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Vol. 14, Issue 5, 2192-2200, May 2003
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* Department of Molecular Genetics, The Ohio State University, Columbus, Ohio
43210;
Department of Biology, University of North Carolina-Chapel Hill, Chapel Hill,
North Carolina 27599; and
Department of Biochemistry and Molecular Biology, Uniformed Services
University of the Health Sciences, Bethesda, Maryland 20814
Submitted October 10, 2002;
Accepted January 16, 2003
Monitoring Editor: Richard McIntosh
 |
ABSTRACT |
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 TOP ABSTRACT INTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONACKNOWLEDGMENTSREFERENCES |
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-tubulin
fusion in living cells by time-lapse spinning-disk confocal microscopy. We
find that tubulin is excluded from interphase nuclei, enters the nucleus
seconds before the mitotic spindle begins to form, and is removed from the
nucleoplasm during the M-to-G1 transition. Our data indicate that
regulation of intranuclear tubulin levels plays an important, perhaps
essential, role in the control of mitotic spindle formation in A.
nidulans. They suggest that regulation of protein movement into the
nucleoplasm may be important for regulating mitotic onset in organisms with
intranuclear mitosis. |
INTRODUCTION |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONACKNOWLEDGMENTSREFERENCES |
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). Microtubules assemble and are organized into a functional
spindle within the nuclear envelope. Organisms with intranuclear mitosis make
up much of the earth's biomass and include species that are enormously
beneficial to humans and others that are serious human or agricultural
pathogens. It is consequently important to understand mitotic spindle
formation in these organisms.
In fungi such as Aspergillus nidulans, mitotic spindle formation
is regulated very precisely (Oakley and
Morris, 1983; Jung et
al., 1998, and earlier references therein). No microtubules
are present in the nucleus in interphase, and the spindle assembles rapidly in
the nucleus at the onset of mitosis. The nuclear envelope does not break down
in mitosis (Robinow and Caten,
1969).
Experiments to date suggest two quite different but nonmutually exclusive
models (Figure 1) for how
spindle formation is initiated in organisms with closed spindles. One model
(Figure 1, a–c) is
suggested by work in Schizosaccharomyces pombe by Masuda et
al. (1992) and Masuda and
Shibata (1996). In this model,
spindle assembly is controlled by a cell-cycle-regulated change in the ability
of the fungal microtubule-organizing center, the spindle pole body (SPB), to
nucleate microtubule assembly. Tubulin dimers would pass freely through the
nuclear envelope, but there would be no microtubule assembly in interphase
because there are no active microtubule nucleation sites within the nucleus.
At the onset of mitosis the nucleation sites at the SPB are activated and
spindle formation occurs. In S. pombe, this may be facilitated by the
penetration of the SPB into a fenestra in the nuclear envelope at mitosis
(Ding et al.,
1997).
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Another model is shown in Figure 1,
d–f. In this model, the SPB is capable of nucleating
microtubule assembly throughout the cell cycle. Spindle formation is regulated
by the concentration of tubulin in the nucleus, which is, in turn, regulated
at the level of tubulin movement across the nuclear envelope. Tubulin is
excluded from the nucleus at interphase, allowed to enter at the onset of
mitosis and removed as the daughter nuclei transit from M to G1. To
date, there is no direct evidence for this model, but Wu et al.
(1998) found that in A.
nidulans the activity of the NIMA kinase is required for the
translocation of the cdc2/cyclin B complex into the nucleus at the onset of
mitosis and, thus for initiation of mitotic entry. These data suggest that
mitotic onset could be triggered by transport of key proteins into the
nucleus.
As mentioned, these models are not mutually exclusive. Both mechanisms might operate redundantly. It is also important to note that each of these mechanisms would be affected by cytoplasmic microtubule dynamics. In A. nidulans, for example, the rapid disassembly of cytoplasmic microtubules, which occurs as cells approach mitosis, should greatly increase the size of the tubulin pool available for incorporation into the assembling spindle.
To evaluate the model shown in Figure 1, d–f, we have attempted to determine whether tubulin is excluded from interphase nuclei and enters at the onset of mitosis. This attempt was complicated by the facts that 1) in interphase the cytoplasmic microtubule array is extensive and it is normally difficult to observe free tubulin in the cytoplasm, and 2) the G2-to-M transition is very rapid in A. nidulans and it is difficult to get an accurate picture of the process from static immunofluorescence images.
We have circumvented these difficulties in two ways. First, we have used
benomyl to disassemble cytoplasmic microtubules at interphase. This creates a
large pool of free tubulin that can be readily observed by immunofluorescence.
We find that tubulin is substantially excluded from interphase nuclei, but is
present in mitotic nuclei. Second, we have observed a green fluorescent
protein (GFP) -tubulin fusion by spinning-disk confocal microscopy.
Tubulin levels are low in interphase nuclei and there is a rapid movement of
tubulin into the nucleoplasm seconds before spindle formation begins. Tubulin
is removed from the nucleoplasm at the end of mitosis. These data provide a
strong indication that regulation of intranuclear tubulin levels plays an
important, perhaps essential, role in the regulation of mitotic spindle
formation.
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MATERIALS AND METHODS |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONACKNOWLEDGMENTSREFERENCES |
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).
Three strains were used for GFP-tubulin observations: GFP-tub7, LO716, and
L0770. GFP-tub7 carries pyroA4, wA2, pyrG89, and
the green fluorescent protein fused to the tubA (-tubulin)
gene (GFP-tubA) (Han et
al., 2001). GFP-tubA is under the control of the
regulatable alcA promoter. LO716 is a diploid formed by fusion of
GFP-tub7 with strain R21 (yA2, pabaA1). It, thus, carries an
inducible GFP-tubA fusion as well as a wild-type tubA. LO770
was created by crossing GFP-tub7 with a strain that carries nimT23.
Its genotype is wA2, nimT23, pyrG89 and it carries
the GFP-tubA insertion at the tubA gene.
For observations of GFP fluorescence, strains were grown in two ways. In
some experiments they were grown in four-well Lab-Tek-chambered cover glasses
with covers (Nalge Nunc, Naperville, IL). Each well contained 6 x
104 conidia (spores) inoculated into 750 µl of induction medium.
Induction medium consisted of minimal medium
(Pontecorvo et al.,
1953) with 50 mM fructose substituted for d-glucose and
with appropriate supplements for nutritional markers. Threonine (6.25 mM) was
used to induce GFP-tubA expression in most experiments, but 1.56,
3.13, or 12.5 mM was used in some experiments. Growth was robust under these
conditions and nuclei were observed to complete mitosis normally. In most
experiments, including all experiments in which GFP-tubA levels were
quantified, strains were grown as follows. Conidia were inoculated into 1%
low-melting temperature agarose containing minimal medium with 50 mM fructose
and 1 mM threonine at 42°C to a concentration of 1 x
107/ml. (For reasons that are not clear, the optimum threonine
concentration was lower under these conditions than in liquid cultures.) Two
pieces of Scotch Magic tape were placed 
35 mm apart on a slide, and 5
µl of the molten spore suspension was placed on the slide between the
pieces of tape. A 24- x 40-mm coverslip was quickly placed on the spore
suspension and pressed down gently such that it rested on the tape. The
agarose spread into a thin layer that did not contact the tape. This
construction was placed in a Petri dish with moist paper and chilled at
4°C for 15 min to allow the agarose to solidify. It was then incubated at
30°C overnight (in the same moist chamber) before observation. Conidia
near the edge of the agarose germinated and grew robustly.
Microscopy
Immunofluorescence micrographs were taken on a standard microscope (Carl
Zeiss, Thornwood, NY) with a 100x Neofluor objective (1.30 numerical
aperture) by using T-MAX 400 film (Eastman Kodak, Rochester, NY). The film was
developed in T-MAX developer and negatives were scanned into Adobe Photoshop
with a Polaroid SprintScan 35 Plus scanner. Time-lapse GFP images were taken
with a 100x Plan Apochromatic objective (1.40 numerical aperture) on an
Eclipse TE300 inverted microscope (Nikon, Tokyo, Japan) equipped with an
Ultraview spinning-disk confocal system (PerkinElmer Life Sciences, Boston,
MA) controlled by Ultraview software. Time-lapse series were saved as 16-bit
tiff files. For quantitation of GFP-tubA fluorescence, the tiff files
were imported into MetaMorph (Image 1). Composite figures were prepared using
CorelDraw 8 (Macintosh).
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RESULTS |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONACKNOWLEDGMENTSREFERENCES |
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) have shown that benomyl
at 2.4 µg/ml causes a rapid disassembly of microtubules in A.
nidulans. We have observed that the tubulin from benomyl-disassembled
microtubules distributes through the cytoplasm and can be seen easily by
immunofluorescence microscopy with the anti 
-tubulin antibody Tu27B.
Observations of material stained with 4,6-diamidino-2-phenylindole (DAPI) and
Tu27B revealed that in interphase germlings there were voids in the
-tubulin staining that precisely coincided with the DAPI-stained nuclei
(Figure 2, a and b). In A.
nidulans, chromosome condensation occurs during mitosis and can be
observed by DAPI staining (Jung et
al. 1998, and earlier references therein). We observed that
in benomyl-treated germlings with condensed chromosomes the tubulin was not
excluded from the nucleoplasm (Figure 2, c
and d). Rather, the tubulin staining through and around the
chromosomes was indistinguishable from that of the remainder of the germlings.
These results suggest that tubulin is substantially excluded from interphase
nuclei, but not from mitotic nuclei.
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To determine whether these initial observations were statistically
significant, we treated germlings for 30 min with benomyl and then fixed and
stained them with Tu27B and DAPI. We then scored germlings for chromosome
condensation and for whether there was a tubulin-deficient void corresponding
to the nucleoplasm. We scored a total of 1252 germlings in three separate
experiments (Table 1). At the
incubation temperature (37°C), the cell cycle for the strain used is
<90 min, and 30 min of treatment with benomyl should result in 38% of
germlings being blocked in mitosis (5% already in mitosis when the benomyl was
added plus 33% that enter mitosis in one-third of the cell cycle). We found
that, as expected, 38% of the germlings had nuclei with condensed
chromatin. As Table 1 shows,
there was a remarkable correlation between chromosomal condensation and the
presence of tubulin in the nucleoplasm. In all 472 of the germlings with
nuclei with condensed chromosomes, tubulin was present in the nucleoplasm at
levels at least as high as that in the surrounding cytoplasm. In all 756 of
the germlings with interphase nuclei (noncondensed chromatin), tubulin
fluorescence in the nucleoplasm was visibly much lower than that in the
surrounding cytoplasm. In 24 of the germlings, chromatin was partially
condensed and there was an intermediate level of tubulin in the nucleoplasm.
Because the benomyl block was in place for much less than a single cell cycle,
we can infer with confidence that the nuclei with partially condensed
chromatin were entering mitosis rather than exiting. Because intermediate
tubulin levels correlate with partially condensed chromatin, we can also
deduce that tubulin movement into the nucleoplasm occurs at the same time that
condensation occurs, in prophase.
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To confirm and expand these experiments we observed a GFP -tubulin
fusion protein in living hyphae. For brevity, we will refer to tubulin dimers
containing the GFP -tubulin fusion protein as GFP-tubulin. The
brightness of the GFP-tubulin fluorescence varied to some extent among hyphae,
but microtubule arrays were clear. There was also a general GFP-tubulin
fluorescence in the cytoplasm. Because tubulin dimers are known to exist in
equilibrium with microtubule polymer, we are confident that the general
GFP-tubulin fluorescence is due to tubulin dimers in the cytoplasm. In these
hyphae, nuclei could be distinguished as volumes with reduced GFP-tubulin
fluorescence. These volumes could be reliably identified as nuclei because of
their sizes, shape, number, and location and the fact that, in many cases,
cytoplasmic microtubules extended from a point on the nuclear surface (the
SPB) (our unpublished data).
We observed nuclei entering mitosis in two ways. For most experiments, we took advantage of the facts that there are multiple nuclei in each cell (or, more properly, hyphal segment) in A. nidulans and that nuclei in a hyphal segment enter mitosis at almost, but not quite, the same time. The nucleus at one end of the hyphal segment generally goes into mitosis first, the one next to it goes in second, and the other nuclei enter sequentially until all nuclei are eventually in mitosis. We searched for hyphae in which nuclei at one end of a hyphal segment were in the early stages of mitosis and in which nuclei at the other end were still in G2. We then took time-lapse images of the G2 nuclei entering mitosis (Figure 3) by using a spinning-disk confocal microscope. The spinning-disk confocal microscope was particularly useful for these studies because it removed most of the signal from GFP-tubulin above and below the nucleus, and it allowed us to capture images rapidly and with little bleaching.
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In a second approach, we used a temperature-sensitive mutation to
synchronize cells. We prepared a strain (LO770) carrying nimT23 and
the GFP -tubulin fusion. nimT23 is a temperature-sensitive
mutation in the phosphatase that regulates p34cdc2 activity. It
blocks the cell cycle in late G2 at 42°C and is rapidly
reversible, allowing entry into mitosis shortly after a shift to a permissive
temperature (O'Connell et al.,
1992; Martin et al.,
1997). To observe entry into mitosis, we grew germlings overnight
at 32°C and then blocked cells in G2 by incubating at 43°C
for 3 h. We released the block by placing the chambers containing the
germlings on the microscope stage at room temperature (24°C). We then
took time-lapse images of cells entering and carrying out mitosis. The two
approaches gave similar results, but the first approach allowed us to acquire
data more rapidly.
As mentioned, interphase nuclei were visible as GFP-tubulin–deficient volumes (Figure 3a, nuclei 3 and 4, for example). They became particularly obvious as cytoplasmic microtubules began to disassemble before mitosis increasing the pool of cytoplasmic tubulin (Figure 3a, nuclei 3 and 4). As nuclei approached mitosis, there was a sudden movement of tubulin into the nucleoplasm (Figure 3, and accompanying movie Ovetne.mov) followed shortly by the initiation of mitotic spindle formation (Figure 3, f and k, arrows).
To estimate the timing and extent of tubulin movement into the nucleus, we quantified GFP-tubulin fluorescence in 10 nuclei in eight hyphae and measured the changes in these values over time. We initially considered measuring GFP-tubulin intensity in the nucleus relative to an area identical in size and shape in the cytoplasm. We noted, however, that GFP-tubulin intensity was not uniform through the cytoplasm (Figures 3 and 4). We presume that the lack of uniformity is due, at least in part, to organelles such as mitochondria and vesicles that exclude tubulin. Because the cytoplasmic GFP-tubulin intensity depended on the region of the cytoplasm chosen, this approach did not provide an objective measure of cytoplasmic GFP-tubulin levels. We also noted that variation in GFP-tubulin intensity among hyphae and differences in exposure conditions among experiments affected the absolute GFP fluorescence values. To circumvent these potential problems, we determined the GFP-tubulin fluorescence of the nuclei relative to the background outside the hypha and to the average GFP-tubulin fluorescence of the entire hypha under consideration. We first determined the average GFP-tubulin signal intensity for the hypha over a period starting before the tubulin began to move into the nucleus and ending with spindle formation. This average was assigned the value 100. (In instances in which two nuclei entered mitosis, the end point was the formation of the second spindle.) We next determined the average signal intensity over the same period for a background region identical in size and shape to the hypha. The signal intensity of the background was remarkably uniform so the positioning of the background region did not alter the average intensity significantly. The average background intensity was assigned the value 0. A region was then selected within each nucleus and the average intensity of the region was measured at each time point. This approach provides an objective measurement of the GFP-tubulin levels in the nucleus relative to the hyphal GFP-tubulin levels. One point that needs to be noted, however, is that because tubulin is almost certainly excluded from mitochondria, vesicles, and nuclei, the cytosolic GFP-tubulin is confined to a volume somewhat smaller than the entire hypha and the cytosolic GFP-tubulin levels are higher than the hyphal average. If, for example, the organelles that exclude GFP-tubulin make up 20% of the hyphal volume, all the GFP-tubulin will be in the remaining 80% of the hyphal volume. The true cytosolic GFP-tubulin levels would thus be 1.00/0.80 = 1.25 or 125% of the hyphal average. The approach we have used, thus, provides an accurate way of measuring the relative concentrations of intranuclear GFP-tubulin in interphase and mitotic nuclei and the changes in these concentrations as nuclei enter and exit mitosis. This approach does not, however, provide an accurate estimate of nuclear GFP-tubulin relative to the surrounding cytosol. Rather, the values we obtain will tend to overstate nuclear GFP-tubulin concentrations (mitotic and interphase) to some extent. Thus, if the GFP-tubulin level in an interphase nucleus was 50% of the hyphal average, it might actually be 30% of the level in the surrounding cytosol.
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The results for the nuclei shown in
Figure 3 are quantified in
Figure 4. In this example, the
GFP-tubulin fluorescence level in the interphase nuclei was <50% of the
level of the average hyphal intensity. As each nucleus entered mitosis, there
was a rapid increase in the GFP-tubulin fluorescence. In less than one minute
the nuclear GFP-tubulin fluorescence exceeded that of the hyphal average and
the mitotic spindle became visible in the next image which was taken 15 s
later.
In the 10 nuclei in which GFP-tubulin fluorescence was quantified, the interphase nuclear GFP-tubulin fluorescence averaged 42.8 ± 8.0% (mean ± SD) of the hyphal average. Because the spinning-disk confocal microscope removes most, but not all, out of focus fluorescence, this value probably represents an overestimate of the amount of GFP-tubulin in the nucleus. The maximum GFP-tubulin fluorescence in the nuclei reached 126 ± 16.1% of the average hyphal intensity before spindles became visible. The intranuclear tubulin level thus increases by a factor of three at mitotic onset. The time from interphase until the nuclei filled with GFP-tubulin was 1.68 ± 0.99 min. The time from interphase until spindle formation was observed was 2.25 ± 1.25 min. The time from the nuclei being filled until visible spindle formation was 0.58 ± 0.43 min (34.5 ± 25.5 s). Because some spindles probably began to form out of the plane of focus, and were not immediately visible, this value is probably an overestimate.
At the end of mitosis, it was apparent that GFP-tubulin was removed from nuclei (Figure 5). G1 nuclei are smaller than G2 nuclei, and move rapidly around the hyphae and in and out of the plane of focus so it was not possible to track telophase/G1 nuclei well enough to determine a precise time course for GFP-tubulin removal. Nuclei were clearly visible, however, as tubulin-deficient volumes by 4 min after mitotic spindle breakdown.
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DISCUSSION |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONACKNOWLEDGMENTSREFERENCES |
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Two general models for the mechanism of movement of tubulin into and out of
the nucleoplasm have occurred to us. One is that there is an active transport
mechanism that moves tubulin into or out of the nucleus. For example, tubulin
could be moved into the nucleus by an active transport mechanism that would be
inactive in interphase and would be activated at the G2-to-M
transition. A search (Columbia University Bioinformatics Center,
http://cubic.bioc.columbia.edu/)
revealed no nuclear localization sequences in any of the A. nidulans
- and -tubulin sequences. Any active transport mechanism would,
thus, require that tubulin form a complex with one or more proteins that have
nuclear localization signals. This has been shown to occur for 
-tubulin
(Pereira et al.,
1998). A variant of this model is that tubulin is actively removed
from interphase nuclei by a nuclear export mechanism that is inactivated at
mitotic onset and reactivated at the G2-to-M transition.
A second model is that the nuclear envelope becomes more permeable at the G2-to-M transition. Tubulin and other proteins would, thus, be free to diffuse into the nucleoplasm during mitosis. Although electron microscopy demonstrates that the nuclear envelope does not break down in mitosis in A. nidulans, it is certainly possible that its permeability characteristics could be altered at the onset of mitosis.
The notion that the nuclear envelope becomes transiently permeable in mitosis is attractive because it provides an economical mitotic regulatory mechanism. Tubulin, the cdc2/cyclin B complex (see below) and other proteins required for mitosis might be excluded from interphase nuclei. At mitotic onset a single event, the alteration of nuclear envelope permeability, would allow these proteins to flood into the nucleoplasm while mitotic inhibitor proteins might diffuse out. At the end of mitosis the nuclear envelope would again become impermeable and proteins involved in mitosis would be removed. This model suggests that mitosis in fungi is more similar to that of higher organisms than previously thought and that the presence of a morphologically intact nuclear envelope in mitosis may be less significant than previously thought.
At present, there is insufficient data to determine which of these models is correct. Our GFP-tubulin quantitation data might seem to suggest that the movement into the nucleus is active. In all 10 nuclei examined, the GFP-tubulin in the nucleus reached a level higher than that of the remainder of the hypha before spindle formation (mean value 126 ± 16.1% of the average hyphal value). If tubulin were diffusing passively into the nucleoplasm, one would not expect the GFP-tubulin concentration in the nucleoplasm to exceed that of the cytoplasm. As mentioned, however, the cytosolic GFP-tubulin concentration must be higher than the hyphal average because GFP-tubulin is excluded from many organelles and the hyphal average would include the cytosolic GFP-tubulin as well as the regions from which it is excluded. It is, thus, quite possible that the GFP-tubulin level in the nucleus is simply reaching the cytosolic level, not exceeding it, and that the movement is passive.
Information on whether the nuclear envelope becomes permeable at mitotic
onset is limited at present. An obvious experiment would be to determine
whether GFP or other fluorescent proteins (not coupled to tubulin or any other
protein) enter the nucleus at mitotic onset. GFP is not excluded from the
nucleoplasm of interphase nuclei of A. nidulans
(Fernandez-Abalos et al.,
1998), however, and neither is DS red fluorescent protein (our
unpublished data). In the fungi Fusarium ventricilloides and
Magneporthe grisea, however, another fluorescent protein, ZsGreen, is
excluded from interphase nuclei and enters at mitosis
(Bourett et al.,
2002). In A. nidulans, a GFP fusion to a portion of the
putative transcription factor stuA that carries a nuclear
localization signal localizes to the nucleoplasm in interphase, and at mitosis
it leaves the nucleoplasm (Suelmann et
al., 1997). Finally, interphase nuclei of living A.
nidulans hyphae are visible by phase contrast microscopy, but they become
nearly invisible in mitosis (Robinow and
Caten, 1969). This indicates that the refractive index of the
nucleoplasm changes upon mitotic entry to match that of the surrounding
cytoplasm. Although limited, these data are all consistent with the
possibility that the nuclear envelope becomes permeable in mitosis.
Regardless of the mechanism of tubulin movement into the nucleus, it seems that there must be a rapid, active mechanism to remove tubulin from the nucleoplasm at the end of mitosis. It is difficult to see how it could be removed by passive diffusion. The removal mechanism could involve proteolytic degradation of tubulin or export back to the cytoplasm.
Because movement of tubulin into and out of the nucleoplasm is timed so precisely with respect to the cell cycle, we can infer that it is controlled by the cell cycle regulatory machinery. At present, we know, from our observations with GFP-tubulin, that it occurs downstream of cdc2 activation. nimT is required for cdc2 activation and at the nimT23 block point cdc2 is inactive and tubulin is excluded from the nucleoplasm. When the block is released by a shift to permissive temperature, tubulin moves into the nucleoplasm and the spindle assembles shortly afterward.
Could the changes in intranuclear tubulin levels be artifactual? Our
evidence suggests strongly that this is not the case. First, two quite
different procedures, immunofluorescence with an antibody against
-tubulin and time-lapse imaging of GFP-tubulin, provide mutually
supportive evidence that there is a rapid influx of tubulin into the nucleus
at the onset of mitosis and removal at the M-to-G1 transition.
Second, the apparent exclusion of tubulin from interphase nuclei is not simply
due to the fact that a large fraction of the tubulin dimers is tied up in
cytoplasmic microtubules because even when cytoplasmic microtubules are
disassembled completely by benomyl, tubulin levels in interphase nuclei are
low. Third, the exclusion is not simply a displacement of tubulin by
chromatin. Our observations with immunofluorescence reveal that tubulin is
uniformly distributed through mitotic nuclei and is not displaced by mitotic
chromatin, which is more condensed than interphase chromatin. In addition,
when GFP-tubulin moves into nuclei, there is a brief period before spindle
formation when it is distributed uniformly through the nucleoplasm and is
clearly not displaced by chromatin. Finally, GFP alone is not excluded from
interphase nuclei (Fernandez-Abalos et
al., 1998), so tubulin is responsible for the behavior of the
GFP-tubulin, not the GFP moiety.
It is important to note that three proteins or protein complexes important
to mitosis, the tubulin dimer, the cdc2/cyclin B complex
(Wu et al., 1998),
and the NIMA kinase (De Souza et
al., 2000; De Souza and Osmani, unpublished data) have now
been shown to enter the nucleoplasm at the G2- to-M transition. It
is tempting to speculate that this may be true for other proteins required for
mitosis and that regulation of protein movement across the nuclear envelope
may be a general regulatory mechanism for the G2-to-M and
M-to-G1 transitions in A. nidulans and other organisms
with intranuclear mitosis.
Could similar mechanisms operate in other organisms? In many fungi and
protists, mitosis is intranuclear and microtubules are only present in the
nucleus for the short period required for mitosis (mitosis in protists
reviewed by Heath, 1980). In
these organisms it is quite possible that regulation of the movement of
tubulin and other proteins into and out of the nucleus is an important switch
for regulating the progression into and out of mitosis. In other fungi,
including Saccharomyces cerevisiae, some microtubules are present in
the nucleus for most of the cell cycle
(Byers and Goetsch, 1975;
Heath, 1994). There is, thus,
no sudden switching on of tubulin movement into the nucleus. In these fungi,
however, there is an increase in microtubule number and length as nuclei pass
through the cell cycle, and this could be regulated by a more gradual change
of tubulin levels in the nucleus.
Our findings may establish an evolutionary context for recent findings on
the roles of importins and the GTPase Ran in spindle formation. Importins are
involved in transport of proteins into the nucleus
(Mattaj and Englmeier, 1998;
Weis, 1998;
Gorlich and Kutay, 1999). They
form complexes with cargo proteins in the cytoplasm and transport them through
nuclear pores. Once in the nucleus the cargo proteins are released by
interaction of importins with RanGTP. It has been known for some time that in
many animal cells chromatin can stabilize microtubules
(Zhang and Nicklas, 1995, and
earlier references therein). More recently it has been shown that RanGTP
promotes spindle formation (Nakamura
et al., 1998; Ohba
et al., 1999; Wilde
and Zheng, 1999). The likely mechanism is that proteins that are
required for spindle formation, including those that stabilize microtubules,
are bound to importins, such that they are inactive. RanGTP causes their
release and this activates them allowing them to promote spindle formation
(Gruss et al., 2001;
Nachury et al., 2001;
Wiese et al., 2001).
The RanGTP concentration is highest near chromatin because its guanine
nucleotide exchange factor, RCC1, is associated with chromatin
(Ohtsubo et al.,
1989). An obvious question raised by these findings is why the
nuclear transport machinery is used in the regulation of spindle formation.
Because mitosis is intranuclear in many protists, it is likely that it was
intranuclear in ancestral eukaryotes. One possibility raised by our finding is
that tubulin is actively transported into the nucleus in A. nidulans
by nuclear transport proteins. If nuclear transport is important to the
regulation of spindle formation in organisms with intranuclear spindles, it
follows that when nuclear envelope breakdown evolved, nuclear transport
proteins retained an important role in the regulation of spindle
formation.
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ACKNOWLEDGMENTS |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONACKNOWLEDGMENTSREFERENCES |
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Footnotes |
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Abbreviations used: DAPI, 4,6-diamidino-2-phenylindole; GFP, green fluorescent protein; SPB, spindle pole body.
Online version of this article contains video material for some figures.
Online version available at
www.molbiolcell.org. 
Present address: Department of Physiology and Biophysics, University of
Washington, 1959 NW Pacific St., Seattle, Washington, 98195.
¶ Corresponding author. E-mail address: oakley.2{at}osu.edu.
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REFERENCES |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONACKNOWLEDGMENTSREFERENCES |
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De Souza, C.P.C., Osmani, A.H., Wu, L.-P., Spotts, J.L., and Osmani, S.A. (2000). Mitotic histone H3 phosphorylation by the NIMA kinase in Aspergillus nidulans. Cell 102, 293–302.[CrossRef][Medline]
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