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Figure 5.
Ultrastructural analyses of the
distribution of microinjected NF subunits. The panels present immuno-EM
analyses of the distribution of biotinylated NF-H after its
microinjection into differentiated NB2a/d1 cells, as described in
Materials and Methods. Top, Biotinylated subunits
(arrows) were readily localized along peripherally
situated NFs within central axonal segments within 2 hr after injection
but were excluded in large part from the bundle (indicated by
arrowheads along the right
side of the micrographs). By 18 hr, however,
biotinylated subunits were dispersed throughout the bundles. The
accompanying graph presents the relative distribution of biotinylated
NF-H subunits within individual and bundled NFs at 2, 6, and 18 hr
after injection; note the progressive association of biotinylated
subunits with bundled NFs. Bottom, Comparative
analyses of the distribution of biotinylated N-H subunits within
bundles and individual NFs within proximal and distal regions of the
axonal shaft at 2 and 6 hr after microinjection; for these analyses we
analyzed regions within the proximal and distal halves of the axonal
shaft, excluding the hillock and the growth cone. Note the presence of
some gold particles (arrows) within the centrally
situated bundle (denoted by arrowheads along
right side of the micrograph) within
proximal segments at 2 hr after injection versus their absence in
distal segments as well as in central segments (e.g., 2 hr micrograph
in top panel). Note that not all NFs within
bundles display even labeling within the proximal shaft at 2 hr; some
NFs apparently remain unlabeled. The accompanying graph presents the
relative distribution of biotinylated NF-H subunits within bundles in
proximal and distal axonal shafts at 2 and 6 hr after injection; note
that the vast majority of bundle-associated biotinylated subunits
is localized within the proximal half of the axonal shaft at 2 hr after
injection but is distributed equally between proximal and distal
regions by 6 hr after injection.
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We next considered the mechanism(s) by which biotinylated NF subunits
associated with NF bundles. Should bundled NFs undergo axonal
transport, we would expect biotinylated subunits to accumulate initially, or in greater concentration, at the proximal-most end of the
bundle and to do so only later along the more distal portion of bundled
NFs. By contrast, should bundled NFs represent essentially nonmoving
NFs (e.g., "stationary" NFs; Nixon and Logvinenko, 1986) and should
NFs in these bundles be replaced by subunit exchange (Takeda et
al., 1994), we would expect biotinylated subunits to accumulate
simultaneously at similar concentrations along the length of the
bundle. This latter possibility is feasible because immunofluorescent
analyses (Jung et al., 1998) (see also Fig. 4B)
demonstrated that biotinylated subunits were present at similar concentrations along the entire axonal length substantially before their association within bundled NFs, as revealed herein by
ultrastructural analyses (Fig. 5). We therefore compared the
distribution of biotinylated NF-H within segments from the proximal
half of the shaft (taking care to exclude the hillock and most proximal
segment) with segments from the distal half of the shaft (taking care
to exclude the distal-most segment and the growth cone). Biotinylated
subunits in proximal and distal segments of the axonal shaft were
scored as associated with bundles when they appeared on or adjacent to bundles (Fig. 5) or otherwise were considered not to be associated with
bundles. At 2 hr after injection 76% of those biotinylated subunits
that were associated with bundles were confined to the proximal portion
of bundles. By 6 hr after injection, however, those biotinylated
subunits that were associated with bundles were distributed evenly
between the proximal and distal aspects of bundles (Fig. 5). Not all
NFs within bundles displayed even labeling within the proximal shaft at
2 hr, and some NFs remained unlabeled (Fig. 5); only 45 ± 14% of
NFs within bundles was labeled at this early time (n = 5 axons). By contrast, nearly all filaments were labeled with
anti-biotin by 18 hr (Fig. 5). This delay was not attributable merely
to steric inhibition of newly transporting subunits entering bundles,
because in some sections the center of the bundle was labeled more
strongly than its edges (see Prox. Shaft at 2 hr; Fig. 5).
Although these findings do not exclude the possibility that
biotinylated subunits exchange with preexisting NFs in bundles, the
observed anterograde progression of the appearance of biotinylated
subunits within bundles suggests that bundled NF also may undergo
transport, albeit more slowly than nonbundled NFs. This conclusion is
supported further by our use of biotinylated NF-H rather than NF-L in
these ultrastructural analyses, because NF-H would have been more
likely to undergo more rapid exchange than NF-L (Takeda et al.,
1994).
The temporal distribution of newly transporting subunits also was
probed by the transfection of differentiated cells with a construct
encoding GFP-tagged NF-M (Yabe et al., 1999). After overnight
incubation to allow for the accumulation of GFP-tagged subunits (Yabe
et al., 1999), the cells were fixed and processed for RT97
immunoreactivity. As with microinjected biotinylated subunits,
GFP-tagged NF-M initially was observed along the periphery of axonal
neurites and did not colocalize with the more centrally situated RT97
immunoreactivity (Fig.
6A). After an
additional 2-4 hr of incubation, however, prominent GFP fluorescence
colocalized with RT97 immunoreactivity (Fig. 6A).
These data indicate that newly transporting NF subunits exhibit a
delayed association with bundled NFs.
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Figure 6.
GFP-tagged NF subunits exhibit a delayed
association with bundled NFs. A, Images of cells
transfected 16-24 hr previously with eGFP-NF-M and then fixed and
immunostained with RT97. Arrowheads in the GFP images
denote the neurite hillock. As described in Materials and Methods (see
also Yabe et al., 1999), 16-24 hr of further incubation was required
after transfection for the accumulation of sufficient eGFP-tagged NF-M
for detection. Cells were fixed either as soon as GFP
immunofluorescence was detected (Early) within axons or
2-4 hr later (Later). GFP fluorescence initially was
observed along the periphery of axonal neurites and did not colocalize
with RT97. Colocalization of GFP and RT97 is revealed by
yellow-orange immunofluorescence in
merged images. Note that GFP fluorescence initially is concentrated
along the periphery of axonal neurites and does not exhibit appreciable
colocalization with the centrally situated RT97-labeled bundle. By
contrast, GFP fluorescence colocalizes with the RT97-labeled
bundle 2 hr later. The accompanying graph presents the ratio of GFP
within the center/on the periphery; data are pooled from multiple axons
when GFP first was detected versus 2-4 hr later, as described in
Materials and Methods. Note the significant increase in relative
fluorescence within the center of the axon during this additional
incubation. B, Fluorescent images of the same
transfected cell taken 2 hr apart. Note in the initial image that NF
subunits had distributed along the length of axonal neurite, yet
relatively intense fluorescence within the central aspect of the axonal
shaft was confined to the proximal-most region. An image of the same
neurite 2 hr later revealed that this centrally located intense
fluorescence extended more distally along the shaft
(arrows). The accompanying graph presents compiled data
that were obtained from 10 transfected cells of the relative intensity
of GFP fluorescence along the center of the axon when first detected
and then 2 hr later. Note the presence of a significant
(p < 0.05) increase in intensity within
central segments in this 2 hr interval.
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Auto-fluorescence of GFP-tagged subunits precluded the necessity of
fixation before observation and therefore also afforded the ability to
observe dynamic alteration in the distribution of newly transporting NF
subunits repeatedly within the same axonal neurites. Such analyses
demonstrated that NF subunits had distributed along the length of
axonal neurites within 2 hr (Fig. 6B). In addition,
relatively intense fluorescence was observed within the central aspect
of the proximal-most region of the axonal shaft. Subsequent
observations of the same axonal neurite revealed this centrally located
intense fluorescence to undergo a progressive proximal-distal
accumulation (Fig. 6B). The intense centrally situated GFP fluorescence is consistent with the relatively large number of NFs within the centrally situated bundle as observed in
ultrastructural analyses and in RT97 immunofluorescence (see above).
These data are consistent with our above speculation that bundled NFs
indeed undergo axonal transport but do so more slowly than do
peripherally situated individual NFs. These data do not, however,
exclude the possibility that subunit and/or filament exchange also
occurs along the length of bundled NFs.
The distribution of newly transported NF subunits also was
monitored by pulse-chase radiolabeling. We reasoned that, if bundled NFs undergo relatively slower transport/turnover than do individual NFs, then radiolabeled subunits initially would be recovered
predominantly within individual NFs and only later would be associated
with bundled NFs. Cultures were radiolabeled with
[35S]methionine for 15 min, followed by
replacement of the medium with fresh medium lacking radiolabel, and
were incubated for a total of 4 hr (Shea et al., 1990). Cultures were
harvested at these times and fractionated under conditions that
separate axonal neurites from perikarya (Shea et al., 1993). These
fractions, respectively, were fractionated further to generate
Triton-soluble and Triton-insoluble material. Finally, Triton-insoluble
material from axonal neurites was sedimented over a 1 M
sucrose cushion to separate bundled and individual NFs (see Fig. 3). As
in previous studies (Shea et al., 1990) radiolabeled NF subunits were
associated with axonal Triton-insoluble structures within 15 min.
However, virtually all detectable radiolabeled subunits were retained
by the sucrose cushion at this time (Fig.
7). By contrast, when cultures were
harvested after 4 hr in the absence of additional radiolabel, virtually
all radiolabeled subunits were recovered within the material that
sedimented through the sucrose cushion. Because bundled but not
individual NFs sedimented through this cushion (see Fig. 2), these
findings indicate that radiolabeled subunits exhibited a delayed
association with bundled NFs and therefore suggest that bundled NFs
undergo slower turnover than do individual NFs within axonal neurites.
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Figure 7.
Endogenous axonal NF subunits exhibit a
delayed association with NF bundles. The panels present immunoblot and
autoradiographic analyses of cytoskeletons derived from perikaryal
(Soma) and axonal preparations from cells that were
pulse-labeled for 15 min and immediately harvested or cultures from
which radiolabeled medium was replaced with medium lacking radiolabel
and in which incubation was continued for a total of 4 hr, as
indicated. Axonal material recovered at the interface and material that
sedimented through a 1 M sucrose cushion
(pellet) are indicated. The relative migration of
molecular weight standards is indicated on the left
side of the figure. Immunoreactive and immunoprecipitated
species corresponding to NF-H, NF-M, and NF-L are indicated also. The
accompanying immunoblots (probed with R39) confirm the presence of NF
subunits in cytoskeletons derived from all fractions. Note that
radiolabeled NF subunits are associated with perikaryal and axonal NFs
within 15 min of radiolabeling, yet within axonal neurites are
recovered virtually entirely from the interface and not from the
pellet. Note further that by 4 hr the majority of radiolabeled subunits
is associated with the pellet. Because bundled NFs sedimented through
sucrose under these conditions yet individual NFs were retained at the
interface, these data indicate that NF subunits associate with
individual NFs within axons before their association with bundled
NFs.
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Bundled NFs are relatively resistant to perturbation
in situ
Finally, should bundled NFs indeed represent population(s) of NFs
that are undergoing relatively more NF-NF interactions in situ than those that are situated more peripherally, the
individual NFs, we reasoned that bundled NFs may be more resistant to
treatments that induce perturbation of the axonal cytoskeleton.
Previous studies have demonstrated differential distribution of axonal constituents after the stretching of axonal fibers (Ochs et al., 1989).
Significant NF immunoreactivity was dispersed into the bead-like
swellings, or varicosities, that were generated along the axon after
experimental stretching, whereas longitudinal filamentous profiles
remained within the compressed center of the axonal shaft (Ochs et al.,
1989). Although this stretching technique is not directly applicable to
cultured neuronal cells, a similar morphological profile is generated
after short-term treatment of NB2a/d1 cells with
microtubule-depolymerizing drugs such as colchicine and nocodazole (Shea and Beermann, 1994; Jung et al., 1998). Surviving neurites exhibit an overall thinning in caliber and exhibit periodic
varicosities along their length (Fig. 8)
(see also Shea and Beermann, 1994). We therefore considered that
nocodazole treatment may foster differential distribution of axonal NFs
in a manner analogous to that observed after the stretching of fibers.
Should bundled NFs represent a population of NFs that have undergone
increased NF-NF associations, we reasoned that they would tend to
remain centrally located along the axonal shaft, whereas individual NFs
may distribute more readily within resultant varicosities. To test this
possibility, we examined the distribution of SMI-31, SMI-32, and RT97
immunoreactivity after treatment of differentiated NB2a/d1 cells with
nocodazole for 2 hr (Fig. 8A). Immunoreactivity
toward these antibodies partitioned differentially within resultant
beads and thinned areas of the axonal shaft. Approximately fourfold
more SMI-31 and SMI-32 immunoreactivity was localized within
varicosities as opposed to adjacent areas of the axonal shaft. By
contrast, only ~1.5-fold more RT97 immunoreactivity distributed
within varicosities as opposed to the shaft. These data suggest that
RT97 selectively labels a population of NFs that are relatively
resistant to redistribution. Because RT97 selectively labels bundled
NFs, these data support the interpretation that bundled NFs are
relatively more stable than individual NFs in situ. This
possibility was probed further by monitoring the distribution of
microinjected NF-H within axonal neurites treated as above with
nocodazole at 2 hr after injection (at which time the majority of
injected subunits is associated with individual, peripherally situated
NFs) and at 18 hr after injection (by which time some injected subunits
have associated with bundled NFs). We reasoned that, should bundled NFs
indeed be more resistant than individual NFs to redistribution after
nocodazole treatment, biotinylated subunits should localize within
nocodazole-induced varicosities to a greater extent at 2 hr after
injection than at 18 hr after injection. When microinjected cells were
treated with nocodazole at 2 hr after injection, ~3.5-fold more
biotin immunoreactivity distributed within varicosities versus adjacent areas of the shaft. Conversely, only approximately twofold more biotin
immunoreactivity partitioned within beads versus shafts when nocodazole
treatment was performed 18 hr after injection (Fig.
8B). These findings indicate that a portion of
microinjected subunits attained increased resistance to
nocodazole-induced redistribution at a time by which some subunits have
associated with bundled NFs and suggest that bundled NFs are indeed
relatively more stable than individual NFs in situ.
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Figure 8.
Nocodazole alters axonal neurite
morphology and the distribution of axonal NF immunoreactivity.
A, Immunofluorescent analyses of the distribution of
SMI-31 and RT97 immunoreactivity with and without treatment with
nocodazole for 2 hr, as indicated. Corresponding phase-contrast images
are presented; note that nocodazole induces overall thinning of the
axonal shaft (white arrows) with periodic varicosities
or beads. The accompanying plot profiles present the
distribution of NF immunoreactivity within select regions of axonal
neurites; these regions are presented as insets within
the graphs and are indicated by arrows in the
appropriate phase-contrast image. Note that relatively more RT97
immunoreactivity is retained within shafts than is SMI-31
immunoreactivity. The accompanying bar graph presents a comparison of
the relative distribution of NF immunoreactivity within varicosities
versus adjacent areas of the shaft; SMI-32 immunoreactivity (not
presented in micrographs) also was included in these analyses. Note
that approximately fourfold more SMI-31 and SMI-32 immunoreactivity was
localized within varicosities as opposed to the adjacent areas of
the axonal shaft, whereas only ~1.5-fold more RT97
immunoreactivity distributed within varicosities as opposed to the
shaft. B, Biotin immunoreactivity in cells treated with
nocodazole at 2 and 18 hr after injection, as indicated. Corresponding
phase-contrast images are presented also. The accompanying plot
profiles present the distribution of biotin immunoreactivity within
select regions of axonal neurites; these regions are presented as
insets within the graphs and are indicated by
arrows within the immunofluorescent images. Note that
relatively more immunoreactivity is retained within shafts at 18 hr
after injection. The accompanying bar graph presents the relative
distribution of biotin immunoreactivity within varicosities versus
adjacent areas of the shaft. Note that, when microinjected cells were
treated with nocodazole at 2 hr after injection, ~3.5-fold more
biotin immunoreactivity distributed within varicosities versus adjacent
areas of the shaft. Conversely, only approximately twofold more biotin
immunoreactivity partitioned within beads versus shafts when nocodazole
treatment was performed 18 hr after injection.
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| |
DISCUSSION |
We present evidence herein that axonal NFs consist of at least two
populations that can be distinguished on the basis of differential NF-NF associations and C-terminal phosphorylation. NFs rich in site-specific C-terminal phosphorylation were organized preferentially into bundles. When we monitored the spatial and temporal distribution of newly transported subunits by microinjection, transfection, and
radiolabeling, our analyses further demonstrate that bundled NFs either
do not undergo transport or undergo transport at a rate slower than
that of individual NFs.
Differential transport of individual versus bundled NFs may represent
the morphological equivalent of the broadening of the transport wave
routinely observed for radiolabeled NF subunits in situ
(Nixon and Logvinenko, 1986; Watson et al., 1989; Lasek et al., 1992,
1993; Jung and Shea, 1999). Retardation of the transport of bundled NFs
is likely to result from a preponderance of phosphate-dependent and/or
other physical filament-filament interactions among the bundled NFs
(Leterrier and Eyer, 1987; Shaw and Hou, 1990; Gotow and Tanaka, 1994;
Nagakawa et al., 1995; Leterrier et al., 1996). Previous studies
demonstrated that NFs move apart on disruption of axonal membrane
integrity, indicating that such NFs are not cross-linked physically
(Brown and Lasek, 1993); however, these investigators pointed out that
their findings remained consistent with the data of Shaw and Hou (1990)
in which some NFs were recovered as bundles. Our recovery in
cytoskeletal preparations of both individual and bundled NFs, with
distinct phospho-dependent immunoreactivity, supports and unifies the
conclusions of these previous studies. We consider it likely that
peripherally situated NFs and centrally situated bundled NFs represent
a continuum in situ rather than discrete populations and
that individual NFs within these experimentally separable categories
are also likely to display broad ranges of transport and turnover
rates. This possibility is supported by the demonstration that
phosphorylated and nonphosphorylated NFs can exist side by side within
axons of cultured neurons (Brown, 1998). Further studies directed
toward elucidating any potential differences among NFs within these two
broadly characterized "populations" are warranted.
If, as our data and that of previous studies suggest, bundled NFs
represent a population that is undergoing relatively more NF-NF
interactions than are individual NFs, these findings are consistent
with the notion that NF-NF interactions may compete with the
interactions of NFs with their transport motor and foster increased
residence time of some NFs within axons (Lasek et al., 1992; Nixon,
1993, 1998). This interpretation is also consistent with the
observation that NFs containing the most highly phosphorylated NF-H
variants undergo axonal transport at the slowest rates (Lewis and
Nixon, 1988), whereas poorly phosphorylated NFs travel at faster rates
(Jung et al., 2000) and with the observed slowing of NF axonal
transport and increase in C-terminal NF phosphorylation after the
inhibition of phosphatase activity in situ (Jung et al.,
1999). In this regard, newly transporting NF subunits are associated
in situ either directly or indirectly with the motor protein
kinesin (Yabe et al., 1999, 2000). Furthermore, this association is
disrupted by site-specific C-terminal NF phosphorylation such that
RT97-immunoreactive NF subunits do not interact with kinesin (Yabe et
al., 2000). These findings are consistent with the interpretation that
bundled NFs (which are preferentially reactive with RT97) are
associated less frequently with their motor complex than are individual
NFs (for review, see Shea and Yabe, 2000). Relatively rapid
translocation of some NF subunits along the axonal length (i.e.,
individual NFs) is consistent with in vivo observation of
the transport of radiolabeled NF subunits at rates 100 times greater
than the bulk of translocating subunits (Lasek et al., 1993).
Observation of faster-moving individual NFs and slower-moving bundled
NFs is also consistent with the recent demonstration by Wang et al.
(2000) of the rapid axonal transport of NFs interrupted by pauses. In
these analyses substantial gaps were observed in the axonal NF array.
These gaps as a whole did not translocate, although some NFs traversed
the gaps, demonstrating NF transport over a broad range of rates. It is
possible that the faster-moving NFs correspond to the individual NFs in
our analyses, whereas the slower-moving NFs (i.e., surrounding the
gaps) correspond to the bundled NFs in our analyses. The periodic
pausing exhibited by the faster-moving NFs may correspond further to a
period of limited interaction with other NFs that could, if prolonged
sufficiently, lead to bundling. Further investigations in this regard
will be of interest.
Some NF subunits enter axons and undergo transport while still in
Triton-soluble form (Jung et al., 1998), and such subunits include
precursors for NF assembly (Shea et al., 1990, 1997; Shea, 1994; Jung
et al., 1998; Yabe et al., 1999). Although our findings of differential
transport rates for axonal NFs are in agreement with existing polymer
transport models (for review, see Baas and Brown, 1997), they also
leave open the possibility that monomeric/oligomeric NF-H subunits
selectively may incorporate at least to some degree into preexisting
NFs and/or assemble onto the ends of elongating NFs. Several lines of
evidence support this possibility, including (1) detection of NF-H
immunoreactivity by immuno-EM analyses in the absence of any
filamentous profiles within neurons in situ (Gotow and
Tanaka, 1994); (2) transport of monomeric/oligomeric NF-M in the
absence of axonal NFs (Terada et al., 1994); (3) slow axonal transport
of some NF subunits in punctate, apparently nonfilamentous form (Yabe
et al., 1999); and (4) selective accumulation of newly transported NF-H
into Triton-insoluble structures within the distal-most segment of
growing axons (Yabe et al., 1997). Incorporation of NF subunits into
bundled NFs may occur via exchange of individual NFs and/or subunits
along the length of bundled NFs. Our observation of stronger labeling
of proximal bundled NFs before distal NFs leads us to speculate that
the bundle is indeed translocating, albeit more slowly than surrounding
individual NFs. However, whether bundled NFs undergo transport or not
does not exclude the possibility that the exchange of NF subunits still
may occur along the length of bundled NFs.
Controversy exists regarding three major aspects of NF biology: (1)
whether axonal NFs constitute a single population or multiple populations, (2) whether NFs are cross-linked in situ, and
(3) whether axonal NFs undergo turnover within the axonal cytoskeleton by replacement with new NFs or instead by incorporation of newly translocating subunits into existing, nonmoving NFs. To the extent that
one can extrapolate findings in a cell culture model to the situation
in situ, the data of the present study address each of these
controversies. In summary, our data support the interpretations that
axonal NFs consist of multiple populations that undergo axonal transport although over a wide range of rates and that some, but not
all, NFs apparently are cross-linked physically in situ. The data at present do not allow us to resolve whether axonal NFs undergo
replacement by newly transporting polymers, by subunit exchange, or by
both. The two NF populations observed in these growing axons may
represent the developmental precursor of a stationary population of NFs
that is replaced along its length by newly transported NFs (Nixon,
1993, 1998). Furthermore, establishment of a stationary cytoskeleton
may favor the exchange of NF subunits with nonmoving NFs as a major
mode of NF turnover (Hirokawa et al., 1997). Although such a scenario
in essence can be envisioned as a logical developmental exaggeration of
the NF populations observed herein, clarification of this issue
requires further investigation of NF dynamics in situ.
| |
FOOTNOTES |
Received Oct. 19, 2000; revised Dec. 12, 2000; accepted Dec. 15, 2000.
This research was supported by the National Science Foundation and the
National Institute of Child Health and Human Development. We thank our
colleagues Anthony Brown and Ralph Nixon for their helpful comments on
this manuscript, Ron Liem for his generous gift of neurofilament cDNA,
Daniela Ortiz for technical assistance, and Louise Trakimas for
excellent assistance with electron microscopy.
Correspondence should be addressed to Dr. Thomas B. Shea, Center for
Cellular Neurobiology and Neurodegeneration Research, Department of
Biological Sciences, University of Massachusetts-Lowell, One
University Avenue, Lowell, MA 01854. E-mail: Thomas_Shea{at}uml.edu.
| |
REFERENCES |
-
Anderton BH,
Breinburg D,
Downes MJ,
Green PJ,
Tomlinson BE,
Ulrich J,
Wood JN,
Kahn J
(1982)
Monoclonal antibodies show that neurofibrillary tangles and neurofilaments share antigenic determinants.
Nature
298:84-86[Medline].
-
Baas PW,
Brown A
(1997)
Slow axonal transport: the polymer transport model.
Trends Cell Biol
7:380-384[Medline].
-
Brady ST
(1993)
Motor neurons and neurofilaments in sickness and health.
Cell
73:1-3[ISI][Medline].
-
Brown AT
(1997)
Visualization of single neurofilaments by immunofluo-rescence microscopy of splayed axonal cytoskeletons.
Cell Motil Cytoskeleton
38:133-145[ISI][Medline].
-
Brown AT
(1998)
Contiguous phosphorylated and nonphosphorylated domains along axonal neurofilaments.
J Cell Sci
111:455-467[Abstract].
-
Brown A,
Lasek RJ
(1993)
Neurofilaments move apart freely when released from the circumferential constraint of the axonal plasma membrane.
Cell Motil Cytoskeleton
26:313-324[ISI][Medline].
-
Chin SSM,
Liem RKH
(1989)
Expression of rat neurofilament proteins NF-L and NF-M in transfected non-neuronal cells.
Eur J Cell Biol
50:475-490[ISI][Medline].
-
Galbraith JA,
Reese TS,
Schlief ML,
Gallant PE
(1999)
Slow transport of unpolymerized tubulin and polymerized neurofilament in the squid giant axon.
Proc Natl Acad Sci USA
96:11589-11594[Abstract/Free Full Text].
-
Gotow T,
Tanaka J
(1994)
Phosphorylation of neurofilament H subunit as related to arrangement of neurofilaments.
J Neurosci Res
37:691-713[ISI][Medline].
-
Hirokawa N,
Terada S,
Funakoshi T,
Takeda S
(1997)
Slow axonal transport: the subunit transport model.
Trends Cell Biol
7:384-388[Medline].
-
Jung C,
Shea TB
(1999)
Regulation of neurofilament axonal transport by phosphorylation in optic axons in situ.
Cell Motil Cytoskeleton
43:230-240.
-
Jung C,
Yabe J,
Wang F-S,
Shea TB
(1998)
Neurofilaments are present within axonal neurites before incorporation into Triton-insoluble structures.
Cell Motil Cytoskeleton
40:44-58[ISI][Medline].
-
Jung C,
Yabe JT,
Lee S,
Shea TB
(2000)
Hypophosphorylated neurofilament subunits undergo axonal transport more rapidly than more extensively phosphorylated subunits in situ.
Cell Motil Cytoskeleton
47:120-129[ISI][Medline].
-
Koehnle TJ,
Brown A
(1999)
Slow axonal transport of neurofilament protein in cultured neurons.
J Cell Biol
144:447-458[Abstract/Free Full Text].
-
Lasek RJ
(1986)
Polymer sliding in axons.
J Cell Sci Suppl
5:161-179[Medline].
-
Lasek RJ,
Paggi P,
Katz MJ
(1992)
Slow axonal transport mechanisms move neurofilaments relentlessly in mouse optic axons.
J Cell Biol
117:607-616[Abstract/Free Full Text].
-
Lasek RJ,
Paggi P,
Katz MJ
(1993)
The maximum rate of neurofilament transport in axons: a view of molecular transport mechanisms continuously engaged.
Brain Res
616:58-64[ISI][Medline].
-
Leterrier J-F,
Eyer J
(1987)
Properties of highly viscous gels formed by neurofilaments in vitro. A possible consequence of specific inter-filament cross-bridging.
Biochem J
245:93-101[ISI][Medline].
-
Leterrier J-F,
Käs J,
Hartwig J,
Vegners R,
Janmey P
(1996)
Mechanical effects of neurofilament cross-bridges.
J Biol Chem
271:15687-15694[Abstract/Free Full Text].
-
Lewis SE,
Nixon RA
(1988)
Multiple phosphorylated variants of the high molecular mass subunit of neurofilaments in axons of retinal cell neurons: characterization and evidence for their differential association with stationary and moving neurofilaments.
J Cell Biol
107:2689-2701[Abstract/Free Full Text].
-
Nagakawa T,
Chen J,
Zhang Z,
Kanai Y,
Hirokawa N
(1995)
Two distinct functions of the carboxyl terminal tail domain of NF-M upon neurofilament assembly: cross-bridge formation and longitudinal elongation of filaments.
J Cell Biol
129:411-429[Abstract/Free Full Text].
-
Nixon RA
(1993)
The regulation of neurofilament protein dynamics by phosphorylation: clues to neurofibrillary pathology.
Brain Pathol
3:29-38[ISI][Medline].
-
Nixon RA
(1998)
The slow transport of cytoskeletal proteins.
Curr Opin Cell Biol
10:87-92[ISI][Medline].
-
Nixon RA,
Lewis SE
(1986)
Differential turnover of phosphate groups on neurofilament subunits in mammalian neurons in vivo.
J Biol Chem
261:16298-16301[Abstract/Free Full Text].
-
Nixon RA,
Logvinenko KB
(1986)
Multiple fates of newly synthesized neurofilament proteins: evidence for a stationary neurofilament network distributed nonuniformly along axons of retinal ganglion cell neurons.
J Cell Biol
102:647-659[Abstract/Fre
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ABSTRACT
INTRODUCTION
