Microtubule dynamics and the role of molecular motors in Neurospora crassa

Microtubule dynamics and the role of molecular motors in Neurospora crassa

Available online at www.sciencedirect.com Fungal Genetics and Biology 45 (2008) 683–692 www.elsevier.com/locate/yfgbi Microtubule dynamics and the r...

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Available online at www.sciencedirect.com

Fungal Genetics and Biology 45 (2008) 683–692 www.elsevier.com/locate/yfgbi

Microtubule dynamics and the role of molecular motors in Neurospora crassa Maho Uchida



, Rosa R. Mourin˜o-Pe´rez b, Michael Freitag c, Salomon Bartnicki-Garcı´a b, Robert W. Roberson a

a School of Life Sciences, Arizona State University, Tempe, AZ 85287-4501, USA Departamento de Microbiologı´a, Centro de Investigacio´n Cientı´fica y Educacio´n Superior de Ensenada, Ensenada, BC, Mexico c Department of Biochemistry and Biophysics, Oregon State University, Eugene, OR 97331-7305, USA

Received 29 August 2007; accepted 19 October 2007 Available online 26 October 2007

Abstract Live-cell imaging methods were used to study microtubule dynamics in the apical regions of leading hyphae and germ tubes of Neurospora crassa expressing b-tubulin-GFP. Microtubule polymerization rates in hyphae of N. crassa were much faster than those previously reported in any other eukaryotic organism. In order to address the roles of motor proteins in microtubule dynamic instability in N. crassa, the microtubule-motor mutant strains, Dnkin and ro-1, were examined. Polymerization and depolymerization rates in leading hyphae of these strains were reduced by one half relative to the wild type. Furthermore, microtubules in germ tubes of wild type and microtubule-motor mutants exhibited similar dynamic characteristics as those in hyphae of mutant strains. Small microtubule fragments exhibiting anterograde and retrograde motility were present in leading hyphae of all strains and germ tubes of wild-type strains. Our data suggest that microtubule motors play important roles in regulating microtubule dynamic instability in leading hyphae but not in germ tubes.  2007 Elsevier Inc. All rights reserved. Keywords: Live-cell imaging; Green-fluorescent protein; Kinesin; Dynein; Dynamic instability

1. Introduction Hyphae are the characteristic tubular cellular element of filamentous fungi. These cellular elements branch frequently and elongate through cytoplasmic mechanisms that sustain polarized transport of secretory vesicles to the hyphal apex and regulate exocytosis at the apical plasma membrane. Thus, hyphal expansion is restricted primarily to the hyphal apex resulting in a highly polarized mode of growth (Bartnicki-Garcı´a et al., 1989; Gow, 1994; Bartnicki-Garcı´a, 2002; Harris and Momany, 2004; Harris et al., 2005). It is through the regulated mechanisms of polarized hyphal growth that fungi undergo cell morphogenesis and interact with their environment.


Corresponding author. E-mail address: [email protected] (M. Uchida).

1087-1845/$ - see front matter  2007 Elsevier Inc. All rights reserved. doi:10.1016/j.fgb.2007.10.013

In order to gain a better understanding of the mechanisms involved in polarized hyphal growth, studies with light and electron microscopy have been used to investigate cytoplasmic organization and dynamics (e.g., Roberson and Fuller, 1988; Lo´pez-Franco et al., 1994; Riquelme et al., 1998; Fischer-Parton et al., 2000). From these works, the view of the Spitzenko¨rper has emerged as a dynamic, pleomorphic, apical body composed of a dense cytoskeletal ground matrix, within which secretory vesicles, ribosomes, signaling molecules, and other elements are arranged spatially and temporally. The Spitzenko¨rper appears to play crucial roles in regulating hyphal extension and morphogenesis and likely serves as an exocytotic regulatory apparatus that receives, distributes, and organizes secretory vesicles and signaling molecules (Bartnicki-Garcı´a, 2002). The involvement of microtubules in polarized hyphal growth has been investigated by use of inhibitors that interfere with microtubule function (e.g., Howard and Aist,


M. Uchida et al. / Fungal Genetics and Biology 45 (2008) 683–692

1977, 1980; Rupes et al., 1995; McDaniel and Roberson, 2000; Horio and Oakley, 2005; Uchida et al., 2005), and mutant strains that disrupt the function of tubulin or microtubule-motor proteins (e.g., Oakely and Morris, 1980; Plamann et al., 1994; Seiler et al., 1997, 1999; Wu et al., 1998; Xiang et al., 1999; Riquelme et al., 2000, 2002). Based on these studies and others from animalbased systems, it is believed the microtubules provide the basis for long distance transport of secretory vesicles from the Golgi apparatus to the Spitzenko¨rper (Seiler et al., 1997; Read and Hickey, 2001; Bartnicki-Garcı´a, 2002). However, a comprehensive understanding of the integrated mechanisms that regulates the rate of apical extension is not fully elucidated. The development and use of fungal strains of Aspergillus nidulans (Fernandez-Abalos et al., 1998; Han et al., 2001; Horio and Oakley, 2005; Sampson and Heath, 2005), Magnaporthe grisea (Czymmek et al., 2005), and Neurospora crassa (Freitag et al., 2004; Mourin˜o-Pe´rez et al., 2006) expressing cytoskeletal proteins tagged with green-fluorescent protein (GFP) have proven invaluable in understanding the dynamic behavior of microtubules and associated proteins in living cells. The most common form of microtubule behavior in eukaryotic cells is dynamic instability. Dynamic instability provides a means for microtubules to reassemble into different structural organizations during cell cycle, growth, and development. Five stages of dynamic instability are recognized: (1) polymerization (growth: the net addition of tubulin heterodimers to the plus end), (2) depolymerization (shrinkage: the net loss of tubulin heterodimers from the plus end, (3) catastrophe (the transitional state between growth and shrinkage), (4) rescue (the transitional state between shrinkage and growth), and (5) pause (a brief stationary state between catastrophe and rescue) (Mitchison and Kirschner, 1984; Heald and Nogales, 2002; Howard and Hyman, 2003). In this study, live-cell imaging methods were employed to investigate apical microtubule dynamics in mature leading-hyphae and germ tubes of N. crassa expressing btubulin-GFP. To study the effects of microtubule-motor mutation on microtubule dynamics, kinesin (Dnkin, kinesin null mutant strain; Seiler et al., 1997) and dynein (ro-1, deficient in one of the heavy chains of cytoplasmic dynein; Plamann et al., 1994) mutants of N. crassa were chosen. 2. Materials and methods 2.1. Strains, media, and growth conditions A list of N. crassa strains used in this study is given in Table 1. These strains were maintained at 25 C on Vogel’s complete medium with 2% (w/v) sucrose. 2.2. Crosses of Bm1+-sgfp+ N. crassa with microtubulerelated motor protein mutants Strains were crossed routinely on Petri dishes with synthetic crossing medium supplemented with 1% (w/v)

Table 1 Neurospora crassa strains used in this study Strain




Reference +


rid mat A his-3 ::Pccg-1-Bml sgfp+ FGSC4351 mat a ro-1 mat a Dnkin RL21SG150 mat a

FGSC Seiler et al. (1997) FGSC

XMF11343 Dnkin his-3+::Pccg-1-Bml+-sgfp+

This study


This study




ro-1 his-3 ::Pccg-1-Bml -sgfp

Freitag et al. (2004)

sucrose and 2% (w/v) agar. For each cross, the N. crassa N2526 strain was first grown on medium for 5 days at 25 C and then fertilized by adding conidia from the second parent (ro-1 or Dnkin). After 14 days of incubation at 25 C, asci from the developed perithecia were collected from the Petri dish cover with distilled water. The ascospores were spread onto Vogel’s minimum medium and incubated for 12 h at 28 C following 2 h heat shock treatment at 60 C. Each developing colony was transferred to 5 ml culture tubes with Vogel minimum medium and incubated for 24 h at 28 C. The colonies were screened to select the ones with the mutation phenotype and GFP expression. 2.3. Digitally enhanced epifluorescence microscopy and image analysis Hyphae and conidia were transferred to water agar for live-cell imaging. Small sections of growing hyphal tips or germlings expressing b-tubulin-GFP were excised and inverted, cell side down, on a cover glass for observation (Hickey et al., 2005). Green FP-tagged microtubules were observed using a standard Eclipse TE300 inverted epifluorescence microscope (Nikon Inc., Instrument Group, Melville, NY) with a Plan Neofluar 100·/1.3 N.A. oil immersion objective lens, a 150 W Xenon bulb, and a blue excitation fluorescence filter (420–495 nm). Time-lapse images were recorded with a low-light level Quantix CCD camera (Roper Scientific, Tucson, AZ) for durations of approximately 2–3 min at 512 · 512 resolution and frame rates of 300–800 msec. For total internal reflection fluorescence (TIRF) microscopy, an IX-70 inverted microscope equipped with a 100·/1.45 Apochromat objective lens (Olympus America Inc., Center Valley, PA) and a krypton/argon laser (Melles Griot, Carlsbad, CA) (488 nm) was used. Images were recorded with a Cascade 512B EMCCD camera (Photometrics, Tucson, AZ) for durations of 2–3 min at 512 · 512 resolution and frame rates of 50–200 msec. For both imaging methods, MetaMorph 6.0/6.1 software (Universal Imaging, Downingtown, PA) was used to control the camera and capture images. Final images were processed using Adobe Photoshop 7.0 (Adobe Systems Inc., San Jose, CA). Analysis of microtubule dynamics was carried out in a similar way as previously described (Dhamodharan and

M. Uchida et al. / Fungal Genetics and Biology 45 (2008) 683–692

Wadsworth, 1995; Dhamodharan et al., 1995; Han et al., 2001). To measure the rates of microtubule dynamics, the original positions of microtubule (+) ends were first marked by drawing a line (= the start line) perpendicular to the ends of microtubules, and then the extension of the (+) end was tracked through individual frames of a time-lapse sequence until the microtubule growth ceased. At this point, a second line (= the finish line) was drawn at the (+) end. The distance between the start and finish lines and time points were calculated using MetaMorph Offline 6.2r6 (Universal Imaging). The rate calculations were done using Microsoft Excel (Microsoft Corp., Redmond, WA). Frequencies of catastrophe and rescue were calculated by dividing the total number of catastrophe or rescue events per sequence by the total time of the sequence. Pausing was recognized as a brief stationary state when microtubule rate of growth or shrinkage was equal to or less than 0.5 lm per frame.


Using the same methodology described above, cell extension rates were measured in the same cells used to study microtubule dynamics. Growth rates for each movie sequence were calculated and then averaged. 3. Results 3.1. Microtubule dynamics in leading hyphae 3.1.1. Wild type Due to the large number of microtubules in hyphae of wild-type N. crassa, it was difficult to monitor the dynamics of individual microtubules using widefield fluorescence microscopy. We, therefore, employed TIRF microscopy to observe microtubule dynamics within the cell cortex. Total internal reflection fluorescence microscopy is an optical technique that uses evanescent waves to excite fluorophores in a restricted region of specimen,

Fig. 1. TIRF microscopy of microtubule polymerization in mature hypha of wild-type strain. The plus end of a single microtubule is indicated by arrows. Elapsed time (sec, msec) is shown in the top right corner of each panel. Scale bar = 5 lm.


M. Uchida et al. / Fungal Genetics and Biology 45 (2008) 683–692

thus only detecting molecules within 200 nm from the interface of the specimen and the coverslip. This technique provided a high signal to noise ratio with low phototoxic effects. Microtubules of wild-type leading hyphae polymerized at a rate of 27.60 ± 7.20 lm min 1 and depolymerized at 53.70 ± 20.40 lm min 1 (Figs. 1 and 2; Table 2; Supplementary movies 1 and 2). Average number of catastrophic events per total time of the sequence was 0.21 s 1 (Table 2). Before undergoing depolymerization, microtubules paused briefly for 1.46 ± 0.82 s 1 (Table 2). Unfortunately, we were not able to determine the frequency of microtubule rescue since these events occurred out of the plane of focus for TIRF microscopy. Short fragments (6.42 lm ± 0.71 lm) of cortical microtubules exhibiting anterograde and retrograde motility (2.31 ± 0.93 lm s 1) were often observed in the hyphal cortex when TIRF microcopy was used (Fig. 3; Supplementary movie 3). Occasionally, these fragments were juxtaposed with and moving along the lengths of established microtubules (data not shown). Severing of these cortical microtubule fragments was also observed (Fig. 4; Supplementary movie 4). A fragment of microtubule (ca 10.10 lm) came into the field of view and severed. The newly formed fragments were rapidly moved away from each other and were no longer in the plane of focus.

3.1.2. Microtubule motor mutants To determine the influence of motor proteins on microtubule dynamics, ro-1 (Fig. 5; Supplementary movie 5) and Dnkin (Fig. 6; Supplementary movie 6) strains were examined. Both mutant strains had fewer microtubules; thus, it was not necessary to use TIRF microscopy for imaging. In addition, we were able to document the frequency of rescue for these mutant strains. The results of both mutant strains were similar in most respects (Table 2). Relative to the microtubules in the wild-type strain, polymerization rates Table 2 Summary of apical microtubule dynamic instability in mature hyphae of wild type, ro-1, and Dnkin Wild type



Polymerization rate (lm min 1)

27.60 ± 7.20 n = 30 (20)

13.86 ± 4.66 n = 55 (20)

14.97 ± 4.06 n = 58 (20)

Depolymerization rate (lm min 1)

53.70 ± 20.40 n = 27 (20)

25.44 ± 10.77 n = 45 (20)

29.58 ± 11.55 n = 48 (20)

Frequency of catastrophe (s 1)




Frequency of rescue (s 1)

Not determined



Duration of pausing (s)

1.46 ± 0.82

7.52 ± 2.90

4.11 ± 1.58

N = number of microtubules measured; numbers in parentheses indicate the number of cells observed, and ± indicates standard deviation.

Fig. 2. TIRF microscopy of microtubule depolymerization in mature hypha of wild-type strain. The plus end of a single microtubule is indicated by arrows. Elapsed time (sec, msec) is shown in the top right corner of each panel. Scale bar = 5 lm.

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Fig. 3. TIRF microscopy of retrograde motility of microtubule fragment motility in wild-type mature hypha. White and black arrows indicate the respective ends of the fragment. Elapsed time (sec, msec) is shown in the top right corner of each panel. Scale bar = 5 lm.

Fig. 4. TIRF microscopy of a microtubule fragment being severed in wild-type mature hypha. The fragment moves into the field of view (b–e; white and black arrows) and is severed as indicated by arrowheads (f). The resulting fragments separate rapidly (g–h). Elapsed time (sec, msec) is shown in the top right corner of each panel. Scale bar = 5 lm.

(ro-1 = 13.86 ± 4.66 lm min 1; Dnkin = 14.97 ± 4.06 lm min 1) and depolymerization rates (ro-1 = 25.44 ± 10.77 lm min 1; Dnkin = 29.58 ± 11.55 lm min 1) were reduced by approximately one half, and frequency of catastrophe was decreased by three-fold. Frequencies of rescue per total time of the sequence in mutant strains were similar (ro-1 = 0.01 s 1; Dnkin = 0.01 s 1). Since the frequency of rescue in the wild-type strain could not be determined, we were not able to assess the effects of motor mutations for this parameter. Microtubules of these mutant strains

paused for a much longer period (ro-1 = 7.52 ± 2.90 s; Dnkin = 4.11 ± 1.58 s) compared to that of wild-type strain (1.46 ± 0.82 s). Microtubule fragments were present in ro-1 and Dnkin strains (data not shown). As in the wild-type strain, these fragments exhibited similar anterograde and retrograde motilities restricted to the cell cortex; however, fragment motility rates in motor mutants were much slower (ro1 = 0.59 ± 0.42 lm s 1; Dnkin = 0.65 ± 0.38 lm s 1) compared to those of wild type.


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Fig. 5. Widefield epifluorescence microscopy of a single microtubule polymerizing (a–c) and depolymerizing (d–f) in ro-1 strain. The plus end of microtubule is indicated by arrows. Elapsed time (sec, msec) is shown in the top right corner of each panel. Scale bar = 5 lm.

3.2. Microtubule dynamics in germ tubes

3.3. Growth rates in leading hyphae and germ tubes

A striking observation was that most elements of microtubule dynamics obtained in growing germ-tubes (Table 3) were indistinguishable among wild-type and microtubule motor mutant strains (Fig. 7). Microtubules of germ tubes polymerized at the rate of 15.63 ± 4.74 lm min 1 and depolymerized at 27.53 ± 8.27 lm min 1. The average catastrophic and rescue events among wild type and microtubule motor mutants were 0.06 s 1 and 0.02 s 1, respectively. Duration of pausing was, however, variable: wild type paused for the longest time (4.36 ± 2.99 s) followed by Dnkin (3.94 ± 2.58 s) and ro-1 (2.81 ± 0.82 s). All the characteristics of microtubule dynamics except the duration of pausing were similar to those of motor mutants in mature hyphae. Microtubule fragments were also observed in wild-type germ tubes but not in microtubule-motor mutants. The average length microtubule fragments was 3.40 ± 1.02 lm. The length of those fragmented microtubules were about half the size of ones found in mature hyphae. The motility rates of microtubule fragments found in wild-type germ tubes were the slowest (0.31 ± 0.22 lm s 1) among the mature hyphae of wild type and microtubule-motor mutants.

To determine if there was any correlation between growth rates and the elements of microtubule dynamic instability, extension rates of leading hyphae and germ tubes were measured (Table 4). The extension rate of leading wild-type hyphae (15.31 ± 0.98 lm min 1) was much faster than those of ro-1 (0.71 ± 0.43 lm min 1) and Dnkin (0.78 ± 0.46 lm min 1) strains. This correlates to the results that wild type exhibits faster microtubule dynamics than microtubule motor mutants in leading hyphae. While the elements of microtubule dynamic instability were similar in most respects in germ tubes, the cell growth rate of Dnkin strain (0.34 ± 0.20 lm min 1) was approximately half of wild-type (0.54 ± 0.23 lm min 1) and ro-1 (0.60 ± 0.25 lm min 1) strains. 4. Discussion 4.1. Microtubule dynamics in wild-type hyphae and germ tubes and the effects of microtubule motor mutation on microtubule dynamics Our results have shown that microtubule polymerization rates in wild-type leading hyphae of N. crassa were much fas-

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Fig. 6. Widefield epifluorescence microscopy of a single microtubule polymerizing (a–c) and depolymerizing (d–f) in Dnkin. The plus end of microtubule is indicated by white arrows. Black arrows indicate microtubule bundling. Elapsed time (sec, msec) is shown in the top right corner of each panel. Scale bar = 5 lm. Table 3 Summary of apical microtubule dynamic instability in germ tubes of wild type, ro-1, and Dnkin Wild type



Polymerization rate (lm min 1)

15.63 ± 4.74 16.52 ± 6.71 16.78 ± 7.39 n = 59 (20) n = 34 (23) n = 31 (20)

Depolymerization rate (lm min 1)

27.53 ± 8.27 26.75 ± 12.45 27.80 ± 12.34 n = 49 (20) n = 29 (23) n = 24 (20)

Frequency of catastrophe (s 1) 0.07 1





Frequency of rescue (s )


Duration of pausing (s)

4.36 ± 2.99 2.81 ± 0.82

3.94 ± 2.58

N = number of microtubules measured; numbers in parentheses indicate the number of cells observed, and ± indicates standard deviation.

ter than in any other eukaryotic cells studied thus far (Han et al., 2001; Sampson and Heath, 2005; Tirnauer et al., 1999; Drummond and Cross, 2000; Shaw et al., 2003; Goncalves et al., 2001) (Fig. 8). Interestingly, microtubule polymerization and depolymerization rates in leading hyphae of both N. crassa and A. nidulans were significantly higher than those reported in the other eukaryotic systems. By comparing microtubule polymerization and depolymerization rates of N. crassa with those of A. nidulans (Sampson and Heath, 2005), polymerization in N. crassa was typically twice as fast, while depolymerization rates in both species were

similar. N. crassa also showed a 10-fold higher frequency of catastrophe events compared to that of A. nidulans (Sampson and Heath, 2005). Mutations of the dynein and kinesin motors had unfavorable effects on microtubule dynamic instability in leading hyphae of N. crassa. This was evident by the fact that polymerization and depolymerization rates were reduced, frequency of catastrophe was reduced, and duration of pausing was longer compared to the same events in wild-type hyphae. The influences of dynein and/or dynactin mutations on microtubule dynamic instability were previously reported in Saccharomyces cerevisiae (Carminati and Stearns, 1997) and germ tubes of A. nidulans (Han et al., 2001). Our results in both kinesin and dynein mutants of leading hyphae provide further evidence that microtubule motors are involved in regulating microtubule dynamics. At present, the specific roles that motors play in influencing microtubule dynamic instability are not fully understood. However, it is reasonable to speculate that microtubule plus-end-binding proteins such as EB1 (Lansbergen and Akhmanova, 2006; Gard and Kirschner, 1987; Morrison et al., 1998; Tirnauer et al., 1999; Rogers et al., 2002) interact with microtubule motors and affect microtubule dynamics. Further studies are required to elucidate these interactions. An unexpected finding of this study was that mutations in microtubule motors did not affect any parameters of


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Fig. 7. Comparison of b-tubulin-GFP distribution in germ tubes of wild type (a) ro-1 (b) and Dnkin (c). In the background of ro-1 mutation, microtubules were disturbed; wild type and Dnkin microtubules were arranged longitudinally along the cell. Arrows (a and c) indicate spindle microtubules. Scale bar = 5 lm.

Table 4 Hyphal growth rates of leading hyphae and germ tubes of wild type, ro-1, and Dnkin Wild type



Leading hyphae (lm min )

15.31 ± 0.98 n = 20

0.71 ± 0.43 n = 21

0.78 ± 0.46 n = 20

Germ tubes (lm min 1)

0.54 ± 0.23 n = 20

0.60 ± 0.25 n = 23

0.34 ± 0.20 n = 20


Dnkin) had reduced polymerization and depolymerization rates, a lower frequency of catastrophe and a longer duration of pausing. Interestingly, these results were analogous to those of ro-1 and Dnkin strains of leading hyphae. The indistinguishable data between wild-type and mutant strains in germ tubes suggest that microtubule motors are not essential for the mechanism of dynamic instability in germ tubes of N. crassa.

N = number of hyphae measured and ± indicates standard deviation.

4.2. The effects of microtubule dynamics and microtubule motor mutation on leading hyphal and germ tube growth microtubule dynamic instability in germ tubes of N. crassa. Relative to wild-type leading hyphae, microtubules in the germ tubes of all the strains (i.e., wild type, ro-1, and

Neurospora crassa exhibits one of the fastest growth rates among filamentous fungi. As opposed to wild-type

Human cancer cell

Microtubule polymerization Microtubule depolymerization

Arabidopsis S. ceravisiae S. pombe A. nidulans germ tube A. nidulans leading hyphae N. crassa germ tube N. crassa leading hyphae 0





Fig. 8. Comparison of microtubule polymerization and depolymerization rates in N. crassa and in the other eukaryotes. The data obtained in this study were compared with previously published data: Aspergillus nidulans (Han et al., 2001; Sampson and Heath, 2005); Schizosaccharomyces pombe (Drummond and Cross, 2000); Saccharomyces ceravisiae (Tirnauer et al., 1999); Arabidopsis (Shaw et al., 2003); human cancer cells (Goncalves et al., 2001).

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hyphae, microtubule motor mutants, ro-1 and Dnkin, are known to grow slow and knobby, which indicate that these motors have an influence in polarized hyphal growth (Seiler et al., 1997; Seiler et al., 1999; Riquelme et al., 2000; Riquelme et al., 2002). Our observations were similar to those previously reported in mature hyphae of N. crassa that extension rates of microtubule motor mutants were much slower than that of wild type. In addition, we have found that both kinesin and dynein mutants displayed strikingly similar microtubule dynamics and hyphal extension rates in leading hyphae. This relationship may suggest that the extension rates of mature hyphae are dependent upon the dynamic characteristics of microtubules, which are then regulated by microtubule motors. In germ tubes, the slow growth observed in Dnkin suggests that kinesin motor is the primary vesicle carrier to the hyphal apex. Dynein motor, however, does not appear to be involved in the growth of germ tubes, as the cell growth rate of ro-1 was same as the wild type. 4.3. Microtubule fragmentation and fragment motility Small microtubule fragments exhibiting anterograde and retrograde motility were present in the cortical regions of all cell types except for microtubule-motor mutants in germ tubes. Rates of fragment motility in wild-type leading hyphae were greater than in motor-mutant strains, suggesting a role of microtubule motors in fragment motility. In wild-type leading hyphae, the speed of fragment translocation was significantly faster than the rates of polymerization and depolymerization of intact microtubules. Therefore, it is unlikely that treadmilling (Rodionov and Borisy, 1997; Shaw et al., 2003; Baas et al., 2005) had a role in this phenomenon. Microtubule-severing proteins such as katanin and spastin, known to produce non-centrosomal microtubules and breaking microtubules into short fragments (Quarmby and Lohret, 1999; Hashimoto, 2003; Baas et al., 2005), have not been identified in N. crassa. Our observations of microtubule fragmentation suggest the presence of these or analogous proteins. 5. Conclusion We studied microtubule dynamics in leading hyphae and germ tubes of N. crassa wild type and microtubule-motor mutant strains expressing b-tubulin-GFP. Our observations of reduced microtubule dynamic rates in hyphae of ro-1 and Dnkin strains suggest a role for molecular motors in regulating these microtubule events. In contrast, the indistinguishable results of germ tubes obtained in wild type and microtubule-motor mutant strains suggest that microtubule motors are not actively involved in regulating microtubule dynamics at this stage of growth. Although we are unsure of the specific roles that the motors play in microtubule dynamics, it is clear that dynein and conventional kinesin promote microtubule dynamics and, consequently, growth of leading hyphae in N. crassa.


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