Department of Biology, York University, 4700 Keele Street, Toronto, Ontario, Canada M3J 1P3
Correspondence
Karina Sampson
ksampson{at}bio.umass.edu
![]() |
ABSTRACT |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
Micrographs showing no evidence of MT fluxing/treadmilling and graphs of cytoplasmic fluorescence intensity gradients are available as supplementary material with the online version of this paper.
Present address: Department of Biology, University of Massachusetts, Amherst, MA 01003, USA.
![]() |
INTRODUCTION |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
In fungal hyphae, the roles of MTs are less clear (Heath, 1990; Hepler et al., 2001
; Srinivasan et al., 1996
). MTs are often seen at the apex (Han et al., 2001
; Hoch & Staples, 1985
; McDaniel & Roberson, 1998
; Minke et al., 1999
; Roberson & Fuller, 1988
; Torralba et al., 1998
), but tip growth after MT depolymerization has been reported (deLucas et al., 1993
; Howard & Aist, 1980
; Jochova et al., 1993
; Peterbauer et al., 1992
; Raudaskoski et al., 1994
; Rupes et al., 1995
; Temperli et al., 1991
; That et al., 1988
; Torralba et al., 1996
). However, anti-MT agents also cause reduced growth rates (Akashi et al., 1994
; Jochova et al., 1993
; Niini & Raudskoski, 1993
; Pedregosa et al., 1995
; Temperli et al., 1991
; Torralba et al., 1996
), altered cell shape (Howard & Aist, 1980
; Rupes et al., 1995
), tip swelling (Rupes et al., 1995
; Torralba et al., 1998
), uncharacteristic branching (Raudaskoski et al., 1994
; Rupes et al., 1995
; Torralba et al., 1998
), aberrant secretion of enzymes (deLucas et al., 1993
; Jochova et al., 1993
; Pedregosa et al., 1995
; Torralba et al., 1996
), the gradual disappearance of the Spitzenkörper (a region of densely packed vesicles; Howard & Aist, 1977
) and abnormal vesicle distribution (Howard & Aist, 1980
; Rupes et al., 1995
; Torralba et al., 1998
). However, as the above studies involved exposure for periods (2 h to 3 days) sufficient to affect the number and/or position of nuclei within apical cells, the effects could be a result of aberrant nuclear populations. Furthermore, the depolymerization of MTs has been reported to disrupt the actin network in Candida albicans (Akashi et al., 1994
), so indirect effects of MT disruption on actin-based systems are also possible. As fungal actin networks are notoriously hard to visualize (Heath, 2000
) and monitor, this possibility may be difficult to verify. In addition, irrespective of the possible roles of apical MTs, there are very few data on the mechanisms by which their populations are generated and maintained in the dynamic growing apex.
Using confocal microscopy, we monitored MT dynamics in a strain of Aspergillus nidulans expressing GFP-tubulin. The aims were to determine how the apical population of MTs is generated and maintained and to what extent they contribute to tip growth.
![]() |
METHODS |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
Confocal microscopy and photobleaching
MT observation.
GFPtubulin was visualized with an Olympus Fluoview 300 confocal microscope, a 40 mW multiline argon ion laser and a 60x oil-immersion (Plan Apo, 1·4 NA) objective. During image acquisition, laser power was set to 13 % to minimize photobleaching of the fluorophore and photodamage to the cell. The confocal aperture was kept in position 3 to increase the depth of field. Images were captured using the Fluoview FV300 tiempo (version 4.1) software at a rate of 1 frame s1 for a minimum of 160 s. MTs were also observed via the GFPtubulin using epifluorescence optics (with a Bl filter set: excitation 450495 nm, dichroic mirror 510 nm, barrier >520 nm). All values given in the text are arithmetic means±SD.
Photobleaching experiments.
GFPtubulin was visualized as above, for 2 s, and then a region of interest (mean area 17·74±9·55 µm2, using the Fluoview FV300 software) was bleached for 60 s using the laser at 100 %. Each cell was only photobleached once and was followed for a minimum of 130 s after bleaching.
Confocal image analysis and processing
Image-Pro Plus software (Media Cybernetics) was used to measure MT length and calculate MT growth rates. MT catastrophe (transition from growth to shrinkage) and rescue (transition from shrinkage to growth) values were defined as the number of incidents recorded over the length of time for which the cell was observed. Each MT within a sequence was monitored and transitions from growth to shrinkage or shrinkage to growth noted. A filament was considered to be growing or shrinking if it showed an obvious change in length for more than 3 s. Image-Pro Plus software was used to make fluorescence intensity measurements.
Measurement of changes in cytoplasmic tubulin subunit gradient over time.
The intensity line-plot function was used to measure changes in cytoplasmic tubulin subunit gradients over time following photobleaching. Ten parallel, longitudinal, regularly spaced line-plots spanning the bleached region were collected from several frames of a time series along the length of hyphae. These lines avoided regions occupied by MTs. The ten line-plots were averaged for each time period and their gradients were calculated using linear regression and then plotted over time.
The cytoplasmic GFPtubulin gradients were also determined in a number of dividing cells. Several frames out of a single time series were processed as above, with the ten parallel line-plot functions being taken down the length of the hypha, between the tip and the nearest spindle.
Measurement of cytoplasmic fluorescence intensity near to nuclei.
The mean fluorescence intensity of the area 1 µm from the spindle of the most apical dividing or interphase nucleus was measured using the mean of five adjacent parallel transverse line-plots taken from the region 1 µm tipward of the spindle or nucleus (see Fig. 6).
|
Growth rate measurements.
Hyphal tips were observed using the Polyvar DIC system, which gave a final magnification of 8500x on the screen, a single pixel representing 0·03 µm. Images were collected at a rate of one frame every 5 min for 1520 min (for drug effects) or one frame every 3 min for 612 min (for mitotic hyphae). Growth rates for each interval were calculated and then averaged.
Drug experiments
Drugs used.
Carbendazim (also known as MBC, methyl benzimidazole-2-yl carbamate; Sigma-Aldrich) was dissolved in DMSO at 10 mg ml1 and then stored at 20 °C in the dark. This stock was diluted with culture medium to 2·5 µg ml1 on the day of the experiment. Latrunculin B (lat B; CN Biosciences) was similarly used at 1 mg ml1 (stock) and 20 µg ml1 (working concentration).
Monitoring MT inhibition and recovery.
Small colonies were mounted in medium in a microchamber, MTs were recorded and then MBC was added. MT dynamics were monitored for 1520 min, the drug solution was then washed out with two to four times the chamber's volume of fresh medium and MT recovery was monitored.
Drug effects on growth rates.
Small colonies were mounted in medium in a microchamber and growth was measured for 15 min each before, during and after drug administration. Controls substituted medium for drug solution.
![]() |
RESULTS |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
Hyphal tips are populated by highly dynamic MTs
Approximate correlations between the numbers of MTs seen via fluorescence microscopy and TEM serial sectioning (unpublished observations) indicate that all MTs were revealed by fluorescence microscopy. Image acquisition did not detectably affect cells; scanned hyphae continued to grow at rates comparable to control hyphae and mitosis proceeded normally. Interphase MTs parallel the hyphal axis, extend to the hyphal tip and are often close to the cell cortex (Fig. 1). Hyphal tips contain 6·4±1·5 (n=7) MTs, with 4·1±0·4 (n=110, from 11 cells) seen in a typical focal plane. Compared with TEM data, the bright fluorescent lines represent single MTs.
|
|
Rearward MT movements were rare (<2 %; Table 1) and consisted primarily of MT sliding; only one of the 448 MTs observed grew rearwards. Rearward and tipward sliding rates were similar [13·1±4·4 µm min1 (n=3) versus 14·9±3·2 µm min1 (n=7)]. Thus, the majority of MTs populating the tip appear to have been nucleated in subapical regions and not at the tip.
The dominant tipward movements of MTs and their rapid depolymerization there predict a tip-high gradient of tubulin subunits. Cytoplasmic fluorescence indicates cytoplasmic tubulin; no autofluorescence was seen in strains of Aspergillus not expressing GFPtubulin (with identical microscope settings; data no shown). The cytoplasm between the tip and the first interphase nucleus displayed a tip-high fluorescence gradient (Fig. 2 and Table 2
). However, rearward flow of subunits was not detected; following photobleaching, the residual gradient in the bleached region was primarily tip-low (Fig. 2
and Table 2
), most consistent with influx of unbleached subunits from the subapical region. During recovery, the gradient returned to tip-high (Fig. 2
and Table 2
), presumably due to renewed input of subunits from apically depolymerizing microtubules. Photobleaching should also reveal MT treadmilling. There was no evidence for MT treadmilling (i.e. translocation of the photobleached region within an MT; see Supplementary Fig. S1 available with the online version of this paper), thus indicating that all MT elongation was due to apical polymerization.
|
|
The behaviour of MTs and their subunits in developing branches was similar to that of growing tips (Tables 1 and 2) except that branches had fewer MT initiations (Table 1
) and possibly a higher rate of MT sliding.
MT dynamics of subapical cells differ from apical cells
The region between nuclei in subapical cells displayed fewer MT initiations than apical cells and little bias of movements towards the tips (Table 1). Unlike hyphal tips, subapical cells rarely recovered cytoplasmic fluorescence after photobleaching (Fig. 3
). On the contrary, photobleaching was commonly accompanied by a loss of fluorescence intensity throughout the cell (Fig. 3
). Interestingly, even though cells supposedly maintain a cytoplasmic continuum through septa (Alexopoulos et al., 1996
), the apical cell apparently did not contribute unbleached subunits to the subapical cell's recovery (Fig. 3
).
|
These changes in MT populations related to the mean distance between the tip and the most apical nucleus, with the distance in medium-MT mitotic hyphae being greater than that in low-MT ones (16·11±3·33 µm, n=14, versus 10·80±5·09 µm, n=12; P<0·006, Student's t-test). Furthermore, the cytoplasmic MTs between subapical nuclei were rarely maintained during mitosis, prior to aster formation (Fig. 4).
|
|
The anti-MT drug MBC depolymerizes tip MTs and reduces hyphal growth rate
To monitor tip growth in the absence of MTs, the anti-MT drug MBC was used. After a 10 min exposure, MTs had depolymerized at the tip but persisted around the nucleus and, by 15 min, all MTs were depolymerized. On washout, MTs returned first to the area surrounding the nucleus and then to the region of the hyphal tip (Fig. 7), with 95 % of cells displaying MTs in both regions within 10 min. In no instance did MT polymerization initiate at the tip, although bright spots were occasionally seen there. MT nucleation occurred close to the nuclei, most probably at the spindle pole bodies (SPBs).
|
|
|
Lat B-treated cells did not recover as well as MBC-treated cells. On washout, 58 % of cells showed signs of recovery within 15 min and only 17 % fully recovered after 30 min. Unlike MBC-treated cells, 75 % of cells recovered with the formation of lateral lumps or swellings (Fig. 8d), which often developed into branches (50 %). This was never observed in MBC-treated cells.
![]() |
DISCUSSION |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
Subapical MT nucleation
SPBs are MT organizing centres (MTOCs) (Heath, 1981). While the location of MT nucleation may change during the cell cycle (Heitz et al., 2001
; Straube et al., 2003
), our results indicate that the SPBs are the only MTOCs within Aspergillus; certainly those responsible for MT production in growing hyphae. MTs always originated close to nuclei and the majority of MT movement was tipward. On MBC treatment, the perinuclear region was the last to lose MTs and the first to recover them on washout. MTs that have their ends capped by an MTOC are more stable than free MTs (Keating et al., 1997
) and therefore would be more resistant to anti-MT drugs. Our results are consistent with the localization of
-tubulin (which is MTOC-associated; Murphy & Stearns, 1996
; Oakley, 2000
) to SPBs and not hyphal tips in Aspergillus (Oakley et al., 1990
). They contrast with reported apical MTOC activity in a basidiomycete (Hoch & Staples, 1985
) and a chytridiomycete (McDaniel & Roberson, 1998
), suggesting taxon-specific diversity in MT organization. However, subapical MTOCs in fungi and other tip-growing organisms are more consistent with obligatory tip-high cytoplasmic Ca2+ gradients (Hepler et al., 2001
; Hyde & Heath, 1997
; Jackson & Heath, 1993
), because high concentrations of Ca2+ typically induce MT depolymerization (Dustin, 1984
).
Approximately 30 % of the MTs between the tip and its closest nucleus had both ends free in the cytoplasm (free MTs; data not shown) and MTs often extended or slid past the most apical nucleus en route to the apex. This indicates that the apical MTs are nucleated by the SPBs of numerous nuclei and then exported tipwards. MT release from centrosomes (SPB equivalents) has been reported (Keating et al., 1997). This could explain how MTs are maintained at the tip in mitotic cells when the SPBs of the most apical nucleus are engaged in mitosis; other nuclei at different mitotic stages (mitosis is asynchronous) would produce MTs for the tip. Supporting this, MTs between nuclei often buckled, presumably as a result of oppositely polarized MTs, emanating from subapical and apical nuclei meeting +-end on. MT buckling was never observed between the tip and its closest nucleus.
Cytoplasmic nucleation of MTs may also occur, as Akashi et al. (1997) reported that 30 % of
-tubulin in Aspergillus is in the cytoplasm and Martin et al. (1997)
found that
-tubulin is not essential for cytoplasmic MT assembly. We saw no evidence of cytoplasmic or septal nucleation. Septal nucleation has been reported in Aspergillus, but only for germlings (Konzack et al., 2005
).
Apical MT dynamics
The dominant form of MT production in Aspergillus is elongation at rates similar to those reported previously (Table 1; Han et al., 2001
). The frequency of MT catastrophe was lower than reported previously, and this is probably underestimated, as those that apparently disappeared on reaching the tip probably depolymerized faster than our range of detection. The mean depolymerization rate is the highest reported and is similarly likely to be underestimated. It is almost double that reported for Aspergillus by Han et al. (2001)
but, while both studies sampled at the same frequency, they worked at 42 °C versus 22 °C. Temperature-dependent changes in MT dynamics have been observed in Aspergillus (Requena et al., 2001
). Furthermore, Han et al. (2001)
also used a different strain of Aspergillus. Dynein and a number of kinesins influence MT dynamics (Han et al., 2001
; Konzack et al., 2005
; Requena et al., 2001
; Rischitor et al., 2004
); the two strains, also of differing ploidy, may differ in motor protein expression levels. Alternatively, if the current strain is faster growing than that used by Han et al. (2001)
(not specified in their report), it would have higher cytoplasmic [Ca2+] at its hyphal tips, since higher growth rates correlate with higher apical cytoplasmic [Ca2+] (Hyde & Heath, 1997
). Elevated [Ca2+] accelerates MT depolymerization (above).
MT subunit transport
The tip-high gradient of tubulin demonstrated by cytoplasmic fluorescence in apical cells is probably the result of both MT depolymerization at the apex (possibly generated by the tip-high cytoplasmic [Ca2+]) and the tipward flow of subunits shown by photobleaching. In contrast, subapical regions of apical cells, especially between nuclei, lacked tipward flow of tubulin and the distribution of subunits varied. Thus, the unknown tubulin subunit distribution mechanism is only active in the tips, with variable subapical distribution possibly being due simply to the level of transcriptional activity of the nearest nuclei.
Subapical cells generally displayed an even distribution of tubulin, with the lack of fluorescence recovery after photobleaching indicating an inactive distribution system. Septal pores supposedly permit cytoplasmic continuity (Alexopoulos et al., 1996), but the absence of tubulin influx into photobleached subapical cells suggests that osmiophilic material within the pore (data not shown) selectively limits molecular flux. Consistent with this, entry of nuclei into mitosis in apical cells appears to be coordinated by a diffusible factor (data not shown) which does not trigger mitosis in the subapical cells, suggesting that the trigger molecule(s) does not traverse the septal pores. Photobleaching may have closed the pores; cell damage can do so (Collinge & Markham, 1985
; Trinci & Collinge, 1973
), but this is unlikely since no harmful effects were noted and mitotic cells completed division normally.
Relationship between MT dynamics and nuclear cycle
As apical nuclei entered mitosis, cytoplasmic MTs were reduced or lost; the rate of new MT formation was reduced and these changes were most marked in smaller cells, indicated by shorter nucleus to tip distances. This relationship is similar to the observations of Horio & Oakley (2005) of MT loss at mitosis, but only in germlings. Both observations are consistent with larger cells maintaining larger total tubulin pools (the concentrations of tubulin were similar in mitotic cells whether MTs persisted or not) and thus being able to sustain both mitotic and cytoplasmic MTs. In cells retaining tip MTs, they often extended past the first mitotic nucleus (arrow, Fig. 4a
), indicating that, although the nucleus influences cytoplasmic MTs, it does not simply generate a zone of depolymerization.
Our observations differ from those of Horio & Oakley (2005), who reported greater retention of tip MTs in mitotic hyphae (but not germlings) and no growth rate change. The studies differ in culture temperatures and media, which may cause the different results, but their combined germling and hyphal data are consistent with the independence between cytoplasmic MT regulation, mitosis and hyphal tip growth discussed below.
MT functions in hyphal growth
Our results indicate that normal cytoplasmic MTs are not essential for hyphal tip growth, as witnessed by slowed but morphologically normal growth in the absence of apical MTs in some mitotic cells and all MBC-treated hyphae. Horio & Oakley (2005) likewise reduced (more than we observed, but the variables cited above and a different carrier for the MBC may explain the differences), but did not eliminate, tip growth with MBC. All these data clearly indicate that, while MTs contribute to the tip growth process, they are not obligatory for either growth or normal tip shape. Similar conclusions have been reached previously (e.g. Heath, 1994
; Heath et al., 2000
, 2003
; Oakley & Rinehart, 1985
; Torralba & Heath, 2001
). Possible MT contributions to tip growth are discussed below.
Organelle positioning
MT-dependent processes that, if disrupted, are likely to affect the multicomponent tip growth process include the transportation and organization of vesicles (Heath & Kaminskyj, 1989; Lehmler et al., 1997
; Seiler et al., 1997
; Steinberg & Schliwa, 1993
, 1995
; Steinberg et al., 1998
; Wedlich-Soldner et al., 2000
; Wu et al., 1998
), nuclei (Heath & Kaminskyj, 1989
; Herr & Heath, 1982
; Kaminskyj et al., 1989
; McKerracher & Heath, 1986
; Meyer et al., 1988
; Oakley & Morris, 1980
; Oakley & Rinehart, 1985
; Steinberg & Schliwa, 1993
; Wedlich-Soldner et al., 2000
; Wu et al., 1998
), vacuoles (Allaway et al., 1997
; Herr & Heath, 1982
; Hyde et al., 1999
; Shepherd et al., 1993
; Steinberg et al., 1998
), mitochondria (Heath & Kaminskyj, 1989
; Herr & Heath, 1982
; Steinberg & Schliwa, 1993
; Wu et al., 1998
) and the endoplasmic reticulum (Wedlich-Soldner et al., 2002
). However, direct structural evidence of MT and organelle interactions is rare (Heath, 1994
). Instead organelle trafficking may involve MTs interacting with other cytoskeletal components such as actin (Heath, 1990
, 1994
, 2000
; Kaminskyj et al., 1989
). Our limited ultrastructural data (not shown) support MTactin interactions in Aspergillus; MTs were in contact with actin-like filaments and not organelles or vesicles. The only direct interaction with an organelle involved a mitochondrion (data not shown). Previous tubulin mutant studies in Aspergillus (Oakley & Rinehart, 1985
) indicated that MTs do not participate in mitochondrial transportation, suggesting that our observed association may be the result of chance. A statistical analysis such as that showing a non-random association between mitochondria and MTs in Uromyces (Heath & Heath, 1978
) would resolve this.
There need be no direct link between MTs and organelles for MTs to influence organelle distribution. The cytoplasm consists of a concentrated mixture of interacting proteins (Heuser, 2003; Luby-Phelps, 1993
; McNiven, 2003
) termed the cytomatrix (McNiven, 2003
). Thus, the dynamics of MTs could affect the position of other proteins and organelles without direct contact. MT activity at the hyphal tip could nudge the cytomatrix, thus ensuring organelles and proteins maintain their proximity to the elongating tip. This model is similar to the polar ejection force model for mitosis, in which Rieder et al. (1986)
proposed that MTs growing away from the spindle poles generated movement by impacting upon components of the cytoplasm. This model could explain how the distribution of vesicles which are not in contact with MTs can be affected by MT disruption (Herr & Heath, 1982
).
Cytoplasm migration and subapical vacuolation
Cytoplasm migrates as the hypha extends to maintain its position relative to the tip. Cytoplasmic migration is an active process comparable to amoeboid movement (Bachewich & Heath, 1999; Heath & Steinberg, 1999
; Kaminskyj & Heath, 1996
). This migration can be accompanied by coordinated vacuolation of subapical regions (Bachewich & Heath, 1999
; Heath & Steinberg, 1999
) to minimize the energy expenditure of hyphal growth (Heath & Steinberg, 1999
; Kaminskyj & Heath, 1996
). MTs are involved in vacuolation and vacuolar transport (Allaway et al., 1997
; Bachewich & Heath, 1999
; Herr & Heath, 1982
; Hyde et al., 1999
; Shepherd et al., 1993
; Steinberg et al., 1998
), but the mechanisms that drive cytoplasmic migration are unknown. The high level of MT activity at the apex may help to drive the cytoplasm tipwards, but they are not obligatory, as hyphae grown in the absence of MTs do not display abnormal apical vacuolation (current study; Bachewich & Heath, 1999
). Bachewich & Heath (1999)
suggested that both actin and MTs contribute to cytoplasmic migration, as the bulk migration of cytoplasm seen on recovery from intracellular acidification was sensitive to both MT and actin disruption.
Conclusion
In this study, MTs proved not to be obligatory for growth. However, the level of MT activity at the tip indicates considerable energy commitment, and reduced MTs reduce growth rate. Thus, MTs maximize growth rates but are not essential for either continued growth or normal tip morphogenesis, and their regulation is independent of tip growth. MTs populate the tip via nucleation in subapical regions and both elongation and transport tipwards. Elongation is facilitated by the tipward migration of MT subunits, a process lacking in subapical cells.
![]() |
ACKNOWLEDGEMENTS |
---|
![]() |
REFERENCES |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
Akashi, T., Kanbe, T. & Tanaka, K. (1994). The role of the cytoskeleton in the polarized growth of the germ tube in Candida albicans. Microbiology 140, 271280.[Medline]
Akashi, T., Yoon, Y. & Oakley, B. R. (1997). Characterization of gamma-tubulin complexes in Aspergillus nidulans and detection of putative gamma-tubulin interacting proteins. Cell Motil Cytoskeleton 37, 149158.[CrossRef][Medline]
Alexopoulos, C., Mims, C. & Blackwell, M. (1996). Introductory Mycology, 4th edn. New York: Wiley.
Allaway, W. G., Ashford, A. E., Heath, I. B. & Hardham, A. R. (1997). Vacuolar reticulum in oomycete hyphal tips: an additional component of the Ca2+ regulatory system? Fungal Genet Biol 22, 209220.[CrossRef][Medline]
Bachewich, C. & Heath, I. B. (1998). Radial F-actin arrays precede new hypha formation in Saprolegnia: implications for establishing polar growth and regulating tip morphogenesis. J Cell Sci 111, 20052016.[Medline]
Bachewich, C. & Heath, I. B. (1999). Cytoplasmic migrations and vacuolation are associated with growth recovery in hyphae of Saprolegnia, and are dependent on the cytoskeleton. Mycol Res 103, 849858.[CrossRef]
Collinge, A. J. & Markham, P. (1985). Woronin bodies rapidly plug septal pores of severed Penicillium chrysogenum hyphae. Exp Mycol 9, 8085.
deLucas, J., Monistrol, M. & Laborda, F. (1993). Effect of antimicrotubular drugs on the secretion process of extracellular proteins in Aspergillus nidulans. Mycol Res 97, 961966.
Dustin, P. (1984). Microtubules, 2nd edn. Berlin, Heidelberg & New York: Springer.
Fiddy, C. & Trinci, A. P. J. (1976). Mitosis, septation, branching and duplication cycle in Aspergillus nidulans. J Gen Microbiol 97, 169184.[Medline]
Geitmann, A. & Emons, A. M. (2000). The cytoskeleton in plant and fungal cell tip growth. J Microsc 198, 218245.[CrossRef][Medline]
Han, G., Liu, B., Zhang, J., Zuo, W., Morris, N. R. & Xiang, X. (2001). The Aspergillus cytoplasmic dynein heavy chain and NUDF localize to microtubule ends and affect microtubule dynamics. Curr Biol 11, 719724.[CrossRef][Medline]
Heath, I. B. (1981). Nucleus associated organelles of fungi. Int Rev Cytol 69, 191221.
Heath, I. B. (1988). Evidence against a direct role for cortical actin arrays in saltatory organelle motility in hyphae of the fungus Saprolegnia ferax. J Cell Sci 91, 4147.
Heath, I. (1990). The roles of actin in tip growth of fungi. Int Rev Cytol 123, 95127.
Heath, I. B. (1994). The cytoskeleton in hyphal growth, organelle movements, and mitosis. In The Mycota. I. Growth, Differentiation and Sexuality, pp. 4365. Edited by J. G. H. Wessels & F. Meinhardt. Berlin: Springer.
Heath, I. B. (2000). Organisation and functions of actin in hyphal tip growth. In Actin: a Dynamic Framework for Multiple Plant Cell Functions, pp. 275300. Edited by C. Staiger, F. Baluska, D. Volkmann & P. Barlow. Dordrecht, Boston & London: Kluwer Academic.
Heath, I. B. & Heath, M. C. (1978). Microtubules and organelle movements in the rust fungus Uromyces phaseoli var. vignae. Cytobiologie 16, 393411.[Medline]
Heath, I. B. & Kaminskyj, S. G. W. (1989). The organization of tip-growth related organelles and microtubules revealed by quantitative analysis of freeze-substituted oomycete hyphae. J Cell Sci 93, 4152.
Heath, I. B. & Steinberg, G. (1999). Mechanisms of hyphal tip growth: tube dwelling amebae revisited. Fungal Genet Biol 28, 7993.[CrossRef][Medline]
Heath, I. B., Gupta, G. & Bai, S. (2000). Plasma membrane-adjacent actin filaments, but not microtubules, are essential for both polarization and hyphal tip morphogenesis in Saprolegnia ferax and Neurospora crassa. Fungal Genet Biol 30, 4562.[CrossRef][Medline]
Heath, I. B., Bonham, M., Akram, A. & Gupta, G. D. (2003). The interrelationships of actin and hyphal tip growth in the ascomycete Geotrichum candidum. Fungal Genet Biol 38, 8597.[CrossRef][Medline]
Heitz, M. J., Petersen, J., Valovin, S. & Hagan, I. M. (2001). MTOC formation during mitotic exit in fission yeast. J Cell Sci 114, 45214532.[Medline]
Hepler, P. K., Vidali, L. & Cheung, A. Y. (2001). Polarized cell growth in higher plants. Annu Rev Cell Dev Biol 17, 159187.[CrossRef][Medline]
Herr, F. B. & Heath, M. C. (1982). The effects of antimicrotubule agents on organelle positioning in the cowpea rust fungus, Uromyces phaseoli var. vignae. Exp Mycol 6, 1524.
Heuser, J. (2003). Whatever happened to the microtrabecular concept? Biol Cell 94, 561596.[CrossRef]
Hoch, H. C. & Staples, R. C. (1985). The microtubule cytoskeleton in hyphae of Uromyces phaseoli germlings: its relationship to the region of nucleation and to the F-actin cytoskeleton. Protoplasma 124, 112122.[CrossRef]
Horio, T. & Oakley, B. R. (2005). The role of microtubules in rapid hyphal tip growth of Aspergillus nidulans. Mol Biol Cell 16, 918926.
Howard, R. J. & Aist, J. R. (1977). Effects of MBC on hyphal tip organization, growth, and mitosis of Fusarium acuminatum, and their antagonism by D2O. Protoplasma 92, 195210.[CrossRef][Medline]
Howard, R. J. & Aist, J. R. (1980). Cytoplasmic microtubules and fungal morphogenesis: ultrastructural effects of methyl benzimidazole-2-ylcarbamate determined by freeze-substitution of hyphal tip cells. J Cell Biol 87, 5564.[Abstract]
Hyde, G. J. & Heath, I. B. (1997). Ca2+ gradients in hyphae and branches of Saprolegnia ferax. Fungal Genet Biol 21, 238247.[CrossRef]
Hyde, G. J., Davies, D., Perasso, L., Cole, L. & Ashford, A. E. (1999). Microtubules, but not actin microfilaments, regulate vacuole motility and morphology in hyphae of Pisolithus tinctorius. Cell Motil Cytoskeleton 42, 114124.[CrossRef][Medline]
Jackson, S. L. & Heath, I. B. (1993). Roles of calcium ions in hyphal tip growth. Microbiol Rev 57, 367382.[Medline]
Jochova, J., Rupes, I. & Peberdy, J. (1993). Effect of the microtubule inhibitor benomyl on protein secretion in Aspergillus nidulans. Mycol Res 97, 2227.
Kaminskyj, S. G. W. & Heath, I. B. (1996). Studies on Saprolegnia ferax suggest the general importance of the cytoplasm in determining hyphal morphology. Mycologia 88, 2037.
Kaminskyj, S. G. W., Yoon, K. S. & Heath, I. B. (1989). Cytoskeletal interactions with post-mitotic migrating nuclei in the oyster mushroom fungus, Pleurotus ostreatus: evidence against a force-generating role for astral microtubules. J Cell Sci 94, 663674.
Keating, T. J., Peloquin, J. G., Rodionov, V. I., Momcilovic, D. & Borisy, G. G. (1997). Microtubule release from the centrosome. Proc Natl Acad Sci U S A 94, 50785083.
Ketelaar, T., de Ruijter, N. C. & Emons, A. M. (2003). Unstable F-actin specifies the area and microtubules direction of cell expansion in Arabidopsis root hairs. Plant Cell 15, 285292.
Konzack, S., Rischitor, P. E., Enke, C. & Fischer, R. (2005). The role of the kinesin motor KipA in microtubule organization and polarized growth of Aspergillus nidulans. Mol Biol Cell 16, 497506.
Lehmler, C., Steinberg, G., Snetselaar, K. M., Schliwa, M., Kahmann, R. & Bolker, M. (1997). Identification of a motor protein required for filamentous growth in Ustilago maydis. EMBO J 16, 34643473.
Luby-Phelps, K. (1993). Effect of cytoarchitecture on the transport and localization of protein synthetic machinery. J Cell Biochem 52, 140147.[Medline]
Martin, M. A., Osmani, S. A. & Oakley, B. R. (1997). The role of gamma-tubulin in mitotic spindle formation and cell cycle progression in Aspergillus nidulans. J Cell Sci 110, 623633.
Mata, J. & Nurse, P. (1998). Discovering the poles in yeast. Trends Cell Biol 8, 163167.[CrossRef][Medline]
McDaniel, D. P. & Roberson, R. W. (1998). -Tubulin is a component of the Spitzenkörper and centrosomes in hyphal-tip cells of Allomyces macrogynus. Protoplasma 203, 118123.[CrossRef]
McKerracher, L. J. & Heath, I. B. (1986). Fungal nuclear behaviour analysed by ultraviolet microbeam irradiation. Cell Motil Cytoskeleton 6, 3547.[CrossRef]
McNiven, M. A. (2003). The solid state cell. Biol Cell 94, 555556.
Meyer, S. L. F., Kaminskyj, S. G. W. & Heath, I. B. (1988). Nuclear migration in a Nud mutant of Aspergillus nidulans is inhibited in the presence of a quantitatively normal population of cytoplasmic microtubules. J Cell Biol 106, 773778.[Abstract]
Minke, P. F., Lee, I. H., Tinsley, J. H., Bruno, K. S. & Plamann, M. (1999). Neurospora crassa ro-10 and ro-11 genes encode novel proteins required for nuclear distribution. Mol Microbiol 32, 10651076.[CrossRef][Medline]
Murphy, S. M. & Stearns, T. (1996). Cytoskeleton: microtubule nucleation takes shape. Curr Biol 6, 642644.[CrossRef][Medline]
Niini, S. & Raudskoski, M. (1993). Response of ectomycorrhizal fungi to benomyl and nocodazole growth-inhibition and microtubule depolymerization. Mycorrhiza 3, 8391.
Oakley, B. R. (2000). -Tubulin. Curr Top Dev Biol 49, 2754.[Medline]
Oakley, B. R. & Morris, N. R. (1980). Nuclear movement is -tubulin-dependent in Aspergillus nidulans. Cell 19, 255262.[CrossRef][Medline]
Oakley, B. R. & Rinehart, J. E. (1985). Mitochondria and nuclei move by different mechanisms in Aspergillus nidulans. J Cell Biol 101, 23922397.[Abstract]
Oakley, B. R., Oakley, C. E., Yoon, Y. & Jung, M. K. (1990). Gamma-tubulin is a component of the spindle pole body that is essential for microtubule function in Aspergillus nidulans. Cell 61, 12891301.[CrossRef][Medline]
Ovechkina, Y., Maddox, P., Oakley, C. E., Xiang, X., Osmani, S. A., Salmon, E. D. & Oakley, B. R. (2003). Spindle formation in Aspergillus is coupled to tubulin movement into the nucleus. Mol Biol Cell 14, 21922200.
Pedregosa, A., Rios, S., Monistrol, I. & Laborda, F. (1995). Effect of the microtubule inhibitor methyl benzimidazol-2-yl carbamate (MBC) on protein secretion and microtubule distribution in Cladosporium cucumerinum. Mycol Res 99, 4348.
Peterbauer, C. K., Heidenreich, E., Baker, R. T. & Kubicek, C. P. (1992). Effect of benomyl and benomyl resistance on cellulase formation by Trichoderma reesi and Trichoderma harzianum. Can J Microbiol 38, 12921297.
Raudaskoski, M., Mao, W. Z. & Yli-Mattila, T. (1994). Microtubule cytoskeleton in hyphal growth. Response to nocodazole in a sensitive and a tolerant strain of the homobasidiomycete Schizophyllum commune. Eur J Cell Biol 64, 131141.[Medline]
Requena, N., Alberti-Segui, C., Winzenburg, E., Horn, C., Schliwa, M., Philippsen, P., Liese, R. & Fischer, R. (2001). Genetic evidence for a microtubule-destabilizing effect of conventional kinesin and analysis of its consequences for the control of nuclear distribution in Aspergillus nidulans. Mol Microbiol 42, 121132.[CrossRef][Medline]
Rieder, C. L., Davison, E. A., Jensen, L. C., Cassimeris, L. & Salmon, E. D. (1986). Oscillatory movements of monooriented chromosomes and their position relative to the spindle pole result from the ejection properties of the aster and half-spindle. J Cell Biol 103, 581591.[Abstract]
Rischitor, P. E., Konzack, S. & Fischer, R. (2004). The Kip3-like kinesin KipB moves along microtubules and determines spindle position during synchronized mitoses in Aspergillus nidulans hyphae. Eukaryot Cell 3, 632645.
Roberson, R. W. & Fuller, M. S. (1988). Ultrastructural aspects of the hyphal tip of Sclerotium rolfsii preserved by freeze substitution. Protoplasma 146, 143149.[CrossRef]
Rupes, I., Mao, W., Astrom, H. & Raudaskoski, M. (1995). Effect of nocodazole and brefeldin-A on microtuble cytoskeleton and membrane organization in the homobasidiomycete Schizophyllum commune. Protoplasma 185, 212221.[CrossRef]
Sawin, K. E. & Nurse, P. (1998). Regulation of cell polarity by microtubules in fission yeast. J Cell Biol 142, 457471.
Seiler, S., Nargang, F. E., Steinberg, G. & Schliwa, M. (1997). Kinesin is essential for cell morphogenesis and polarized secretion in Neurospora crassa. EMBO J 16, 30253034.
Shepherd, V., Orlovich, D. & Ashford, A. (1993). A dynamic continuum of pleiomorphic tubules and vacuoles in growing hyphae of a fungus. J Cell Sci 104, 495507.
Srinivasan, S., Vargas, M. & Roberson, R. (1996). Functional, organizational, and biochemical analysis of actin in hyphal tip cells of Allomyces macrogynus. Mycologia 88, 5770.
Steinberg, G. & Schliwa, M. (1993). Organelle movements in the wild type and wall-less fz;sg;os-1 mutants of Neurospora crassa are mediated by cytoplasmic microtubules. J Cell Sci 106, 555564.
Steinberg, G. & Schliwa, M. (1995). The Neurospora organelle motor: a distant relative of conventional kinesin with unconventional properties. Mol Biol Cell 6, 16051618.[Abstract]
Steinberg, G., Schliwa, M., Lehmler, C., Bolker, M., Kahmann, R. & McIntosh, J. R. (1998). Kinesin from the plant pathogenic fungus Ustilago maydis is involved in vacuole formation and cytoplasmic migration. J Cell Sci 111, 22352246.
Straube, A., Brill, M., Oakley, B. R., Horio, T. & Steinberg, G. (2003). Microtubule organization requires cell cycle-dependent nucleation at dispersed cytoplasmic sites: polar and perinuclear microtubule organizing centers in the plant pathogen Ustilago maydis. Mol Biol Cell 14, 642657.
Temperli, E., Roos, U.-P. & Hohl, H. (1991). Germ tube growth and the microtubule cytoskeleton in Phytophthora infestans. Effects of antagonists of hyphal growth, microtubule inhibitors, and ionophores. Mycol Res 95, 611617.
That, T. C., Rossier, C., Barja, F., Turian, G. & Roos, U.-P. (1988). Induction of multiple germ tubes in Neurospora crassa by antitubulin agents. Eur J Cell Biol 46, 6879.[Medline]
Torralba, S. & Heath, I. B. (2001). Cytoskeletal and Ca2+ regulation of hyphal tip growth and initiation. Curr Top Dev Biol 51, 135187.[Medline]
Torralba, S., Pedregosa, A., Lucas, J. D., Dias, M., Monistrol, I. & Laborda, F. (1996). Effects of the microtubule inhibitor methyl benzimidazole-2-yl carbamate (MBC) on production and secretion of enzymes in Aspergillus nidulans. Mycol Res 100, 13751382.
Torralba, S., Raudaskoski, M. & Pedregosa, A. (1998). Effects of methyl benzimidazole-2-yl carbamate on microtubule and actin cytoskeleton in Aspergillus nidulans. Protoplasma 202, 5464.[CrossRef]
Trinci, A. P. J. & Collinge, A. J. (1973). Occlusion of septal pores of damaged hyphae of Neurospora crassa by hexagonal crystals. Protoplasma 80, 5767.[CrossRef]
Vidali, L. & Hepler, P. (2001). Actin and pollen tube growth. Protoplasma 215, 6476.[CrossRef][Medline]
Wedlich-Soldner, R., Bolker, M., Kahmann, R. & Steinberg, G. (2000). A putative endosomal t-SNARE links exo- and endocytosis in the phytopathogenic fungus Ustilago maydis. EMBO J 19, 19741986.
Wedlich-Soldner, R., Schulz, I., Straube, A. & Steinberg, G. (2002). Dynein supports motility of endoplasmic reticulum in the fungus Ustilago maydis. Mol Biol Cell 13, 965977.
Wu, Q., Sandrock, T. M., Turgeon, B. G., Yoder, O. C., Wirsel, S. G. & Aist, J. R. (1998). A fungal kinesin required for organelle motility, hyphal growth, and morphogenesis. Mol Biol Cell 9, 89101.
Received 3 November 2004;
revised 3 February 2005;
accepted 15 February 2005.
HOME | HELP | FEEDBACK | SUBSCRIPTIONS | ARCHIVE | SEARCH | TABLE OF CONTENTS |
INT J SYST EVOL MICROBIOL | MICROBIOLOGY | J GEN VIROL |
J MED MICROBIOL | ALL SGM JOURNALS |