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Colonization of plant roots by egg-parasitic and
nematode-trapping fungi
Blackwell Science, Ltd
J. J. Bordallo1, L. V. Lopez-Llorca1, H.-B. Jansson1,2, J. Salinas1, L. Persmark2 and L. Asensio1
Departamento de Ciencias Ambientales y Recursos Naturales, Universidad de Alicante, Apartado Correos 99, ES−03080 Alicante, Spain; 2Department of
1
Microbial Ecology, Lund University, Ecology Building, S−223 62 Lund, Sweden.
Summary
Author for correspondence:
L. V. Lopez-Llorca
Tel: +34 96 590 3400, Ext. 3381
Fax: +34 96 590 3815
Email: [email protected]
Received: 13 September 2001
Accepted: 5 December 2001
• The ability of the nematode-trapping fungus Arthrobotrys oligospora and the
nematode egg parasite Verticillium chlamydosporium to colonize barley (Hordeum
vulgare) and tomato (Lycopersicum esculentum) roots was examined, together with
capability of the fungi to induce cell wall modifications in root cells.
• Chemotropism was studied using an agar plate technique. Root colonization was
investigated with light microscopy and scanning electron microscopy, while compounds involved in fungus–plant interactions were studied histochemically.
• Only A. oligospora responded chemotropically to roots. Colonization of barley and
tomato by both fungi involved appressoria to facilitate epidermis penetration. V. chlamydosporium colonized tomato root epidermis and produced chlamydospores. Papillae,
appositions and lignitubers ensheathing hyphae on tomato were also found. Phenolics
(including lignin), protein deposits and callose were present in papillae in both hosts.
Both fungi were still present in epidermal cells 3months after inoculation.
• Nematophagous fungi colonized endophytically monocotyledon and dicotyledon
plant roots. Arthrobotrys oligospora seemed to be more aggressive than V. chlamydosporium on barley roots. Both fungi induced cell wall modifications, but these did
not prevent growth. The response of root cells to colonization by nematophagous
fungi may have profound implications in the performance of these organisms as
biocontrol agents of plant parasitic nematodes.
Key words: chemotropism, Verticillium chlamydosporium, Arthrobotrys oligospora,
root colonization, Hordeum vulgare (barley), Lycopersicum esculentum (tomato),
papillae, endophytic growth.
© New Phytologist (2002) 154: 491–499
Introduction
Nematophagous fungi are common soil inhabitants infecting
living nematodes through different strategies ( Jansson &
Lopez-Llorca, 2001). Plant-parasitic nematodes generally
attack plant roots, therefore the ability of nematophagous
fungi to colonize roots should be a great advantage if the fungi
could be used for biological control. It was found that pea
rhizosphere harbours higher densities of nematode-trapping
fungi and more species of nematophagous fungi than the
root-free soil and the rhizospheres of barley and white
mustard (Persmark & Jansson, 1997). A positive rhizosphere
effect on Arthrobotrys spp. was also reported for citrus
(Gaspard & Mankau, 1986), soybean and tomato (Peterson
& Katznelson, 1965). Fungal parasites of nematode eggs are
© New Phytologist (2002) 154: 491– 499 www.newphytologist.com
commonly found in the rhizosphere of crop plants (Bourne
et al., 1996) and were recently shown to be able to colonize
the roots of barley (Lopez-Llorca et al., 2002).
Directed or chemotropic growth of hyphae is a well-known
phenomenon in many fungi (Carlile, 1983; Jansson et al., 1988;
Allan et al., 1992). The tropism may be due to host-finding
or, in saprophytic fungi, to the attraction towards a suitable
substrate. This phenomenon was explained by tropism towards
oxygen (Robinson, 1973), volatile (Koske, 1982) or non-volatile
compounds (Jansson et al., 1988). Chemotropic growth of
nematophagous fungi hyphae towards nematodes was observed
in detached traps of Dactylella doedycoides (Zachariah, 1981)
and in hyphal tips of the oyster mushroom, Pleurotus ostreatus
(Nordbring-Hertz et al., 1995). Furthermore, zoospores of
the endoparasite Catenaria anguillulae are attracted to their
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492 Research
nematode hosts (Jansson & Thiman, 1992). However, nematodes are also usually attracted towards nematophagous fungi
(Jansson & Nordbring-Hertz, 1988).
Preliminary results on the cell biology of the interactions
between root cells and nematophagous fungi with barley and
the egg parasite Verticillium chlamydosporium (Lopez-Llorca
et al., 2002) indicate that the fungus has an endophytic
behaviour in root cells. This behaviour is a closer relationship
than the saprotrophic capacity, which it has been assumed
that the fungus has in the rhizosphere (Kerry, 2000). These
results prompted us to investigate further the relationships of
nematophagous fungi and roots at a cellular level. This area of
research has been largely neglected despite its importance.
There is an increasing body of knowledge on the possible
indirect biocontrol action of fungal symbionts and antagonists
of plant pathogens by modulation of plant host defence
responses (Poromarto et al., 1998; Sivasithamparam, 1998;
Dumas-Gaudot et al., 2000).
In this paper we describe results on chemotropic growth of
nematode-trapping and egg-parasitic fungi towards roots of
barley and other plants. The ability of the nematode-trapping
fungus Arthrobotrys oligospora and the nematode egg parasite
Verticillium chlamydosporium to colonize barley and tomato
roots was examined together with of the capability of these
fungi to induce cell wall modifications of root cells.
of incubation at 22°C in the dark the tips of primary or secondary roots free from contaminants were used in the experiments.
Materials and methods
Histology and histochemistry
Chemotropism experiments
Fungi Nematode-trapping fungi recently isolated from the
rhizosphere of different plants from Swedish agricultural soils
were used. Arthrobotrys oligospora L 9201, Arthrobotrys musiformis
L 9018, Arthrobotrys superba L 9035 and Monacrosporium
psychrophilum L 9203. In addition A. oligospora ATCC 24927,
which was kept in culture collection for many years in our
laboratory, with frequent subcultivations on common agar
media, was also included. The nematode egg-parasitic fungi
Verticillium suchlasporium (CBS 464.88) and V. chlamydosporium (isolate Sevilla), isolated from Heterodera avenae eggs
(Lopez-Llorca & Duncan, 1986), were also included. The
fungi were grown at 22°C on corn meal agar (CMA, Difco,
Michigan, MI, USA) amended with/without K2HPO4 (2 g l−1).
Spores from 2-to 4-wk-old plates were suspended in sterile
water containing 0.01% Triton X-100 (Sigma, St Louis, MO,
USA) and used in the chemotropism experiments.
Plants Seeds of pea, barley and white mustard were surfacesterilized with chlorine. Pea seeds were soaked for 30 min in
5% NaClO. Barley seeds were soaked for 1 h in water and
thereafter for 1 h in 10% NaClO. White mustard seeds were
soaked for 5 min in 5% NaClO. After the chlorine treatment
the seeds were washed three times in sterile water and spread
on sterilized wetted filter paper in Petri dishes. After 2–10 d
Chemotropism assay The method used was modified after
Jansson et al. (1988). Water agar (WA, 1%) was poured onto
sterilized microscope slides to form a thin layer. Conidia, suspended in 0.01% Triton X-100, were spread on the WA surface,
leaving approximately 150 spores mm−2, and the slides were left
to dry for a few min in the air flow of a laminar flow cabinet.
Excised root tips, approx. 2 cm long, were then placed on the agar
and gently pressed down to ensure full contact. A carefully washed
and sterilized nylon string of the same length was used as a
control. The slides were kept in Petri dishes to prevent the agar
from drying out and incubated at 22°C for 7–15 h, depending
on the fungal species. On average, six to eight root tips per
fungus or control strings were set up. When the germ tubes
measured between 50 and 300 µm (after 7–15 h) and before
secondary hyphae started to emerge, the direction of the
growing hyphae, positive or negative vs the root or string,
was determined. Eighty to 100 germlings per each root tip or
nylon string were scored. The results were tested statistically
using Student’s t-test. Chemotropism was initially determined
to a distance of 1.2 mm, but in a majority of the experiments
directed growth was measured within a distance of 0.4 mm
from the root. Experiments were repeated once or twice.
Fungi For root colonization experiments A. oligospora ATCC
24927 and V. chlamydosporium (isolate V10, kindly provided
by B. Kerry, IACR, Rothamsted, UK) growing on CMA at
25°C in the dark for 1–4wks were used as inoculum.
Plants Barley (Hordeum vulgare L. var. disticum) seeds were
surface-sterilized using 5% NaClO with a drop of commercial
detergent, for 30 min at room temperature and shaking at
120 rpm. The seeds were then rinsed five times (5 min each)
in sterile distilled water and dry blotted onto sterile filter
paper. Five to 10 seeds were plated on germinating medium
at 25°C in the dark. The medium consisted of 1.2% agar
supplemented with glucose (10 g l−1), peptone (0.1 g l−1) and
yeast extract (0.1 g l−1). Since this germination medium was
rich in nutrients, contaminating fungi and bacteria were
readily detected. Only seedlings free from those contaminants
were axenically placed in 50 ml autoclaved tissue culture tubes
(one per tube) with 30 ml of water saturated vermiculite at 5–
10 mm from the surface. Tubes with seedlings were either left
as controls or were inoculated with nematophagous fungi.
Fungi were inoculated with four 5 mm diameter disks placed
10mm deep and mixed with the vermiculite. To avoid root
disturbances fungal inoculation was carried out immediately
before seedling planting. Tubes with seedlings, or seedlings
and fungus, were incubated as described above for up to 3 wks
and were sequentially sampled weekly.
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A different experimental set-up was used for tomato.
Polycarbonate Magenta GA-7 vessels (Sigma) were used.
These made a 7.5 × 7.5 × 20cm chamber. This chamber was
filled with 150 ml vermiculite and 100 ml Gamborg’s B-5
basal salt mixture (Sigma) diluted 10 times, and autoclaved at
121°C for 1 h. Tomato seeds were sterilized in 5% NaClO
and allowed to germinate for 5–7 d in the dark at room
temperature. Seedlings, together with four 5-mm disks cut
from the edge of growing fungal colonies, were added on top
of the vermiculite and incubated in a growth chamber at 25°C
under a 16 h light and 8 h dark cycle. Uninoculated tomato
seedlings served as controls. Since the development of tomato
is much slower than that of cereals, we sampled plants
1 month and 3 months after inoculation.
Light microscopy (LM) At least two barley plants were
selected at each sampling time (1, 2 and 3 wks after planting).
Fresh straight roots were chosen (two per plant). These were
cut in 0.5-cm pieces descending from the base of the stem to
the root apex. Tomato plants were sampled 1 month and
3 months after inoculation. A 0.5-cm segment was sampled
from the roots 0.5 cm below the stem base. Similar samples
were taken from the first secondary root and from secondary
roots 2 cm below the stem base.
Root segments were embedded in OCT ( Jung tissue freezing
medium, Leica, Nusslock, Germany) placed in rubber tubing
(1 mm wall thickness) 1 cm high and wide. The embedded
roots were frozen at −20°C in a Leica CM1510 cryostat and
the frozen blocks containing root segments were kept at
−20°C before sectioning in the same device. Roots were
sectioned longitudinally as 50-µm thick sections. Freshly cut
sections were placed individually on precooled (−20°C)
microscopy slides using cool tweezers. Before staining,
sections were soaked in two or three drops of distilled water
and dislodged from the OCT embedding medium. Sections
were kept in water for 5 min at room temperature. Water
was removed from the sections which were then stained
(see Histochemistry/fluorescence microscopy) and observed.
Sections were stored at 4°C, but never longer than 12 h before
observation. Samples were observed and photographed in
Olympus CH or BH microscopes (Olympus, Tokyo, Japan).
Histochemistry/fluorescence microscopy To further investigate the relationships of root cells and nematophagous fungi,
classical histochemistry protocols for the detection of compounds involved in plant interactions were adapted. They
were applied to root sections colonized with A. oligospora and
V. chlamydosporium as follows. Toluidine Blue O (0.01% w/v;
Panreac, Barcelona, Spain) in 0.1 M potassium dihydrogen
phosphate–NaOH buffer, pH 6, was used for staining of phenolics.
Coomasie Brilliant Blue R-250 (0.1%, w : v) was used in water
for protein staining, followed by destaining in acetic acid–
glycerin (1 : 1). Aniline sulphate (1% w : v, acidified solution in
water) was used for lignin staining. Sudan III (0.5%, w : v) was
used in polyethylene glycol (PEG) 400 (Sigma) for lipid staining.
Ferric chloride (20%, w : v solution in water) was used to stain
tannins and polyphenolics in general. Sirofluor (Biosupplies
Ltd, Parkville, Australia), a stain for callose, was used as a 0.1%
(w/v) solution in water for 15–30 min at room temperature.
Low-temperature scanning electron microscopy (LTSEM)
Specimens were processed as for light microscopy. They were
then ‘cryotrimmed’ in the cryostat to the required level (LopezLlorca & Duncan, 1991) and were frozen in subcooled liquid
N2 in an Oxford cryoSEM model CT 1500C (Oxford
Instruments, Witney, UK) attached to an Hitachi S-3000 N
scanning electron microscope (Hitachi, Tokyo, Japan). Samples
were observed uncoated and frozen (−150°C), then etched
(−90°C) for the time required to remove surface ice. The
samples were Au-coated in the Oxford cryoSEM model CT
1500C sputter. Coated specimens were observed and images
digitally recorded in the S-3000 N scanning electron microscope.
Results
Chemotropism
Both isolates of A. oligospora showed a clear tropism towards
plant roots and typically about 70% of the germ tubes
grew towards the root. It was found that A. musiformis,
M. psychrophilum and the two Verticillium spp. did not grow
tropically towards the roots (Table 1). Directed growth in
A. oligospora was clear to a distance of 0.4 mm from the root
Table 1 Directed growth in germ tubes of different nematophagous fungi at a distance of 0 –0.4 mm from plant roots or nylon string
Fungus
Barley
Pea
Arthrobotrys musiformis L 9018
Arthrobotrys oligospora L 9201
Arthrobotrys oligospora ATCC 24927
Arthrobotrys superba L 9035
Monacrosporium psychrophilum L 9203
Verticillium chlamydosporium (isolate Sevilla)
Verticillium suchlasporium
44.1 ± 3.2
78.0 ± 3.8***
75.9 ± 1.1***
49.5 ± 0.2
53.3 ± 2.1
53.5 ± 9.6
56.0 ± 4.4
57.9
70.9
72.0
51.7
49.5
nd
nd
White mustard
± 4.3
± 5.4**
± 0.8***
± 0.8
± 2.0
65.1
62.9
64.9
52.9
nd
nd
nd
± 3.8
± 6.2
± 3.9***
± 0.6
Nylon string
48.5
nd
47.2
52.1
46.7
46.2
48.2
± 9.1
± 0.8**
± 3.9
± 1.9
± 4.3
± 4.0
**,***Numbers significantly different from 50% at P > 0.01 and P > 0.001, respectively. (n varies between 4 and 18). Number of germ tubes
growing towards roots or nylon string are expressed as percentage of the total numbers of germ tubes ±standard error; nd, not determined.
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Table 2 Directed growth of germ tubes of Arthrobotrys oligospora
L 9201 at different distances from the root surface
Distance from the root
Plant
0– 0.4 mm
0.5 –0.8 mm
0.9 –1.2 mm
Barley
Pea
White mustard
63.3 ± 3.7*
59.4 ± 2.3***
63.3 ± 2.7***
59.0 ± 2.6**
56.7 ± 2.2**
58.0 ± 3.8
57.3 ± 2.1*
55.0 ± 2.2
53.6 ± 0.8
*,**,***Numbers significantly different from 50% at P > 0.05,
P > 0.01 and P > 0.001, respectively (n varies between 5 and 9).
Number of germ tubes growing towards roots are expressed as
percentage of the total numbers of germ tubes±standard error.
and became less evident at increasing distances (Table 2).
No clear differences were observed in the response towards
the different plants tested (Tables 1 and 2). Incubation of
plant roots on the agar surface 24 h before addition of the
spores did not change the directed growth response of the fungi
compared with experiments where the spores and the roots
were applied to the agar surface at the same time (data not
shown).
Root colonization
Arthrobotrys oligospora – barley It was found that A.
oligospora began to penetrate the epidermis of the barley roots
after 2 days (Fig. 1a). Occasionally, root hairs were colonized
by the fungus and an expansion resembling an infection bulb
was found within them (Fig. 1b, arrow). Appressoria were found
both using LM and LTSEM when the fungus spread in the
epidermis. (Fig. 1c,d, arrows). Root necrotic areas (Fig. 2a)
were found 7d after inoculation close to the inoculum. Light
microscopy revealed cell collapse in these areas and deep
staining of epidermal cells (Fig. 2b). Hyphae were always
associated with these necroses (Fig. 2b, arrow). Conversely,
root cells from adjacent nonnecrotic areas did not show these
features (Fig. 2c). Hyphae resembling nematode-trapping
organs were sometimes observed. Vacuolation was common
in the mycelium associated with these zones (Fig. 2c). Downstream from those necrotic regions root decortication was found,
14d after inoculation (Fig. 2d). Extensive hyphal colonization
was found but no necrotic tissue was detected (Fig. 2e, arrow).
Adjacent nondecorticated areas showed healthy (stained)
hyphae in epidermal cells (Fig. 2f, arrow).
Verticillium chlamydosporium–tomato Tomato rhizosphere
appeared different from that of barley (Fig. 3a). Upon root
growth, root cells (both epidermal and cortex) changed from
long into square-rounded cells in older roots with wider cell
wall (results not shown). One month after inoculation, the
fungus colonized the epidermis, producing chlamydospores
(Fig. 3b, arrow) and conidia (Fig. 3c). Coomassie-stained
appositions on tomato cell walls were sometimes associated
with fungal appressoria (Fig. 3d, arrow). Three months after
inoculation the fungus was still present in epidermal cells
(Fig. 3e). Lignitubers ensheathing hyphae in root cells were
found 3months after inoculation (Fig. 3f ).
Arthrobotrys oligospora–tomato Three months after inoculation, tomato roots still showed signs of colonization by A.
oligospora. The fungus was present in epidermal cells (Fig. 4a)
with extensive branching (Fig. 4b). Signs of cell wall penetration (appressoria) of epidermis cells were also found
(Fig. 4c, arrow). Papillae in epidermal cells close to stem
base were sometimes found (Fig. 4d, arrow).
Fig.1 Early colonization of barley roots by
Arthrobotrys oligospora. (a) Epidermal cell
penetration (arrow), 2d after inoculation.
Bar, 30 µm. (b) Root hair colonization
including infection bulb (arrow) 6d after
infection. Bar, 10 µm. (c) Appressorium
(arrow) on epidermal cell wall, 6d after
inoculation. Bar, 30 µm. (d) Cryo-scanning
electron micrograph of epidermis
colonization, 10d after inoculation. Arrow
indicates appressorium. Bar, 20 µm.
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Fig.2 Mid to late colonization of barley roots
by Arthrobotrys oligospora. (a) General view
of root 7d after inoculation. Arrows indicate
necrotic areas. Bar, 3mm. (b) Detail of
necrotic area 14d after inoculation, showing
collapsed and deeply stained epidermal cells.
Arrow indicates unstained hyphae. Bar,
30 µm. (c) Detail of non-necrotic area in
epidermal-cortex area 12d after inoculation.
Note curved vacuolated hypha resembling
traps. Bar, 30 µm. (d) General view of
decorticated area of cortex removal away
from seed, 14d after inoculation. Bar, 1mm.
(e) Detail of (d) showing broad hyphae
(arrow). Bar, 30 µm. (f) Root area adjacent to
decorticated zone showing healthy (stained)
hyphae (arrow). Bar, 30 µm.
Table 3 Histochemical labelling of plant roots colonized by nematophagous fungi
Barley
Tomato
Stain/staining
Labelling found
Verticillium
chlamydosporium
Arthrobotrys oligospora
V. chlamydosporium
Toluidine Blue/phenolics
Aniline/callose
Sirofluor/callose
Aniline sulphate/lignin
Coomassie/proteins
Papillae, cytoplasm
Papillae, cell wall deposits
Papillae, cell wall deposits
Papillae, lignitubers, cell wall deposits
Papillae, lignitubers, cell wall deposits
+
+
+
+
±
+
+
+
+
±
+
nd
nd
+
±
nd, Not determined; +, labelling; ±, occasional labelling.
Histochemistry The results of histochemical labelling of
plant roots colonized by V. chlamydosporium and A. oligospora
are shown in Table 3. No differences were found in labelling
for any plant–fungus combination. Similar structures were
labelled in both hosts. These included cell wall modifications
such as papillae, lignitubers and deposits. Proteins accumulated in papillae, lignitubers (Fig. 5a) and cell wall deposits
(Fig. 3d). Lignin was present in papillae (Fig. 5d) and cell wall
deposits. Phenolics were also present in papillae (Fig. 5b).
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Callose deposition in papillae was revealed by Sirofluor
labelling (Fig. 5e,f ). No cutin (Fig. 5c) or tannins were found
in fungus-induced cell wall modifications.
Discussion
Arthrobotrys oligospora was the only species of the
nematophagous fungi tested that grew chemotropically
towards root tips from all three plants tested. The directed
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Fig.3 Colonization of tomato roots by
Verticillium chlamydosporium (isolate V10).
(a) Three-month-old uninoculated root. Bar,
60 µm. (b–d) One month after inoculation.
(b) External chlamydospores (arrow). Bar,
60 µm. (c) Conidia production (arrow). Bar,
30µm. (d) Appressorium and cell wall protein
apposition (arrow) in epidermal cell. Bar,
30 µm. (e,f) Three months after inoculation.
(e) Hyphal remains (arrows) in epidermal cells.
Bar, 60 µm. (f) Lignituber ensheathing
hyphae in root cell (arrow). Bar, 60 µm.
Fig.4 Tomato roots after 3month’s
colonization by Arthrobotrys oligospora.
(a) Fungus growth in epidermal cell. Bar,
30 µm. (b) Hyphal network in epidermal cells.
Bar, 30 µm. (c) Appressorium (arrow) on
cortex cell wall. Bar, 30 µm. (d) Papillae
(arrow) in epidermal cells close to stem base.
Bar, 30 µm.
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Fig.5 Histochemical detection of compounds
in barley root cell–nematophagous fungi
interactions. Verticillium chlamydosporium
(a,c,d,e) and Arthrobotrys oligospora (b,f).
(a) Proteins (arrow) in lignituber of cortex cell
stained with Coomassie Brilliant Blue. Bar,
30 µm. (b) Phenolics (arrow) in papillae near
vascular system stained with toluidine. Bar,
30 µm. (c) Absence of cutin (arrow) in
papillae of cortex cell treated with Sudan III.
Bar, 30 µm. (d) Lignin in papillae (arrow) close
to vascular tissue stained with aniline
sulphate. Bar, 30 µm. (e,f) Callose deposition
in papillae (arrow) stained with Sirofluor. Bar,
30 µm in (e) and 20 µm in (f).
growth towards roots evident in A. oligospora may explain the
higher abundance of this fungus in rhizosphere rather than in
root-free soil (Persmark & Jansson, 1997). The directed
growth response was strongest closest to the root surface
(0–0.4 mm). At more than 1 mm from the root the tropic
response was lost, indicating that substances released from the
root surface were responsible for the effect. Similar results
were also obtained for chemotropic growth of Cochliobolus
sativus towards barley roots, and partial characterization of
the substances responsible suggested that these were small
molecules, with a molecular mass less than 2000Da, and
possibly common root exudates ( Jansson et al., 1988).
Arthrobotrys oligospora rapidly colonized the barley roots
and reached the cortex but not the vascular tissues. This is the
first report that a nematode-trapping fungus is capable of colonizing root cells. A recent study showed that the egg-parasite
V. chlamydosporium also endophytically colonized barley roots
(Lopez-Llorca et al., 2002) and completed its life cycle inside
the roots. Colonization of barley roots by A. oligospora
induced necrosis in root epidermal cells. Root necrotic areas
were confined only to the initial inoculum of the fungus. No
such events were found in roots colonized by V. chlamydosporium. This may be partly explained by the chemotropic
© New Phytologist (2002) 154: 491– 499 www.newphytologist.com
response of A. oligospora towards roots, but also by its faster
growth rate. Nematophagous fungi spread between barley and
tomato root cells was by means of direct penetration or appressoria. These structures, also observed for V. chlamydosporium
on roots (Lopez-Llorca et al., 2002) and during host penetration,
were not previously described for A. oligospora. Nematodetrapping fungi, like A. oligospora, penetrate the nematode
cuticle with penetration hyphae formed on the trapping organ
and thereafter produce infection bulbs inside the nematode,
from which trophic hyphae emerge (Jansson & NordbringHertz, 1988). The capacity of A. oligospora to produce appressoria may be connected with its teleomorph (Orbilia), which
includes wood decomposing fungi (Pfister, 1997). Barron
(1992) suggested that the nematophagous habit has evolved
from cellulose- and lignin decomposing fungi. In addition to
appressorium formation, we found structures similar to the
infection bulbs that the fungus forms when infecting nematodes ( Jansson & Nordbring-Hertz, 1988) and structures
resembling the three-dimensional traps of A. oligospora.
Further down in the barley root system, A. oligospora caused
decortication but not necroses. Decortication was observed by
A. oligospora but not for V. chlamydosporium. It was probably
caused by intercellular growth of the former in epidermis and
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cortex root cells, compared with the growth habit of the latter,
which is mostly intracellular.
In tomato, root colonization by nematophagous fungi was
restricted compared with that of barley. No necroses or decortication were found irrespective of the nematophagous
fungus inoculated. In most cases, cells containing hyphae of
either V. chlamydosporium or A. oligospora were close to the
root surface, indicating that they were most likely epidermic.
It was difficult to estimate fungus spread to cortex because of
the cell dynamics, which made epidermal and cortex cells similar (see Results Section). Long lignitubers found ensheathing
only V. chlamydosporium hyphae in tomato but not barley
roots could explain the different extent of colonization of
the two hosts by the fungus. Fungal growth rate, although
decreased by such plant cell wall modifications, was not
arrested. This was also observed for several fungal root
pathogens (Rodríguez-Gálvez & Mendgen, 1995; Mims
et al., 2000). However, other factors might be controlling
fungus spread in tomato, since no long lignitubers were found
in A. oligospora-colonized roots.
The differences in colonization between barley and tomato
roots by nematophagous fungi shown in this study could
result from host defence mechanisms (see below) or more
likely from structural differences between monocotyledon
and dicotyledonous plants. Graminaceae cell wall contains
less pectin and extensin than other plants (Fry, 1988). This
may explain why barley was best colonized by nematophagous fungi. Other experiments on wheat colonization by V.
chlamydosporium provided further support to this hypothesis
(J. J. Bordallo, 2001, unpublished). Soils suppressive to the
cereal cyst nematode H. avenae are well known in cereal monocultures world-wide (Stirling, 1991). In these agroecosystems, fungus infection (mainly by V. chlamydosporium) causes
nematode decline. Their occurrence could be explained by the
structure and biology of the nematode host, being more prone
to fungus infection than potato cyst nematodes or root-knot
nematodes. We suggest, however, that the ability of V. chlamydosporium to colonize cereal roots may play an important role
in establishing the fungus to levels sufficient to cause nematode suppression and maintain the fungal inoculum in soil.
Both barley and tomato roots responded to colonization by
nematophagous fungi forming papillae and other cell wall
appositions. These structures of heterogeneous composition
and morphology are induced by pathogenic (Heitefuss, 1997)
and non-pathogenic fungi (Beswetherick & Bishop, 1993; Bao
& Lazarovits, 2001) to different extents. Callose (β-1-3 glucan) which has been described in many fungus–plant interactions and is related to plant resistance (Mims et al., 2000) was
found in cell wall papillae associated with V. chlamydosporium
and A. oligospora hyphae. Protein deposits were detected on
cell walls and lignitubers associated with penetrating hyphae
and hydroxyproline-rich proteins are known to accumulate at
the sites of infection of fungal and bacterial plant pathogens
(O’Connell et al., 1990). Phenolics (including lignin),
thought to play an important role in plant defence responses,
were also present in papillae from roots colonized by nematophagous fungi (Nicholson & Hammerschmidt, 1992).
These compounds have both a structural role, enhancing
mechanical resistance of tissues to penetration and antimicrobial properties. The cell wall modifications observed did not,
however, arrest root colonization by nematophagous fungi,
suggesting that nematophagous fungi may have an indirect
biocontrol effect modulating host defence responses. This
hypothesis, which has yet to be tested for nematophagous
fungi, has already been investigated for bacteria antagonistic
to nematodes (Hasky-Günter et al., 1998), arbuscular mycorrhizae (Dumas-Gaudot et al., 2000) and for fungi antagonistic
to plant pathogens (Sivasithamparam, 1998; Salerno et al.,
2000; Bao & Lazarovits, 2001).
Future studies involving practical use (i.e. application and
delivery strategies) of nematophagous fungi should address
their root colonization behaviour and possible induction of
plant defence mechanisms.
Acknowledgements
This research was funded by a project from the European
Union (FAIR5-PL97-3444, A Strategy of Biomanagement of
Root Knot Nematodes), the Swedish Council for Forestry
and Agricultural Sciences and the Swedish Natural Science
Research Council.
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