The role of tip-localized mitochondria in hyphal growth

Copyright © 2005 Elsevier Inc. All rights reserved.

The role of tip-localized mitochondria in hyphal growth

Natalia N. Levina and Roger R. Lew,

Department of Biology, York University, 4700 Keele Street, Toronto, Ont., Canada M3J 1P3


Received 25 May 2005; accepted 23 June 2005. Available online 7 February 2006.

Abstract
Hyphal tip-growing organisms have a high density of tip-localized mitochondria which maintain a membrane potential based on Rhodamine 123 fluorescence, but do not produce ATP based on the absence of significant oxygen consumption. Two possible roles of these mitochondria in tip growth were examined: Calcium sequestration and biogenesis, because tip-high cytoplasmic calcium gradients are a common feature of tip-growing organisms, and the volume expansion as the tip extends would require a continuous supply of additional mitochondria. Co-localization of calcium-sensitive fluorescent dye and mitochondria-specific fluorescent dyes showed that the tip-localized mitochondria do contain calcium, and therefore, may function in calcium clearance from the cytoplasm. Short-term inhibition of DNA synthesis or mitochondrial protein synthesis did not affect either tip growth, or mitochondrial shape or distribution. Therefore, mitochondrial biogenesis may not occur from the tip-localized mitochondria in hyphal organisms.

Keywords: Tip growth; Calcium; Mitochondria; Hyphal growth; Biogenesis; Neurospora crassa; Saprolegnia ferax

Article Outline
1. Introduction
2. Materials and methods
2.1. Strains and growth conditions
2.2. Preparation for microscopy
2.3. Treatment with fluorescent dyes
2.4. Treatment with inhibitors
2.5. Microscopy and objectives
2.6. Image processing
2.7. Mitochondria isolation
2.8. Fluorometric quantitation of Ca2+ dependence of chlortetracycline fluorescence
3. Results
3.1. Tip-localized mitochondrial energization
3.2. Ca2+ sequestration in tip-localized mitochondria
3.3. Tip-localized mitochondria do not undergo biogenesis
4. Discussion
Acknowledgements
References
1. Introduction
The dominant growth form in fungal organisms is a mycelial structure in which hyphal extension is used to invade new territory. Normally under high hydrostatic pressure (Lew et al., 2004), hyphae undergo continuous expansion solely at the tip, creating a tubular extension in a dynamic process called tip growth. Tip growth occurs in many organisms that have cell walls, often in specialized cells such as pollen tubes, root hairs, and rhizoids. Neuronal growth cones exhibit a similar tip extension, but using an amoeboid mechanism. In all the tip-growing cells examined to date, there appears to be a consistent role for calcium, which is found at a high concentration at growing tips (reviewed by Holdaway-Clarke and Hepler, 2003 and Torralba and Heath, 2000).

Tip-growing cells have a polarized cytological architecture. Vesicles often fill the apex, presumably to supply membrane, and cell wall precursors for the expanding tip. The cytoskeleton has been implicated as an organizing element that maintains the tip-localized vesicles during tip extension (Geitmann and Emons, 2000). Amongst tip-growing organisms, the cytology of fungi has been mapped extensively. In hyphal tips of the ascomycete Neurospora crassa, wall vesicles, which fuse with the membrane at the expanding tip are located in a steep gradient 0–5 μm behind the tip (Collinge and Trinci, 1974). This region is also the site of cell wall synthesis based on the incorporation of radioactive wall precursors (Gooday, 1971). A cytoplasmic Ca2+ gradient extends in a less steep gradient from 0 to 15 μm behind the tip, probably due to diffusion of Ca2+ released at the apex during growth (Silverman-Gavrila and Lew, 2003). The Ca2+ gradient is required for hyphal growth (Silverman-Gavrila and Lew, 2000), it is generated by IP3-activated Ca2+ release (Silverman-Gavrila and Lew, 2001 and Silverman-Gavrila and Lew, 2002) from Ca2+-containing vesicles (Torralba et al., 2001). Putative components of vesicle docking mediators have also been located at the apex (Gupta and Heath, 2000).

Mitochondria are present at a high density in some, but not all, tip-growing cells. During root hair initiation, mitochondria are spatially associated with the initiation bulge (Ciamporova et al., 2003). However, there is no indication of a unique tip-localized mitochondrial population behind the vesicle-filled apex of the root hair (Galway, 2000). In pollen tubes, numerous mitochondria are observed in the sub-apical zone behind the vesicle-filled apex (Pierson et al., 1990 and Uwate and Lin, 1980), which are elongate, compared to spherical in the vacuolated zone (Cresti et al., 1977). The establishment of polarity during neuronal growth is closely associated with mitochondrial location (Mattson, 1999), including high mitochondrial densities at the neuronal growth cone, which may function in energy supply (Chada and Hollenbeck, 2004 and Morris and Hollenbeck, 1993) and calcium clearing (Rumpal and Lnenicka, 2003).

Hyphal organisms have a high density of tip-localized mitochondria. In the oomycete Saprolegnia ferax, mitochondria first appear about 5 μm behind the tip, in the central cytoplasm, shifting to peripheral location about 15 μm behind the tip (Heath and Kaminskyj, 1989). The volume fraction of cytoplasm occupied by mitochondria is highest 5–15 μm behind the tip, which was also observed in Neurospora crassa (Lew, 1999). Zalokar (1959) mapped the distribution of mitochondria and mitochondrial biochemical activity in Neurospora, and found that, while mitochondria are located throughout the hyphal apical region (0–150 μm), cytochrome oxidase and succinic dehydrogenase, which were identified histochemically, are first observed 50 μm behind the tip, and increase to a maximal level 100–150 μm behind the tip. Of necessity, Zalokar’s measurements required fixation, which may alter the natural distribution of mitochondria. With a vibrating oxygen electrode, respiratory activity along growing hyphae was measured with 1–2 μm spatial resolution, and was first observed 15 μm behind the tip (Fig. 1C) (Lew and Levina, 2004), behind the high density of tip-localized mitochondria, measured by quantitation of electron micrographs (Fig. 1C) (Lew, 1999) or mitochondria-specific fluorescent dyes in growing hyphae (Figs. 1A and B). This corroborates Zalokar’s observations, and leads to the research question: What are the roles of the tip-localized mitochondria during hyphal growth in hyphal organisms?



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Fig. 1. Tip-localized mitochondria and respiratory activity along growing hyphae. (A) MitoFluor Red fluorescence imaging of mitochondria in a growing hyphae and quantitative fluorescence intensity transects (see Section 2.6). (B) Rhodamine 123 fluorescence imaging of mitochondria in a growing hyphae and quantitative fluorescence intensity transects. (C) Mitochondrial densities from electron micrographs (squares) and oxygen influx measurements (circles) to show that the tip-localized mitochondria do not respire (data are re-drawn from Lew, 1999 and Lew and Levina, 2004).


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Since the tip-localized mitochondria do not supply ATP during hyphal extension, we explored two alternative functions: Ca2+ clearing and mitochondrial biogenesis. Ca2+ clearance is now accepted as a physiological role of mitochondria in animal cells (Gunter et al., 2004 and Nicholls and Chalmers, 2004). N. crassa has been used extensively in studies of biogenesis (Neupert, 1997), which have focused primarily on the mechanisms responsible for protein import into mitochondria. Only limited research has been done on the location of mitochondrial biogenesis. Luck (1963) used [3H]choline to monitor phospholipid incorporation into mitochondria in a choline-requiring mutant of N. crassa. The distribution of [3H]choline followed a Poisson distribution, indicative of incorporation throughout the mycelium. Tip-localized biogenesis may also occur, since the supply of mitochondria must keep pace with the hyphal volume expansion during tip growth; or, the tip-localized mitochondrial population may be maintained by molecular motors and the cytoskeleton (Westermann and Prokisch, 2002). We explored both alternative mitochondrial roles, Ca2+ clearing and biogenesis, using fluorescence microscopy of growing hyphae.

2. Materials and methods
2.1. Strains and growth conditions
Wild-type (74-OR23-1A) N. crassa was obtained from the Fungal Genetics Stock Center (FGSC 987; School of Biological Sciences, University of Missouri, Kansas City, Missouri, USA) (McCluskey, 2003) and maintained on 2% (w/v) agar slants containing Vogel’s minimal medium (Vogel, 1956) plus 1.5% (w/v) sucrose.

2.2. Preparation for microscopy
Conidia from the slants were sown on Petri dishes containing 1.5% agar and Vogel’s minimal medium (with 1.5% sucrose) or OM (organic medium (w/v): 1% glucose, 0.1% peptone, 0.01% yeast extract, 0.1% KH2PO4, and 0.03% MgSO4·7H2O). After overnight incubation at 28 °C, the cultures were flooded with OM. In preliminary experiments imaging chlortetracycline fluorescence, when a simple salt solution was used (BS: 10 mM Mes, 10 mM KCl, 1 mM CaCl2, 1 mM MgCl2, and 133 mM sucrose, pH adjusted to 5.8 with KOH) the dye fluorescence tended to be diffuse, although tip-localized, possibly due to inefficient dye loading. OM was chosen as the medium of choice because of the better imaging clarity and well-defined structures observed in the hyphae loaded with chlortetracycline.

2.3. Treatment with fluorescent dyes
To load cells with chlortetracycline, 500 μl of OM with 50 μM chlortetracycline was added on the surface of the culture. After chlortetracycline addition, cells stopped growing for 10–15 min, some cells formed multiple tips, then continued to grow with normal hyphal morphology. After the cells had recovered for at least 2 h, chlortetracycline fluorescence was imaged on the confocal microscope using an Argon laser (458 nm excitation line). After scanning, hyphal growth slowed and mitochondrial morphology was altered as detected with Rhodamine 123 (Molecular Probes R-302, Abs/Em 507/529, final concentration 2.5 μM from a 2000× stock in methanol) or MitoFluor Red 589 (Molecular Probes M-22424, Abs/Em 588/622, final concentration 500 nM) fluorescence. Thus repeated scanning of the cells damaged them, so that continuous observation of one cell was difficult. Instead, different cells from the same culture were examined before and after various treatments. The same technique was used with other fluorescent dyes for single or dual dye imaging. After growth recovery of chlortetracycline-treated cells, 1 ml of OM with MitoFluor Red (500 nM final concentration) was added to the plate. Within 15–20 min after addition of the solution, cells resumed normal growth. Rhodamine 123 was used in some experiments to determine whether the mitochondria maintain a membrane potential (Scaduto and Grotyohann, 1999) (Fig. 1B; Figs. 2C–F), by adding the dye (2.5 μM final concentration) to the plate from a 2000× stock in methanol as noted above.


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Fig. 2. Mitochondria potential. Mitochondria distribution was monitored with MitoFluor Red (A, fluorescence; B, brightfield images). The mitochondria potential was monitored with Rhodamine 123 fluorescence (C and E, fluorescence; D and F, brightfield images). Images were taken at the times shown, inhibitors were added at time 0 s. Inhibition of growth by cyanide addition is observed at +30 s. Cyanide inhibition of growth had no immediate effect on mitochondria distribution (A and B). Rhodamine 123 fluorescence declined slightly (C and D). Direct depolarization of the mitochondrial potential with μm valinomycin caused complete dissipation of Rhodamine 123 fluorescence (E and F) and mitochondrial shape change (arrow). Bars = 10 μm. Under normal conditions, tip-localized mitochondria do maintain a potential, an important prerequisite for Ca2+ sequestration.


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2.4. Treatment with inhibitors
Hydroxyurea (Sigma) was added at a final concentration of 20 mM. Hydroxyurea is an inhibitor of ribonucleotide reductase and therefore inhibits both nuclear DNA and organellar DNA synthesis (Heinhorst et al., 1985), and would cause depletion of mitochondrial enzymes, such as cytochrome c oxidase, a respiratory chain enzyme encoded from both mitochondrial and nuclear DNA, and therefore should inhibit mitochondrial biogenesis. Hydroxyurea at 10–20 mM is sufficient to induce cell cycle arrest in filamentous fungi (Garcia-Muse et al., 2003); at 30 mM (the lowest concentration examined), Srivastava et al. (1988) reported that hydroxyurea immediately inhibits DNA synthesis in N. crassa. Chloramphenicol (Sigma) was added from a 100 mg/ml ethanol stock solution at a final concentration of 2 mg/ml. Ethanol, at the same final concentration (2% v/v), was used in growth measurement controls. Chloramphenicol is an antibiotic inhibiting mitochondrial protein synthesis, e.g., cytochrome c oxidase and malate dehydrogenase (Howell et al., 1971). Concentrations of 1–4 mg/ml are reported to inhibit mitochondrial protein synthesis in vitro (Sebald et al., 1968) and in vivo (Howell et al., 1971). We used chloramphenicol to inhibit the replication of mitochondria in hyphae. Cyanide (NaCN) was added at a final concentration of 1 mM. Nocodazole (Sigma) was added from 33 mM stock in DMSO to the final concentration 10 μM. For all inhibitor treatments, we measured their short-term effect (20 min) on apical growth rates and tip-localized mitochondrial morphology and distribution by fluorescent labeling with MitoFluor Red.

2.5. Microscopy and objectives
Observations were made on Olympus Fluoview 300 confocal system with Fluoview software. For most experiments, either 40, 60, or 63× water immersion objectives (all infinity tube length) were used. Multi-argon laser excitation line 488 nm and He–Ne laser excitation line 543 nm were used for visualization of cells simultaneously stained with chlortetracycline and MitoFluor Red, with FITC and Texas Red filter cubes and scanned in a linear sequencing mode. Control experiments with single labeled cells demonstrated that there was no fluorescence ‘leakage’ of chlortetracycline fluorescence through the Texas Red filter cube, nor ‘leakage’ of MitoFluor Red fluorescence through the FITC filter cube.

2.6. Image processing
Image processing and analysis were performed using the public domain ImageJ program (developed at the US National Institutes of Health and available on the Internet at http://rsb.info.nih.gov/ij/). When performed, image processing was limited to linear contrast stretch. Analysis included growth rate measurements and RGB merge of chlortetracycline (green) and mitochondria-specific fluorescent dyes (red) to identify regions of co-localization (yellow). Care was taken to assure that the fluorescent images exhibited well-defined structure, not just a diffuse fluorescence that would cause an erroneous identification of co-localized regions of fluorescence. To measure the fluorescence intensity of mitochondria labeled with MitoFluor Red or Rhodamine 123, transects (4 μm wide) were abutted to the hyphal tip in the center of hyphae, which were selected for medial focus extending from the tip to 30–40 μm behind the tip. The intensities for each column of pixels were summed to obtain total fluorescence intensity in arbitrary units.

2.7. Mitochondria isolation
Conidia were inoculated into liquid Vogel’s minimal medium (106 cells/ml) and grown for 6 h at 28 °C in shaker flasks (250 ml) at 100 rpm until almost 90% of conidia germinated and germ tubes were up to 200 μm length.

Germlings were harvested by centrifugation for 10 min at 1500g and resuspended in approximately 10 ml of homogenization medium (HM) (1:1 v/v) (0.25 M sucrose, 10 mM Na2EDTA, 5 mM MgSO4, 25 mM MES, 2.5 mM dithiothreitol, and 1% BSA, pH 7.0 (KOH)). Pre-chilled glass beads (1:1, acid-washed, 150–212 μm (Sigma G-1145), 106 μm Sigma, G-4649) were added in an amount equal to the volume of the germlings suspension (approximately 10 g total). Germlings were homogenized by grinding with a pestle in a pre-chilled porcelain mortar. The progress of homogenization and appearance of broken cells was monitored under a microscope with a X10 phase-objective, to determine when most cells had been disrupted. Then, beads were washed with additional HM; the final suspension was approximately 35 ml.

The homogenate was centrifuged for 10 min at 100g to pellet beads and cell material. Supernatant was then centrifuged for 30 min at 14,700g. The mitochondrial pellet had a characteristic rust color. Purity was confirmed by microscopic examination using a dark-field condenser. Mitochondria were resuspended with a fine camel hair brush in suspension medium (SM) (0.25 M sucrose, 25 mM MOPS, 50 mM KCl, and 5 mM EGTA, pH 7.2 with KOH); the final protein concentration varied between 7 and 16 mg/ml (in four preparations).

2.8. Fluorometric quantitation of Ca2+ dependence of chlortetracycline fluorescence
The Ca2+ dependence of chlortetracycline fluorescence (final concentration 50 μM) of mitochondria suspended in SM (protein concentration 1.0–1.5 mg/ml) was measured with a Cary Eclipse Fluorescence spectrophotometer (Varian, Canada). The excitation wavelength was 380 nm (5 nm slit), emission was at 540 nm (5 nm slit). Free Ca2+ concentration was calculated based on an iterative algorithm and binding constants for EGTA according to Goldstein (1978).

3. Results
To visualize mitochondria in N. crassa we used fluorescent mitochondrial dyes, Rhodamine 123 and MitoFluor Red. Both dyes labeled cable-like structures, which were highly dynamic and moved forward with the growing tip. The density of mitochondria labeled with fluorescent dyes was higher in the growing tips. When growth was temporarily slowed or stopped as a result of solution change, the tip-high gradient in mitochondrial density was not disrupted, while permanently non-growing cells (not treated with inhibitors) had either diffuse fluorescence still brighter at the tip, or round-shaped structures (data not shown).

3.1. Tip-localized mitochondrial energization
The high density of mitochondria at the hyphal tip is well-established based upon imaging of growing cells and electron microscopy; but oxygen influx measurements indicate that the tip-localized mitochondria do not respire (Fig. 1C). Rhodamine 123 fluorescence is reported to depend upon the presence of a membrane potential in isolated mitochondria (Scaduto and Grotyohann, 1999), therefore, we used Rhodamine 123 to confirm that tip-localized mitochondria maintain a potential. Tip-localized mitochondria fluoresced strongly, while mitochondria in hyphal compartments behind the colony edge were only weakly fluorescent (data not shown). Cyanide was used to de-energize the mitochondria, it caused both growth inhibition and a slight decline of Rhodamine 123 fluorescence (Figs.2C and D), but not MitoFluor Red fluorescence (Figs. 2A and B). Direct dissipation of the mitochondrial potential with μm valinomycin caused disappearance of Rhodamine 123 fluorescence (Figs. 2E and F) and growth to slow to about 10% of the normal rate. In 14/21 experiments, before fluorescence disappeared, the vermiform mitochondria changed shape to disk-like structures (Figs. 2E and F). The appearance of disk-like structures appeared to be related to a slower time for valinomycin to diffuse to the hyphae and inhibit growth. Therefore, the tip-localized mitochondria have a membrane potential, but are not synthesizing ATP at a rate sufficient to cause oxygen influx, while mitochondria behind the tip do synthesize ATP at a rate high enough to require oxygen uptake. Because the tip-localized mitochondria are energized, they may function as a Ca2+-sequestering organelle at the tip. We tested this possibility by using chlortetracycline.

3.2. Ca2+ sequestration in tip-localized mitochondria
To ensure that chlortetracycline can be used to image mitochondrial calcium, we assayed for Ca2+-dependent chlortetracycline fluorescence in vitro (Fig. 3). Chlortetracycline fluorescence begins to increase at about 0.75 mM free [Ca2+], and appears to reach a maximum at about 10 mM. Therefore, in vivo, we expect chlortetracycline to ‘report’ on mitochondrial calcium, if the mitochondria contain high levels of calcium.


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Fig. 3. Ca2+ dependence of chlortetracycline fluorescence. Unenergized mitochondria (1 mg/ml) were incubated with 50 μM chlortetracycline and varying free [Ca2+] concentrations as shown. Experiments from three mitochondria isolations are shown by different symbols (circles, left scale; triangles and squares, right scale; one experiment (squares) is the average of two assays from the same mitochondria isolation). Note that chlortetracycline fluorescence begins increasing at about 0.75 mM, and approaches maximal levels at about 10 mM.


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Dual dye imaging was used to determine the localization of chlortetracycline and mitochondrial dye fluorescence in growing N. crassa hyphal tips (Figs. 4A–H). The fluorescence of both dyes was higher in the hyphal tip, and was associated with elongate structures. When the chlortetracycline and MitoFluor Red fluorescence images were merged, for the majority of the fluorescent structures, there was co-localization, indicating that mitochondria do contain high levels of Ca2+. There were also chlortetracycline-fluorescing structures which were not co-localized with mitochondria, indicating at least two calcium-storing organelles in the growing tips. Another hyphal organism, the oomycete Saprolegnia ferax, is reported to have a high density of tip-localized mitochondria (Lew, 1999), and chlortetracycline fluorescing structures identified as mitochondria (Yuan and Heath, 1991); we observed co-localization of chlortetracycline and MitoFluor Red fluorescence (Figs. 4I–L).


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Fig. 4. Partial co-localization of Ca2+ stores and mitochondria at the growing hyphal apex and behind the tip in Neurospora crassa (A–H) and Saprolegnia ferax (I–L). Dual imaging of chlortetracycline-fluorescing Ca2+ stores (A, E, and I) and MitoFluor Red imaging of tip-localized mitochondria (B, F, and J) were merged using green and red pseudocoloring to reveal partial co-localization of the two structures (C, G, and K). Bright field images of the hyphae are shown in (D), (H), and (I). For Neurospora crassa (A–H), the tip-localized mitochondria and calcium stores are shown in (A)–(D). the same hyphae was then scanned just behind the tip (the branch structure in D and H serves as an internal marker of location) in (E)–(H). Images in (A), (E), (F), (I), and (J) were linear contrast stretched to maximize the dynamic range prior to green/red merging in (C), (G), and (K). Bars = 10 μm.


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To confirm that the mitochondrial membrane potential is required for Ca2+ sequestration, we treated the growing tips with valinomycin to dissipate the potential (cf. Figs. 2E and F) and examined the effect on co-localization (Fig. 5). Prior to complete inhibition of growth, mitochondria changed shape, from vermiform to disk-like structures. The shape changes were probably due to the dissipation of the mitochondrial potential (Figs. 2E and F). Eventually, the MitoFluor Red fluorescence became diffuse. However, in some experiments, chlortetracycline still labeled disk-like structures even though the MitoFluor Red fluorescence had become diffuse. Thus, MitoFluor Red localization in mitochondria may depend upon the mitochondrial potential. The chlortetracycline fluorescence observed after dissipating the mitochondrial potential with valinomycin indicates that much of the mitochondrial Ca2+ is unavailable for release.


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Fig. 5. Ca2+ retention in tip-localized mitochondria after depolarization. Valinomycin was used to dissipate the mitochondrial potential (Figs. 2 D and F). It caused mitochondrial shape to change from vermiform to round, most noticeable in the third column (arrow), and growth slowed considerably. Co-localization of mitochondria and chlortetracycline fluorescence was observed after valinomycin treatment. However, MitoFluor Red fluorescence became diffuse, suggesting its localization in mitochondria is dependent on the potential. In other experiments, chlortetracycline continued to label disk-like structures after valinomycin treatment, indicating that much of the mitochondrial Ca2+ is not free to diffuse from the mitochondria. The images were taken at the times shown, valinomycin was added at time 0 s. RGB merging of chlortetracycline fluorescence (A, green) and MitoFluor Red fluorescence (B, red) is shown in (C). Brightfield images are shown in (D). Bar = 10 μm.


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3.3. Tip-localized mitochondria do not undergo biogenesis
Given the high density of mitochondria at the hyphal tip, it is possible that the tip is the site of localized mitochondrial biogenesis. Attempts to quantify mitochondrial DNA (assuming actively dividing mitochondria would contain higher DNA quantities) failed. The vital DNA-specific dye SYTO13 (Molecular Probes) selectively stained mitochondria based upon co-localization with MitoFluor Red. SYTO13 fluorescent-labeling of mitochondria disappeared after inhibition with cyanide, when it stained multiple 2–3 μm round structures behind the tip (presumed to be nuclei). This occurred concomitant with morphological changes to mitochondria, which became compacted and condensed into the apical region (data not shown). Therefore, we were unable to quantitatively assess whether tip-localized mitochondria in growing hyphae contained higher levels of DNA, an indicator of mitochondrial biogenesis.

The alternative strategy was to inhibit mitochondrial biogenesis by inhibiting either DNA synthesis, or protein synthesis. With a growth rate of 20 μm min−1, volume 0–20 μm behind the tip doubles every minute, so tip-localized mitochondrial biogenesis should be rapid. Hydroxyurea treatment had no effect upon growth rates; mitochondrial shape and distribution were also unaffected (Fig. 6). Inhibition of mitochondrial protein synthesis using chloramphenicol caused an occasional transient inhibition of growth, also observed with control additions of ethanol or mock solution changes, and no effect on mitochondrial shape and distribution (Fig. 7).


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Fig. 6. Mitochondrial biogenesis: Hydroxyurea treatment. Hydroxyurea was applied at time 0. Images of hyphae, MitoFluor Red fluorescence and bright field images, are shown −6, 7, and 25 min after treatment. Open symbols show control treatments with OM alone, closed symbols show hydroxyurea treatments. Hydroxyurea had no effect upon mitochondrial shape and distribution, or growth rate. Each image shows a different hypha. Bar = 10 μm.


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Fig. 7. Mitochondrial biogenesis: Chloramphenicol treatment. Chloramphenicol was applied at time 0. Images of hypha, MitoFluor Red fluorescence and bright field images, are shown −6, 5 min, and 20 min after treatment. Open symbols show control treatments with OM plus ethanol. Closed symbols show chloramphenicol treatments; squares, measurements taken during fluorescence imaging on the confocal microscope; circles, growth measurements taken on a Zeiss Axioskop microscope. Chloramphenicol had no effect upon mitochondrial shape and distribution, transient inhibition of growth rate was occasionally observed, but also observed in control treatments, probably due to ethanol. Each image shows a different hypha. Bar = 10 μm.


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To assess whether the cytoskeleton maintains the tip-high distribution of mitochondria, growing cells were perfused with OM containing 10 μM nocodazole, an inhibitor causing microtubule depolymerization (Steinberg and Schliwa, 1993). After treatment, hyphal growth slowed from 12 down to 8 ± 1.3 μm min−1 after 3–5 min, typically hyphae emerged from the swollen tips formed immediately after nocodazole application. After 15–20 min, the hyphae continued growing but very slowly (2–3 μm min−1), with a narrow morphology. Nocodazole also affected the distribution of MitoFluor Red-labeled mitochondria. The mitochondria stopped moving along with the extending tip and clustered in a fixed position behind the tip. The slow growing tips initially did not have any fluorescent mitochondrial structures, but after 20–30 min single vermiform structures were occasionally visible in the tips (Fig. 8).


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Fig. 8. Mitochondrial distribution: The role of microtubules. Nocodazole was applied at time 0. Images of hypha, MitoFluor Red fluorescence and bright field images, are shown −1, 5, 11, and 24 min after treatment. Open symbols show control treatments with OM. Closed symbols show nocodazole treatments. Disruption of microtubules with nocodazole affects mitochondria distribution causing them to be distributed basal to the tip. Growth is not inhibited completely, but slows considerably. Each image shows a different hypha. Bar = 10 μm.


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4. Discussion
Tip-localized mitochondria are observed in some (fungi, oomycetes, and amoeboidal growth of neuronal growth cones), but not all tip-growing cells (root hairs and pollen). Thus they cannot be considered an obligatory cytological feature of tip growth. In hyphal organisms, tip-high mitochondria densities are observed in Saprolegnia ferax, based upon quantitation with electron microscopy. In N. crassa, the tip-high gradient observed with electron microscopy was confirmed in growing cells using a variety of mitochondrial-specific dyes. The two we used, MitoFluor Red and Rhodamine 123, are both reported to accumulate in mitochondria (Haugland, 2002). MitoFluor Red is reported to accumulate in mitochondria regardless of the membrane potential, while Rhodamine 123 is cationic, and accumulates in mitochondria maintaining a potential. There is a tip-high density of mitochondria whether hyphae were growing in a minimal salt solution plus sucrose (BS) or a nutrient-replete solution (OM). Growth rates are the same in either solution (Lew, 1999), which is expected since both solutions supply glucose to fuel growth, either via extracellular invertase (BS) or directly (OM). The principal difference is that OM will supply amino acids, the likely cause of higher H+ influx observed in hyphae growing in OM compared to BS, probably due to H+/amino acid symport activity (Lew, 1999). In N. crassa growing in BS, oxygen flux measurements showed that the tip-localized mitochondria do not consume oxygen, pointing to a unique role of tip-localized mitochondria, separate from ATP production. We have not performed the oxygen flux measurements on hyphae growing in OM, but expect the same result. We did observe that tip-localized mitochondria exhibit higher Rhodamine 123 fluorescence than mitochondria in hyphal trunks behind the growing edge when growing in OM (data not shown), consistent with a difference in respiratory function. To examine one possible role of tip-localized mitochondria, Ca2+ clearing, we used chlortetracycline.

The partial co-localization of chlortetracycline fluorescence and mitochondrial-specific dyes suggested that chlortetracycline is ’reporting’ high Ca2+ levels in mitochondria. Although the [Ca2+] dependence of chlortetracycline fluorescence has been reported for microsomal membranes (Lew et al., 1986), the lipid composition of mitochondria is very different, with significant levels of cardiolipin, an acidic phospholipid that can bind Ca2+. Therefore, we examined the [Ca2+] dependence of chlortetracycline fluorescence in isolated unenergized mitochondria. Free [Ca2+] of at least 500 μM was required for chlortetracycline fluorescence to occur, fluorescence intensity approached maximal levels at about 10 mM. Mitochondria in State four (not synthesizing ATP) have a potential of about −170 mV (Mitchell and Moyle, 1969), sufficient to accumulate [Ca2+] as high as 140 mM when cytoplasmic [Ca2+] is 200 nM. The sensitivity of Rhodamine 123 fluorescence to the mitochondrial membrane potential is well-established in vitro (Scaduto and Grotyohann, 1999). In vivo, Rhodamine 123 fluorescence indicates that the tip-localized mitochondria do have a membrane potential, consistent with in vitro results. Therefore, the mitochondria are competent to accumulate Ca2+. However, recent measurements of mitochondrial free [Ca2+] are about 3–4 μM (reviewed by Nicholls and Chalmers, 2004), contradicting the chlortetracycline quantitation, which suggests that at least 750 μM free [Ca2+] is required for fluorescence to increase. Chlortetracycline may be ’reporting’ on the total mitochondrial Ca2+ store, and cannot be considered a direct quantitative reporter of mitochondrial free [Ca2+]. This is consistent with the chlortetracycline fluorescence observed after the mitochondrial potential is dissipated by valinomycin, suggesting that much of the mitochondrial Ca2+ is not ’free’ but instead unavailable for release after it is sequestered. Co-localization of chlortetracycline and MitoFluor Red was observed in another hyphal organism, the phylogenetically distant oomycete Saprolegnia ferax, confirming a previous report (Yuan and Heath, 1991). Both organisms exhibit similar hyphal morphology and growth rates, as well as a high density of tip-localized mitochondria (Lew, 1999), also observed in the related oomycete Pythium ultimum (Grove et al., 1970). We can conclude that tip-localized mitochondria do play a role in Ca2+ sequestration during tip growth of hyphal organisms. The converse, that tip-localized mitochondria can function as a redundant source of Ca2+ to induce the vesicle fusion required for hyphal expansion cannot be discounted.

With a growth rate of about 20 μm min−1, hyphal extension involves a large and continuous increase in cellular volume at the hyphal tip. Mitochondrial density at the tip is about 30% of cell volume, basally they comprise about 15% of cell volume (Lew, 1999). It is easy to infer that tip-localized mitochondrial densities are so high because mitochondrial biogenesis at the tip supplies mitochondria for the basal regions behind the tip as it grows. Attempts to demonstrate this directly, by quantifying DNA per mitochondria, failed because we were unable to visualize mitochondrial DNA during hyphal growth. Indirect attempts relied upon the known inhibitors of DNA synthesis (hydroxyurea) and organellar protein synthesis (chloramphenicol). If tip-localized mitochondria are undergoing biogenesis, we expected to see rapid effects on growth and mitochondria, within the time frame of volume-doubling of the tip region. The tip-localized mitochondria occur in a zone from 0–20 μm. During growth at about 20 μm min−1, volume-doubling of this region would occur every minute or so. If mitochondrial biogenesis is occurring in the tip region, the tip-high mitochondrial density should decline by about 50% per minute when biogenesis is inhibited. Inhibition of DNA synthesis or organellar protein synthesis had no effect on growth or mitochondrial morphology within 20 min. Therefore, we conclude that tip-localized mitochondria are not undergoing significant biogenesis. Luck (1963) had already documented that mitochondrial biogenesis occurs throughout the mycelium of N. crassa. Our results exclude tip-localized mitochondria biogenesis as an additional source of mitochondria during cellular growth.

The tip-high mitochondria distribution must be maintained by the cytoskeleton, both microtubules (Fuchs et al., 2002) and molecular motors (Fuchs and Westermann, 2005 and Westermann and Prokisch, 2002). Disruption of the molecular motors that move along microtubules decreases growth rate and affects morphogenesis (Seiler et al., 1999). We used disruption of microtubules to corroborate the role of the cytoskeleton in maintaining mitochondria distribution. Mitochondria moved away from the hyphal tip, but growth continued, for at least 60 min at a lower rate. This indicates that tip-high mitochondria distributions are the norm during growth, but are not essential for growth. The decline in growth rate may be because tip-localized mitochondria contribute significantly to growth. However, disruption of the microtubules will affect many other processes that may contribute to maximize growth rates.

To summarize, hyphal organisms maintain a high density of tip-localized mitochondria during hyphal extension. These mitochondria appear to be unique, in that they do not function in ATP production, but do play a role in Ca2+ sequestration and must play a role in maintaining the tip-high Ca2+ gradient required for tip growth (Silverman-Gavrila and Lew, 2003). Their role cannot be considered obligatory, since even after disruption of the distribution of tip-localized mitochondria by depolymerizing the microtubules, growth can still continue.

Acknowledgments
We are grateful for the technical assistance of Ms. Karen Rethoret, and critical comments of Dr. I. Brent Heath.

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Funded by a Discovery Grant from the Natural Sciences and Engineering Research Council of Canada.


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