Rust fungi of New Zealand - An introduction, and list of recorded species

Rust fungi of New Zealand - An introduction, and list of recorded species
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Author(s): McKenzie EHC
Source: NEW ZEALAND JOURNAL OF BOTANY Volume: 36 Issue: 2 Pages: 233-271 Published: JUN 1998
Times Cited: 9 References: 128 Citation Map
Abstract: An overview of the rust fungi (Basidiomycota, Teliomycetes, Uredinales) is presented as an introduction towards a new rust mycoflora for New Zealand. All species recorded from New Zealand are listed, together with details on their host plants, a reference to the first New Zealand record of each unique rust/host combination, and a separate alphabetical list of host plants and the rust fungi which parasitise them. New Zealand has a depauperate rust flora consisting of 234 recorded species, of which 54% are native. Of 22 genera recorded from New Zealand only five genera (Hamaspora, Kuehneola, Phragmidium, Puccinia, Uromyces) and three form genera (Aecidium, Caeoma, Uredo) contain native species. Only five genera (Phragmidium, Melampsora, Puccinia, Uromyces, Uromycladium) and two form genera (Aecidium, Uredo) are represented by more than two species. Melampsora contains mainly adventive species; approximately half of the Phragmidium, Puccinia, and Uromyces species and all the Uromycladium species are adventive. Some 95% of the Uredo species and all the Aecidium species are native. Only eight native rusts have spread to exotic hosts; Uredo puawhananga commonly infects some exotic, cultivated Clematis species, while Puccinia lagenophorae is sometimes troublesome on cultivars of Bellis perennis. None of the 17 exotic rusts infecting native plants is of economic or conservation concern.The widespread rusts in New Zealand are often adventive species. Only 5% of adventive rusts are confined to the South Island, but 30% are confined to the North Island. This inequity probably reflects the warmer conditions in the north, and the fact that adventive species are often of tropical origin. Of the native species, 34% occur only in the South Island and just 14% are restricted to the North Island. Since 1945, on average, more than one new adventive rust has been found per year. Most of them are of northern temperate origin, but often considered to be introduced from Australia by trans-Tasman airflows.
Document Type: Review
Language: English
Author Keywords: Uredinales; New Zealand; rusts; fungi; checklist; host list
KeyWords Plus: PLANT-DISEASE RECORDS; CHATHAM-ISLANDS; PATHOGENS; AUSTRALIA
Reprint Address: McKenzie, EHC (reprint author), Landcare Res, Herbarium PDD, Private Bag 92170, Auckland, New Zealand
Addresses:
1. Landcare Res, Herbarium PDD, Auckland, New Zealand
Publisher: SIR PUBLISHING, PO BOX 399, WELLINGTON, NEW ZEALAND
Subject Category: Plant Sciences
IDS Number: 100MK
ISSN: 0028-825X

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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.


Corresponding author. Fax: +1 416 736 5698.

Expressed sequence tag analysis of the soybean rust pathogen Phakopsora pachyrhizi

Published by Elsevier Inc.

Expressed sequence tag analysis of the soybean rust pathogen Phakopsora pachyrhizi

Martha Lucia Posada-Buitrago1 and Reid D. Frederick,

USDA-Agricultural Research Service, Foreign Disease-Weed Science Research Unit, 1301 Ditto Avenue, Fort Detrick, MD 21702, USA


Received 5 November 2004; accepted 10 June 2005. Available online 15 November 2005.

Abstract
Soybean rust is caused by the obligate fungal pathogen Phakopsora pachyrhizi Sydow. A unidirectional cDNA library was constructed using mRNA isolated from germinating P. pachyrhizi urediniospores to identify genes expressed at this physiological stage. Single pass sequence analysis of 908 clones revealed 488 unique expressed sequence tags (ESTs, unigenes) of which 107 appeared as multiple copies. BLASTX analysis identified 189 unigenes with significant similarities (Evalue < 10−5) to sequences deposited in the NCBI non-redundant protein database. A search against the NCBI dbEST using the BLASTN algorithm revealed 32 ESTs with high or moderate similarities to plant and fungal sequences. Using the Expressed Gene Anatomy Classification, 31.7% of these ESTs were involved in primary metabolism, 14.3% in gene/protein expression, 7.4% in cell structure and growth, 6.9% in cell division, 4.8% in cell signaling/cell communication, and 4.8% in cell/organism defense. Approximately 29.6% of the identities were to hypothetical proteins and proteins with unknown function.

Keywords: Phakopsora pachyrhizi; Genome analysis; cDNA sequencing; Expressed sequence tags; Gene expression

Article Outline
1. Introduction
2. Materials and methods
2.1. Fungal isolate and growth conditions
2.2. cDNA library construction
2.3. DNA sequencing
2.4. Data handling
3. Results
3.1. EST analysis
3.2. Gene families
4. Discussion
Acknowledgements
References
1. Introduction
Soybean rust causes significant yield loss to soybean crops in Asia, Africa, Australia, and nearly all tropical countries in the Eastern Hemisphere where soybeans are grown have reported its occurrence (AVRDC, 1987 and AVRDC, 1992). Recent findings of soybean rust in Hawaii in 1994 (Killgore and Heu, 1994), Zimbabwe in 1998 (Levy, 2003), Nigeria in 1999 (Akinsanmi and Ladipo, 2001), South Africa in 2001 (Pretorius et al., 2001), Paraguay in 2001 (Morel, 2001), Brazil and Argentina in 2002 (Rossi, 2003 and Yorinori et al., 2002), and Bolivia in 2003 (Yorinori et al., 2005) demonstrate that Phakopsora pachyrhizi is spreading to new geographic regions. Rust is considered to be a major threat to soybean production in the United States (Sinclair, 1989), especially with the identification of P. pachyrhizi in Louisiana in November 2004 (Schneider et al., 2005). In Brazil, this disease was estimated to cost growers approximately $1.2 billion (USD) in 2003 alone: $500 million in direct yield losses to the disease and $700 million resulting from inappropriate use of fungicides (Yorinori et al., 2005). If P. pachyrhizi becomes established in the continental USd, serious yield losses are likely to occur. It has been estimated that yield losses could exceed 10% in most of the United States with up to 50% yield loss in the Mississippi Delta and southeastern states (Yang et al., 1991).

Four single resistances genes, Rpp1–4 (for resistance to P. pachyrhizi), have been described that impart resistance to some isolates of P. pachyrhizi (Bromfield and Hartwig, 1980, Hartwig, 1986, Hartwig and Bromfield, 1983 and McLean and Byth, 1980). However, no soybean lines have been found with broad-spectrum resistance to all isolates of P. pachyrhizi, and all of the commercial soybean cultivars currently grown in the US are susceptible to soybean rust. In countries where rust has become problematic to commercial production, control strategies have relied on the use of fungicides; however, most growers in the US currently do not apply fungicides to soybeans. The increased costs associated with multiple applications of fungicides might be prohibitive for some growers in the US, and there are concerns about the potential negative effects to the environment if fungicides are applied to such large production acreage.

Soybean rust is caused by two closely related species of fungi, P. pachyrhizi Sydow and P. meibomiae (Arthur) Arthur, which are differentiated based upon morphological characteristics of the telia (Ono et al., 1992). Sequence analysis of the internal transcribed spacer region of the ribosomal RNA genes revealed approximately 80% similarity between these two Phakopsora species; however, only a few nucleotide differences were observed among isolates of P. pachyrhizi or P. meibomiae (Frederick et al., 2002). Unlike most other rust pathogens, both Phakopsora species infect and produce disease symptoms on a wide range of host plants. P. pachyrhizi naturally infects 31 species in 17 genera of Leguminosae, and it has been found to infect 60 species in other genera under controlled conditions (Rytter et al., 1984 and Sinclair and Hartman, 1996). Similarly, P. meibomiae infects 42 species in 19 genera of Leguminosae, and it can infect 18 species in another 12 genera following artificial inoculation (Sinclair and Hartman, 1996). On soybeans, P. pachyrhizi is the more aggressive pathogen and causes considerably more yield loss compared to P. meibomiae.

Phakopsora pachyrhizi produces three types of spores. The urediniospore is the most common spore type and is found throughout the growing season on soybeans and other legume hosts. Urediniospores are produced in large quantities, easily wind disseminated, and multiple spore cycles occur throughout the growing season. Telia and teliospores have been observed on infected plants late in the season in Asia as well as in greenhouse studies (Bromfield, 1984 and Yeh et al., 1981). Teliospore germination and the subsequent production of basidiospores have been reported, but only under laboratory conditions (Saksirirat and Hoppe, 1991). As no alternate host has been identified, there has been no further characterization of the life cycle.

Most of the published research on soybean rust has focused on monitoring disease development, evaluating yield losses, modeling epidemics, host range studies, developing risk assessment models, and screening for sources of resistance. In addition, there have been several reports on the basic biology of the fungus, including histological studies using susceptible lines and those containing single resistance genes (Bonde et al., 1976, Hartwig and Bromfield, 1983 and Sinclair and Hartman, 1996). The infection process employed by P. pachyrhizi consists of several distinct steps: attachment of the spore to the host surface, spore germination, formation of the appressorium, penetration through the cuticle, and invasive growth within the host plant (Bonde et al., 1976). Understanding these processes at both the biochemical and molecular levels is essential for developing new methods of disease management.

Here, we report the first assessment of gene expression at a critical stage of the P. pachyrhizi life cycle: urediniospore germination. This study identifies transcripts present in germinating urediniospores and provides insight into the biochemical processes that occur at this developmental stage. Some of the genes expressed display a high degree of similarity to genes described in other fungi and plants, but the majority corresponded to unclassified genes or genes of unknown function. A preliminary report of this work has been given (Posada and Frederick, 2002).

2. Materials and methods
2.1. Fungal isolate and growth conditions
The P. pachyrhizi isolate Taiwan 72-1 (TW 72-1) used in this study was maintained at the USDA-ARS Foreign Disease-Weed Science Research Unit (FDWSRU) Plant Pathogen Biosafety Level 3 Containment Facility at Ft. Detrick, MD (Melching et al., 1983) under the appropriate USDA Animal and Plant Health Inspection Service (APHIS) permit. TW 72-1 was propagated by spray inoculation onto soybean plants, and urediniospores were harvested from infected leaves 10–14 days following inoculation and at subsequent intervals using a mechanical harvester (Cherry and Peet, 1966). Urediniospores were maintained under liquid nitrogen. Frozen urediniospores were heat shocked at 42 °C for 5 min, and 300 mg of spores was germinated in 300 ml distilled water in a sterile 13 in. × 9 in. Pyrex baking dish for 16 h at room temperature. The fungal tissue was collected using a spatula, frozen in liquid nitrogen, and used for RNA extractions.

2.2. cDNA library construction
Total RNA was isolated from germinating spores of P. pachyrhizi isolate TW 72-1 using the ToTally RNA kit (Ambion, Austin, TX, USA), and the poly(A)+ mRNA was purified using an OLIGOTEX mRNA purification kit (Qiagen, Valencia, CA, USA) according to the manufacturer’s instructions. A unidirectional cDNA library was constructed in the plasmid pSPORT1 using the Superscript Plasmid System for cDNA synthesis and Cloning (Invitrogen, Carlsbad, CA, USA) following the manufacturer’s protocol. The titer of the library was approximately 20,000 colonies, and 5000 individual colonies were transferred to 96-well microtiter plates containing Luria broth with 15% (v/v) sterile glycerol. The plates were archived by storing in a freezer at −80 °C, and 908 clones were sent for sequencing.

2.3. DNA sequencing
Prior to sequencing, all colonies were checked for the presence of an insert by colony-PCR using the SP6 and T7 primers. The PCR products were separated by electrophoresis using 1.5% agarose gels. DNA was prepared for sequencing reactions using a Qiagen BioRobot 9600 and a Beckman Biomek 2000. Purified plasmid DNA was sequenced from the 5′ end with the M13 reverse primer using an Applied Biosystems (ABI) PRISM big dye terminator kit (Perkin-Elmer) and an ABI Applied Biosystems 3700 DNA analyzer at the USDA Agricultural Research Service, Eastern Regional Research Center, Nucleic Acids Facility (ARS-ERRC-NAF) in Wyndmoor, PA.

2.4. Data handling
Raw sequence data were retrieved electronically from the USDA-ARS-ERRC-NAF using the file transfer protocol (ftp) for subsequent processing and analysis. The sequence data were imported into the computer software package Chromas 2.13 (Technelysium Pty, Helensvale, Australia) and manually trimmed of vector sequences. Ambiguous base calls were corrected by manually inspecting the sequence electropherograms, and the edited sequences were used in similarity searches.

Each cDNA sequence was queried against the current non-redundant (nr) protein database at the National Center for Biotechnology Information (NCBI, Bethesda, MD, USA) using the BLASTX algorithm and the NCBI EST database using the BLASTN algorithm (Altschul et al., 1997). In both cases, the default BLAST parameters were used. The redundancy of the 908 cDNA sequences was determined by comparing all sequences with one another using the program FastA (Wisconsin Package, Genetic Computer Group, Madison, WI, USA).

3. Results
3.1. EST analysis
The cDNA clones were checked by PCR, and 99% were found to contain inserts ranging in size from 350 to 3000 bp. A total of 908 clones were sequenced from the 5′ end of the cDNA inserts. The single pass sequencing runs generated an average of 650 nucleotides of readable sequences after manual editing.

All ESTs were assembled into a database and compared using the FastA program (Wisconsin Package, Genetic Computer Group, Madison, WI, USA) to identify redundant clones. A total of 488 unique ESTs were identified of which 381 appeared only once and 107 were represented by multiple clones at frequencies ranging from 2 to 142. The frequency of redundant ESTs is shown in Fig. 1. The sequences of the P. pachyrhizi EST clones were submitted to NCBI as dbEST IDs 28583523–28584357 and GenBank Accession Nos. DN739461–DN740295.



Full-size image (3K)


Fig. 1. The frequency of occurrence of EST clones derived from germinating P. pachyrhizi urediniospores. The number of EST clones is shown above each of the number of occurrences.


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The BLASTX algorithm (Altschul et al., 1997) was used to translate each edited EST into the six possible reading frames for comparison with data in the current nr protein database at the NCBI. A total of 431 ESTs displayed significant similarity to sequences in the NCBI database, while 477 ESTs did not exhibit significant similarity to the database entries. ESTs with similarity scores of Evalue < 10−5 were grouped according to their putative function (Table 1), according to the Expressed Gene Anatomy Database (EGAD) categories developed by The Institute for Genomic Research (TIGR, Rockville, MD, USA).


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Table 1.

EST clones displaying similarity (BLASTX, Evalue < 1E−05) to proteins in the non-redundant protein NCBI database, grouped into functional categories according to expressed gene anatomy database

Clone Accession No. Description Species Evalue No. of clones Organism
1. Cell division
1.1. DNA synthesis/replication
Pp0906 NP_595357 Checkpoint rad 3 Schizosaccharomyces pombe 9.00E−88 1 Yeast
Pp1817 T41457 DNA repair protein rad 18 S. pombe 4.00E−17 1 Yeast
Pp0244 NP_593482 Exonuclease II S. pombe 2.5E−44 1 Yeast
Pp2018 CAB91747 Related to syntaxin 12 Neurospora crassa 4.00E−14 1 Filamentous fungus


1.2. Apoptosis
Pp0322 AF316601 Metacaspase S. pombe 9.00E−50 1 Yeast


1.3. Cell cycle
Pp1417 AAA34617 G1 cyclin S. cerevisiae 8.00E−12 1 Yeast
Pp1017 AJ272133 Cyclin A. nidulans 7.00E−06 3 Filamentous fungus


1.4. Chromosome structure
Pp0437 P62792 Histone H4 Phanerochaete chrysosporium 3.70E−42 1 Filamentous fungus
Pp0729 P62792 Histone H4 P. chrysosporium 3.70E−45 1 Filamentous fungus
Pp1936 AAA35311 Histone H2A-α S. pombe 6.00E−33 3 Yeast
Pp1709 PN0142 Histone H2B N. crassa 5.00E−39 2 Filamentous fungus
Pp1628 A35072 Non-histone chromosomal protein NHP6A S. cerevisiae 4.00E−19 1 Yeast
Pp1812 S78076 Non-histone chromosomal protein NHP6B S. cerevisiae 2.00E−23 1 Yeast


2. Cell signaling/cell communication
2.1. Cell adhesion
Pp0813 Q28983 Zonadhesin Sus scrofa 1.00E−09 2 Mammal


2.3. Effectors/modulators
Pp0839 NP_593464 Calmodulin kinase I homolog S. pombe 5.00E−34 2 Yeast
Pp2023 AAA21544 Casein kinase-1 S. pombe 7.00E−48 1 Yeast
Pp0948 T18359 Nik-1 protein (histidine kinase) N. crassa 1.00E−39 1 Filamentous fungus
Pp0229 T45137 Phosphoprotein phosphatase catalytic chain 2B S. pombe 2.00E−06 1 Yeast
Pp1003 D84555 Probable protein kinase Arabidopsis thaliana 5.00E−27 1 Plant
Pp0424 T11657 RhoGDP dissociation inhibitor S. pombe 3.00E−30 1 Yeast
Pp1001 NP_596024 RhoGAP GTPase activating protein S. pombe 5.00E−12 1 Yeast
Pp1337 NP_594429 Probable phosphatidylinositol-4-phosphate kinase S. pombe 1.00E−51 1 Yeast


3. Cell structure and growth
3.1. Cytoskeletal
Pp1318 CAC17476 α-Tubulin Ustilago maydis 3.10E−87 1 Filamentous fungus
Pp1432 CAC83953 β-Tubulin Uromyces viciae-fabae 3.00E−98 1 Filamentous fungus
Pp1440 Q90631 Kinectin Gallus gallus 5.00E−07 1 Bird
Pp0920 AB018696 RanBPM Xenopus laevis 6.00E−05 1 Amphibian
Pp0414 U92845 Kinesin motor protein U. maydis 2.00E−37 1 Filamentous fungus


3.2. Growth and sporulation
Pp0432 XP_330886 Conidiation-specific protein 6 N. crassa 8.60E−16 7 Filamentous fungus
Pp0926 AAA33573 Conidiation protein N. crassa 1.00E−06 1 Filamentous fungus
Pp0122 CAD10036 Deacetylase Filobasidiella neoformans 5.00E−39 3 Filamentous fungus
Pp1605 A59290 Csm1 (class V chitin synthase with a myosin motor-like domain) Magnaporthe grisea 3.00E−07 1 Filamentous fungus
Pp1209 AAO49384 Class V chitin synthase Fusarium oxysporum 5.70E−88 1 Filamentous fungus


3.3. Others
Pp0941 EAA57250 Hypothetical protein MG08219.4 M. grisea 1.00E−08 2 Filamentous fungus
Pp0223 BAB13330 N-Acetylglucosaminidase Emericella nidulans 4.00E−25 1 Filamentous fungus
Pp1112 NP_014463 Sortilin homolog S. cerevisiae 1.00E−52 1 Yeast
Pp0811 NP_595238 Putative vacuolar protein; β-catenin family S. pombe 2.00E−27 1 Yeast


4. Cell/organism defense
4.1. Apoptosis
Pp1737 I49285 Defender against death protein 1 Mus musculus 1.00E−26 1 Mammal


4.2. Stress response
Pp0611 CAC20378 14-3-3-like protein Hypocrea jecorina 7.00E−92 1 Filamentous fungus
Pp0528 AAK15159 Heat-induced catalase Pleurotus sajor-caju 2.00E−82 4 Filamentous fungus
Pp1848 1908431A Heat-shock protein A. thaliana 1.00E−62 1 Plant
Pp1303 NP_596091 hsp16 (heat-shock protein 16) S. pombe 1.00E−19 1 Yeast
Pp1121 AAN75572 Copper chaperone TahA Trametes versicolor Trametes versicolor 3.00E−10 1 Filamentous fungus
Pp1929 CAD21425 Related to stress response protein rds1p N. crassa 7.00E−34 3 Filamentous fungus
Pp2004 BAA77283 DyP (peroxidase) Galactomyces geotrichum 3.00E−07 1 Filamentous fungus
Pp1616 T49477 Phenol hydroxylase related protein N. crassa 2.00E−16 1 Filamentous fungus


5. Gene/protein expression
5.1. RNA synthesis
5.1.1. RNA polymerases
Pp0946 P29035 Probable RNA-directed RNA polymerase (2Aprotein) (RNA replicase) Tomato aspermy virus 3.00E−08 1 Virus
Pp2029 NP_049325 Replicase Pea early browning virus 1.00E−07 1 Virus


5.1.2. RNA processing (e.g., spliceosomal, helicases)
Pp0519 O42861 Probable helicase S. pombe 4.00E−22 1 Yeast
Pp1810 S22646 Splicing factor U2AF homolog M. musculus 9.00E−42 1 Mammal
Pp1327 AAF37551 RNA-binding motif protein 8 Homo sapiens 4.00E−25 1 Mammal


5.1.3. Transcription factors
Pp1348 NP_010680 Transcription factor; Spt3p S. cerevisiae 1.00E−25 1 Yeast
Pp1504 AAA79367 TATA-binding protein Pneumocystis carinii 2.00E−95 1 Filamentous fungus
Pp0237 NP_011561 Transcription factor Tfc4p S. cerevisiae 3.40E−17 1 Yeast
Pp2041 Q00659 Sulfur metabolite repression control protein E. nidulans 1.00E−16 1 Filamentous fungus


5.2. Protein synthesis
5.2.1. Post-translational modification/targeting
Pp1724 S34655 Polyubiquitin 5 P. chrysosporium 9.00E−91 2 Filamentous fungus
Pp0936 T06053 Probable ubiquitin-dependent proteolytic protein A. thaliana 2.00E−35 1 Plant


5.2.2. Post-translational modification/trafficking
Pp1547 T39383 t-Complex protein 1, α-subunit S. pombe 1.00E−44 1 Yeast
Pp0719 NP_596649 Putative cytochrome C oxidase copper chaperone protein S. pombe 1.00E−12 2 Yeast
Pp0105 2113205A DNA J-like protein S. pombe 3.00E−19 1 Yeast


5.2.3. Protein turnover
Pp0126 CAA09863 Putative tripeptidyl peptidase I M. musculus 3.00E−06 1 Mammal
Pp1331 BAC56232 Tripeptidyl peptidase A A. oryzae 3.70E−39 2 Filamentous fungus
Pp1031 CAC39600 Prolidase A. nidulans 2.00E−38 1 Filamentous fungus


5.2.4. Ribosomal proteins
Pp1147 XP_326286 40S ribosomal protein S22 (S15A) (YS24) N. crassa 3.70E−74 1 Filamentous fungus
Pp1420 P05736 60S ribosomal protein L2 (YL6) (L5) (RP8) S. cerevisiae 2.00E−68 1 Yeast
Pp2011 T40111 14p-like ribosomal protein S. pombe 2.00E−12 1 Yeast


5.2.5. tRNA synthesis/metabolism
Pp1213 P46655 Cytosolic glutamyl-tRNA synthetase S. cerevisiae 6.00E−64 1 Yeast


5.2.6. Translation factors
Pp2028 S43861 Translation elongation factor eEF-1 α-chain Podospora anserina 1.00E−104 1 Filamentous fungus
Pp0835 NP_595367 eIF3 p48 subunit eIF3/signalosome component S. pombe 2.60E−43 1 Yeast
Pp0206 T48731 Probable translation initiation factor N. crassa 2.00E−100 1 Filamentous fungus
Pp0317 NP_015366 Tif5p S. pombe 1.00E−34 1 Yeast
Pp1107 P32186 Elongation factor eEF-1 α-chain Puccinia graminis 6.20E−82 1 Filamentous fungus
Pp2027 NP_502791 ADP-ribosylation factor-like protein (21.3 kDa) (4P563) Caenorhabditis elegans 7.40E−15 2 Nematode


6. Metabolism
6.1. Amino acid
Pp0134 M10139 3-Dehydroshikimate dehydratase N. crassa 9.00E−31 2 Filamentous fungus
Pp0425 AAN31488 DAHP synthase Phytophthora infestans 1.00E−74 1 Oomycete
Pp1503 NP_289154 DAHP synthetase, tyrosine repressible Escherichia coli 4.00E−40 1 Bacteria
Pp1336 NP_009808 DAHP synthase (is feedback-inhibited by tyrosine) S. cerevisiae 1.00E−37 1 Yeast
Pp1502 NP_012612 Tryptophan 2,3-dioxygenase S. cerevisiae 2.00E−12 1 Yeast
Pp0744 NP_592942 Phospho-2-dehydro-3-deoxyheptonate aldolase S. pombe 7.70E−29 1 Yeast
Pp1343 O94225 Homocitrate synthase, mitochondrial precursor Penicillium chrysogenum 3.00E−116 1 Filamentous fungus
Pp1727 T39244 Probable phospho-2-dehydro-3-deoxyheptonate aldolase S. pombe 3.00E−25 4 Yeast
Pp0806 AAO27751 Monooxygenase Fusarium sporotrichioides 1.00E−24 1 Filamentous fungus


6.2. Cofactor
Pp1931 CAB85691 Riboflavin aldehyde-forming enzyme Agaricus bisporus 2.00E−11 2 Filamentous fungus


6.3. Energy/TCA cycle
Pp1004 P34728 ADP-ribosylation factor F. neoformans 5.00E−108 1 Filamentous fungus
Pp2036 AAK18073 Aldehyde dehydrogenase ALDH15 E. nidulans 3.00E−38 1 Filamentous fungus
Pp1816 BAA09832 Isobutene-forming enzyme and benzoate 4-hydroxylase Rhodotorula minuta 8.00E−37 1 Yeast
Pp0727 CAA67613 Mitochondrial carrier protein S. cerevisiae 2.00E−13 1 Yeast
Pp1722 NP_035016 NADH dehydrogenase (ubiquinone) 1 α subcomplex 4 M. musculus 5.00E−08 2 Mammal
Pp1517 NP_594397 Putative isocitrate dehydrogenase (NADP+) S. pombe 2.00E−15 1 Yeast
Pp2040 T50403 Probable succinate dehydrogenase membrane anchor subunit precursor S. pombe 3.00E−24 1 Yeast
Pp1317 AAN74818 Fum15p Gibberella moniliformis 1.30E−16 1 Filamentous fungus
Pp2033 NP_172773 Putative cytochrome P450 monooxygenase A. thaliana 2.00E−16 1 Plant
Pp0837 NP_593578 Putative mitochondrial carrier S. pombe 2.90E−11 1 Yeast
Pp0127 CAC81058 Mitochondrial F1 ATP synthase β-subunit A. thaliana 1.30E−16 1 Plant
Pp0236 CAC81058 Mitochondrial F1 ATP synthase β-subunit A. thaliana 9.90E−16 1 Plant


6.4. Lipid
Pp1703 AAK26619 Acetyl-CoA acetyl transferase Laccaria bicolor 2.00E−39 1 Filamentous fungus
Pp0546 AAK26620 Acetyl-CoA acetyltransferase L. bicolor 7.00E−23 1 Filamentous fungus
Pp1511 CAB55552 Fox2 protein Glomus mosseae 2.00E−28 1 Filamentous fungus
Pp0337 XP_325309 Glycerol-3-phosphate dehydrogenase precursor related protein N. crassa 1,4E−92 1 Filamentous fungus
Pp0913 AAK63186 Probable acyl-CoA dehydrogenase G. intraradices 1.00E−37 2 Filamentous fungus
Pp1714 AAK63186 Probable acyl-CoA dehydrogenase G. intraradices 7.00E−33 3 Filamentous fungus
Pp0121 T40135 Probable involvement in ergosterol synthesis S. pombe 3.00E−33 1 Yeast
Pp1910 AAQ72469 SCS7p (oxidoreductase) Pichia pastoris 3.00E−09 3 Yeast
Pp0804 AAF27123 Putative glycerolkinase A. thaliana 4.00E−33 2 Plant


6.5. Sugar/glycolysis
Pp1827 AAA34858 6-Phosphofructo-2-kinase S. cerevisiae 4.00E−21 2 Yeast
Pp1140 Q24319 Dolichyl-diphosphooligosaccharide–protein glycosyltransferase Drosophila melanogaster 1.00E−12 1 Insect
Pp1235 AAB22823 Fructose-2,6-biphosphatase S. cerevisiae 2.70E−20 1 Yeast
Pp1735 CAC48025 Mutanase (α-1,3 glucanase) E. nidulans 3.00E−23 1 Filamentous fungus
Pp1048 CAC48025 Mutanase (α-1,3 glucanase) E. nidulans 2.00E−14 2 Filamentous fungus
Pp1033 XP_323561 Neutral trehalase N. crassa 1.00E−83 1 Filamentous fungus
Pp0133 CAA20128 Phosphomannomutase (predicted) S. pombe 6.00E−43 1 Yeast
Pp1501 ZP_00110197 COG0235: Ribulose-5-phosphate 4-epimerase, related epimerases and aldolases Nostoc punctiforme 1.00E−40 1 Bacteria
Pp1306 AAC17104 Endo-1,3(4)-β-glucanase Phaffia rhodozyma 1.00E−30 1 Filamentous fungus


6.6. Transport
Pp0917 CAD21006 ABC transporter (ATP-binding cassette transporter) F. neoformans 3.00E−68 1 Filamentous fungus
Pp1541 AAC08353 Calcium/proton exchanger N. crassa 6.00E−10 1 Filamentous fungus
Pp1713 T40789 Clathrin light chain S. pombe 1.00E−17 2 Yeast
Pp1340 CAA05841 Plasma membrane (H+) ATPase U. viciae-fabae 1.00E−105 1 Filamentous fungus
Pp1536 T38039 Probable nuclear transport factor 2 S. pombe 1.00E−26 1 Yeast
Pp1524 NP_594553 Putative membrane protein required for ER-Golgi transport S. pombe 4.00E−10 1 Yeast


6.7. Nucleotide
Pp2031 BAD00051 Ribonuclease T2 A. bisporus 2.00E−29 1 Filamentous fungus
Pp1845 XP_322797 Ribonucleoside-diphosphate reductase large chain N. crassa 1.00E−90 1 Filamentous fungus
Pp0547 AAN73281 UPL-1 Giardia intestinalis 4.10E−06 1 Protozoa


6.8. Protein modification
Pp0618 BAB56108 Carboxypeptidase Aspergillus nidulans 4.00E−21 1 Filamentous fungus
Pp1631 NP_253798 Lactoylglutathione lyase Pseudomonas aeruginosa 3.00E−13 1 Bacteria
Pp0643 AAA20876 Pepsinogen Aspergillus niger 1.00E−77 3 Filamentous fungus
Pp1341 CAC28786 Related to UDP-acetylglucosamine-peptide N-glucosaminyltransferase N. crassa 7.00E−44 1 Filamentous fungus
Pp1032 NP_035322 Proteasome activator subunit 3 M. musculus 1.00E−11 3 Mammal
Pp0747 AAG05190 ATP-dependent Clp protease proteolytic subunit P. aeruginosa 5.00E−22 2 Bacteria
Pp0915 AAB19394 Aspartate aminotransferase S. cerevisiae 2.00E−47 1 Yeast


6.9. Other metabolism
Pp1221 CAD79489 Glyoxal oxidase 2 Ustilago maydis 8.20E−28 1 Filamentous fungus
Pp0235 CAD79489 Glyoxal oxidase 2 Ustilago maydis 2.00E−34 1 Filamentous fungus
Pp1324 AAF02494 Alcohol oxidase 1 Pichia methanolica 1.00E−23 1 Yeast
Pp0218 T46646 Pyridoxine (Vitamin B6) biosynthesis protein pdx1 Cercospora nicotianae 5.00E−26 1 Filamentous fungus


7. Transposon
Pp0944 NP_921277 Transposase Tn10 Oryza sativa 7.00E−77 1 Plant


8. Unclassified
Pp0116 T30954 Hypothetical protein C44E4.6 C. elegans 6.00E−15 1 Nematode
Pp0115 AAA65309 pB602L African swine fever virus 1.00E−07 2 Virus
Pp1326 NP_780783 Hypothetical protein CTC00065 Clostridium tetani 5.10E−14 1 Bacteria
Pp0529 NP_754280 Transthyretin-like protein precursor E. coli 2.00E−17 1 Bacteria
Pp0819 NP_267806 Hypothetical protein L98109 Lactococcus lactis 7.00E−07 1 Bacteria
Pp0308 CAB85694 Hypothetical protein A. bisporus 7.60E−15 12 Filamentous fungus
Pp0103 AAK25792 Putative Egh16H1 precursor isoform A B. graminis f. sp. hordei 7.00E−12 4 Filamentous fungus
Pp0104 JC4750 gEgh 16 protein B. graminis f. sp. hordei 2.00E−32 36 Filamentous fungus
Pp0417 JC4750 gEgh 16 protein B. graminis f. sp. hordei 3.00E−34 142 Filamentous fungus
Pp0730 AAK25793 Putative Egh16H1 precursor isoform B B. graminis f. sp. hordei 5.00E−11 1 Filamentous fungus
Pp1039 JC4750 gEgh 16 protein B. graminis f. sp. hordei 3.00E−26 1 Filamentous fungus
Pp1043 JC4750 gEgh 16 protein B. graminis f. sp. hordei 3.00E−32 1 Filamentous fungus
Pp0326 CAD10781 Pentahydrophobin Claviceps purpurea 7.90E−06 1 Filamentous fungus
Pp0927 NP_758766 Hypothetical protein Erwinia amylovora 2.00E−24 1 Filamentous fungus
Pp1044 AAK52794 MAS3 protein M. grisea 9.00E−09 1 Filamentous fungus
Pp1429 AF264035 MAS1 protein M. grisea 4.00E−20 1 Filamentous fungus
Pp1610 AAK52794 MAS3 protein M. grisea 5.00E−05 1 Filamentous fungus
Pp0119 EAA55479 Hypothetical protein MG09286.4 M. grisea 1.10E−10 1 Filamentous fungus
Pp0222 EAA49745 Hypothetical protein MG09736.4 M. grisea 4.00E−06 1 Filamentous fungus
Pp0612 EAA48468 Hypothetical protein MG00126.4 M. grisea 3.30E−16 1 Filamentous fungus
Pp1045 EAA53245 Hypothetical protein MG07522.4 M. grisea 1.40E−31 1 Filamentous fungus
Pp2038 EAA51058 Hypothetical protein MG04818.4 M. grisea 2.00E−25 2 Filamentous fungus
Pp0225 XP_330149 Hypothetical protein N. crassa 3.40E−20 1 Filamentous fungus
Pp0534 XP_328580 Hypothetical protein N. crassa 1.30E−07 1 Filamentous fungus
Pp0704 XP_322643 Predicted protein N. crassa 1.10E−11 1 Filamentous fungus
Pp0809 XP_328793 Hypothetical protein N. crassa 5.20E−10 1 Filamentous fungus
Pp0829 XP_328520 Hypothetical protein N. crassa 9.60E−09 1 Filamentous fungus
Pp0925 XP_331047 Hypothetical protein N. crassa 1.00E−35 1 Filamentous fungus
Pp1006 XP_322643 Predicted protein N. crassa 1.00E−12 1 Filamentous fungus
Pp1013 XP_324202 Predicted protein N. crassa 2.60E−09 2 Filamentous fungus
Pp1027 XP_327468 Hypothetical protein N. crassa 2.00E−59 1 Filamentous fungus
Pp1325 XP_326398 Hypothetical protein N. crassa 2.70E−29 1 Filamentous fungus
Pp1411 XP_328221 Hypothetical protein N. crassa 1.70E−06 1 Filamentous fungus
Pp1615 XP_324693 Hypothetical protein N. crassa 3.00E−30 1 Filamentous fungus
Pp1835 XP_327028 Hypothetical protein N. crassa 5.00E−11 1 Filamentous fungus
Pp1925 CAD21504 conserved hypothetical protein N. crassa 5.00E−09 1 Filamentous fungus
Pp2010 XP_324370 Predicted protein N. crassa 1.00E−16 1 Filamentous fungus
Pp0334 NP_054890 Post-synaptic protein CRIPT; HSPC139 protein H. sapiens 8.80E−16 1 Mammal
Pp0748 BAA91611 Unnamed protein product H. sapiens 9.00E−09 1 Mammal
Pp0346 NP_001009405 PTPL1-associated RhoGAP 1 Rattus norvegicus 1.70E−06 1 Mammal
Pp1446 AAP78751 Ac1147 R. norvegicus 7.10E−34 1 Mammal
Pp1514 Q94480 VEG136 protein Dictyostelium discoideum 2.00E−31 1 Mycetozoan
Pp0516 T23541 Hypothetical protein K09C8.4 C. elegans 1.00E−05 1 Nematode
Pp0826 NP_917657 P0410E01.14 (hypothetical protein) O. sativa 7.00E−14 1 Plant
Pp1134 AAB49498 183 kDa protein Odontoglossum ringspot virus 3.00E−05 1 Virus
Pp1439 XP_332134 Hypothetical protein N. crassa 1.00E−39 1 Filamentous fungus
Pp0444 T37512 Hypothetical protein SPAC11D3.01c S. pombe 6.00E−14 1 Yeast
Pp0522 T38996 Hypothetical protein SPAC637.04 S. pombe 2.00E−07 1 Yeast
Pp0924 NP_596150 Hypothetical zinc finger protein S. pombe 2.50E−06 1 Yeast
Pp0931 NP_595085 Hypothetical glycine-rich protein S. pombe 6.80E−06 1 Yeast
Pp1019 NP_595449 Conserved hypothetical protein S. pombe 8.00E−21 1 Yeast
Pp1106 NP_595642 Hypothetical protein S. pombe 1.00E−07 2 Yeast
Pp1238 T41411 Hypothetical protein SPCC576.01c S. pombe 1.20E−06 1 Yeast
Pp1704 EAA50939 Hypothetical protein MG04698.4 M. grisea 1.00E−16 1 Filamentous fungus
Pp0406 EAA55557 Hypothetical protein MG01208.4 M. grisea 2.50E−44 1 Filamentous fungus
Pp1626 EAA49354 Hypothetical protein MG01012.4 M. grisea 3.00E−41 1 Filamentous fungus
Full-size table

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The best BLASTX score is reported for redundant clones (Table 1). A total of 189 putative genes were identified, of which 28.6% shared similarity to proteins from yeast, 50.8% to protein sequences from other fungi, while the rest exhibited similarity to proteins from a wide variety of organisms including bacteria, plants, mammals, insects, nematodes, and other invertebrates. The P. pachyrhizi cDNA library contained a broad range of genes, predominantly encoding putative proteins involved in primary metabolism, gene/protein expression, and cell structure (Table 1; Fig. 2). The ESTs with significant similarity to hypothetical proteins or proteins with unknown function were placed into the unclassified proteins category (Table 1; Fig. 2). The EST sequences with significant similarities (Evalue ≤ 10−15) to fungal and plant ESTs are shown in Table 2. Two different homologs of gEgh16, a protein expressed by Blumeria graminis f. sp. hordei during appressorium formation, were the most abundant ESTs in the P. pachyrhizi EST library (Table 1).


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Full-size image (42K)


Fig. 2. Classification of the 189 unique P. pachyrhizi ESTs from the germinating urediniospore library. The ESTs with significant matches (BLASTX Evalue < E−5) to the non-redundant database were classified into functional Expressed Gene Anatomy Database categories as described in Table 1. The percentage of ESTs in each of the eight categories is shown.


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Table 2.

EST clones displaying similarity (BLASTN, Evalue < 1E−15) to entries in the NCBI EST database

Clone Accession No. Description E value Organism
Pp0116 BI191959 l3h06fs.r1 Fusarium sporotrichioides Tri 10 overexpressed cDNA library F. sporotrichioides cDNA clone l3h06fs 5′, mRNA sequence 1.10E−24 Filamentous fungus
Pp0206 AU011975 AU011975 S. pombe late log phase cDNA S. pombe cDNA clone spc06169, mRNA sequence 4.90E−16 Yeast
Pp0406 BU060702 Fgr-C_1_H20_T3 Carbon-starved mycelia G. zeae cDNA, mRNA sequence 6.00E−35 Filamentous fungus
Pp0437 CB012207 Lb12C03 mycelium of L. bicolor grown for 3 weeks L. bicolor cDNA 5′, mRNA sequence 7.00E−32 Filamentous fungus
Pp1004 CF883485 Tric088xm20.b1 Trichoderma reesei mycelial culture, Version 6 October 2003 H. jecorina cDNA clone tric088xm20, mRNA sequence. 1.00E−51 Filamentous fungus
Pp1027 BU060160 Fgr-C_0_M05_T7 Carbon-starved mycelia G. zeae cDNA, mRNA sequence 0 Filamentous fungus
Pp1147 BG279541 b3h06np.r1 N. crassa sexual cDNA library, Uni-zap vector system N. crassa cDNA clone b3h06np 5′, mRNA sequence 1.00E−152 Filamentous fungus
Pp1318 CF190146 k7i06j2.r1 C. neoformans strain B3501 C. neoformans var. neoformans cDNA clone k7i06j2 5′, MRNA sequence 1.30E−21 Filamentous fungus
Pp1326 CF847171 psHB042xA02f USDA-IFAFS: expression of P. sojae genes during infection and propagation_sHB P. sojae cDNA clone sHB042A02 5′, mRNA sequence 3.00E−121 Oomycete
Pp1420 BQ110457 VD0108A10 VD01 Verticillium dahliae cDNA, mRNA sequence 1.00E−114 Filamentous fungus
Pp1432 AW324553 Basidiome and primordium cDNA libraries A. bisporus cDNA 5′ similar to β-tubulin, mRNA sequence 2.50E−46 Filamentous fungus
Pp1446 BU038322 LIT000228 root-induced cDNA library from L. bicolor L. bicolor cDNA, MRNA sequence 1.00E−124 Filamentous fungus
Pp1504 CF641217 D37_B10 Filamentous Forced Diploid Ustilago maydis cDNA 3′, mRNA sequence 5.00E−42 Filamentous fungus
Pp1547 CB898049 tric013xf01 Trichoderma reesei mycelial culture, Version 3 April H. jecorina cDNA clone tric013xf01, mRNA sequence 2.30E−51 Filamentous fungus
Pp1709 CF639134 D11_G01 Filamentous Forced Diploid U. maydis cDNA 3′, mRNA sequence 4.00E−27 Filamentous fungus
Pp1744 AI211414 p0b02a1.r1 A. nidulans 24 h asexual developmental and vegetative cDNA lambda zap library E. nidulans cDNA clone p0b02a1 5′, mRNA sequence 3.20E−28 Filamentous fungus
Pp1811 AW333990 S29A2 AGS-1 P. carinii cDNA 3′, mRNA sequence 4.80E−68 Filamentous fungus
Pp1843 CF644300 K19_B10 Filamentous Forced Diploid U. maydis cDNA 3′, mRNA sequence 3.00E−116 Filamentous fungus
Pp0127 BQ800593 EST 7628 Veraison Grape berries SuperScript Plasmid Library Vitis vinifera cDNA clone PT011A12 3′, mRNA sequence 6.90E−54 Plant
Pp0207 BQ464782 HU01I02T HU Hordeum vulgare subsp. vulgare cDNA clone HU01I02 5-PRIME, mRNA sequence 0 Plant
Pp0236 BQ907430 P006B08 Oryza sativa mature leaf library induced by M. grisea O. sativa cDNA clone P006B08, mRNA sequence 1.30E−53 Plant
Pp0404 CB643819 OSJNEb04L15.r OSJNEb O. sativa (japonica cultivar-group) cDNA clone OSJNEb04L15 3′, mRNA sequence 0 Plant
Pp0611 CA522045 KS11039D12 KS11 Capsicum annuum cDNA, mRNA sequence 1.10E−32 Plant
Pp0630 CD879728 AZO4.106C24F011012 AZO4 Triticum aestivum cDNA clone AZO4106C24, mRNA sequence 1.40E−15 Plant
Pp0713 CD879049 AZO4.104E06F010929 AZO4 T. aestivum cDNA clone AZO4104E06, mRNA sequence 2.10E−17 Plant
Pp0729 BI123652 I026P65P Populus leaf cDNA library Populus tremula x Populus tremuloides cDNA, mRNA sequence 7.50E−39 Plant
Pp0910 BQ908773 T015B01 Oryza sativa mature leaf library induced by M. grisea O. sativa cDNA clone T015B01, MRNA sequence 2.80E−45 Plant
Pp1219 CD878534 AZO4.102P17F011002 AZO4 T. aestivum cDNA clone AZO4102P17, mRNA sequence 3.00E−17 Plant
Pp1724 CA126740 SCVPLR1006B09.g LR1 Saccharum officinarum cDNA clone SCVPLR1006B09 5′, mRNA sequence 1.00E−80 Plant
Pp1729 CA253801 SCRLFL4105G02.g FL4 S. officinarum cDNA clone SCRLFL4105G02 5′, mRNA sequence 2.80E−91 Plant
Pp1848 CF811551 NA760 cDNA non-acclimated Bluecrop library Vaccinium corymbosum cDNA 5′, mRNA sequence 7.20E−24 Plant
Pp1924 BU672690 TR51 Leaf rust-infected wheat T. aestivum/P. triticina mixed EST library cDNA clone TR51, mRNA sequence 8.00E−35 Plant
Full-size table

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3.2. Gene families
Among the 908 ESTs analyzed, 18 potential gene families were identified by sequence similarity. Predicted function of these gene families could be ascribed to four groups, whereas 14 of the putative gene families did not show any significant similarity to entries in the databases. Eleven families contained two members, and four of them had three members. The remaining three putative gene families consisted of four, five, and nine members, respectively. The latter gene family contained four distinct homologs of gEgh16, one for the putative gEgh16 precursor isoform A, and one for the putative gEgh16 precursor isoform B from B. graminis f. sp. hordei. In addition, this group had one homolog for MAS1 and two homologs for MAS3 from Magnaporthe grisea. Three gene families showed similarity to homologs for DAHP synthase, conidiation protein 6 from Neurospora crassa, and the non-histone chromosomal proteins from Saccharomyces cerevisiae.

4. Discussion
Within the past decade, EST analyses have been conducted for several filamentous fungi and oomycetes such as: Agaricus bisporus (Ospina-Giraldo et al., 2000), Aspergillus flavus and Aspergillus parasiticus (OBrian et al., 2003), Aspergillus nidulans (Sims et al., 2004), B. graminis (Thomas et al., 2001), Cryphonectria parasitica (Dawe et al., 2003), Fusarium graminearum (Trail et al., 2003), Heterobasidium annosum (Abu et al., 2004 and Karlsson et al., 2003), M. grisea (Ebbole et al., 2004 and Kim et al., 2001), Mycosphaerella graminicola (Keon et al., 2000), N. crassa (Nelson et al., 1997 and Zhu et al., 2001), Pleurotus ostreatus (Lee et al., 2002), Schizophyllum commune (Guettler et al., 2003), Sclerotinia sclerotiorum (Li et al., 2004), Trichoderma reesi (Diener et al., 2004 and Steen et al., 2003), Ustilago maydis (Austin et al., 2004 and Nugent et al., 2004), Verticillium dahliae (Neuman and Dobinson, 2003), and Phytophthora infestans (Kamoun et al., 1999, Qutob et al., 2000 and Randall et al., 2005). In addition to the fungal ESTs available in the dbEST database at the NCBI, another EST database exists with sequences from 14 different phytopathogenic fungi and oomycetes (Soanes et al., 2002). In this study, we investigate the molecular genetics in the obligate soybean rust pathogen P. pachyrhizi. A total of 908 randomly chosen EST clones were sequenced and analyzed to identify which genes are expressed in germinating urediniospores. A relatively low level of redundancy was found among the P. pachyrhizi EST clones, similar to what has been observed in EST analyses from other filamentous fungi (Keon et al., 2000, Lee et al., 2002, Ospina-Giraldo et al., 2000, Thomas et al., 2001 and Trail et al., 2003). More than 52% of the EST clones showed no significant similarity to the entries in the public protein databases, which highlights the paucity in our knowledge of gene expression in filamentous fungi. The 432 P. pachyrhizi sequences that showed significant matches to sequences in the databases were classified into eight functional categories following the EGAD. Although proteins with unknown function or hypothetical proteins were the most prevalent, proteins involved in metabolism and in protein and gene expression were highly represented (Table 1).

Among the 908 cDNA clones, 488 unique ESTs were identified. These unigenes represent approximately 4–5% of the total 8000–12,000 expressed genes that are estimated in filamentous fungi (Kupfer et al., 1997 and Martinez et al., 2004). The remaining 420 sequences correspond to redundant cDNAs that form clusters ranging from 2 to 142 ESTs. Some P. pachyrhizi genes appear to be highly expressed during urediniospore germination, especially the two EST clones Pp0104 and Pp 0417, which share similarity to gEgh16 from B. graminis and appeared 36 and 142 times, respectively, among the clones sequenced in the library (Table 1). The function of gEgh16 is unknown (Justesen et al., 1996).

When the P. pachyrhizi ESTs were queried against the dbEST at NCBI, only 18 ESTs showed significant similarity (Evalue ≤ 10−15) to fungal or yeast entries, while 14 ESTs showed significant similarity to plant entries (Table 2). The low number of P. pachyrhizi ESTs with similarity to other fungi is due to the lack of gene expression studies that have been conducted in fungi. Two P. pachyrhizi EST clones, Pp1147 and Pp1420, have significant similarity to ribosomal proteins from N. crassa and S. cerevisiae, respectively, and these two ESTs also have high similarity to ESTs from other filamentous fungi. The EST Pp1027 shows significant similarity to a hypothetical protein from N. crassa and A. nidulans (Evalue < 10−30) and 93% identity (Evalue = 0) to an EST from Gibberella zeae, which suggests that it is a conserved gene.

Spore germination is an essential developmental stage in the life cycle of all filamentous fungi. It is a highly regulated process that responds to environmental stimuli via signaling cascades that are amenable to genetic and biochemical inquiry (Osherov and May, 2000 and Osherov and May, 2001). Three important steps can be distinguished during spore germination. First, the dormancy is broken in response to appropriate environmental conditions. Second, isotropic growth occurs, involving water uptake and the resumption of numerous metabolic activities. Third, polarized growth takes place and a germ tube is formed from which new mycelium originates (d’Enfert, 1997). Unlike most filamentous fungi in which low-molecular mass nutrients such as sugars, amino acids, and inorganic salts are required for conidial germination (Osherov and May, 2001), P. pachyrhizi urediniospores are capable of germinating on the surface of water. For some fungi, contact with a solid surface is required for conidial germination (Thomas et al., 2001). It is interesting to note that two ESTs identified in this analysis, Pp1527 and Pp0839, share very high similarity to Ca2+/calmodulin-dependent protein kinase and calmodulin kinase I, respectively. The expression of calmodulin is induced by contact with a hard surface in both Colletotrichum gloeosporioides and M. grisea (Kim et al., 1998, Kim et al., 2000 and Liu and Kolattukudy, 1999). The expression of these calmodulin kinase homologs suggests that a similar calcium-signaling pathway may regulate urediniospore germination in P. pachyrhizi.

In fungi, the cell wall undergoes significant modification during spore germination. Three P. pachyrhizi ESTs showed similarity to enzymes involved in the dissolution and formation of the cell wall. EST clones Pp0122, Pp0922, and Pp1605 share similarity to chitin deacetylase, acetylxylan esterase, and chitin synthase (csm1), respectively, from M. grisea. The csm1 gene product contains a myosin motor-like domain (Park et al., 1999). In A. nidulans, its homolog CsmA has an important role in polarized cell wall synthesis and maintenance of cell wall integrity, and the myosin motor-like domain has been shown to be required for these functions (Horiuchi et al., 1999).

DNA and RNA synthesis do not appear to be necessary during the early stages of spore germination, whereas protein synthesis is required (Osherov and May, 2001). They suggest that dormant conidia contain a pre-existing pool of mRNA and ribosomes that are primed for rapid activation and translation in the presence of nutrients. Our results indicate that increased protein synthesis activity occurs during spore germination in P. pachyrhizi. Three different homologs for translation initiation factors and two homologs for elongation factors were identified, as well as several genes involved in post-translational modification, protein modification, and metabolism of amino acids (Table 1). In their model of spore germination, DNA and RNA synthesis are required in the later stages of spore germination for hyphal development (Osherov and May, 2001). As the germination of the P. pachyrhizi urediniospores was asynchronous in our experiment, sequences similar to genes involved in both the early and later spore germination processes were found among the P. pachyrhizi ESTs.

Several putative gene families were identified among the ESTs analyzed in this study. The main group consists of nine different ESTs: four ESTs, Pp0104, Pp0417, Pp1033, and Pp1039, are homologs of gEgh16; Pp0103 is a homolog of the putative gEgh16 precursor isoform A; Pp0730 is a homolog of the putative gEgh16 precursor isoform B from B. graminis f. sp. hordei; Pp1429 is a homolog for MAS1; and two ESTs, Pp1044 and Pp1610, are homologs for MAS3 from M. grisea. The EST clones Pp0104 and Pp0417, which are similar to gEgh16, are highly redundant in the P. pachyrhizi library suggesting that they are highly expressed during urediniospore germination. The function of the gEgh16 protein has not been determined in B. graminis f. sp. hordei, but it is highly expressed during germ tube formation and hyphal growth. There is evidence that gEgh16 is a member of a gene family in B. graminis f. sp. hordei (Justesen et al., 1996). Although the function of MAS1 and MAS3 are unknown in M. grisea, the genes encoding for these proteins are expressed during appressorium formation (Choi and Dean, 2000).

Another potential gene family in P. pachyrhizi is comprised of five ESTs similar to DAHP synthase (Pp0323, Pp0425, Pp0744, Pp1336, and Pp1503). DAHP synthase catalyzes the first step in the shikimate pathway that leads to the biosynthesis of aromatic amino acids. In N. crassa and Escherichia coli, three isozymes of DAHP synthase have been characterized and each one is regulated by the three aromatic amino acids. In A. nidulans and S. cerevisiae, two DAHP synthase encoding genes have been described, and the enzymes are differentially regulated by tyrosine and phenylalanine (Hartmann et al., 2001 and Künzler et al., 1992). In addition to the DAHP synthase homologs, a P. pachyrhizi EST clone (Pp0134) was found to share similarity to dehydroshikimate dehydrogenase, which is also part of the shikimate pathway. It has been shown that quinate and shikimate, two metabolic intermediates of the shikimate pathway, can be metabolized by a variety of fungi as alternative carbon sources (Keller and Hohn, 1997).

Two ESTs (Pp1628 and Pp1812) share similarity to the non-histone chromosomal proteins NHP6A and NHP6B, respectively, from S. cerevisiae. NHP6A and NHP6B are high mobility group proteins, which are members of a family of heterogeneous chromatin-associated DNA-binding proteins in eukaryotic cells (Masse et al., 2002 and Yen et al., 1998). NHP6A is a member of the subclass HMG1/2 proteins that contain the HMG DNA-binding domain and are present at approximately 1 molecule per 2–3 nucleosomes (Kuehl et al., 1984). These proteins have been implicated in chromatin remodeling, DNA replication, transcription, and recombination (Giavara et al., 2005), and it will be interesting to determine their role in P. pachyrhizi ediniopsore germination.

In this study, approximately 39% of the unique ESTs appeared to be related to previously characterized genes. This highlights the scarcity of genomic information available from pathogenic fungi. The EST projects have been shown to be a valid and fast way to gain information on components that regulate vital processes in pathogenic fungi and the interaction with their hosts. In 2002, a Phakopsora genome sequencing project, funded by the U.S. Department of Agriculture-Agricultural Research Service and the Department of Energy (DOE), was initiated at the DOE-Joint Genome Institute to generate draft quality sequence of P. pachyrhizi and P. meibomiae. The ESTs identified in this study, along with the analyses of the cDNA libraries from P. pachyrhizi infected soybeans, will aid in the annotation of genes from the Phakopsora genome project. These data will facilitate our understanding of the biology and the evolution of obligate fungal pathogens and will also advance our efforts to develop effective means for soybean rust control.

Acknowledgments
We thank Connie Briggs at the USDA-ARS-ERRC-NAF for sequencing the EST clones. We are grateful to Drs. Seogchan Kang, William Schneider, Morris Bonde, Paul Tooley, and Douglas Luster for critical review of the manuscript. This work was supported by USDA-ARS CRIS Projects 1920-22000-23-00D and 1920-22000-27-00D. M.L.P.B. was supported by an USDA-ARS Administrator’s Postdoctoral Fellowship.

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Corresponding author. Fax: +1 301 619 2880.
1 Present address: DOE-Joint Genome Institute, Lawrence Berkeley National Laboratory, 2800 Mitchell Drive, Walnut Creek, CA 94598, USA.

In vitro differentiation of haustorial mother cells of the wheat stem rust fungus, Puccinia graminis f. sp. tritici, triggered by the synergistic acti

Copyright © 2003 Elsevier Science (USA). All rights reserved.

In vitro differentiation of haustorial mother cells of the wheat stem rust fungus, Puccinia graminis f. sp. tritici, triggered by the synergistic action of chemical and physical signals

Nicola Wiethölter, Susanne Horn, Katrin Reisige, Ursula Beike and Bruno M. Moerschbacher,

Department of Plant Biochemistry and Biotechnology, Westfälische Wilhelms-Universität Münster, Hindenburgplatz 55, 48143, Münster, Germany


Received 17 July 2002; accepted 19 September 2002. ; Available online 11 March 2003.

Abstract
Biotrophic plant pathogenic fungi often develop a sophisticated series of infection structures for non-destructive host tissue penetration. In vitro, early infection structures of rust fungi—germ tube, appressorium, substomatal vesicle, infection hyphae—can easily be induced, but in vitro differentiation rates of late infection structures—haustorial mother cells (hmc), haustoria—are low at best. Under appropriate conditions (humid atmosphere), a combination of physical (mild heat shock) and chemical signals (trans-2-hexen-1-ol) induced the in vitro differentiation of hmc in the wheat stem rust fungus, Puccinia graminis f. sp. tritici. Around two thirds of the in vitro differentiated germlings developed up to three hmc which were cytologically identical to hmc formed in planta. Efficient in vitro differentiation of hmc will allow us to analyse in molecular detail the processes involved in the induction and differentiation of this critically important developmental stage of the economically important plant pathogenic rust fungi.

Author Keywords: Puccinia graminis f. sp. tritici; Wheat stem rust fungus; Haustorial mother cells; Biotrophic fungi; Leaf alcohol; Trans-2-hexen-1-ol; In vitro differentiation; Infection structures

Article Outline
1. Introduction
2. Materials and methods
2.1. Origin of fungal material
2.2. Induction of infection structures
2.3. Staining and microscopy
3. Results
4. Discussion
Acknowledgements
References
1. Introduction
Rust fungi are obligately biotrophic plant pathogens that are highly specialised for growth and development on and in their respective host plant tissues. Their biotrophic nature requires careful host tissue penetration and colonisation in order to prevent premature recognition by the host cells and the ensuing triggering of induced resistance mechanisms such as hypersensitive host cell death. Uredospore germlings of most rust fungi enter their host tissues via the natural openings of the plant stomates, thus preventing host tissue wounding (Mendgen et al., 1996).

Uredospores germinating on a plant surface produce a germ tube which tightly adheres to the cuticle, apparently a prerequisite for oriented germ tube growth towards the stomates (Dickinson, 1969; Maheshwari and Hildebrandt, 1967; Wynn, 1976). Upon reaching a stoma, tip growth of the germ tube is arrested and an appressorium is formed on top of the stoma ( Allen, 1923 and Allen, 1926). The appressorium containing the two fungal nuclei which emerged from the spore is separated from the germ tube by a septum, and a first round of synchronised mitoses occurs in the appressorium. From the appressorium, a narrow penetration peg grows through the stomatal opening and develops into a vesicle in the substomatal cavity.

The cell wall of penetration peg and vesicle differs from the cell wall of germ tube and appressorium (Harder et al., 1986; Littlefield and Heath, 1979). The cytoplasmic content of the appressorium is transferred into the substomatal vesicle where a second round of synchronised mitoses occurs, leading to a total of eight nuclei. Infection hyphae emanating from the vesicle start growing in the intercellular spaces of the host tissue, and pairs of fungal nuclei migrate into these hyphae ( Allen, 1923; Heath and Heath, 1976; Staples et al., 1975). When the tip of an infection hypha reaches the cell wall of an epidermal host cell, tip growth is arrested and a haustorial mother cell is formed. The haustorial mother cell usually containing 2–4 fungal nuclei is separated from the infection hypha by a septum ( Heath and Heath, 1975; Heath et al., 1996). From the haustorial mother cell, a narrow haustorial neck grows through the host plant’s cell wall and develops into a haustorium in the periplasmic space of the host cell. A second change in cell wall characteristics occurs upon differentiation of haustorial mother cells ( Chong and Harder, 1982; Chong et al., 1985 and Chong et al., 1986). Infection hyphae branch just proximal to the haustorial mother cell septum, and the branches develop new haustorial mother cells at their tips upon host mesophyll cell encounters. The infection hyphae of the rust mycelium remain extracellular throughout host tissue colonisation and eventual fungal sporulation. Only the terminally differentiated haustoria reach into the pericellular space of the host cells.

Quite apparently, the sophisticated series of infection structures—germ tube, appressorium, substomatal vesicle, infection hyphae, haustorial mother cells, haustoria—are a prerequisite for the obligately biotrophic rust fungi to penetrate, colonise, and feed from the host plant tissue with a minimum of host cell perturbation. The unique process of infection structure differentiation can be expected to be an absolute conditio sine qua non for the successful rust development and as such, to provide promising targets for the development of novel fungicides.

Rust uredospores readily germinate in the presence of liquid water, producing a germ tube tightly adhering to any hydrophobic surface. Also, the differentiation of infection structures can be induced in vitro when appropriate chemical or physical signals, such as an appropriately structured hydrophobic surface, are provided (Dickinson, 1969; Heath and Perumalla, 1988; Hoch and Staples, 1987; Mendgen et al., 1996; Read et al., 1992). Unlike the in vivo situation, where a series of independent signals can be assumed to regulate the development of infection structures on and in the host leaf ( Heath, 1997; Mendgen, 1982), a single trigger may induce the consecutive differentiation of appressorium, substomatal vesicle, infection hyphae and, albeit often at low frequency only, haustorial mother cells in vitro ( Deising et al., 1991; Heath and Perumalla, 1988). The cowpea rust fungus, Uromyces vignae, has even been reported to build some rare haustoria in vitro upon appropriate chemical triggers (Heath, 1990).

Topographical signals are less effective in triggering the differentiation of infection structures of rust fungi specialised on monocotyledonous host plants, such as the wheat stem rust fungus (Allen et al., 1991; Read et al., 1997). Instead, a number of other physical and chemical signals such as a mild heat shock ( Dunkle and Allen, 1971; Emge, 1958; Maheshwari et al., 1967), organic compounds ( Macko et al., 1978), host epicuticular waxes ( Daniels, 1996; Grambow, 1977; Grambow and Grambow, 1978; Grambow and Riedel, 1977), or leaf volatiles ( Daniels, 1996; Grambow, 1977; Grambow and Riedel, 1977) have been reported to trigger the sequential in vitro development of appressoria, substomatal vesicles, and infection hyphae of the wheat stem rust fungus. Recently, a combination of surface ridges of appropriate sizes and spacings with trans-2-hexen-1-ol was shown to act synergistically in inducing these infection structures (Collins et al., 2001). However, the wheat stem rust fungus does not easily differentiate haustorial mother cells in vitro. We here report on the reproducible, high frequency differentiation of haustorial mother cells of the wheat stem rust fungus in vitro, by the synergistic action of a physical and a chemical signal.

2. Materials and methods
2.1. Origin of fungal material
Uredospores of the wheat stem rust fungus Puccinia graminis f. sp. tritici Eriks. & Henn., race 32, were collected from fully susceptible wheat plants Triticum compactum L. cv. Little Club. Uredospores were frozen in liquid nitrogen and stored at −70 °C. Spores were reactivated for 3 min at 43 °C.

2.2. Induction of infection structures
For the induction of infection structures of the wheat stem rust fungus (appressoria, substomatal vesicles, infection hyphae and haustorial mother cells), 0.5 mg of uredospores were distributed with a brush in the lid of a Petri dish (Ø 6 cm, Greiner, Frickenhausen). Five milliliters of trans-2-hexen-1-ol (Aldrich, Taufkirchen) (0.5 mM in doubly distilled water) was filtered (pore Ø 0.22 μm, cellulose ester membrane, Fisherbrand, Schwerte; 30 ml Luer Lock syringe, Plastipak, Becton–Dickinson, Heidelberg) into the bottom of the Petri dishes. The spore-containing lids were replaced on the Petri dish bottoms and sealed with parafilm (American National Can, Baltimore, Maryland) to prevent evaporation. The Petri dishes were placed in a temperature controlled chamber (Heraeus, Hanau) at 23 °C where the uredospores started to germinate. When the average length of the germ tubes had reached the size of the spore diameter (approx. 50–60 min) the germlings were given a mild heat shock at 30 °C for 2 h (Emge, 1958; Maheshwari et al., 1967), followed by further incubation at 23 °C. All incubation steps were carried out in darkness.

2.3. Staining and microscopy
Fungal infection structures were analyzed using an Olympus BX 40 and a Leica DM RBE microscope both equipped with epifluorescence, and an Olympus IX 50 inverted microscope with phase contrast optics. The filter module U-MNV (excitation filter BP 400–410, dichroic mirror DM 455, barrier filter BA 455 nm) was used for fluorescence microscopy using the Olympus BX 40. Fluorescence microscopy using the Leica DM RBE was carried out with the excitation filter BP 450–490 nm, the dichroic mirror RKP 510 nm and the barrier filter LP 515 nm. Microphotographs were taken on Agfa CT Precisa slide film. Cell walls of the fungus were stained using Calcofluor White (Sigma, Taufkirchen) (Maeda and Ishida, 1967). For this procedure, a stock solution (2.5 mg ml−1 in water) was diluted 1:20 before use. The fungus was incubated with this staining solution for 30 s and washed 10 times with water. Fungal nuclei could be stained afterwards using DAPI (4′,6-Diamin-2′-phenylindol-dihydrochloride; Sigma, Taufkirchen) (0.01 μgml−1 DAPI in water, 10 s) then washed 10 times with water (Butt et al., 1989).

3. Results
Germlings of the wheat stem rust fungus, Puccinia graminis f. sp. tritici, were treated with trans-2-hexen-1-ol, either as a volatile when the germlings were grown in a humid atmosphere, or in dissolved form when the germlings were grown immersed in a liquid medium. In both cases, trans-2-hexen-1-ol induced the differentiation of an appressorium, a substomatal vesicle, and one or two infection hyphae within 24 h. Fig. 1 shows that in a humid atmosphere, induction by trans-2-hexen-1-ol also led to the formation of haustorial mother cells in about 10% of the germlings within about three days. In contrast, no haustorial mother cells were formed when the germlings were grown submerged in a trans-2-hexen-1-ol solution (data not shown).



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Fig. 1. Time course of in vitro differentiation of infection structures by Puccinia graminis f. sp. tritici uredospores induced by the application of the volatile leaf alcohol trans-2-hexen-1-ol (0.5 mM). Around 20% of the sporelings produced infection structures within 24 h, and about half of these formed haustorial mother cells. Symbols represent appressoria (•), substomatal vesicles (▪), infection hyphae (), and haustorial mother cells (). Data given are means ± SD of three independent experiments, with a minimum of 80 sporelings counted per time point in each experiment.


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In vitro differentiated haustorial mother cells of the wheat stem rust fungus appeared as small (15 μm in length), granular structures at the end of the infection hypha, from which they were clearly separated by a septum (Fig. 2E). Their cell walls appeared thicker than those of the infection hypha, and the fluorescent brightener Calcofluor White bound more strongly, leading to bright fluorescence under UV-light. Invariably, two nuclei were observed in each haustorial mother cell (Fig. 2E). These typical characteristics easily allowed their unequivocal identification, e.g. as compared to stress-induced terminal swellings of infection hyphae ( Fig. 2D).


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Fig. 2. In vitro differentiation of infection structures including haustorial mother cells by the wheat stem rust fungus Puccinia graminis f. sp. tritici after application of a mild heat shock (2 h, 30 °C) and trans-2-hexen-1-ol (0.5 mM) in a humid atmosphere. (A) and (B) represent fully differentiated germlings using bright field and phase contrast optics, respectively (gt, germ tube; ap, appressorium; sv, substomatal vesicle; ih, infection hypha; hmc, haustorial mother cell) (bar: 15 μm). When fungal structures were stained with DAPI (E, F) and Calcofluor (C–F), nuclei, septa, and cell wall alterations were visible under fluorescent light, and haustorial mother cells were clearly distinguishable from terminal swellings of infection hyphae (D). When differentiation was induced by trans-2-hexen-1-ol alone, appressoria and subsequent infection structures formed at the end of a long and often branched germ tube (C). When trans-2-hexen-1-ol was combined with a heat shock, infection structures were formed much faster at the end of a short germ tube only (A, B, D–F). When germlings were immersed in liquid culture medium after infection structures had formed (A, F), an occasional outgrowth of haustorial mother cells was observed (F).


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One of the most effective ways of inducing infection structure differentiation in the wheat stem rust fungus is a mild heat shock given shortly after germination. This treatment is active with germlings growing submerged in liquid or in a humid atmosphere, but no haustorial mother cells formed even after prolonged times of incubation (data not shown). However, we found that a combination of the chemical signal trans-2-hexen-1-ol (0.5 mM) and the physical signal of a mild heat shock (30 °C, 2 h) effectively triggered the induction of haustorial mother cells at high frequency, when the germlings were grown in a humid atmosphere. Fig. 3 gives the time course of appearance of the different infection structures induced.


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Fig. 3. Time course of in vitro differentiation of infection structures by Puccinia graminis f. sp. tritici uredospores induced by a mild heat shock (2 h, 30 °C, grey bar) and the application of the volatile leaf alcohol trans-2-hexen-1-ol (0.5 mM). Appressoria (•) started to appear during the heat shock. Substomatal vesicles (▪) and infection hyphae () were first observed 2 and 8 h after the end of the heat shock, respectively. First haustorial mother cells () were formed between 21 and 45 h after the heat shock. Data given are means ± SD of three independent experiments, with a minimum of 80 germlings counted per time point in each experiment.


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Tip growth of the germ tube was stopped by the heat shock so that the germlings had a compact shape (Figs. 2A, B, D–F). Under these conditions, about 60% of the germinated uredospores had differentiated appressoria, substomatal vesicles, and infection hyphae 24 h after germination. Haustorial mother cells emerged 24–48 h after germination, and a maximum hmc-rate of 40% was reached 72 h after spore germination (Fig. 3), after which time no further increase was observed.

Often, haustorial mother cells developed at the tip of both infection hyphae of a rust germling. Some infection hyphae branched just proximal to the haustorial mother cell septum, and a third haustorial mother cell might then form at the tip of the branch (Figs. 2A and B). Fungal growth and development then stopped; we never observed more than three haustorial mother cells on a single germling. In order to test whether failure of further growth could be attributed to exhaustion of the nutrient pool of the uredospores, we dissolved the trans-2-hexen-1-ol in axenic culture medium (Fasters et al., 1993) instead of water, and inverted the Petri dishes at different times after germination, bringing the germlings into contact with the nutrient solution. As a control, Petri dishes with trans-2-hexen-1-ol in water were inverted at the same times. No substantial further growth was observed even in the axenic culture medium. Immersion of the germlings at earlier times after germination (7, 12, or 48 h) prevented or stopped further development of haustorial mother cells.

In axenic medium, we occasionally observed outgrowth of a fungal hypha from the differentiated haustorial mother cell (Fig. 2F). We never observed the differentiation of haustoria from the haustorial mother cells induced in vitro. Failure to resume sustained growth may be due to the trans-2-hexen-1-ol present in the medium, as growth did resume in pure culture medium. However, without the addition of trans-2-hexen-1-ol, no haustorial mother cells were formed.

4. Discussion
In vitro differentiation of haustorial mother cells has been reported previously for several rust species, including the bean rust fungus, Uromyces appendiculatus (Maheshwari et al., 1967), the cowpea rust fungus, U. vignae (Heath and Perumalla, 1988; Stark-Urnau and Mendgen, 1993), and the broad bean rust fungus, Uromyces viciae-fabae (Deising et al., 1991). In these cases, a single inductive signal (oil-collodion membranes or scratched polyethylene membranes) led to the formation of haustorial mother cells in 10–25% of the germlings.

In vitro induction of haustorial mother cell differentiation of the wheat stem rust fungus appears to be strictly dependent on (i) the development in a humid atmosphere—no haustorial mother cells were observed when the fungus grew submerged in water or liquid axenic medium—and (ii) on the presence of a suitable topographical signal (Read et al., 1997) or of a volatile chemical inductor, such as the leaf alcohol trans-2-hexen-1-ol. Under such conditions of a single inductive signal, haustorial mother cell differentiation was sporadic and occurred at low frequency only. A reproducible high frequency induction of haustorial mother cells was reached by the combination of a mild heat shock—which given alone induced appressoria, substomatal vesicles, and infection hyphae, but never haustorial mother cells—with the volatile inductor trans-2-hexen-1-ol. Under these conditions, about two thirds of the germlings developed infection structures, and about two thirds of these differentiated up to three haustorial mother cells. The high frequency of induction combined with the formation of multiple haustorial mother cells per germling allows an unprecedented high number of haustorial mother cells to be differentiated in vitro.

In planta, haustorial mother cells of the wheat stem rust fungus are oval, long and slender (Allen, 1923; Niks, 1986). They are separated from the infection hypha by a septum, they are characterised by an optically dense appearance, and they contain two nuclei ( Allen, 1923). The cell wall of in planta grown haustorial mother cells of the wheat stem rust fungus is more complex than that of the infection hyphae, containing additional layers which are also present in the septum, leading to increased stainability with Calcofluor ( Chong et al., 1985). Haustorial mother cells of the wheat stem rust fungus formed in vitro exhibited all of these typical characteristics.

Although the morphogenetically active signals inducing infection structure differentiation in different rust fungi differ in detail, the genetic program induced appears to share similarities. In vitro, a single trigger usually induces the sequential differentiation of appressoria, substomatal vesicles, and infection hyphae at high frequency, while the differentiation rarely extends to the building of haustorial mother cells. In planta, however, those rust fungi that do not penetrate closed stomata in the dark, e.g. Puccinia graminis f. sp. tritici, differentiate an appressorium upon encounter of a closed stoma, but the development of further infection structures is arrested until the stoma opens (Wynn and Staples, 1981). Moreover, the subsequent development of the infection hyphae appears to respond to additional host factors, as e.g. the intercellular infection hyphae of the oat crown rust fungus have been shown to grow directly into the mesophyll of an infected oat leaf, while they grow parallel to the leaf surface in an infected wheat leaf ( Moerschbacher et al., 1990). Clearly, the morphogenetic program of sequential infection structure differentiation is naturally triggered and regulated by a number of host derived signals.

The concept of multiple recognition extends to the differentiation of haustorial mother cells and, most likely at least, also of haustoria (Heath, 1997). Surface signals from the plant cell walls have been implicated in these differentiation steps ( Fasters et al., 1993; Heath, 1990; Mendgen, 1978 and Mendgen, 1982). While the exact nature of these signals acting in planta is not yet known, the wheat stem rust fungus appears to be a suitable object to study in vitro the combined action of different signals in the triggering of infection structure differentiation. It has been shown that the volatile leaf alcohol trans-2-hexen-1-ol acts synergistically with an inductive factor from epicuticular waxes of the host leaf (Grambow, 1977; Grambow and Grambow, 1978; Grambow and Riedel, 1977), and with topographical signals mimicking a gramineaceous stoma ( Collins et al., 2001). We are currently studying the combined action of all three of these factors on rust differentiation in vitro. We have shown in this study that a mild heat shock combined with the volatile inductor leads to the differentiation of haustorial mother cells. It will be difficult to identify the presumably chemical in planta equivalent of the thermal signal as haustorial mother cells did not develop in submerged culture so that application of putative signal molecules is difficult. We are currently developing methods to reproducibly deposit known amounts of putative non-volatile chemical inductors on appropriate surfaces so that we can study their morphogenetic effects on rust development in a humid atmosphere.

Acknowledgements
We gratefully acknowledge many fruitful discussions with Dr. Nick Read, University of Edinburgh, where some preliminary experiments to this study were carried out.

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Corresponding author. Fax. +49-251-832-83-71

Cloning and Characterization of a cDNA of cro rI from the White Pine Blister Rust Fungus Cronartium ribicola*1

Cloning and Characterization of a cDNA of cro rI from the White Pine Blister Rust Fungus Cronartium ribicola*1

Xueshu Yu, Abul K. M. Ekramoddoullah2, Doug W. Taylor and Nina Piggott

Pacific Forestry Centre, Canadian Forest Service, Natural Resources Canada, 506 West Burnside Road, Victoria, British Columbia, V8Z 1M5, Canada


Accepted 8 October 2001. ; Available online 4 March 2002.

Abstract
White pine blister rust (WPBR) is caused by the fungus Cronartium ribicola which has five spore stages on two unrelated hosts, the five-needle pines and Ribes spp. Recently, during the molecular analysis of the proteins and genes involved in host–pathogen interaction, the WPBR fungal protein Cro rI was identified in infected white pine tissues. To further characterize Cro rI, an expression cDNA library from poly(A)+ mRNA of C. ribicola axenic mycelial culture was constructed and immunoscreened and the cDNA was cloned. Sequence analysis indicated an open reading frame of 462 bases, which encodes a protein of 153 amino acid residues with a molecular mass of 16.7 kDa and a predicted isoelectric point (pI) of 8.93. Based on the N-terminal amino acid sequences of Cro rI, the secreted portion of Cro rI protein should be 136 amino acids long with several putative posttranslational modification sites and a molecular mass of 14.8 kDa. The predicted pI for the secreted portion was 9.34. The predicted N-terminal signal peptide was 17 amino acids long. The N-terminal 42-amino acid sequence of the predicted mature protein (secreted portion) was identical to the amino terminal sequence of Cro rI that was previously determined. Southern blot hybridizations indicated that the C. ribicola genome contained at least two copies of the cro rI gene. Isolation of the genomic PCR fragment, which was approximately 400 bp longer than the cDNA, and subsequent cloning and sequencing analyses confirmed that there were three introns within the coding regions. Western immunoblot analyses revealed that Cro rI protein accumulated in large amounts only in the infected white pine tissues while no trace was detectable in the alternate Ribes stage or the five different spores, suggesting a critical role of Cro rI in the haploid stage of the fungus (in pine). The translocation of Cro rI was only found to occur in cankered trees, and not in the young infected seedlings. The implications of Cro rI in pathogenesis are discussed.

Author Keywords: amplification of genomic DNA by PCR; gene structure; secretory signal; translocation; spore; susceptible tree

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*1 The nucleotide sequence data reported in this paper have been submitted to the GenBank Nucleotide Sequence Database under the Accession Nos. AF232039, AF232040, AF232041, AF232042, and AF232043.

2 To whom correspondence should be addressed. Fax: (250) 363-0775. E-mail: aekramoddoul@pfc.forestry.ca.

Plasma Membrane H+-ATPase Activity in Spores, Germ Tubes, and Haustoria of the Rust FungusUromyces viciae-fabae

Regular Article
Plasma Membrane H+-ATPase Activity in Spores, Germ Tubes, and Haustoria of the Rust FungusUromyces viciae-fabae*1

Christine Struck, Matthias Hahn and Kurt Mendgen 1

Lehrstuhl für Phytopathologie, Fakultät für Biologie, Universität Konstanz, 78 434, Konstanz, Germany


Accepted 9 December 1995. ; Available online 19 April 2002.

Abstract
Using plasma membrane-enriched vesicles, the properties of the H+-ATPase (EC 3.6.1.35) from the rust fungusUromyces viciae-fabaewere studied. The enzyme is strictly Mg2+-dependent and is inhibited by vanadate. The pH-optimum is at 6.7. By Western blot analysis using a monoclonal antibody against corn plasma membrane H+-ATPase a polypeptide of approximately 104 kDa could be detected. The vanadate-sensitive H+-ATPase activity of microsomal vesicles obtained from different stages of rust development was determined. Uredospores had only a very low enzyme activity (1.9 μmol Pi × mg−1protein × h−1). In germ tubes the ATPase activity was about twofold higher (4.0 μmol Pi × mg−1protein × h−1). An eightfold higher ATPase activity (16.1 μmol Pi × mg−1protein × h−1) was found in microsomal vesicles from haustoria which had been isolated from rust-infectedVicia fabaleaves. These results suggest, that the electrochemical gradient generated by the H+-ATPase of haustoria plays an important role for their function, possibly by promoting nutrient uptake from host cells.

Author Keywords: ATPase; biotrophic fungi; haustorium; nutrient uptake; plant pathogen

*1 This paper was accepted under the editorship of Robert Brambl.

Introductory mycology

Author Alexopoulos, Constantine John, 1907-
Title Introductory mycology / C.J. Alexopoulos, C.W. Mims, M. Blackwell.
Publication Details New York : Wiley, 1996.
589.2 5 /4
Edition 4th ed.
Description x, 869 p. : ill. ; 25 cm.
ISBN 0471522295 (cloth : alk. paper)
9780471522294
Subject Mycology.
Other Author Mims, Charles W.,
Blackwell, Meredith.

Development of an immunomagnetic capture-reverse transcriptase-PCR assay for three pineapple ampeloviruses.

J Virol Methods. 2008 Nov 6; : 18996414 (P,S,G,E,B,D) [Cited?]Development of an immunomagnetic capture-reverse transcriptase-PCR assay for three pineapple ampeloviruses.

[My paper] C F Gambley, A D W Geering, J E Thomas
Department of Primary Industries and Fisheries, Horticulture and Forestry Science, 80 Meiers Road, Indooroopilly, Queensland 4068, Australia; School of Integrative Biology, The University of Queensland, St. Lucia, Queensland 4072, Australia.
A semi-automated, immunomagnetic capture-reverse-transcription PCR (IMC-RT-PCR) assay for the detection of three pineapple-infecting ampeloviruses, Pineapple mealybug wilt-associated virus-1, -2 and -3, is described. The assay was equivalent in sensitivity but more rapid than conventional immunocapture RT-PCR. The assay can be used either as a one- or two-step RT-PCR and allows detection of the viruses separately or together in a triplex assay from fresh, frozen or freeze-dried pineapple leaf tissue. This IMC-RT-PCR assay could be used for high throughput screening of pineapple planting propagules and could easily be modified for the detection of other RNA viruses in a range of plant species, provided suitable antibodies are available.

Banana contains a diverse array of endogenous badnaviruses

Banana contains a diverse array of endogenous badnaviruses
Andrew D. W. Geering1, Neil E. Olszewski2, Glyn Harper3, Benham E. L. Lockhart4, Roger Hull3 and John E. Thomas1

1 Department of Primary Industries and Fisheries, 80 Meiers Road, Indooroopilly, Queensland 4068, Australia
2 Department of Plant Biology, University of Minnesota, St Paul, MN 55108, USA
3 John Innes Centre, Colney Lane, Norwich NR4 7UH, UK
4 Department of Plant Pathology, University of Minnesota, St Paul, MN 55108, USA


Correspondence
Andrew D. W. Geering
andrew.geering@dpi.qld.gov.au

Banana streak disease is caused by several distinct badnavirus species, one of which is Banana streak Obino l'Ewai virus. Banana streak Obino l'Ewai virus has severely hindered international banana (Musa spp.) breeding programmes, as new hybrids are frequently infected with this virus, curtailing any further exploitation. This infection is thought to arise from viral DNA integrated in the nuclear genome of Musa balbisiana (B genome), one of the wild species contributing to many of the banana cultivars currently grown. In order to determine whether the DNA of other badnavirus species is integrated in the Musa genome, PCR-amplified DNA fragments from Musa acuminata, M. balbisiana and Musa schizocarpa, as well as cultivars ‘Obino l'Ewai’ and ‘Klue Tiparot’, were cloned. In total, 103 clones were sequenced and all had similarity to open reading frame III in the badnavirus genome, although there was remarkable variation, with 36 distinct sequences being recognized with less than 85 % nucleotide identity to each other. There was no commonality in the sequences amplified from M. acuminata and M. balbisiana, suggesting that integration occurred following the separation of these species. Analysis of rates of non-synonymous and synonymous substitution suggested that the integrated sequences evolved under a high degree of selective constraint as might be expected for a living badnavirus, and that each distinct sequence resulted from an independent integration event.

The GenBank/EMBL/DDBJ accession numbers reported in this paper are AY189378–AY189383, AY189384–AY189392, AY189444–AY189453, AY189393–AY189400, AY189401–AY189419, AY189420–AY189435 and AY189436–AY189443

First record of Nematospora coryli in Australia and its association with dry rot of Citrus

First record of Nematospora coryli in Australia and its association with dry rot of Citrus

Roger G. Shivas A , E , Malcolm W. Smith B , Thomas S. Marney A , Toni K. Newman B , Debra L. Hammelswang B , Anthony W. Cooke C , Ken G. Pegg C and Ian G. Pascoe D

A Plant Science, Queensland Department of Primary Industries and Fisheries, 80 Meiers Road, Indooropilly, Qld 4068, Australia.
B Horticulture and Forestry Science, Queensland Department of Primary Industries and Fisheries, Bundaberg Research Station, 49 Ashfield Road, Kalkie, Qld 4670, Australia.
C Horticulture and Forestry Science, Queensland Department of Primary Industries and Fisheries, 80 Meiers Road, Indooropilly, Qld 4068, Australia.
D Primary Industries Research Victoria, Department of Primary Industries, Private Bag 15, Ferntree Gully Delivery Centre, Vic. 3156, Australia.
E Corresponding author. Email: roger.shivas@dpi.qld.gov.au


Abstract
Nematospora (Eremothecium) coryli was isolated from Citrus and identified for the first time in Australia. This insect-transmitted yeast was associated with dry rot in cultivated and native Citrus fruits. Although N. coryli is known as a serious seed pathogen of many tropical and sub-tropical plants, evidence is presented that it has been present and undetected in Queensland for at least ninety years.

Australasian Plant Pathology 34(1) 99–101

Submitted: 1 July 2004 Accepted: 26 July 2004 Published: 22 March 2005

Full text DOI: 10.1071/AP04075

© CSIRO 2005

Fungi defence against quarantine threats

Fungi defence against quarantine threats
7/05/2008 10:36:00 AM
A key diagnostic tool which could help speedily resolve threats to Australian quarantine or export trade crises has been developed by Queensland and international scientists.
Qld Department of Primary Industries principal plant pathologist Dr Roger Shivas has co-authored the book, ‘Fungi of Australia: The Smut Fungi’ with German mycologist and world expert Dr Kálmán Vánky.

The book details long-term research into Australian smut fungi in the first comprehensive guide of these important plant pathogens for almost 100 years.

The book and a CD provide interactive keys that allow quick and accurate identification of all known species of smut fungi in Australia.

The CD was co-authored with DPI&F senior research scientist Dr Dean Beasley who has crafted over 1000 images in the first ever diagnostic key of this kind for a group of plant pathogens.

Dr Shivas anticipates that the book will become an essential resource for resolving quarantine and trade issues, as well as identifying smut fungi and the diseases they cause.

He said the rapid and accurate identification of new and unusual smut fungi will allow industry to move quickly to stop new invasions.

Dr Shivas says that the book and CD take the taxonomy of fungi into the 21st Century.

“Now anyone with basic training in plant health will be able to quickly and with reasonable confidence identify all the Australian species of smut fungi.

“There are 296 species of smut fungi in Australia from covered smut which attacks barley to the recently arrived sugarcane smut.

“Smuts are parasites usually of cereals and grasses that form black powdery masses of spores which spread the disease in the air, seeds and soil.” Dr Shivas said.

Smut fungi can cause diseases in cereal crops that could devastate yields if left untreated.

Using effective fungicidal seed treatments and the development of smut resistant varieties has reduced the importance of smut diseases in recent years.

Some exotic smut fungi still threaten Australian crops however.

“The seriousness of the threat posed by smuts to Australia’s billion dollar wheat industry was highlighted in 2004 when a shipment of Australian wheat was rejected by an importing country because it allegedly contained spores of the Karnal bunt smut fungus.

“The issue was only resolved when common contaminant spores were shown to have been confused with those of Karnal bunt.”

Dr Shivas said that event convinced him that an easy-to-use and reliable means of identifying smut fungi was needed.

* The book and CD are available from CSIRO Publishing, PO Box 1139, Collingwood VIC 3066 (sales@publish.csiro.au).

SOURCE: Queensland Country Life

Identification and systematics of rust fungi in Queensland

Topic: Identification and systematics of rust fungi in Queensland
Supervisors: Tanya Scharaschkin (NRS), Roger Shivas (QDPI&F), Andrew Geering (QDPI&F)
If interested, contact: Dr Tanya Scharaschkin, School of Natural Resource Sciences. Email – t.scharaschkin@qut.edu.au
Duration: Mid November 2008 to mid February 2009 (start and end dates are flexible)
Description:
The rust fungi (Uredinales) are destructive pathogens of a range of plants including cereals, legumes, forest trees and native plants. They have caused famines and destroyed the economies of entire countries. Rusts usually attack the leaves of plants where they can produce up to five different types of spores. Most rusts are limited to specific host plant families, genera or even species.
There are approximately 7,000 species of rust fungi worldwide, with over 500 species reported from Australia. The QDPI&F Plant Pathology Herbarium (Indooroopilly) has a large collection of specimens of rust fungi from Australia. Amongst these specimens are many unidentified and undescribed species of rust fungi, particularly those found on native Australian plants.
Traditional taxonomy of the rusts has been based primarily on the morphology of teliospores. An opportunity exists for a student to work with this collection under the supervision of experienced mycologists and molecular biologists. The student will use morphological and molecular techniques to gain an understanding of the diversity and phylogeny of rust fungi in Queensland. This project has potential to result in publications (e.g., description of new species) and could be extended into an honours research project.

DNA replication cycle in parthenogenetically developing eggs of the starfish Asterina pectinifera

DNA replication cycle in parthenogenetically developing eggs of the starfish Asterina pectinifera

Author(s): Nomura A, Nemoto S
Source: DEVELOPMENT GROWTH & DIFFERENTIATION Volume: 40 Issue: 4 Pages: 377-386 Published: AUG 1998
Times Cited: 4 References: 16 Citation Map
Abstract: Starfish oocytes artificially activated by a calcium ionophore will develop normally if the formation of polar bodies is suppressed. In the present paper, schedules of the DNA replication period (S phase) of these parthenogenotes were explicitly timed using 5-bromo-2'-deoxyuridine (BrdU) and anti-BrdU monoclonal antibody. Their schedule of S phase was identical to that of fertilized eggs. Consequently an S phase regulation system is triggered even in parthenogenotes raised by dual treatment of egg activation and polar body suppression. The S phase schedule of parthenogenotes confirms the temporal pattern of chromosome duplication, observed by other researchers, leading to tetraploid parthenogenotes. The S phase determination also provides a basis for argument concerning the number of centrioles participating in parthenogenetic development. If polar body formation of activated eggs was not suppressed, the first S phase was normal, but the second S phase did not recur on time. A rigidly regulated system of DNA replication cycle, which should be an essential prerequisite for parthenogenesis, thus requires the content of polar bodies.

Biology of Wolbachia

Biology of Wolbachia
Author(s): Werren JH
Source: ANNUAL REVIEW OF ENTOMOLOGY Volume: 42 Pages: 587-609 Published: 1997
Times Cited: 414 References: 132 Citation Map
Abstract: Wolbachia are a common and widespread group of bacteria found in reproductive tissues of arthropods. These bacteria are transmitted through the cytoplasm of eggs and have evolved various mechanisms for manipulating reproduction of their hosts, including induction of reproductive incompatibility, pathenogenesis, and feminization. Wolbachia are also transmitted horizontally between arthropod species. Significant recent advances have been made in the study of these interesting microorganisms. In this paper, Wolbachia biology is reviewed, including their phylogeny and distribution, mechanisms of action, population biology and evolution, and biological control implications. Potential directions for future research are also discussed.