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The Journal of Neuroscience, July 15, 2002, 22(14):5982-5991
Microtubule Reconfiguration during Axonal Retraction Induced by
Nitric Oxide
Yan
He,
Wenqian
Yu, and
Peter W.
Baas
Department of Neurobiology and Anatomy, Drexel University College
of Medicine, Philadelphia, Pennsylvania 19129
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ABSTRACT |
Axonal retraction is induced by different types of physiological
cues and is responsible for the elimination of mistargeted axons. There
is broad agreement that alterations in the cytoskeleton underlie axonal
retraction. The prevailing view is that axonal retraction involves a
wholesale depolymerization of microtubules and microfilaments. However,
axons retracting physiologically display a very different morphology
than axons induced to retract by experimental depolymerization of
microtubules. Experimental depolymerization of microfilaments actually
prevents retraction rather than causing it. We have proposed an
alternative hypothesis, namely that axonal retraction involves a
backward retreat of cytoskeletal elements rather than their wholesale
depolymerization. In the present study, we sought to test this
hypothesis with regard to microtubules. When a donor of nitric oxide
was applied to cultured chick sensory neurons, the majority of axons
retracted dramatically within 30-60 min. Retracting axons were
characterized by an enlarged distal region, a thin trailing remnant,
and sinusoidal bends along the shaft. Quantitative immunofluorescence
analyses showed no detectable loss of microtubule mass during
retraction, even with regard to the most labile microtubules. Instead,
microtubules were reconfigured into coiling and sinusoidal bundles to
accommodate the shortening of the axon. Stabilization of microtubules
by taxol did not prevent the retraction, even at concentrations of the drug that actually caused microtubule levels to increase. The retractions induced by nitric oxide were remarkably similar to those
observed when motor proteins are manipulated, suggesting that these
retractions may result from alterations in the activities of the motors
that configure microtubules.
Key words:
microtubule; axon; neuron; retraction; nitric oxide; actin
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INTRODUCTION |
The normal functioning of the
nervous system depends on precise connections that wire individual
neurons together into an orderly network. Establishment of the neural
network involves both progressive events, which include neurogenesis
and axonal outgrowth, and regressive events, which include neuronal
death and axonal retraction (Cowan et al., 1984 ; Bernstein and
Lichtman, 1999). Selective retraction of newly formed axons occurs
extensively in the developing nervous system, where it functions as an
alternative mechanism to neuronal death to eliminate inappropriate
axonal projections. Examples of such selective axonal retraction are seen in the withdrawal of all but one axon from each muscle fiber (Gan
and Lichtman, 1998) and the withdrawal of occipital neuronal axons
projecting to the spinal cord (Stanfield et al., 1982). In addition,
the normal growth and pathfinding of most axons involve alternate bouts
of retraction and elongation (Kaethner and Stuermer, 1992; Godement et
al., 1994), and a variety of external cues have been identified that
can cause axons to retract (for review, see Tessier-Lavigne and
Goodman, 1996). It is known that axonal growth involves the elaboration
of the cytoskeletal arrays that provide architectural support for the
axon, and there is broad agreement that the cytoskeleton must undergo
alterations to elicit retraction. However, the nature of these
alterations remains largely unknown.
The prevailing view is that cytoskeletal elements depolymerize
extensively during physiological axonal retraction (Song and Poo,
1999). This view is largely based on pharmacological studies showing
that wholesale microtubule depolymerization can indeed cause axons to
retract (Yamada et al., 1970; Daniels, 1973). However, axons retracting
because of microtubule loss display a thin beaded morphology
that is very different from axons retracting physiologically, which
generally thicken distoproximally and show sinusoidal bends rather than
beads along their length (Bandtlow et al., 1990; Bastmeyer and
Stuermer, 1993; Dent and Meiri, 1998; Nagashima et al., 1999). Furthermore, pharmacological depletion of microfilaments actually prevents axons from retracting rather than causing them to retract (Solomon and Magendantz, 1981; Joshi et al., 1985). We have proposed a
different model for how the cytoskeleton is altered during
physiological axonal retraction. In our model, instead of extensive
depolymerization, microtubules and microfilaments are reconfigured such
that they retreat backward in the retracting axon. Our model is based
on recent studies that indicate that molecular motors in neurons are
capable of configuring and translocating cytoskeletal polymers in much
the same way that motors function in the mitotic spindle (Baas and
Ahmad, 2001). In the present study, we have focused specifically on
microtubules and performed a battery of tests to ascertain whether
physiological axonal retraction involves wholesale depolymerization or
reconfiguration of the microtubule polymers. Nitric oxide, widely
recognized to induce axonal retractions during the development of the
vertebrate nervous system (Cramer et al., 1998; Ernst et al., 2000),
was used to induce retractions of axons in cultures of chick sensory neurons.
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MATERIALS AND METHODS |
Cell culture. Cultures of chick sensory neurons were
prepared as described previously (Yu and Baas, 1995; Ahmad et al.,
2000). Briefly, dorsal root ganglia were dissected from embryonic day 10 chicks and digested in 0.25 mg/ml trypsin and collagenase (both from
Worthington Biochemical, Lakewood, NJ) at 37°C for 15 min. After
neutralization of the enzymes with 10% fetal bovine serum (FBS;
Hyclone, Logan, UT), the ganglia were triturated into a single-cell
suspension with a Pasteur pipette, which had been fire polished to
obtain a narrow opening. Cells were then diluted in a modified
L-15-based medium containing Leibovitz's L-15 (Invitrogen, Grand Island, NY), 0.6% glucose (Sigma, St. Louis, MO), 2 mM L-glutamine (Invitrogen), 100 U of penicillin and 100 µg of streptomycin per milliliter (Sigma), 0.6% methylcellulose (Dow Chemical, Midland, MI),
10% FBS, and 25 ng/ml nerve growth factor (Upstate Biotechnology, Lake
Placid, NY). Cells were plated onto prephotoetched glass coverslips
(Bellco Glass, Vineland, NJ), which had been attached to the bottom of
a 35 mm plastic Petri dish into which had been drilled a 1 cm hole. The
photoetched marks on the coverslip assist in relocating individual
cells during the experimental procedures. Cells were kept at 37°C in
normal air. By 16-20 hr after plating, neurons had generated
distinguishable axons that were suitable for the experiments.
Experimental treatments. The nitric oxide donor
3-(2-hydroxy-1-methyl-2-nitrosohydrazino)-N-methyl-1-propanamine
(NOC-7) was purchased from Calbiochem (La Jolla, CA). A 100 mM stock solution of NOC-7 in 100 mM NaOH was frozen in aliquots. At the time of the experiment, aliquots of the NOC-7 stock solution were thawed quickly, diluted to 1 mM in the modified
L-15-based medium but without NGF, and warmed to 37°C. The medium
from the cultures was removed and replaced with the medium deficient in
NGF but containing NOC-7. Cultures were then incubated for various
times depending on the particular experiment. Nocodazole (prepared from a 10 mM stock solution in DMSO and diluted to a
final concentration of 10 µg/ml) and taxol (prepared from a 10 mM stock solution in DMSO and diluted to a final
concentration of 100 nM) were prepared in
complete medium and applied to cells in the same manner. When NOC-7 was
added to taxol-treated cultures, the medium contained NGF during the
first 30 min in taxol alone, and then the NGF was removed during
subsequent treatment with taxol and NOC-7 together.
Sample preparation for fluorescence microscopy. To prevent
detachment of axons from the relatively poorly adherent plain glass coverslip, cells were simultaneously fixed and extracted for
fluorescence microscopy using a method described previously (Ahmad et
al., 2000). Effective extraction of free tubulin was demonstrated by the marked diminution of fluorescence intensity for tubulin in nocodazole-treated cultures. Briefly, cells were fixed and extracted for 15 min in a solution containing 60 mM
1,4-piperazinediethanesulfonic acid, 25 mM HEPES,
10 mM EGTA, and 2 mM
MgCl2, pH 6.9, as well as 4% paraformaldehyde,
0.15% glutaraldehyde, and 0.2% Triton X-100. Cells were then rinsed
extensively in PBS, pH 7.4. To stain for microtubules alone, fixed
cells were exposed to a blocking solution containing 1% bovine serum
albumin in PBS for 30 min and then incubated for 1 hr at room
temperature with a 1:100 dilution of a monoclonal anti- -tubulin
antibody that had been conjugated directly to Cy3 (Sigma). After
extensive rinsing in PBS, the cultures were mounted in a medium that
reduces photobleaching (0.212% N-propylgallate in 50%
glycerol and 50% PBS). To double-label microtubules and microfilaments, the same staining protocol was followed except that the
incubation was with a combination of the tubulin antibody and a 1:40
dilution of Alexa Fluor 488 phalloidin (Molecular Probes, Eugene, OR).
To double-label tyrosinated and total microtubules, cells were first
incubated at 4°C overnight with a rat anti-tyrosinated tubulin
antibody named YL 1/2 (Serotec, Raleigh, NC) at a 1:100 dilution. The
next day, the cultures were exposed for 1 hr to an FITC-conjugated goat
anti-rat IgG (1:200) at 37°C and then exposed, after extensive
rinsing, to the Cy3-conjugated -tubulin antibody (1:100) for 1 hr at
room temperature. The FITC secondary antibody (Jackson ImmunoResearch,
West Grove, PA) was found to have minimal cross-reactivity with
mouse antigens in control experiments.
Light microscopy. For phase-contrast microscopy, cells were
observed with an Axiovert 200M inverted microscope (Zeiss,
Munchen-Hallbergmoos, Germany), and images were acquired using a
charge-coupled device (Zeiss AxioCam b/w) and AxioVision 3.0 software
(Zeiss). All of the fluorescence work was performed on the same
microscope, except for the experiment involving tyrosinated
microtubules. The FITC and Cy3 filters on our Axiovert 200M have
relatively narrow excitation and emission spectra, which minimizes
potential bleed-through between the channels. Nevertheless, because Cy3
and FITC have slightly overlapping spectra, we also empirically
confirmed that there was no detectable bleed-through of our samples
visualized at our microscope and software settings. As with the
phase-contrast work, fluorescence images were captured by the Zeiss
AxioCam running on the AxioVision 3.0 software.
For the experiments involving double-labeling of tyrosinated and
-tubulin, our goal was to obtain a ratio image, each pixel of which
displays the ratio of fluorescence intensity between the two channels.
To accomplish this, we used the Zeiss Pascal laser-scanning confocal
microscope. The tyrosinated tubulin (FITC) and -tubulin (Cy3) scans
were captured sequentially (rather than simultaneously), using
configuration and laser attenuation settings that were kept the same
for all imaged cells. The pinhole of the confocal was maximally opened,
because we did not obtain optical sections for this application. Any
bleed-through between the FITC and Cy3 channels was eliminated by
activating only the appropriate excitation wavelength and
photomultiplier (and leaving that of the other channel off). The
fluorescence ratio per pixel of the FITC and Cy3 images was calculated
and presented as a third image by using the ratio function of the
laser-scanning microscope 5 Pascal software (version 2.8). All
images were processed in the same way so that they could be compared
quantitatively. The ratio images were exported into NIH Image (version
1.62; NIH freeware) and were assigned pseudocolors that represent the
relative ratio values. We chose the pseudocolor scale termed
"fire-1," in which the highest intensity is shown in white, the
lowest intensity is shown in black, and intermediate intensities are
shown in shades of yellow and red.
Quantification of fluorescence intensity. In the
single-label experiments, the total microtubule mass in the retracted
axons was compared with that in the control axons. Microtubule mass was
quantified by measuring the fluorescence intensity of -tubulin staining. Fluorescence images of both the retracted axons and control
axons were captured at the same camera and microscope settings.
Overexposure and underexposure were avoided by using the color-coded
live window in the AxioVision 3.0 software. The total fluorescence
intensity of each observed axon was measured as arbitrary fluorescence
units (AFU) using the NIH Image 1.62 software. Because axons of
different lengths were included in our analyses, we calculated the
fluorescence intensity per unit length (AFU per micrometer) of the
axons, which was obtained by dividing the total AFU by the length of
the same axon. The means of the fluorescence intensity per unit length
of the retracted axons and control axons were compared using the
two-tailed, unpaired Student's t test. A p value
of <0.05 was used as the standard for statistical significance in this
study. All statistical analyses and graphing were done using Prism 3.0 software (GraphPad Software, San Diego, CA). For the experiments
involving taxol, images were displayed in the same quantitative
pseudocolor format used to display the ratio images (as noted above).
So that all of the quantitative studies could be compared relative to
one another, the mean for the control microtubule levels in each
experiment was adjusted to an AFU of 100, and other values were
adjusted accordingly.
Final processing of all images was done using Adobe PhotoShop 6.0 (Adobe Systems, San Jose, CA).
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RESULTS |
Morphology of axons exposed to nitric oxide
Nitric oxide is a physiological factor that is released as a
gas within the nervous system. Several lines of evidence indicate that
nitric oxide is a key factor for inducing large-scale retractions of
mistargeted axons during the pruning of the developing nervous system
(Cramer et al., 1998; Mize et al., 1998). In a recent study, Ernst et
al. (2000) reported that donors of nitric oxide introduced into the
medium of cultures of chick sensory neurons release nitric oxide gas,
and that this results in the retraction of a portion of the axons.
These studies primarily used the nitric oxide donor known as
3-morpholino-sydnonimine (SIN-1), which was shown to induce
retractions at concentrations that did not typically induce cell death
or any detectable nonspecific damage to the neurons. We found that
NOC-7, another potent nitric oxide donor (Ernst et al., 2000), was even
more effective at inducing retractions and even less apt to cause
nonspecific damage at fairly high concentrations (as assessed using the
criteria of Ernst et al., 2000). In addition, we found that axonal
retractions were more robust and dramatic if the neurons were cultured
on plain glass as opposed to plastic or a substrate such as polylysine
or laminin (Ahmad et al., 2000). All of the quantitative studies
reported here were performed using NOC-7, but a small number of
qualitative studies with SIN-1 produced essentially the same results.
Dissociated chick sensory neurons generate multiple neurites in
culture, all of which are axonal as indicated by their length, uniform
diameter, and expression of axonal markers (Smith, 1998). We selected
for our studies axons of medium thickness, each with a total length of
~50-200 µm, which sometimes included one to four primary axonal branches.
After the addition of NOC-7, four different types of axonal
behaviors were observed and were defined as follows: "axonal
retraction" was defined as the shortening of the axon by >5 µm,
"growth-cone collapse only" was defined as the shortening of the
axon by <5 µm but with the collapse of the growth cone from a broad
structure to a narrow structure, "axonal elongation" was defined as
the continued growth of the axon, and "no change" was defined as
axons that do not show any of the above three behaviors; these axons showed no significant change in length or growth-cone morphology during
the treatment. Notably, all of these four behaviors were represented in
control cultures treated with NOC-7 carrier only, but the proportions
of axons in each category changed on exposure of the cultures to the
nitric oxide donor. In control cultures, 30.6% of 50 axons from 15 neurons retracted, 2% showed growth-cone collapse only, 65.3%
elongated, and 2% showed no change. After 30 min of exposure to NOC-7,
67% of 273 axons from 100 neurons retracted, 19% showed growth-cone
collapse only, 10% elongated, and 4% showed no change. These data are
summarized in Figure
1D. Measurement of
axonal length showed that axons retracted an average of 41 ± 2.8 µm (mean ± SEM) during the first 30 min in NOC-7, whereas axons
of control cells elongated an average of 2.3 ± 5.8 µm in 30 min. (Rates of axonal growth in control cultures were much faster
during the next 30 min, suggesting that the exchange of medium within
itself stalled the growth somewhat.) Axons continued to retract with
additional time in NOC-7, but given that retraction was so dramatic in
neurons exposed for only 30 min, we performed all of our cytoskeletal
studies on neurons at this time point.
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Figure 1.
The morphology of axons induced to retract by
NOC-7. Cultures of chick sensory neurons were treated with 1 mM NOC-7 (nitric oxide donor). The majority of the axons in
the culture showed significant retraction in response to the NOC-7
treatment over the first 30 min, as shown here. Three examples are
shown in A-C, wherein the first panel of
each pair shows the axons immediately before the addition of NOC-7, and
the second panel of each pair shows the axons 30 min
after the addition of NOC-7. The morphology of the retracted axons is
characterized by a retraction bulb (arrows), a trailing
remnant (white arrowheads), and sinusoidal bends along
the axonal shaft (black arrowheads). D,
The percentages of different axonal behaviors after NOC-7 treatment,
compared with the percentages for control cultures (for more details,
see Results). Scale bar, 20 µm.
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Our studies focused specifically on the axons that showed retraction
during the first 30 min of NOC-7 treatment. Phase-contrast images of
these axons show that most of them are characterized by an enlarged
distal region, a thin trailing remnant, and sinusoidal bends along the
shaft of the axon (Fig. 1). This retraction morphology resembles that
of axons retracting in vivo, which are also characterized by
a distal enlargement termed the "retraction bulb" (Riley, 1981; Gan
and Lichtman, 1998). The trailing remnant was not included in the
length measurements.
Changes in cytoskeletal configuration during axonal retraction
In a first set of experiments on the cytoskeleton, we wanted
to obtain a global sense of cytoskeletal configuration in retracting axons. For these experiments, after axons retracted in response to
NOC-7, cultures were extracted and fixed simultaneously, as outlined in
Materials and Methods, and then double-labeled for microtubules and
microfilaments. The axons stained for both of these cytoskeletal
elements in both control and NOC-7-treated axons (Fig.
2). In control axons, microtubules
appeared as a continuous dense bundle along the length of the axon that
oftentimes splayed apart in the region of the growth cone. Staining for
microfilaments was generally dim along the length of the axon but was
bright in the peripheral regions of the growth cone. In the axons
induced to retract by NOC-7, the distal enlarged region was
particularly brightly stained for microtubules. Microtubules showed
coiling and sinusoidal bending both in the distal enlargement and along the shaft, suggesting a backward retreat and reconfiguration of microtubules to accommodate shortening of the axon. Microfilaments were
redistributed into a clump in the distal region of the axonal shaft, as
would be expected if the peripheral growth-cone microfilaments folded
inward during growth-cone collapse. Interestingly, these microtubule and microfilament reconfigurations were also observed in
axons just starting to show signs of retraction, even before a
detectable reduction in the length of the axon. Figure
2B shows such an axon, which displays changes in
microtubule and microfilament configuration that are similar to those
seen for an axon that has retracted substantially (Fig. 2C),
both of which are notably different from control axons showing no
retraction (Fig. 2A). Qualitatively, there appears to
be no detectable reduction in the total fluorescence intensity of
either microtubules or microfilaments during axonal retraction.
Additional dramatic examples of the coiling and bending of microtubules
during axonal retraction are shown in Figure
3.
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Figure 2.
Reconfiguration of cytoskeletal components
in axons exposed to NOC-7. Double fluorescence labeling of microtubules
and microfilaments reveals reconfiguration of both cytoskeletal
components in axons exposed to NOC-7. For each axon, the first
fluorescence image shows microtubules, the second
image shows microfilaments, and the third image
shows a color overlay in which microtubule staining is
red and microfilament staining is green.
A, In a control axon, microtubules appear as a tight
straight bundle, and microfilaments are concentrated in the growth
cone. B, This axon was fixed immediately after the
growth cone had collapsed in response to NOC-7, before any retraction
of the axonal shaft occurred. Bending of the microtubules and
redistribution of the microfilaments are already observed at this
stage. C, This axon was fixed after retraction was well
underway. Microtubules are reconfigured into coiled and sinusoidal
bundles, and microfilaments are concentrated in the most distal region
of the retracted axon. B, C, Phase-contrast images
before and after retraction are shown at a lower magnification. Scale
bars, 20 µm.
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Figure 3.
Microtubule reconfiguration in axons induced to
retract by NOC-7. Single-label studies for microtubules in retracting
axons are shown here. Phase-contrast images are shown (at a lower
magnification) above corresponding -tubulin images.
Numerals represent the minutes after NOC-7 addition to
the culture. A, Filamentous microtubule bundles are
present in the retraction bulb (arrows).
B, Microtubules have accumulated in a distal-proximal
manner in the retracted region (the arrow shows the
brightest region at the tip of the retraction bulb). The
coiling of microtubule bundles in the shaft corresponds to the contour
of the axon (arrowhead). C, Sinusoidal
bending of microtubule bundles accommodates the shortening of the axon
(arrows). Scale bars, 20 µm.
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Quantitative analyses of microtubule levels in retracting and
control axons
To quantitatively assess microtubule levels in control and
retracting axons, we obtained images of axons under identical
microscope, camera, and software settings and determined the
fluorescence intensity per micrometer length of the axon. We also
included in these studies axons that had undergone retraction in
response to treatment with nocodazole, a potent microtubule
depolymerizing agent. Nocodazole was applied at 10 µg/ml for 30 min
before simultaneous extraction and fixation. Figure
4 shows that the morphology of the axons
retracting in response to nocodazole is markedly different from that of
the axons retracting in response to NOC-7. Nocodazole-induced axonal
retraction is characterized by the formation of multiple bead-like
enlargements along the shaft and shortening and withering of the axon.
Also, in some cases, nocodazole treatment induced the formation of fine
lateral extensions from the axonal shaft [as has been observed
previously in some studies in which axons were treated with
antimicrotubule drugs (Bray et al., 1978; Joshi et al., 1986)].
Microtubule staining was very dim in these axons and showed no
filamentous appearance (Fig. 4, last panel).
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Figure 4.
Retraction of axons in response to nocodazole.
Axons treated for 30 min with nocodazole retracted with a morphology
distinct from that of axons retracting in response to NOC-7. Treatment
with 10 µg/ml nocodazole induced axons to retract with a morphology
characterized by bead formation along the axon (arrow),
outgrowth of fine lateral extensions (arrowhead), and
the withering of the axonal shaft. Immunofluorescence staining of
-tubulin revealed a dramatic depletion of microtubules in axons
treated with nocodazole. Scale bar, 20 µm.
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Figure 5 shows the results of
quantitative analyses on the levels of microtubules in control axons
and axons induced to retract by either NOC-7 or nocodazole. Two values
were calculated for each of the two types of retracting axons: (1) one
was obtained by dividing the total fluorescence intensity by the axonal
length after retraction, and (2) the other was obtained by dividing the total fluorescence intensity by the axonal length before retraction. Figure 5A (together with the figure legend) explains how
these quantitative measurements were obtained. In NOC-7-treated
cultures, the mean AFU per micrometer calculated for the length after
retraction is significantly higher than that of control axons, whereas
the mean AFU per micrometer calculated for the original length is essentially the same as that of the control axons (Fig. 5B).
In contrast, nocodazole-treated cultures showed a significant reduction in microtubule levels to less than half of control levels (Fig. 5C). These results show that there is significant
microtubule depolymerization during axonal retraction induced by
nocodazole, but that there is no detectable loss of microtubules during
axonal retraction induced by NOC-7. The nocodazole experiment also
demonstrates that our cofixation-extraction method indeed extracted
free tubulin effectively, which is necessary for accurate assessment of
polymer levels.
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Figure 5.
Quantitative data on microtubule levels in axons
induced to retract by treatment with either NOC-7 or nocodazole. The
method used to quantify microtubule levels is shown in
A. Phase-contrast images of a representative neuron show
two axons before and after 30 min of treatment with 1 mM
NOC-7. Both axons retracted significantly. For each axon, the original
length (LO) and length after retraction
(LR) were measured. After simultaneous
fixation/extraction to preserve microtubules while releasing free
tubulin, neurons were immunostained for -tubulin. Total fluorescence
intensity within the axon (It) and background
fluorescence intensity in a region alongside the axon (Ib) were
measured in AFU. The difference between It and Ib represents the
fluorescence intensity from the axon, adjusted for any nonspecific
background fluorescence. To enable us to compare the microtubule levels
in axons of different lengths, we calculated for each axon the
fluorescence per micrometer of axonal length (AFU per micrometer) by
dividing the (It  Ib) by either LO and
LR for treated axons (see the equations) and by
LO only for control axons. B, Quantification
of NOC-7-treated axons compared with controls. C,
Quantification of nocodazole-treated axons compared with controls.
B, C, The first bar in each
graph shows control microtubule levels in parallel
cultures prepared identically at the same time as the experimental
cultures for the immunofluorescence analyses. The second
bar in each graph shows the microtubule levels
(per micrometer) in drug-treated axons, calculated for LO.
The third bar in each graph also shows
microtubule levels in treated axons, but calculated for LR.
Separate control axons were analyzed for each experiment, and their
mean AFU per micrometer values were normalized to 100 so that data from
the different experiments could be compared. The two-tailed Student's
t test (*p < 0.05) shows a
significant difference between control axons and NOC-7-treated axons
when the calculations are performed using LR. However,
there is no significant difference between the control and the
NOC-7-treated axons when LO is used. In the case of the
nocodazole-treated axons, there is a significant difference between
controls and the treated axons calculated in both ways. Thus, there is
no detectable microtubule loss in axons induced to retract by NOC-7; in
contrast, there is dramatic microtubule loss in axons induced to
retract by nocodazole. Scale bar, 20 µm.
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Quantitative analyses on levels of microtubules enriched with
tyrosinated tubulin during axonal retraction
Previous studies have established that axons contain populations
of stable microtubule polymer and labile microtubule polymer (Baas and
Black, 1990), and that the most distal region of the axon contains
polymer that is potentially even more labile than that found elsewhere
in the axon (Ahmad et al., 1993). The labile polymer undergoes more
rapid bouts of subunit turnover, and this is true regardless of whether
there is net assembly or disassembly of microtubules. With time after
assembly, the -tubulin that comprises the microtubules is gradually
detyrosinated such that older, more stable regions of the microtubules
contain less tyrosinated tubulin, whereas the more labile regions of
the microtubules contain higher levels of tyrosinated tubulin (for
discussion, see Baas et al., 1991). The fact that we observed no
reduction of microtubule mass during axonal retraction suggests that
both the stable and labile classes of polymer are not undergoing any
significant loss. Nevertheless, because the labile polymer is normally
undergoing more rapid dynamics, it would be more likely to be lost than
the stable polymer. Indeed, previous qualitative observations suggest that there may be a partial loss of tyrosinated microtubules at the
tips of axons retracting in response to other factors (but also see Fan
et al., 1993; Fritsche et al., 1999). Therefore, to be particularly
cautious, we focused one set of analyses specifically on the most
labile polymer in the distal retracting regions of the axons. Because
the distal regions of axons often stain more brightly for total
tubulin, as well as tyrosinated tubulin, it is necessary to obtain a
ratio image that reflects the amount of tyrosinated tubulin relative to
the total tubulin in the microtubules (for details, see Yu et al.,
1994). We examined  25 control axons, all of which consistently showed
an enrichment in the ratio of tyrosinated/total tubulin in the
microtubules occupying the distal region of the axon. This enrichment
was also observed in 25 of 25 axons undergoing retraction in response
to NOC-7, confirming that even the most labile microtubules are not
undergoing any significant loss during retraction. These results are
shown in quantitative pseudocolor in Figure
6.
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Figure 6.
Analyses on the ratio of tyrosinated/total tubulin
in microtubules during axonal retraction. A, Control
axons. B, Axons retracting in response to treatment for
30 min with NOC-7. The first panel of each set shows
staining for tyrosinated tubulin, whereas the second
panel shows staining for total -tubulin. Tyrosinated tubulin
staining is brighter distally, but so too is total tubulin. The
third panel of each set shows a ratio image of
tyrosinated/total tubulin (see Materials and Methods and Results). The
ratio image is displayed in a pseudocolor scale in which
white is the most intense and black is
the least intense. (There is no white in these axons; hence,
yellow is the brightest.) Note that the ratio is clearly
highest distally in the control axons, indicating a concentration of
microtubules particularly rich in tyrosinated tubulin. This same distal
concentration is observed in retracting axons, indicating no loss in
the distal enrichment of these tyrosinated tubulin-rich microtubules.
Scale bar, 20 µm.
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Effects of taxol treatment on axonal retraction induced
by NOC-7
The results described thus far show that there is no detectable
decrease in microtubule levels during axonal retractions induced by
NOC-7, but that the microtubules change their configuration from a
straight bundle into arrays of coiled and bended microtubules. The
simplest interpretation of these results is that the microtubules do
not undergo significant depolymerization during retraction but instead
retreat backward. However, on the basis of the results presented thus
far, we cannot completely dismiss the possibility that the microtubules
are depolymerized in a wholesale manner and then repolymerized into
coiled and bended arrays. To investigate this further, we exposed the
axons for 30 min to 100 nM taxol, a drug that stabilizes
microtubules against disassembly and promotes excess assembly. At this
relatively low concentration, the taxol treatment in and of itself did
not alter the morphology of the axons, which continued to grow normally
over the 30 min of treatment (Fig.
7A). Quantitative
immunofluorescence analyses showed that the average microtubule levels
within these axons increased by ~30%. When taxol-treated axons were
exposed to NOC-7 for 30 min (in the continuous presence of taxol), the
axons showed dramatic retraction. The retractions occurred with the
characteristic morphology of axons retracting to NOC-7 in the absence
of taxol, with a classic retraction bulb, trailing remnant, and
sinusoidal bends along the shaft (Fig. 7A) [in some other
culture systems, higher doses of taxol have been reported to inhibit
axonal retractions, but available evidence suggests that this may be
caused by other effects of the drug rather than by the stabilization of
microtubules (George et al., 1988; McNeil et al., 1999)].
Quantification of microtubule levels showed that the retracted axons
have levels of microtubules that are statistically identical to those
treated with taxol only (i.e., ~30% higher than controls).
Fluorescence images are shown in quantitative pseudocolor in Figure
7B, and the data are displayed graphically in Figure
7C. These results demonstrate that axonal retraction does
not require microtubule depolymerization to occur, and that the same
kind of microtubule reconfiguration occurs during retraction when
microtubules are prevented from depolymerizing. Moreover, they show
that these changes can occur even when the polymer levels are actually
increasing above normal.
View larger version (23K):
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|
Figure 7.
Axonal retraction induced by NOC-7 in the presence
of taxol. A, Phase-contrast images of an axon before any
treatment, after 30 min in taxol alone, and after 30 min of treatment
with NOC-7 in the continued presence of taxol. Note that the taxol
treatment did not prohibit axonal growth when applied without the
NOC-7, nor did it prohibit axonal retraction after the NOC-7 was added.
B, Microtubule immunofluorescence studies of three
different axons: (1) a control axon, (2) an axon treated for 30 min
with taxol alone, and (3) an axon treated with taxol for 30 min and
then treated with both taxol and NOC-7 for an additional 30 min. Axons
are shown in a quantitative pseudocolor scale in which
white is the most intense and black is
the least intense. Note that the taxol-treated axon is brighter than
the control axon. The taxol/NOC-7-treated axon is even brighter yet
(per micrometer). C, Quantitative studies on
fluorescence intensity, in arbitrary fluorescence units. Taxol
treatment alone increased microtubule levels by ~30% above control
levels. Axons retracting in response to NOC-7 (in the continued
presence of taxol) displayed a much higher yet fluorescence intensity
per micrometer when calculated for length after retraction
(LR). However, when calculated for original
length (LO), the intensity per micrometer was
indistinguishable from that of the axons treated with taxol alone.
Thus, taxol treatment elevated total microtubule levels above controls,
but this stimulation of microtubule assembly did not prohibit axonal
retraction in response to NOC-7. The two-tailed Student's
t test (*p < 0.05) shows a
significant difference between control axons and axons treated with
taxol and a significant difference between control axons and axons
treated with taxol followed by NOC-7. There was also a significant
difference between axons exposed only to taxol and axons exposed to
taxol followed by NOC-7, if calculations were performed using
LR. However, there was no significant difference between
axons exposed only to taxol and axons exposed to taxol followed by
NOC-7 if calculations were performed using LO. Scale bars,
20 µm.
|
|
| |
DISCUSSION |
There has been great interest over the past few decades in how the
cytoskeletal arrays of the axon are elaborated during its growth. This
topic has been of particular interest both because of its importance
and because of a great deal of heated controversy (for review and
discussion, see Baas, 2002). Early models maintained that the
cytoskeletal elements are transported down the axon in the form of
assembled polymers and that this transport is fueled by molecules that
we would today refer to as molecular motor proteins. This view was
challenged in the mid-1980s by workers who felt that it was more
reasonable that the polymers were stationary and that their subunit
components were somehow transported down the axon. In this alternate
model, the elaboration of the cytoskeletal arrays resulted from the
polymerization of new cytoskeletal elements along the length of the
axon during its growth. In recent years, the transport of cytoskeletal
polymers has been directly observed (Dent et al., 1999; Roy et al.,
2000; Wang et al., 2000) but so too has local polymer assembly (Dent et
al., 1999; Kabir et al., 2001). It appears that the most satisfactory
model includes both polymer transport and local assembly dynamics as
key contributors to the elaboration of the axonal cytoskeletal arrays.
In recent years, neuroscientists have begun to focus on the retraction
of axons as an event that is equally noteworthy during the development of the nervous system. However, to date, most consideration of the
cytoskeletal mechanisms involved in axonal retraction has been limited
to the general concept that the polymers most probably depolymerize to
accommodate the shortening of the axon during retraction.
As outlined in the introductory remarks, there are reasons to question
this conclusion. Although it is true that pharmacologically depleting
an axon of its microtubules can cause it to retract, the morphological
features of the retraction are not at all similar to those of axons
retracting in the developing nervous system or to axons retracting in
the culture dish in response to physiological cues. Although some
physiological cues appear to cause local actin depolymerization within
growth cones (Fan et al., 1993) (for information regarding what
underlies their collapse, see Fritsche et al., 1999), it is unlikely
that there would be wholesale actin depolymerization within the axonal
shaft. It has been shown repeatedly that axonal retraction is an
actomyosin-based process and that depleting microfilaments from axons
actually prohibits their retraction rather than causing it (Solomon and
Magendantz, 1981; Joshi et al., 1985; Ahmad et al., 2000). In addition,
recent studies have even called into question whether microfilaments
are always depolymerized in growth cones during their collapse; these
studies suggest that the microfilaments can also be debundled and
reconfigured, with no significant net diminution in polymer levels
(Fournier et al., 2000; Zhou and Cohan, 2001). Our qualitative
observations suggest that microfilaments are folded inward during
growth-cone collapse in response to nitric oxide, whereas other studies
using nitric oxide indicate that there is some local depolymerization
of actin as well (Ernst et al., 2000). It is our impression, however,
that phalloidin staining favors actin bundles over debundled filaments,
and, hence, we feel that additional studies are merited to fully
resolve this issue. Our focus was on microtubules.
Our analyses on microtubules resulted in four major findings. First, as
in previous studies using physiological inducers of axonal retraction,
the morphology of the retracting axons was characterized by an enlarged
distal region, a thin trailing remnant, and sinusoidal bends along the
axonal shaft. There were no signs of the kind of beading and withering
observed in axons depleted of microtubules with nocodazole. Second,
fluorescence imaging of microtubules revealed that in the retracting
axons, microtubules are still abundant but reconfigured into coiling
and sinusoidal arrays to accommodate the shortening of the axon.
Quantification of microtubule levels showed no detectable diminution in
microtubule levels during retraction. Third, there was no reduction in
the retracting axons of the distal concentration of the more labile microtubule polymer enriched in tyrosinated tubulin, indicating that
even the polymer turning over most rapidly is not lost during retraction. Finally, pretreatment of the axons with 100 nM
taxol, a drug that stabilizes microtubules against disassembly and
actually caused a 30% increase in microtubule levels in our studies,
did not abolish axonal retraction induced by the nitric oxide donor. Thus, not only is there no loss in microtubule levels during
retraction, but we can also conclude that the microtubules do not
undergo any wholesale depolymerization followed by repolymerization to produce the newly configured coiled and bent microtubule arrays, because the same response is obtained under conditions in which polymer
levels are actually rising.
Together, all of these observations demonstrate that there is no
wholesale depolymerization of microtubules during the axonal retractions that we have observed, but instead that the pre-existing microtubules retreat backward. We cannot dismiss the possibility that
there might be some depolymerization of microtubules during retraction,
especially in the latter stages of axonal retraction, as the
microtubules are ultimately chased back into the cell body. Another
factor that should be considered is the ongoing addition of
microtubules to the axon by anterograde transport from the cell body;
this transport may or may not persist during axonal retraction but
could potentially offset a small amount of polymer that does in fact
depolymerize during retraction. Despite these caveats, our data clearly
show that the early stages of axonal retraction cannot be explained in
terms of polymer loss but instead appear to be the result of forces
that impinge on the microtubule array to reconfigure it. Consistent
with this conclusion, we reported previously that axons retract with
similar characteristics when at least one microtubule-associated motor
protein is experimentally manipulated (Ahmad et al., 2000).
Specifically, we showed that inhibition of cytoplasmic dynein causes
axons to retract with no observable loss of microtubules but with
bending and coiling of the axon and the microtubules that were similar
(but not identical) to the bending and coiling observed in the present
study in response to nitric oxide. On this basis, and based on the fact
that retraction itself is dependent on myosin-driven forces, we have
proposed a new model for axonal retraction based on reconfiguration of cytoskeletal polymers by motor-driven forces (Baas and Ahmad, 2001).
In our previous study, we suggested that dynein-driven forces between
the microtubule and microfilament arrays may attenuate the myosin-based
contractility of the axon. If this is correct, physiological signals
that induce retraction might function by uncoupling cytoplasmic dynein
from either the microtubules or the actin. Alternatively, the signals
may simply increase the contractility of the myosin so that it
overwhelms the dynein forces (e.g., by enhancing phosphorylation of
myosin regulatory light chain) (Billuart et al., 2001). Another
possibility is that there are additional motors, specialized kinesins,
that contribute directly to the reconfiguration of the microtubules.
Some support for this possibility comes from our observations that the
microtubules begin to show coiling and bending even before there is any
detectable actomyosin-based shortening of the axon. Additional support
comes from the fact that the retractions observed with nitric oxide were not completely identical to those observed with dynein inhibition; the coiling of microtubules was more dramatic with the nitric oxide,
suggesting that additional motor forces may be involved.
Of course, it is possible that different physiological signals elicit
their effects by different pathways; some may affect myosins, and
others may affect cytoplasmic dynein or particular kinesins. We also
cannot eliminate the possibility that some inducers of retraction might
cause large-scale depolymerization of microtubules; our studies have
focused exclusively on nitric oxide, and there is certainly a wide
variety of physiological factors that can induce axons to retract. To
the best of our knowledge, however, there are no observations of axons
retracting physiologically that display characteristics of widespread
or large-scale microtubule depolymerization, and the results of
previous analyses on other types of retracting axons, although not
quantitative, are entirely consistent with the conclusions of our study
(Miller et al., 1994; McNeil et al., 1999). We believe that the next
important step in understanding cytoskeletal reconfiguration during
axonal retraction is to elucidate how the biochemical pathways affected
by physiological cues alter the forces generated by molecular motor proteins.
| |
FOOTNOTES |
Received Jan. 23, 2002; revised March 7, 2002; accepted April 16, 2002.
This work was supported by two grants to P.W.B. from the National
Institutes of Health. We thank Gianluca Gallo and Paul Letourneau of
the University of Minnesota for helpful advice. We also thank Dan
Buster and Doug Baird of our institution for assistance, advice, and discussions.
Correspondence should be addressed to Peter W. Baas, Department of
Neurobiology and Anatomy, Drexel University College of Medicine, 2900 Queen Lane, Philadelphia, PA 19129. E-mail peter.w.baas{at}drexel.edu.
| |
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Visualization of Microtubule Growth in Cultured Neurons via the Use of EB3-GFP (End-Binding Protein 3-Green Fluorescent Protein)
J. Neurosci.,
April 1, 2003;
23(7):
2655 - 2664.
[Abstract]
[Full Text]
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C. Badorff and S. Dimmeler
NO Balance: Regulation of the Cytoskeleton in Congestive Heart Failure by Nitric Oxide
Circulation,
March 18, 2003;
107(10):
1348 - 1349.
[Full Text]
[PDF]
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TOP
ABSTRACT
INTRODUCTION
