Ni2+ induces changes in the morphology of vacuoles, mitochondria and microtubules in Paxillus involutus cells


Author for correspondence: Sandra Tuszyńska Tel: +61 2 93852014 Fax: +61 2 93851558 Email:


  • • Organelles of ectomycorrhizal fungi are known to respond to changes in the extracellular environment. The response of vacuoles, mitochondria and microtubules to short-term nickel (Ni2+) exposure were investigated in hyphal tip cells of a Paxillus involutus from a heavy metal-rich soil.
  • • Vacuoles, mitochondria and microtubules were labelled with Oregon Green® 488 carboxylic acid diacetate, 3,3′-dihexyloxacarbocyanine iodide (DiOC6(3)) and anti-α-tubulin antibodies, respectively; hyphae were treated with NiSO4 in the range of 0–1 mmol l−1 and examined microscopically.
  • • Untreated hyphal tip cells contained tubular vacuole and mitochondrial networks. Ni2+ caused loss of organelle tubularity and severe microtubule disruption that were exposure-time and concentration dependent. Fine tubular vacuoles thickened and eventually became spherical in some hyphae, tubular mitochondria fragmented and microtubules shortened and aggregated into patches in most hyphae. Tubular vacuoles reformed on NiSO4 removal and tubular mitochondria in the presence of NiSO4 suggesting cellular detoxification.
  • • These results demonstrate that Ni2+ induces changes in organelle and microtubule morphology. Recovery of tubular organelles to pretreatment morphology after Ni2+ exposure suggests cellular detoxification of the metal ion.


Ectomycorrhizal fungi such as Paxillus involutus are commonly found in heavy metal rich soils (Turnau et al., 1993, 1994; Hartley et al., 1997) where their hyphal tip cells are directly exposed to the soil solution containing metal ions. Hyphal tip cells are especially designed for nutrient uptake (Jennings & Lysek, 1996). Many heavy metal cations are essential in various cellular processes, for example nickel (Ni2+) is a component of many enzyme systems (Hausinger, 1997). However, at elevated concentrations it becomes toxic and is capable of inhibiting enzyme function (Palumbo et al., 2001) as well as protein, RNA and DNA synthesis (Huang et al., 1994). Nickel also induces lipid peroxidation, reactive oxygen species generation (Huang et al., 1994) and mutagenesis (Fletcher et al., 1994) often leading to apoptosis (Lee et al., 1998; Kim et al., 2002).

Cellular perturbations caused by elevated metal concentrations are likely to affect organelles and may be reflected by changes in organelle morphology. In healthy cells various organelles take on tubular shapes, forming interconnected networks (Allen, 1995; Bachewich & Heath, 1997, 1999; Cole et al., 1997, 2000; Hyde et al., 1999). Membrane tubulation results in high surface area to volume ratio which is thought to be important in efficient ATP production and distribution and in formation of proton gradients for ion transport across membranes (Allen, 1995). Such networks are sensitive to intracellular ionic imbalance. For example, the tubular and motile vacuoles and mitochondria of ectomycorrhizal and other fungi vesiculate or fragment when exposed to pH variations, fixatives, antibiotics, cytoskeletal inhibitors and increased cation concentrations (Bachewich & Heath, 1997, 1999; Cole et al., 1997, 2000; Hyde et al., 1999). Disruption of organelle tubularity can thus be considered as a cytotoxic response. In fact, fragmentation of tubular mitochondria is one of the first signs of apoptotic cell death (Zamzami et al., 1996; Frank et al., 2001; Jagasia et al., 2005). Changes in mitochondrial morphology are tightly correlated with their function and are, therefore, a good indication of cellular stress response (Brocard et al., 2003).

Formation of organelle networks as well as changes in their morphology and dynamics depend on membrane fusion-fission processes (Bone et al., 1998; Sesaki & Jensen, 1999) and cytoskeletal components (Steinberg et al., 1998; Bachewich & Heath, 1999; Hyde et al., 1999; Suelmann & Fischer, 2000; Koster et al., 2003). Motor proteins attach to membranes exerting forces to tubulate them along cytoskeletal elements which serve as tracks on which organelle extension is possible (Hirokawa et al., 1998; Koster et al., 2003). Cytoskeleton disruption has been shown to cause vesiculation of tubular vacuoles and fragmentation of tubular mitochondria in various fungi (Steinberg et al., 1998; Bachewich & Heath, 1999; Hyde et al., 1999; Fuchs et al., 2002; Riquelme et al., 2002; McDaniel & Roberson, 2000). Cytoskeletal components are also sensitive to changes in the intracellular ionic environment and various cations have the capacity to depolymerize or stabilize them (Lin & Chou, 1990; Li et al., 1996; Bachewich & Heath, 1999). Cation-induced disruption of the cytoskeletal components may therefore lead to changes in organelle morphology. This paper investigates short-term effects of Ni2+ on tubular vacuoles and mitochondria and on the microtubule (MT) a cytoskeleton of an ectomycorrhizal Paxillus involutus from heavy metal-rich soil.

Materials and Methods

Fungal culture

Paxillus involutus (Batsch: Fr.) Fr. was isolated in Chrzanow, Poland, from a soil containing a total of 9795 mg kg−1 zinc (Zn), 1131.1 mg kg−1 lead (Pb), 123.91 mg kg−1 copper (Cu), 51.09 mg kg−1 cadmium (Cd), and 25 mg kg−1 Ni. Stock cultures were grown in the dark for 2–4 wk at 23°C on modified Melin–Norkrans agar (Marx, 1969) in Petri dishes. Cultures prepared for experiments were grown between two cellophane sheets placed on Melin–Norkrans agar as described in Cole et al. (1997).

Organelle labelling and heavy metal treatment

Hyphal wedges (maximum width, 5 mm) were excised from the growing edge of four replicate colonies using a razor blade; one wedge per colony was used for each treatment. Each wedge was placed in the well of a microscope slide in a solution of either 40 µmol l−1 Oregon Green® 488 carboxylic acid diacetate (Cat. no. O6151; Molecular Probes, Inc., Eugene, OR, USA) to label vacuoles, or 0.55 µmol l−1 3,3′-dihexyloxacarbocyanine iodide (DiOC6(3)) (Cat. no. D-273; Molecular Probes Inc.) to label mitochondria (Campbell, 1983; Cole et al., 1997). After 30 min the fluorescent probe solution was carefully removed by a Pasteur pipette and replaced with a solution of NiSO4 in the range of 0.25–1 mmol l−1 in reverse osmosis (RO) water (0.1 ml). Controls were placed in RO water, or in 1 mmol l−1 K2SO4 (to separate any effects of Ni2+ from those of sulphate). Hyphal wedges were treated for up to 3 h; all solutions were changed every 10–15 min. One hundred randomly chosen hyphal tip cells (with free access to the treatment solution) were examined by fluorescence microscopy for changes in vacuole morphology every 15 min and mitochondrial morphology every 30 min. For each tip cell one of three defined morphological states (fine tubular, thick tubular or spherical) was scored for vacuoles and one of two states (tubular or fragmented) for mitochondria, as described later.


Vacuoles were labelled with Oregon Green® and hyphal wedges treated with 0.25, 0.5 or 1 mmol l−1 NiSO4 for 30 min. Mitochondria were labelled with DiOC6(3) and wedges treated with 1 mmol l−1 NiSO4 for 1 h. At the end of the treatment period the NiSO4 solution was replaced with RO water to test for recovery of organelles. All solutions were changed every 10–15 min. Some wedges were left in the treatment solution to test for any recovery without removal of NiSO4. One hundred hyphal tips in each wedge (n = 4) were examined for changes in organelle morphology every 30 min using fluorescence microscopy.

Freeze substitution and immunocytochemistry

Four fungal colonies were placed in either 0, 0.25 or 1 mmol l−1 NiSO4, or 1 mmol l−1 K2SO4 (as a sulphate control) for 1–3 h. The following methods for freeze substitution and immunofluorescence were modified from Neuhaus et al. (1998). Wedges were excised from the growing edge of each colony, plunged into liquid propane (−185°C), transferred into methanol (−85°C) for 30 min and warmed to room temperature. Methanol was then replaced with phosphate buffered saline (PBS), rinsing for 15 min. The wedges were incubated in 5% skimmed milk powder and 3% bovine serum albumin (BSA) in PBS buffer for 60 min. Next they were incubated with a primary monoclonal anti-α-tubulin antibody clone B-5-1-2 (T-516; Sigma Chemical Co. Ltd, St Louis, MO, USA) in PBS + 1% Tween 20 +1% BSA (PBST) 1 : 1000 for a further 60 min at 37°C. The wedges were then washed in PBST three times for 10 min and incubated with antimouse IgG fluorescein isothiocyante (FITC) conjugate (AD12B; Silenius Laboratory, Hawthorn, Victoria Australia) in PBST (1 : 400) for 60 min at 37°C. The unbound antibody was washed off using PBST, as described earlier, and finally rinsed in PBS only before mounting in Citifluor (Leica Microsystems Pty. Ltd, North Ride, NSW, Australia) followed by microscopic examination. One hundred hyphal tip cells were scored in each wedge (n = 4).


All material was examined with a Zeiss Axiophot microscope fitted with differential interference contrast (DIC) and epifluorescence optics using filter combination BP450-490, FT510 and LP515-585, and a ×63/1.2WC apochromatic water immersion objective. Images were captured with an Image Point CCD camera (Photometrics, Tucson, AZ, USA) and processed on a Dell Precision 420 PC computer using Axio Vision 3.1 software (Carl Zeiss, Jena, Germany).


To check the concentration of Ni2+ in the external solution throughout the experiment, samples were collected every 30 min from the 1 mmol l−1 NiSO4 treatments and from recovery solutions, where the metal ion solution had been removed, and analysed by a Optima 3000 DV Inductively Coupled Plasma (ICP) Emission Spectrometer (Perkin Elmer, Norwalk, CT, USA). The instrument was calibrated with three single element calibration standards (1% HNO3 matrix).

Data collected from studies on organelles were arcsine-transformed, expressed as percentages and analysed by a two-factor analysis of variance, testing the effect of exposure time and NiSO4 concentration as well as the interaction between the two variables.


Effects of NiSO4 on vacuole morphology and recovery

Untreated hyphal tip cells of P. involutus contain a dynamic reticulum of interconnected tubular and spherical vacuoles. Fine tubular vacuoles are most abundant within the apex of the tip cell (Fig. 1a,b). Tubules extend, retract and fuse with other tubules. They also dilate and contract by peristalsis-like movements. Some small spherical vacuoles/vesicles are also present in some tip cells and these travel along the hyphae, in saltatory movements, to fuse with other parts of the reticulum. Larger spherical or ellipsoid vacuoles are found further away from the tip. They are relatively immotile but change shape and form tubular bridges that connect to other spherical vacuoles or extend into fine tubules. This vacuole morphology has been defined as fine tubular vacuoles.

Figure 1.

Fluorescence and corresponding differential interference contrast (DIC) images of vacuoles labelled with Oregon Green® 488 carboxylic acid diacetate, demonstrating the effects of 0.5 mmol l−1 NiSO4 on their morphology and recovery from 0.5 h exposure. (a–f) Vacuoles of hyphal tip cell: (a,b) in reverse osmosis (RO) water (controls) they are mostly fine and tubular, forming a reticulum (ft); (c,d) in NiSO4 at 0.5 h exposure, some are tubular and thick (tt) and some are spherical (sv); (e,f) in NiSO4 at 1 h exposure they are spherical (sv) or ellipsoid (ev). (g–j) Vacuoles in recovering hyphal tip cells: (g,h) 1 h after removal of NiSO4, most are large and ellipsoid (ev) and some are tubular and fine (ft); (i,j) 4 h after removal of NiSO4 most are tubular and fine (ft), forming a reticulum. Bars, 10 µm.

When hyphae were treated with NiSO4 the diameter of fine tubular vacuoles started to increase, so they became thicker (Fig. 1c,d) and less motile. This effect was more pronounced with increasing NiSO4 concentration and/or with longer exposure, which caused vacuoles to fill up the width of the hyphae and fine tubular vacuoles were no longer present. Vacuoles in this state were defined as thick tubular vacuoles. In some hyphae, thick tubular vacuoles formed into spherical vacuoles or larger ellipsoid vacuoles and any tubular vacuoles were lost (Fig. 1e,f). This vacuole morphology was defined as spherical vacuoles.

Paxillus involutus hyphae recovered fine tubular vacuoles when NiSO4 was removed. In some hyphae large ellipsoid or spherical vacuoles extended to produce one or several tubules that progressively became longer and finer (Fig. 1g,h). Alternatively large spherical and ellipsoid vacuoles separated into clusters of small motile vacuoles, some of which formed chains parallel with the hyphal axis and joined to form fine tubular vacuoles. Thick tubular vacuoles progressively extended into fine tubular vacuoles and a tubular reticulum reformed (Fig. 1i,j).

Inductively coupled plasma (ICP) analysis revealed that during a 3-h experiment with 1 mmol l−1 NiSO4 treatment, the concentration of the cation in the solution was approximately constant throughout. Control hyphae treated with RO water or 1 mmol l−1 K2SO4 contained fine tubular vacuoles throughout the 60-min treatment period. Nickel sulphate had an effect on vacuoles that was both time and concentration dependent, and this was quantified (Fig. 2). The proportion of hyphae with fine tubular vacuoles began to progressively decline 15 min after the onset of treatment with NiSO4 (Fig. 2a). This was concurrent with an increasing proportion of hyphae containing thick tubular vacuoles or spherical vacuoles across the NiSO4 treatments (Fig. 2b,c).

Figure 2.

Proportion of hyphal tip cells with a particular vacuole morphology: (a) fine tubular vacuoles, (b) thick tubular vacuoles and (c) spherical vacuoles in either 0 (open squares), 0.25 (closed squares), 0.50 (triangles) or 1 mmol l−1 NiSO4 (closed circles), or 1 mmol l−1 K2SO4 (open circles) during a 3-h exposure. The effect was concentration- and exposure time-dependent (P < 0.0001). Data represent arcsine-transformed percentages from a field of a 100 hyphae in four different colonies for each treatment. Standard error was in the range of ± 0.01–0.8.

Inductively coupled plasma analysis showed that during the recovery period after NiSO4 removal, Ni2+ levels fell to 0.4 mmol l−1 after 0.5 h and were below detection limits thereafter. The extent and time taken for recovery were dependent on NiSO4 concentration and exposure time during treatment. Once NiSO4 was removed, recovery was time-dependent. Following a 0.5 h treatment with 0.25 mmol l−1 NiSO4, 77% hyphae contained fine tubular vacuoles, 18% thick tubular vacuoles and 5% spherical vacuoles (time 0, Fig. 3). During the first hour after removal of the treatment solution, the percentage of hyphae with fine tubular vacuoles continued to decline and that of hyphae with thick tubular vacuoles, or spherical vacuoles continued to rise. Hyphae then began to reform fine tubular vacuoles and by 2.5 h over 96% hyphae had reformed them (Fig. 3).

Figure 3.

Proportion of hyphal tip cells with a particular vacuole morphology: (a) fine tubular vacuoles, (b) thick tubular vacuoles and (c) spherical vacuoles after 0.5 h exposure to 0 (open squares), 0.25 (closed squares) and 0.5 mmol l−1 NiSO4 (triangles). Time 0 indicates end of NiSO4 treatment and transfer to reverse osmosis (RO) water. Data represent arcsine-transformed percentages from a field of 100 hyphae in four different colonies for each treatment. Standard error was in the range of ± 0.02–0.38.

Exposure to 0.5 mmol l−1 NiSO4 for 0.5 h resulted in 37% hyphae with fine tubular vacuoles; 55% hyphae with thick tubular vacuoles and 8% with spherical vacuoles (Fig. 3). The decline in hyphae with tubular vacuoles continued for 1.5 h after removal of NiSO4. Recovery began, thereafter, as an increase in the percentage of hyphae with fine tubular vacuoles concurrent with a decline in hyphae with thick tubular or spherical vacuoles. Four hours after NiSO4 removal 96% of hyphae reformed fine tubular vacuoles (Fig. 3). Hyphae did not recover fine tubular vacuoles from 1 h exposure to 1 mmol l−1 NiSO4 treatment within 4 h after removal of the solution (unpublished).

Effects of NiSO4 on mitochondrial morphology and recovery

In untreated hyphal tip cells mitochondria are motile, tubular and branched forming a loosely interconnected network (Fig. 4a,b). Tubular mitochondria extend along the hyphae and small mitochondrial fragments often travel along the hypha to fuse with other parts of the reticulum. Short tubular mitochondria are abundant in the apex of tip cells, these are less frequent with distance from the apex and instead form very long filaments. This morphology has been defined as tubular mitochondria (Fig. 4a,b).

Figure 4.

Fluorescence and corresponding differential interference contrast (DIC) images of mitochondria labelled with 3,3′-dihexyloxacarbocyanine iodide (DiOC6(3)), the effects of 100 mmol l−1 NiSO4 on their morphology and recovery from 1 h exposure. (a–f) Mitochondria of hyphal tip cells: (a,b) in reverse osmosis (RO) water most are tubular (tm), forming a network; (c,d), in NiSO4 at 1 h exposure some are tubular (tm) and some are fragmented (fm); (e,f) in NiSO4 after 3 h exposure most are fragmented (fm). (g–j) Mitochondria in recovering hyphae: (g,h) 1 h after removal of NiSO4 some are fragmented (fm) and some are tubular (tm); (i,j) 3 h after removal of NiSO4 most are tubular (tm), forming a reticulum. Bars, 10 µm.

Nickel sulphate treatment caused tubular mitochondria to become shorter, until most appeared as fluorescent spots of different sizes (Fig. 4c–f). This mitochondrial morphology was defined as fragmented mitochondria.

Recovery of tubular mitochondria from exposure to NiSO4 was seen as fusion of mitochondrial fragments into tubules and increase in mitochondrial frequency especially in the apex until they resembled tubular mitochondria of control hyphae (Fig. 4g–j).

As with vacuoles, the effects of NiSO4 on mitochondria were concentration- and exposure time-dependent. There was no change in mitochondrial morphology in hyphae treated with RO water and 1 mmol l−1 K2SO4 at any time during the 3-h treatment. There was a decrease in the percentage of hyphae containing tubular mitochondria at 1 h in 0.25 mmol l−1 NiSO4 and at 0.5 h in 0.5 and 1 mmol l−1 NiSO4 (Fig. 5a). The proportion of hyphae with tubular mitochondria continued to decline progressively in all treatments with time until there were 43%, 31% and 17% hyphae with tubular mitochondria remaining at 3 h in 0.25, 0.5 and 1 mmol l−1 NiSO4, respectively (Fig. 5a).

Figure 5.

(a) Proportion of hyphal tip cells with tubular mitochondria in either 0 (open squares), 0.25 (closed squares), 0.50 (triangles) and 1 mmol l−1 NiSO4 (closed circles) or 1 mmol l−1 K2SO4 (open circles) during a 3 h exposure. (b) Proportion of hyphal tip cells with tubular mitochondria after 1 h exposure to 1 mmol l−1 NiSO4. Time 0 indicates end of exposure after which NiSO4 treatment was removed (closed circles) or not removed (open circles). Effects of nickel on tubular mitochondria and recovery were exposure time- and concentration-dependent (P < 0.0001). Data represent arcsine-transformed percentages from populations of 100 hyphae in four different colonies for each treatment. Standard error was in the range ± 0.02–0.76.

Recovery of tubular mitochondria occurred after removal of NiSO4 but also when NiSO4 additions were withheld at 1 h but the solution was not removed. Recovery was time-dependent (P < 0.0001). Following a 1-h treatment with 1 mmol l−1 NiSO4 close to 75% hyphae contained tubular mitochondria (Fig. 5b). Their proportion continued to decline for 0.5 h after removal of NiSO4 and for 1 h after NiSO4 additions were withheld but the solution was not removed. Recovery then began as a progressive increase in the percentage of hyphae with tubular mitochondria until close to 100% hyphae contained them by 3 h (Fig. 5b).

Effects of NiSO4 on microtubules

Control P. involutus hyphae contained long microtubules (MTs) which were grouped into bundles. Most of them were positioned parallel with the hyphal axis and were evenly distributed across the width of the hypha (Fig. 6a). This MT morphology was not affected by a 3-h exposure to 1 mmol l−1 K2SO4 (unpublished). By contrast, treatment with 0.25 mmol l−1 NiSO4 for 3 h caused aggregation of MT bundles to one side and substantially decreased the amount of MTs in the apex of 90% (± 3.54) of hyphal tip cell (Fig. 6b). Three-hour treatment with 1 mmol l−1 NiSO4, caused shortening of MTs which aggregated and separated into large, uneven patches. Individual MTs were not distinguishable within these patches in 95% (± 3.49) of hyphal tip cells (Fig. 6c). The MT patches were also usually concentrated on one side of the hypha. Very few MTs remained in the apex of these hyphae (Fig. 6c).

Figure 6.

Fluorescence images showing microtubules (MTs) labelled with anti-α-tubulin antibodies in hyphal tip cells: (a) in reverse osmosis (RO) water for 3 h, MTs are straight and parallel with the hyphal axis (arrowhead); (b) in 0.25 mmol l−1 NiSO4 for 3 h, MTs are wavy, bundled and aggregated to one side of the hypha and are scarce in the apex (arrowheads); (c) in 1 mmol l−1 NiSO4 for 3 h, MTs are short and aggregated into large patches concentrated mostly at one side of the hypha and are scarce in the apex (arrowheads). Bars, 10 µm.


Effects on vacuoles and mitochondria are caused by Ni2+

In control P. involutus hyphal tip cells, fine tubular vacuoles form a very dynamic interconnected reticulum especially in the apex. Similarly, the mitochondria are tubular and form a loosely interconnected network. Exposure to NiSO4 caused thickening of the fine tubular vacuoles and eventual formation of spherical vacuoles in some hyphae, as well as fragmentation of mitochondria. The effects were governed by both exposure time and NiSO4 concentration and were likely to be Ni2+ and not SO42– induced since K2SO4 had no such effects on the organelles. These finding appear to be the first to demonstrate Ni2+ effects on fungal organelles.

The experimental system was designed to test short-term organelle reaction to a range of sublethal Ni2+ concentrations which were kept constant, as confirmed by ICP analysis. Judging from the progressive organelle response, Ni2+ was most likely taken up by the hyphal tip cells which are adapted for rapid nutrient uptake (Jennings & Lysek, 1996). Nickel uptake is thoroughly documented in other fungal cells (Mohan et al., 1984; Joho et al., 1995; Nishimura et al., 1998; Dönmez & Aksu, 2001) and it has been localized within Hebeloma crustuliniforme ectomycorrhizas (Brunner & Frey, 2000). Vacuole vesiculation observed during Ni2+ exposure also occurs by exposure to acids, cytoskeletal drugs, fixatives and fluorescent probes (Wilson et al., 1990; Cole et al., 1997, 2000; Bachewich & Heath, 1999; Hyde et al., 1999), suggesting that this is a general response to toxins.

The fragmentation of mitochondria seen here probably occurs by excessive mitochondrial fission as in other cell types (Frank et al., 2001; Sugioka et al., 2004). Mitochondrial fragmentation could be a response to Ni2+ toxicity, which is known to inhibit mitochondrial enzymes, Ca2+ uptake, ATP, respiration and electron transport (Ligeti et al., 1981; Messer et al., 2000; Pochet et al., 2003). Nickel also induces DNA damage and reactive oxygen species generation leading to apoptosis (Lee et al., 1998; Qu et al., 2001; Kim et al., 2002; Pulido & Parrish, 2003). Mitochondrial fragmentation is often associated with apoptosis, causing the release of apoptotic proteins (Zamzami et al., 1996; Jagasia et al., 2005). However, inhibition of mitochondrial fragmentation also inhibits apoptosis (Frank et al., 2001; Sugioka et al., 2004). It is possible that mitochondrial fragmentation occurs either to generate more mitochondria to compensate for damage in some of the organelles, or to generate enough energy for apoptosis of damaged cells (Brocard et al., 2003). Nickel-induced mitochondrial fragmentation seen in P. involutus may therefore indicate apoptotic behaviour in this fungus, which is a feature of various other fungal cells (Cheng et al., 2003; Phillips et al., 2003; Coyle et al., 2004; Chen & Dickman, 2005).

Ni2+ effects on the cytoskeleton

Microtubules in P. involutus are straight, long, parallel with the hyphal axis and evenly distributed across the width of hyphal tip cells. Nickel caused shortening of MTs and aggregation into patches. This appears to be the first report of Ni2+ effects on MTs in fungal cells. In animal cells MTs are a Ni2+ target, enhancing their polymerization, making them shorter, severely aggregated, bundled, redistributed and stabilized (Lin & Chou, 1990; Li et al., 1996). Nickel has this effect by enhancing acetylated α-tubulin levels or by interacting with MT-associated proteins (MAPs) (Li et al., 1996). These results suggest that Ni2+ effects on MTs are related to the disruption of organelle tubularity and motility in P. involutus. Microtubule disruption causes vacuole vesiculation and mitochondrial fragmentation in other fungal cells (Steinberg et al., 1998; Bachewich & Heath, 1999; Hyde et al., 1999; McDaniel & Roberson, 2000; Khalaj et al., 2001; Fuchs et al., 2002). Interestingly MT disruption also triggers apoptosis through the mitochondrial pathway (Chen et al., 2002; Giacca, 2005.

Instead of MTs, various fungal cells rely on the actin cytoskeleton for organelle motility and morphology (Morris & Hollenbeck, 1995; Simon et al., 1995; Gupta & Heath, 1997; Bachewich & Heath, 1998, 1999; Suelmann & Fischer, 2000). Nickel is known to disrupt actin (Strzelecka-Gołaszewska et al., 1978; Simchowitz & Cragoe, 1990; Pulcinelli et al., 1998; Dalledonne et al., 1999), and actin disruption can lead to apoptosis via the mitochondrial pathway (Gourlay & Ayscough, 2005). Thus, it can not be ruled out that the effects on organelle morphology observed here were not associated with Ni2+ effects on actin; this requires future investigation.

Recovery of organelle networks from Ni2+

Paxillus involutus hyphal tip cells were capable of Ni2+ detoxification, as demonstrated by the return of organelle tubularity. While vacuoles reformed a tubular reticulum after removal of Ni2+, mitochondria did so at twice the concentration as well as in presence of Ni2+. In fact, recovery of tubular mitochondria occurred when vacuoles remained spherical or ellipsoid. Such results stress the importance of mitochondrial ‘health’ in cell survival and further support the fact that mitochondrial function relies on its shape and distribution (Brocard et al., 2003; Logan, 2003). The return of mitochondrial function probably required removal of the possibly accumulated cytosolic Ni2+. During recovery of Saprolegnia ferax hyphae from acid treatment the tubular vacuole reticulum undergoes fusion and expansion into a large ellipsoid form similar to that observed here. Such change in vacuole morphology is suggested to restore cytoplasmic homeostasis from cellular ion imbalance and osmotic changes, by vacuoles sequestering the harmful agent (Bachewich & Heath, 1999). However, there is no solid evidence for vacuolar sequestration in ectomycorrhizal fungi (Orlovich & Ashford, 1993; Turnau et al., 1994).

Other common ways to detoxify the cytoplasm are through metal-binding compounds such as superoxide dismutase, glutathione and metallothioneins, which have been isolated from various P. involutus strains in response to Cd2+ and Cu2+ (Howe et al., 1997; Schützendübel & Polle, 2002; Jacob et al., 2001). However, the mechanisms of Ni2+ detoxification have yet to be investigated in this strain.

Recovery of tubular organelle networks suggests that membrane fusion and tubulation processes resume once the metal stress is removed, they also indicate the importance of these networks in proper cell growth and function, especially with regard to mitochondria (Karbowski & Youle, 2003; Jagasia et al., 2005). The protocols developed here provide an experimental system for investigating short-term heavy metal cytotoxicity and detoxification based on organelle and cytoskeletal response.


The author would like to thank Anne Ashford for her expert advice, Katarzyna Turnau for providing the fungal culture, methods for culture growth and Louise Cole for help with immunocytochemistry protocols and providing some of the antibodies, Danielle Davies, Steve Bonser and Bettye Rees for critically assessing the manuscript, Jerzy Jankowski and Dorothy Yu for help with ICP analysis and the Australian Research Council for grants.