Journal of Applied Microbiology ISSN 1364-5072
ORIGINAL ARTICLE
Effect of chitosan on hyphal growth and spore germination
of plant pathogenic and biocontrol fungi
J. Palma-Guerrero, H.-B. Jansson, J. Salinas and L.V. Lopez-Llorca
Laboratory of Plant Pathology, Multidisciplinary Institute for Environmental Studies (MIES) Ramón Margalef, Department of Marine Sciences and
Applied Biology, University of Alicante, Alicante, Spain
Keywords
biocontrol fungi, chitosan, confocal laser
microscopy, electron microscopy, hyphal
growth, root-pathogenic fungi, spore
germination.
Correspondence
H.-B. Jansson, Laboratory of Plant Pathology,
Multidisciplinary Institute for Environmental
Studies (MIES) Ramón Margalef, Department
of Marine Sciences and Applied Biology,
University of Alicante, 03080 Alicante, Spain.
E-mail: [email protected]
2007 ⁄ 0749: received 11 May 2007, revised
18 July 2007 and accepted 8 August 2007
doi:10.1111/j.1365-2672.2007.03567.x
Abstract
Aims: To investigate the toxic effect of chitosan on important root pathogenic
and biocontrol fungi (nematophagous, entomopathogenic and mycoparasitic).
Methods and Results: We have used standard bioassays to investigate the effect
of chitosan on colony growth and developed bioassays to test spore germination. The results showed that the root pathogenic and mycoparasitic fungi
tested were more sensitive to chitosan than nematophagous and entomopathogenic fungi. Chitosanases (and perhaps related enzymes) are involved in the
resistance to chitosan. Two fungi, one sensitive to chitosan, Fusarium oxysporum f. sp. radicis-lycopersici, and one less sensitive, Pochonia chlamydosporia,
were selected for ultrastructural investigations. Transmission electron microscopy revealed differences in the ultrastructural alterations caused by chitosan
in the spores of the plant pathogenic fungus and in those of the nematophagous fungus. Confocal laser microscopy showed that Rhodamine-labelled
chitosan enters rapidly into conidia of both fungi, in an energy-dependent
process.
Conclusions: Nematophagous and entomopathogenic fungi are rather resistant
to the toxic effect of chitosan. Resistance of nematophagous and entomopathogenic fungi to chitosan could be associated with their high extracellular chitosanolytic activity. Furthermore, ultrastructural damage is much more severe in
the chitosan sensitive fungus.
Significance and impact of the study: The results of this paper suggest that
biocontrol fungi tested could be combined with chitosan for biological control
of plant pathogens and pests.
Introduction
Chitin is the second most abundant polymer in nature
after cellulose (Cohen-Kupiec and Chet 1998) and occurs
in various organisms, e.g. crustaceans, insects, nematodes
and most fungi. Chitin is a major waste product from the
seafood industry. Chitosan is a partly deacetylated form
of chitin, and consists of polymers of b-1,4-glucosamine
subunits, with molecular weight up to 400 kDa. It is environmentally safe and non-toxic to higher organisms
(Kumar 2000). Chitosan and its derivatives display antibiotic activity against microorganisms, both bacteria (Rabea
et al. 2003; Liu et al. 2004; Tikhonov et al. 2006) and
fungi (Bell et al. 1998; Saniewska 2001; Park et al. 2002;
Rabea et al. 2003; Tikhonov et al. 2006). Chitosan is not
toxic to plants, but it can enhance plant resistance in
seeds (Benhamou et al. 1994; Lafontaine and Benhamou
1996), fruits (Benhamou 2004) or leaves (Trotel-Aziz
et al. 2006) and reduce disease caused by fungal pathogens.
Chitosan can be produced by chemical or microbial ⁄ enzymatic treatments (Tsai et al. 2002). Besides,
fungi can turn chitin in their cell walls to chitosan. This
is achieved by means of chitin deacetylase activity and
this may block host recognition and degradation of fungal cell walls by plant chitinases in biotrophic fungi
ª 2007 The Authors
Journal compilation ª 2007 The Society for Applied Microbiology, Journal of Applied Microbiology 104 (2008) 541–553
541
Effect of chitosan on fungi
J. Palma-Guerrero et al.
(Deising et al. 1995). Fungi, including biocontrol agents,
produce chitinolytic enzymes necessary for growth and
their antagonistic behaviour, e.g. nematophagous eggparasitic fungi for penetration of nematode eggshells
(Tikhonov et al. 2002), entomopathogenic fungi for
penetration of insect cuticles (Kang et al. 1998) and
mycoparasitic fungi for infection of host hyphae
(Guthrie and Castle 2006). Nematophagous and entomopathogenic fungi are fairly closely related and have
similar mode of action (Lopez-Llorca and Jansson
2007). Furthermore, some of these invertebrate pathogens have the capacity also to parasitize other fungi and
colonize plants endophytically (Jansson and Lopez-Llorca
2004). As far as we know, no information is available
on the effect of chitosan on biocontrol fungi. Therefore,
we wanted to investigate the compatibility of biocontrol
agents with chitosan, with the purpose of future development of a combined application of these in biological
control programmes.
The fungitoxic effect of chitosan has mostly dealt with
colony growth inhibition of plant-pathogenic fungi and
the associated ultrastructural changes in the hyphae (Laflamme et al. 1999). Much less is known about the effects
of chitosan on spore germination. In the current work,
we have compared the effects (both on hyphal growth
and spore germination) of chitosan on economically
important root-pathogenic and various biocontrol fungi
(entomopathogenic, nematophagous and mycoparasitic).
We have also studied the utrastructural damage of conidia (of one chitosan-sensitive and one chitosan-insensitive
fungus) caused by chitosan using transmission electron
microscopy. The uptake of chitosan in fungal spores has
been investigated using Rhodamine-labelled chitosan and
confocal laser microscopy.
Materials and methods
Cultures and growth conditions
Phytopathogenic, nematophagous, entomopathogenic and
mycoparasitic fungi were used in this study (Table 1).
Fungi were routinely grown on corn meal agar (CMA,
Becton Dickinson and Company, Sparks, MD, USA) at
24C in the dark, except for Fusarium oxysporum f. sp.
radicis-lycopersici (Forl), Verticillium dahliae, Rhizoctonia
solani and the Oomycete Pythium ultimum, which were
grown on potato dextrose agar (PDA, Oxoid, Basingstoke,
Hampshire, UK).
Table 1 Cultures used in this work, origin and suppliers
Plant pathogens
Species (isolates)
Origin (Supplier)
Fusarium oxysporum f. sp.
radicis-lycopersici (Forl)
Gaeumannomyces graminis
var. tritici (Ggt)
Pythium ultimum
Rhizoctonia solani
Unknown (Dr R. González, SIA, Spain)
Verticillium dahliae
Nematophagous
Pochonia chlamydosporia (P.c. 123)
P. chlamydosporia (P.c.
P. chlamydosporia (P.c.
P. chlamydosporia (P.c.
P. chlamydosporia (P.c.
P. chlamydosporia (P.c.
P. chlamydosporia (P.c.
P. chlamydosporia (P.c.
P. chlamydosporia (P.c.
P. rubescens
Paecilomyces lilacinus
Entomopathogenic
Mycoparasitic
542
BK69)
BK132)
BK309)
BK392)
BK399)
4624)
64)
75)
Beauveria bassiana
Lecanicillium cf. psalliotae
Trichoderma harzianum
T. atroviride
Perth, Australia (Dr K. Sivasithamparam, University
of Western Australia, Australia)
Unknown (CBS-101588)
Unknown (Dr F. Schauer, Greifswald University,
Germany)
Córdoba, Spain (Dr R. Jiménez Dı́az, Instituto de
Agricultura Sostenible, CSIC, Spain)
Sevilla, Spain (Collection at Plant Pathology Laboratory,
University of Alicante, Spain)
New Zeeland (Dr B. Kerry, Rothamsted Research, UK)
Kenya (Dr B. Kerry)
Zimbabwe (Dr B. Kerry)
Cuba (Dr B. Kerry)
China (Dr B. Kerry)
Australia (Dr K. Sivasithamparam)
Tarragona, Spain (University of Alicante, Spain)
Valladolid, Spain (University of Alicante, Spain)
Scotland (University of Alicante, Spain)
Unknown (Dr J. Jimenez, Universidad Politecnica,
Valencia, Spain)
Alicante, Spain (University of Alicante, Spain)
Alicante, Spain (University of Alicante, Spain)
Unknown (Dr J. Jimenez)
Sweden (Dr M. Rey, NBT, Spain)
ª 2007 The Authors
Journal compilation ª 2007 The Society for Applied Microbiology, Journal of Applied Microbiology 104 (2008) 541–553
J. Palma-Guerrero et al.
Effect of chitosan on fungi
Preparation of chitosan
Chitosanase activity assay
Chitosan (T8s), with a deacetylation degree of 79Æ6% and
a molecular weight of 70 kDa, was from Marine BioProducts GmbH (Bremerhaven, Germany). T8s chitosan was
dissolved in 0Æ25 mol l)1 HCl under continuous stirring
and pH was adjusted to 5Æ6 with 1 mol l)1 NaOH as
described in Benhamou et al. (1994). The resulting solution was dialyzed for salt removal. Dialyzed chitosan was
autoclaved (120C, 20 min) prior to be used for fungal
growth or spore germination assays.
Rhodamine-labelled T8s chitosan (Rhodamine-T8s
chitosan) was a kind gift from Dr V. Tikhonov (Laboratory of Physiologically Active Biopolymers, A.N.Nesmeyanov Institute of Organoelement Compounds, Russian
Academy of Sciences, Moscow, Russia). After dialysis, the
labelled chitosan was subjected to ultrafiltration through
3 kDa ultramembranes in an Amicon Cell (Omega, Pall
Corporation, Ann Arbor, MI, USA) to remove free Rhodamine. The ultrafiltrate was sterilized by filtration through
0Æ2 lm pore-size syringe filter (Albet, Barcelona, Spain).
Halos of substrate clearing were observed around colonies of some fungi growing on chitosan-amended media.
The halo radii were measured daily. An estimation of
fungal chitosan degrading activity (CDA) was carried out
using the following index: CDA = 1 ) (colony radius ⁄
chitosan clearing halo radius), to compare CDA in all
fungi as described in Olivares-Bernabeu and LopezLlorca (2002).
To test that chitosan clearing halos occurred as a result
of the enzymatic (chitosanase) activity of the fungi, we
proceeded as follows: three 1 cm2 agar squares from the
halo (around the colony) of P. chlamydosporia (isolate
P.c. 123) growing in chitosan-amended CMA (at
2 mg ml)1) were taken per plate, with three plates sampled per treatment. These squares were placed in a 1Æ5 ml
Eppendorf tube and homogenized with 500 ll of
0Æ05 mol l)1 Tris–HCl buffer (pH = 7Æ5). After 1 h shaking at 4C, the tubes were centrifuged at 19 300 g for
10 min at 4C and the supernatants collected. Chitosanase
activity was measured using a modification of the fluorometric method described by Tikhonov et al. (2004). The
reaction mixture contained 100 ll of FITC (fluorescein
isothiocyanate)-chitosan (4 mg ml)1), 300 ll 0Æ1 mol l)1
acetate buffer (pH = 4Æ7), 50 ll of enzyme solution, and
distilled water up to 1 ml. After incubation at 37C for
90 min, the reaction was terminated by adding 50 ll of
10 mol l)1 sodium hydroxide. Tubes were centrifuged at
20 800 g for 15 min, and 25 ll of supernatant was added
to 2 ml 0Æ5 mol l)1 Tris–HCl buffer (pH = 8Æ5). As a
control we used 1 cm2 squares of culture media from
plates of P. chlamydosporia (isolate P.c. 123) growing in
CMA or HCl-amended CMA, from the same area used in
the culture media with chitosan. A further control was
performed by treating the supernatants from chitosanamended culture media at 100C for 30 min to inactivate
the enzymes. The experiment was carried out twice.
Effect of chitosan on fungal growth
An experiment to evaluate the effect of chitosan on fungal
growth was carried out on PDA plates amended
with chitosan at different concentrations (0Æ5, 1 and
2 mg ml)1) as in Laflamme et al. (1999). Unamended
PDA plates served as controls. A further control consisted
of 0Æ25 mol l)1 HCl neutralized (pH 5Æ6) amended media.
These media were prepared with the same volume of neutralized HCl used to prepare 2 mg ml)1 of chitosan. These
controls aimed at ruling out fungal inhibition because of
the NaCl produced when adjusting HCl pH with NaOH.
Fungi used in this experiment included three plant pathogenic fungi (Forl, Gaeumannomyces graminis var. tritici
(Ggt), and V. dahliae), a plant pathogenic Oomycete
(Py. ultimum) and three biocontrol fungi (Pochonia chlamydosporia (isolate P.c. 123), Pochonia rubescens and
Beauveria bassiana). In a second experiment, we evaluated
the effect of the nutrient content of chitosan-amended
media as follows: chitosan (2 mg ml)1) was added to
PDA, CMA and WA (1% water agar; Panreac, Barcelona,
Spain). This was carried out for all fungi and the Oomycete described in Table 1. Three plates per chitosan concentration in both experiments were inoculated in the
centre with a plug (5 mm diameter) from the edge of a
10–15 day-old-colony of each fungus to be tested. Two
colony radii were measured daily for each plate, until controls reached the edge of the plate. Growth percentage was
the growth in each plate amended with chitosan respect to
that of the corresponding control plate. The experiments
were performed twice.
Effect of chitosan on spore germination
Conidia used for these experiments were collected from
2-week-old cultures of fungi growing in the media
described above. The conidia were collected from the
plates with 1 ml sterile distilled water, passed through a
glass wool filter to remove hyphae, diluted, counted and
immediately used in the bioassay. Spore germination
assays were carried out on multi-well microscope slides
containing 10 wells (ICN Biomedicals, OH, USA). Each
well was filled with 25 ll of the chitosan solutions at
0Æ0001, 0Æ001, 0Æ01, 0Æ05, 0Æ1, 0Æ5 and 1 mg ml)1, sterilized by filtration, together with conidia (to a final concentration of 106 conidia ml)1) of the fungi to be tested.
ª 2007 The Authors
Journal compilation ª 2007 The Society for Applied Microbiology, Journal of Applied Microbiology 104 (2008) 541–553
543
Effect of chitosan on fungi
J. Palma-Guerrero et al.
Assays with sterile distilled water or 0Æ25 mol l)1 HCl
neutralized (pH = 5Æ6), were used as controls. After
24 h, numbers of germinated and non-germinated conidia were counted in a light microscope and the spore
germination percentage calculated. A spore was considered germinated when the germ tube length was 1Æ5
times the spore diameter (Plascencia-Jatomea et al.
2003). For all experiments, three wells per chitosan concentration were prepared, three areas per well were
recorded (each containing 20–40 conidia), and the experiment was performed twice. Conidia for each individual
experiment (control and chitosan treatments) were collected from a single plate. Conidia of B. bassiana and
V. dahliae did not germinate in water; therefore, the
experiments with these fungi were performed in 0Æ25%
PDB instead of water. Spore germination percentage was
the germination percentage in each chitosan treatment
compared with that of the corresponding control. Chitosan-treated and untreated conidia were plated on PDA
to check their viability.
Transmission electron microscopy
Conidia of F. oxysporum f. sp. radicis lycopersici and
P. chlamydosporia (isolate P.c. 123), either untreated or
treated with chitosan overnight, were fixed by overnight
immersion in 2Æ5% (v ⁄ v) glutaraldehyde (Sigma, St. Louis,
MO, USA) in 1 mol l)1 phosphate buffer (pH = 7Æ5) at
4C. Conidia were then postfixed with 0Æ1% osmium
tetroxide (Fluka, St. Gallen, Switzerland) in the same buffer for 30 min at room temperature (c. 22C). Samples
were dehydrated in a graded ethanol series and embedded
in Epon 812 (Electron Microscopy Sciences, Fort, WA,
USA). Ultrathin sections, 50-70 nm thick, were cut with a
diamond knife and collected on Formvar-coated copper
grids. They were stained with uranyl acetate and lead
citrate and examined in a Zeiss EM10 transmission
electron microscope operated at 60 kV.
Confocal laser microscopy
Rhodamine-chitosan was used to label conidia and trace
chitosan using confocal microscopy. Conidia of Forl and
P. chlamydosporia (P.c. 123) were treated with
0Æ1 mg ml)1 Rhodamine-T8s chitosan for 15 min at room
temperature (c. 22C). Conidia were washed with distilled
water by centrifugation at 19 300 g for 5 min three times
to remove unbound Rhodamine-labelled chitosan.
Labelled conidia were finally examined in a confocal
microscope (Leica DM IRBE2) using a laser with 543 nm
excitation and 555–700 nm emission wavelengths.
Sodium azide (Sigma) and low temperature (4C) were
used to test if the uptake of chitosan in conidia was an
544
active process (Hoffmann and Mendgen 1998; Atkinson
et al. 2002). Conidia were either treated with 75 mmol l)1
sodium azide or kept at 4C for half an hour before treating them with 0Æ1 mg ml)1 Rhodamine-T8s chitosan for
15 min at 4C. Conidia were then washed three times
(5 min each) with distilled water by centrifugation at
19 300 g at 4C, to remove unbound Rhodamine-labelled
chitosan. Conidia were kept at 4C until examined in the
confocal microscope.
Statistical analyses
Statistical analyses of the data were performed using SPSS
13Æ0 for Windows. To test possible differences between
chitosan treatments and controls we carried out raw data
(e.g. colony diameters) analyses. To study differences
between chitosan treatments (e.g. one fungus with several
treatments, or several fungi with the same treatment) we
analyzed percentages respect to control to normalize the
data. One-way anova analyses were used in every case,
assuming a P-value of 0Æ05. After that, post hoc tests were
performed using the method of the least significant difference (LSD), for detecting significant differences between
treatments.
Results
Effect of chitosan on fungal growth
Plant pathogenic (Forl, Ggt, Py. ultimum and V. dahliae)
and biocontrol fungi (P. chlamydosporia isolate P.c. 123,
P. rubescens and B. bassiana) showed clear differences
when growing in chitosan-amended PDA at different
concentrations. Except for Ggt, chitosan markedly
reduced the radial growth of the plant pathogenic fungi
and the Oomycete in PDA, especially at 2 mg ml)1
(Table 2). Forl and Py. ultimum were the most sensitive
to chitosan, with growth percentages of 22Æ8% and
23Æ7%, respectively, at the highest chitosan concentration
(2 mg ml)1). Ggt appeared to be only slightly affected by
high chitosan concentration (Table 2). For V. dahliae,
chitosan improved growth at 0Æ5 and 1 mg ml)1, but at
2 mg ml)1 there was growth reduction (Table 2). The
nematophagous fungus P. chlamydosporia (isolate P.c.
123) was only slightly affected by chitosan at 1–
2 mg ml)1. P. rubescens did not show differences from
the control growing in 0Æ5 and 1 mg ml)1 chitosanamended medium. However, at 2 mg ml)1 P. rubescens
showed significant growth increase respect to the control
(110Æ6%). The entomopathogenic fungus B. bassiana
showed the largest increase in growth for all fungi tested
in chitosan-amended media, especially at the highest
concentrations.
ª 2007 The Authors
Journal compilation ª 2007 The Society for Applied Microbiology, Journal of Applied Microbiology 104 (2008) 541–553
J. Palma-Guerrero et al.
Effect of chitosan on fungi
Table 2 Effect of chitosan concentration on
growth rates (%) of plant pathogenic fungi
and biocontrol fungi
Chitosan concentration (mg ml)1)
0Æ5
Plant pathogenic fungi
Fusarium oxysporum f. sp.
radicis-lycopersici
Gaeumannomyces graminis
var. tritici
Pythium ultimum
Verticillium dahliae
Biocontrol fungi
Pochonia chlamydosporia
(P.c. 123)
P. rubescens
Beauveria bassiana
a
b
61Æ4 ± 4Æ2a*
102Æ6 ± 2Æ7a
b
99Æ5 ± 2Æ4a
131Æ2 ± 12Æ0a*
c
b
100Æ9 ± 1Æ7a
b
100Æ9 ± 5Æ3a
166Æ7 ± 2Æ6a*
d
1
2
a
61Æ9 ± 11Æ9a*
a
22Æ8 ± 2Æ1b*
102Æ6 ± 3Æ0a
b
a
63Æ4 ± 3Æ1b*
113Æ8 ± 8Æ8b
a
d
d
b
92Æ7 ± 1Æ5b*
23Æ7 ± 3Æ2c*
45Æ7 ± 2Æ2c*
c
c
88Æ1 ± 2Æ0b*
100Æ9 ± 4Æ6a
188Æ9 ± 5Æ3b*
80Æ6 ± 1Æ0b*
b
e
e
f
110Æ6 ± 4Æ1a*
185Æ5 ± 1Æ5b*
% growth rates relative to corresponding control plates. Superscripts on the left show differences between fungi growing under the same conditions (columns). Superscripts on the right
show differences for each fungi growing under different conditions (rows). Values with different
letters are significantly different (P < 0Æ05). Values with an asterisk show differences from the
control (P < 0Æ05). Each value represents the mean of three replicates.
The comparison of the effect of chitosan at 2 mg ml)1
in the three different media (PDA, CMA and WA) for all
fungi and the Oomycete showed that plant pathogens and
mycoparasitic fungi were more affected by chitosan than
nematophagous and entomopathogenic fungi (Table 3).
Ggt was an exception, since it showed growth rates similar to those of nematophagous and entomopathogenic
fungi. The plant pathogens Forl, Py. ultimum and V. dahliae were more affected by chitosan in CMA and WA
than in PDA (Table 3). Ggt showed values close to 90%
in the three media. R. solani showed values close to 70%
in PDA and CMA, but did not grow in WA amended
with chitosan. The nematophagous fungi (P. chlamydosporia, nine isolates, P. rubescens and Paecilomyces lilacinus)
were, in general, less affected by chitosan than plant pathogenic fungi (Table 3). Different behaviour was observed
in the media tested depending on the isolate, with growth
increase in PDA amended with chitosan for some fungi
(e.g. P.c. 4624, P.c. 75 and P. rubescens), and growth inhibition in CMA or WA amended with chitosan for most
of them. In all cases, growth inhibition was much less
than that observed in the plant pathogenic fungi
(Table 3). The entomopathogenic fungi B. bassiana and
L. cf. psalliotae showed results similar to the nematophagous fungi. B. bassiana showed the largest differences
between the three media, with 185Æ5% growth in PDA
amended with chitosan, and 68Æ4% growth in WA
amended with chitosan. The mycoparasitic fungi Trichoderma harzianum and Trichoderma atroviride were sensitive to chitosan in all culture media tested (Table 3). No
differences were observed in the HCl-amended media
respect to the control for all fungi (data not shown).
Fungal degradation of chitosan
Halos were observed around the colonies of some fungi
growing in chitosan-amended media, suggesting that
these fungi were able to degrade the chitosan (Fig. 1).
The degrading halos were observed in 2- or 3-day old colonies, and increased in size with that of the colony. The
CDA indices recorded for all fungi growing in the three
chitosan-amended media are summarized in Table 4. The
highest CDA indices were found for nematophagous
fungi. In PDA, chitosan clearing halos were only observed
for the nematophagous fungi and the plant pathogenic
fungus V. dahliae. The highest CDA value (0Æ62) in PDA
was observed for P. rubescens. In CMA, halos were
observed for all biocontrol fungi and V. dahliae. As in
PDA, P. rubescens was the fungus with the highest CDA
value of all fungi tested (0Æ7) in CMA. For B. bassiana,
T. harzianum and T. atroviridae the halos were of the
same size as the colony diameter (CDA value of 0). In
WA, halos were observed for most nematophagous fungi
and entomopathogenic fungi. In this medium, the mycoparasitic fungi did not grow at 2 mg ml)1 of chitosan,
and subsequently no halos were recorded. P.c. 4624 was
the nematophagous fungus with the lowest CDA index
for all media tested.
When the chitosanase enzymatic assay was carried out
with agar from the halo of the nematophagous fungus
P. chlamydosporia (isolate P.c. 123) high fluorescence
(11 907 UF per lg of protein) was detected. In contrast,
a very low fluorescense background was detected in
media without chitosan-amendment (167 UF per lg of
protein).
ª 2007 The Authors
Journal compilation ª 2007 The Society for Applied Microbiology, Journal of Applied Microbiology 104 (2008) 541–553
545
Effect of chitosan on fungi
J. Palma-Guerrero et al.
Table 3 Effect of chitosan (at 2 mg ml)1) in
different culture media on growth rates of
plant pathogenic and biocontrol fungi
Culture media
PDA
Plant paghogenic fungi
Fusarium oxysporum f. sp.
radicis-lycopersici
Gaeumannomyces graminis
var. tritici
Pythium ultimum
Rhizoctonia solani
Verticillium dahliae
Nematophagous fungi
Pochonia chlamydosporia
(P.c. 123)
P. chlamydosporia (P.c. BK69)
P. chlamydosporia (P.c. BK132)
P. chlamydosporia (P.c. BK309)
P. chlamydosporia (P.c. BK392)
P. chlamydosporia (P.c. BK399)
P. chlamydosporia (P.c. 4624)
P. chlamydosporia (P.c. 64)
P. chlamydosporia (P.c. 75)
P. rubescens
Paecilomyces lilacinus
Entomopathogenic fungi
Beauveria bassiana
Lecanicillium cf. psalliotae
Mycoparasitic fungi
Trichoderma harzianum
T. atroviride
CMA
a
22Æ8 ± 2Æ1a*
b
92Æ7 ± 1Æ5a*
b
c
23Æ7 ± 3Æ2a*
72Æ7 ± 4Æ0a*
d
45Æ7 ± 2Æ2a*
c
e
80Æ6 ± 1Æ0a*
b,e,f,g
2Æ7a
0Æ9a
0Æ0a*
4Æ3a*
9Æ0a
7Æ3a*
8Æ7a
1Æ5a*
4Æ1a*
4Æ7a,b
b
a
f
104Æ1
100Æ0
h
90Æ9
e,i
84Æ1
f,g,j
107Æ5
k
126Æ9
b,g,h,i
89Æ9
k
122Æ9
f,j
110Æ6
f,g,j
102Æ7
g
l
j
±
±
±
±
±
±
±
±
±
±
185Æ5 ± 1Æ5a*
108Æ6 ± 0Æ0a*
m
3Æ7 ± 0Æ0a*
30Æ30 ± 4Æ7a*
a
WA
a
0 ± 0b*
a
0 ± 0b*
90Æ2 ± 0Æ7a,b*
b,c,d
86Æ2 ± 3Æ2b*
a
1Æ6 ± 1Æ6b*
67Æ4 ± 3Æ5a*
d
4Æ8 ± 0Æ8b*
89Æ3 ± 6Æ7a*
89Æ6
91Æ9
f,h
79Æ9
i
46Æ5
g
96Æ8
j
101Æ6
f
85Æ0
e
92Æ7
e,g
93Æ6
g
96Æ3
e
±
±
±
±
±
±
±
±
±
±
0Æ8b*
1Æ6b*
4Æ0b*
5Æ4b*
2Æ1a*
1Æ6b
2Æ4a*
0Æ9b*
2Æ8b*
2Æ5a
89Æ3 ± 0Æ7b*
79Æ0 ± 1Æ7b*
a
b,e,f
77Æ8 ± 9Æ0a*
b
85Æ9
66Æ9
i
46Æ2
j
34Æ1
c,e
81Æ6
k
94Æ0
g
64Æ9
e
77Æ8
k,l
93Æ1
m
108Æ6
g,h
b
f,h
h
d,l
d
5Æ0 ± 0Æ8a*
28Æ5 ± 0Æ9a*
k
1Æ3 ± 1Æ3b*
a
0 ± 0b*
a
0 ± 0c*
±
±
±
±
±
±
±
±
±
±
1Æ5c*
3Æ1c*
2Æ1c*
3Æ4c*
2Æ4b*
1Æ0c*
1Æ1b*
1Æ0c*
3Æ0b*
2Æ5b
68Æ4 ± 2Æ2c*
89Æ4 ± 2Æ0c*
a
1Æ9 ± 1Æ9a*
a
0 ± 0b*
Growth rates relative to corresponding control plates. Superscripts on the left show differences
between different fungi growing in the same culture media (columns). Superscripts on the right
show differences for each fungi growing in different culture media (rows). Values with different
letters are significantly different (P < 0Æ05). Values with an asterisk show difference from the
control (P < 0Æ05). Each value represents the mean of three replicates.
Effect of chitosan on fungal spore germination
Spores were clearly more sensitive to chitosan than
hyphae. For the two plant pathogenic and the two mycoparasitic fungi tested a chitosan concentration of
0Æ01 mg ml)1 completely inhibited spore germination
(Table 5). Spore germination for nematophagous and
entomopathogenic fungi was less affected at 0Æ01 mg ml)1
chitosan. Pae. lilacinus, B. bassiana and some isolates of
P. chlamydosporia did not show differences respect to control. P. chlamydosporia isolate P.c. 4624 and the entomopathogenic fungus L. cf. psalliotae showed the same
sensitivity to the one displayed by plant pathogenic and
mycoparasitic fungi (Table 5). When chitosan concentration was increased to 1 mg ml)1, spore germination of the
nematophagous and entomopathogenic fungi was more
affected, but still not completely inhibited (Table 5). No
differences were observed in the HCl treatment respect to
the control for any of the fungi (data not shown).
546
Non-germinated conidia plated on PDA did not
develop colonies, confirming that they were non-viable.
The only exception were conidia of the entomopathogenic
fungus L. cf. psalliotae, which did not germinate in presence of chitosan, but produced colonies when plated on
PDA.
Effect of chitosan on ultrastructure of fungal spores
Examination of ultrathin sections of the conidia from the
plant pathogenic fungus F. oxysporum f. sp. radicis-lycopersici exposed to chitosan (0Æ01 mg ml)1) showed that
they had suffered severe damage. Cytoplasm appeared
completely disorganized, plasma membranes retracted and
most cell contents were lost (Fig. 2a and b). Conidia of
the nematophagous fungus P. chlamydosporia (isolate
P.c. 123) treated with chitosan at 0Æ01 and 1 mg ml)1
showed a mixture of damaged and non-damaged cells at
both concentrations. Electron micrographs showed that
ª 2007 The Authors
Journal compilation ª 2007 The Society for Applied Microbiology, Journal of Applied Microbiology 104 (2008) 541–553
J. Palma-Guerrero et al.
(a)
Effect of chitosan on fungi
incubated at 4C, prior to Rhodamine–chitosan labelling,
fluorescence could only be detected on the surface of the
conidia (Fig. 3c–f). For P. chlamydosporia conidia were
always covered by an extracellular material (Fig. 3b, d
and f). Conidia of neither Forl nor P.c. 123 untreated
with Rhodamine–chitosan showed any autofluorescence
(data not shown).
Discussion
(b)
(c)
Figure 1 Chitosan degrading halos around colonies of the nematophagous fungus Pochonia chlamydosporia (isolate P.c. 123) in culture
media of decreasing content (left to right) amended with 2 mg ml)1
chitosan. (a): PDA, (b) CMA, (c) WA.
affected conidia had the cytoplasm retracted, but still kept
their structure (Fig. 2c–f). Furthermore, a material was
detected surrounding the conidia of P. chlamydosporia
(Fig. 2f).
Tracing of Rhodamine-chitosan in fungal spores
We could see fluorescence inside the conidia of Forl and
P. chlamydosporia (isolate P.c. 123) 15 min after treatment with Rhodamine–chitosan at 0Æ1 mg ml)1 (Fig. 3a
and b). When conidia were treated with sodium azide or
There were great variations in tolerance to chitosan
between the different fungi tested. The plant pathogenic
fungus F. oxysporum f. sp. radicis-lycopersici was the most
sensitive fungus to chitosan. This agrees with previous
results on colony growth of F. oxysporum (Benhamou
et al. 1994; Laflamme et al. 1999). Py. ultimum and
R. solani showed a result similar to Forl. The growth of
V. dahliae increased in PDA amended with either 0Æ5 or
1 mg ml)1 of chitosan. Inhibition was only observed
when the chitosan concentration was raised to 2 mg ml)1.
This suggests that V. dahliae can use chitosan as nutrient
at low concentrations. Xu et al. (2007) also studied the
effect of chitosan on hyphal growth of nine plant-pathogenic fungi and found variation between fungi tested, but
also showed that V. dahliae was the most tolerant of the
plant pathogens tested.
Nematophagous and entomopathogenic fungi were
much less inhibited by chitosan than the plant pathogenic
fungi or Oomycete tested. For most of them, growth in
chitosan-amended media was similar to that in unamended media, or even higher (e.g. in PDA). As for plant
pathogens, growth inhibition by chitosan increased with
the decrease in nutrient concentration of the growth
media used. Growth inhibition of these biocontrol fungi
was, however, significantly lower than that of plant pathogenic fungi. Mycoparasitic fungi were the only exception
among the three types of biocontrol fungi tested, since
they were as sensitive to chitosan as the plant pathogenic
fungi and the Oomycete Py. ultimum.
The low effect of chitosan on nematophagous and
entomopathogenic fungi seemed to be related, at least
partly, with their ability to degrade chitosan. These fungi
showed the highest chitosan-degrading activity, according
to the size of their substrate degradation halos. Extracts
from the halo around P. chlamydosporia (isolate P.c. 123)
were able to degrade FITC-chitosan. This confirmed that
the chitosan-clearing halo occurred as a result of true
enzymatic activity. The fact that for B. bassiana, L. cf.
psalliotae, T. harzianum and T. atroviride no halo was
observed on PDA amended with chitosan could be due to
halos of the same diameter as the colony. From our own
experience, this often happened on CMA amended with
chitosan. However, PDA colonies were more dense and
ª 2007 The Authors
Journal compilation ª 2007 The Society for Applied Microbiology, Journal of Applied Microbiology 104 (2008) 541–553
547
Effect of chitosan on fungi
J. Palma-Guerrero et al.
Table 4 Chitosan degrading activity (CDR indices) of all the fungi in different culture media amended with chitosan (at 2 mg ml)1)
Culture media
Plant paghogenic fungi
Fusarium oxysporum f. sp. radicis-lycopersici
Gaeumannomyces graminis var. tritici
Pythium ultimum
Rhizoctonia solani
Verticillium dahliae
Nematophagous fungi
Pochonia chlamydosporia (P.c. 123)
P. chlamydosporia (P.c. BK69)
P. chlamydosporia (P.c. BK132)
P. chlamydosporia (P.c. BK309)
P. chlamydosporia (P.c. BK392)
P. chlamydosporia (P.c. BK399)
P. chlamydosporia (P.c. 4624)
P. chlamydosporia (P.c. 64)
P. chlamydosporia (P.c. 75)
P. rubescens
Paecilomyces lilacinus
Entomopathogenic fungi
Beauveria bassiana
Lecanicillium cf. psalliotae
Mycoparasitic fungi
Trichoderma harzianum
T. atroviride
PDA
CMA
WA
–
–
–
–
–
–
–
–
–
–
–
–
–
a,b
0Æ35 ± 0Æ04a (23)
a
0Æ39
0Æ22
c
0Æ25
d
0Æ47
a,e
0Æ40
a,b
0Æ39
f
0Æ07
b,c
0Æ28
d,e
0Æ44
g
0Æ62
h
0Æ1
c
±
±
±
±
±
±
±
±
±
±
±
0Æ02a
0Æ02a
0Æ07a
0Æ01a
0Æ03a
0Æ04a
0Æ01a
0Æ10a
0Æ01a
0Æ03a
0Æ02a
–
–
–
–
(18)
(9)
(14)
(13)
(21)
(17)
(23)
(23)
(9)
(6)
(21)
a
0Æ31 ± 0Æ05a (23)
b
0Æ61
0Æ67
b,d
0Æ59
b,c
0Æ67
b,c
0Æ63
d,e
0Æ53
f
0Æ18
b
0Æ60
e
0Æ50
b,c
0Æ70
a
0Æ25
c
h
±
±
±
±
±
±
±
±
±
±
±
0Æ04b (4)
0Æ01b (3)
0Æ06b (3)
0Æ05b (6)
0Æ05b (11)
0Æ03b (3)
0b (13)
0Æ03b (3)
0Æ03b (3)
0Æ07a (2)
0Æ06b (3)
g
0 ± 0a
0Æ35 ± 0a (14)
g
0±0
0±0
g
a
0Æ62
0Æ63
–
b
0Æ37
–
c
0Æ04
c
0Æ04
–
a
0Æ62
a
0Æ67
d
0Æ08
a
± 0Æ03b (14)
± 0Æ01c (6)
± 0Æ02c (6)
± 0c (21)
± 0Æ01c (20)
± 0Æ02c (6)
± 0Æ02a (2)
± 0a (21)
e
0 ± 0a
0Æ05 ± 0b (17)
c
–
–
Chitosan degrading activity (CDR) index Maximum value (day when was obtained). – = halo not observed. Superscripts on the left show differences between different fungi growing in the same culture media (columns). Superscripts on the right show differences for each fungi growing in
different culture media (rows). Values with different letters are significantly different (P < 0Æ05). Each value represents the mean of three
replicates.
with more aerial mycelium than on CMA, thus making
the observation of the halo difficult.
Nematophagous fungi showed the highest ability to
degrade chitosan. This is being confirmed for all isolates,
which have a worldwide distribution, of P. chlamydosporia
tested. These fungi have high chitinolytic activities necessary for degradation of the nematode eggshells (Tikhonov
et al. 2002). The same is true for the entomopathogenic
fungi, which have to breach the cuticle of their hosts,
formed by chitin and protein, and in which chitinolytic
activities have been characterized (St Leger et al. 1996;
Nahar et al. 2004). Therefore, one possible explanation to
the lower effect of chitosan on the growth of nematophagous and entomopathogenic fungi may be that their
chitosan degrading enzymes (including chitinases, deacetylases, deaminases, etc.) are better adapted to degrade the
chitosan, thus preventing them from the toxic effect of
chitosan to fungal cells. This hypothesis will be tested in
future studies. These fungi may even be using chitosan as
nutrient in some cases. Although chitinolytic and chitosanolytic activities have been found in plant pathogenic
548
and mycoparasitic fungi (Shimosaka et al. 1993; Nogawa
et al. 1998), we could not detect degrading halos in the
plant pathogenic fungi or Oomycete tested (except for
V. dahliae). For the mycoparasitic fungi tested the halo of
chitosan degradation was as large as the colony diameter
in chitosan-amended CMA. Since these fungi were clearly
inhibited by chitosan, this suggests that the chitosanolytic
activities of these fungi were insufficient to protect them
against the toxic effect of the chitosan, or that chitosan
degradation product may have caused the inhibition.
Conidial germination was more sensitive to chitosan
than hyphal growth. In the plant pathogenic and mycoparasitic fungi tested, 0Æ01 mg ml)1 chitosan was enough
to arrest conidial germination irreversibly. Conidia of
nematophagous and entomopathogenic fungi were again
the least sensitive to chitosan and consequently their conidia germinated at 0Æ01 mg ml)1 chitosan. For some isolates, germination was similar to untreated controls.
Furthermore, germination was not completely inhibited
by increasing chitosan concentration up to 1 mg ml)1.
P. chlamydosporia isolate P.c. 4624 was the only one that
ª 2007 The Authors
Journal compilation ª 2007 The Society for Applied Microbiology, Journal of Applied Microbiology 104 (2008) 541–553
J. Palma-Guerrero et al.
Effect of chitosan on fungi
Table 5 Spore germination (%) of plant pathogenic and biocontrol
fungi at 0Æ01 and 1 mg ml)1 of chitosan
Chitosan concentration (mg ml)1)
0Æ01
Plant pathogenic fungi
Fusarium oxysporum f.
sp. radicis-lycopersici
Verticillium dahliae
Nematophagous fungi
Pochonia chlamydosporia
(P.c. 123)
P. chlamydosporia (P.c. BK69)
P. chlamydosporia (P.c. BK132)
P. chlamydosporia (P.c. BK309)
P. chlamydosporia (P.c. BK392)
P. chlamydosporia (P.c. BK399)
P. chlamydosporia (P.c. 4624)
P. chlamydosporia (P.c. 64)
P. chlamydosporia (P.c. 75)
P. rubescens
Paecilomyces lilacinus
Entomopathogenic fungi
Beauveria bassiana
Lecanicillium cf. psalliotae
Mycoparasitic fungi
Trichoderma harzianum
T. atroviride
1
a
0 ± 0a*
a
0 ± 0a*
a
0 ± 0a*
a
0 ± 0a*
b
18Æ5 ± 5Æ2a*
c
63Æ7
42Æ2
c,d
74Æ8
d
91Æ0
d
95Æ3
a
0
d
94Æ7
d
97Æ1
c
59Æ8
d
100Æ7
e
±
±
±
±
±
±
±
±
±
±
11Æ0a*
6Æ8a*
18Æ7a*
10Æ4a
10Æ6a
0a*
6Æ9a
8Æ5a
17Æ5a*
19Æ7a
d
98Æ5 ± 3Æ6a
a
0 ± 0a*
a
a
0 ± 0a*
0 ± 0a*
b
6Æ4 ± 2Æ0b*
d,e
65Æ6
55Æ1
e
78Æ9
e
74Æ1
d,e
68Æ7
a
0
d,e
75Æ9
c,f
38Æ7
c
31Æ6
c
30Æ0
d,f
b
±
±
±
±
±
±
±
±
±
±
12Æ0a*
11Æ2b*
11Æ5a*
14Æ9a*
9Æ5b*
0a*
22Æ2a*
19Æ8b*
6Æ8b*
10Æ8b*
11Æ2 ± 4Æ1b*
a
0 ± 0a*
a
a
0 ± 0a*
0 ± 0a*
Spore germination relative to corresponding controls. Superscripts on
the left show differences between different fungi growing under the
same chitosan concentration (columns). Superscripts on the right
show differences for each fungi growing at different chitosan concentrations (rows). Values with different letters are significantly different
(P < 0Æ05). Values with an asterisk show difference from the control
(P < 0Æ05). Each value represents the mean of three replicates.
showed the same sensitivity to chitosan as the plant pathogenic fungi tested. The fact that this was the nematophagous fungus with the lowest chitosanase activity could
explain its sensitivity to chitosan. In our experiments,
spore germination tests were performed with conidia
immersed in aqueous solutions of chitosan and this may
explain the higher sensitivity of the conidia compared
with Plascencia-Jatomea et al. (2003) who added conidia
to Czapek agar medium amended with 1–5 mg ml)1
chitosan.
Transmission electron microscopy showed that chitosan-treated conidia of F. oxysporum f. sp. radicis-lycopersici were severely damaged. The extensive cell membrane
disruption could account for the arrest of spore germination. This ultrastructural damage in conidia is similar to
that found for the hyphae of this fungus, including alterations of the plasma membrane, cell-wall thickening and
cytoplasm aggregation (Laflamme et al. 1999). The disruption of the plasma membrane and the consequences
for the cells are similar also in bacteria (Liu et al. 2004)
and in the Oomycete Phytophthora capsici (Xu et al.
2007). The lower sensitivity of P. chlamydosporia conidia
to chitosan was reflected in the better preservation of
their ultrastructure and the presence of a mixture of
damaged and non-damaged conidia, even at 1 mg ml)1
chitosan. The material surrounding the conidia of
P. chlamydosporia treated with either Rhodamine-labelled
or unlabelled chitosan (Figs 2f, 3b, d and f) is probably
chitosan binding to the extracellular glycoprotein covering
the conidial surface of P. chlamydosporia (Lopez-Llorca
et al. 2002). It appears that the chitosan needs to bind to
the conidial surface of both Forl and P. chlamydosporia to
be able to enter the cells (unpublished data). Whether
this binding is due to hydrophobic, charge or other
mechanisms is presently unknown. The importance of cell
surface charge and hydrophobicity for chitosan binding
and toxicity has been discussed for bacteria (Liu et al.
2004), as well as for hyphal growth and zoospore release
of Phy. capsici (Xu et al. 2007).
Confocal microscopy analysis using Rhodamine-labelled
chitosan showed that T8s chitosan, a polymer of 70 kDa,
could enter the conidia of both Forl and P. chlamydosporia in less than 15 min. Furthermore, the entry of chitosan in the condia was inhibited by either exposure to
sodium azide or 4C pre-incubation. This suggests that
the phenomenon could be an energy-dependent process
rather than one involving passive diffusion (Hoffmann
and Mendgen 1998; Atkinson et al. 2002). Chitosan was
able to enter in the conidia of both fungi, but transmission electron microscopy showed that the two fungi were
not affected in the same way. We therefore propose that
the difference in the effect of chitosan in both fungi could
be explained after the entry of chitosan into the cell. Once
chitosan enters the conidia, it causes membrane disorganization leading to loss of the cellular content in Forl,
whereas in P. chlamydosporia the effect is much smaller,
maintaining the cellular content. A possible explanation
could be that P. chlamydosporia is able to detoxify the
chitosan or dispose of it using, for instance, ABC transporters (Andrade et al. 2000). We have shown in this
paper that there is a relation between the capacity of the
fungi to degrade chitosan and their sensitivity to it. This
suggests that chitosanase activity (and ⁄ or other factors)
could play an important role in protecting the nematophagous and entomopathogenic fungi against chitosan
cell toxicity. Xu et al. (2007) suggested that multiple
modes of action of (oligo)chitosan exist. We are currently
investigating this hypothesis.
The work in this paper on the differential effects of
chitosan on biocontrol fungi and plant pathogens could
help to develop new strategies of biological control based
on these two agents from natural origin.
ª 2007 The Authors
Journal compilation ª 2007 The Society for Applied Microbiology, Journal of Applied Microbiology 104 (2008) 541–553
549
Effect of chitosan on fungi
J. Palma-Guerrero et al.
(a)
(b)
(c)
(d)
(e)
(f)
Acknowledgements
We thank Dr V. Tikhonov (A.N. Nesmeyanov Institute
of Organoelement Compounds, Russian Academy of Sciences) for providing the fluorescently labelled chitosans
and for his advice with the enzymatic assay. We also
550
Figure 2 Effect of chitosan on the ultrastructure of Fusarium oxysporum f. sp. radicis-lycopersici, Forl (a, b) and Pochonia
chlamydosporia (isolate P.c. 123) (c–f) conidia.
(a) Forl control (untreated). Note plasma
membrane pressed to the cell wall and the
structured cytoplasm. (b) Chitosan-treated
(0Æ01 mg ml)1) Forl conidia. Note plasma
membrane retraction, cytoplasm aggregation
and cell content disorganization. (c) P.c. 123
control (untreated). (d) Non-damaged
chitosan-treated (0Æ01 mg ml)1) conidia. (e)
Slightly damaged conidia, cytoplasm retracted
but still keeping its structure. (f) Chitosan-treated conidia (1 mg ml)1). Mixture of slightly
damaged and non-damaged conidia. Note
extracellular material adhering the conidia.
Abbreviations: CW = Cell wall; PM = Plasma
membrane; Cy = Cytoplasm; LB = Lipid body,
EM = Extracellular material. Scale bar = 500 nm
in a and b, and 250 nm in c–f.
wish to thank many colleagues, especially Dr B. Kerry
(Rothamsted Research, UK), for providing fungal strains
for our experiments. Useful advice and comments of
Dr N.D. Read (University of Edinburgh, UK) are greatly
appreciated. We also thank Dr D. Blome (Marine
Bioproducts GmbH, Bremerhaven, Germany) for the
ª 2007 The Authors
Journal compilation ª 2007 The Society for Applied Microbiology, Journal of Applied Microbiology 104 (2008) 541–553
J. Palma-Guerrero et al.
Effect of chitosan on fungi
(a)
(b)
3·45 µm
10·51 µm
(c)
(d)
3·76 µm
8·48 µm
(e)
(f)
10·01 µm
4·51 µm
Figure 3 Confocal laser microscopy of conidia of Fusarium oxysporum f. sp. radicis-lycopersici (Forl) (Fig. 3a, c and e) and Pochonia chlamydosporia (isolate P.c. 123) (Fig. 3b, d and f). (a and b) Conidia of Forl (a) and P.c 123 (b) treated with 0Æ1 mg ml)1 Rhodamine-chitosan for 15 min. The
highest intensity of fluorescence was observed inside the conidia. (c and d) Conidia of Forl (c) and P.c. 123 (d) pre-incubated at 4C for 30 min
and treated with 0Æ1 mg ml)1 Rhodamine-chitosan for 15 min. The highest intensity of fluorescence was observed on the conidial surface. (e and
f) Conidia of Forl (e) and P.c. 123 (f) pre-treated with 75 mmol l)1 sodium azide for 30 min and treated with 0Æ1 mg ml)1 Rhodamine-chitosan
for 15 min. The highest intensity of fluorescence was observed on the surface of the conidia. Inserts show the fluorescence intensity (arbitrary
units) measured along the bars positioned over the conidia.
ª 2007 The Authors
Journal compilation ª 2007 The Society for Applied Microbiology, Journal of Applied Microbiology 104 (2008) 541–553
551
Effect of chitosan on fungi
J. Palma-Guerrero et al.
kind gift of the T8s chitosan used in the experiments
presented in this paper. Dr E. Tzortzakakis, (Plant
Protection Institute, National Agricultural Research
Foundation, Heraklion, Greece), is greatly acknowledged
for skilful technical assistance in the early phases
of this project. This study was supported by the
Spanish Ministry of Education and Science (AGL200405808 ⁄ AGR).
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