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Chapter 2: Isolation, Molecular Characterization and Predatory
Activity of an Indian Isolates of Nematode-Trapping Fungi
2.1 INTRODUCTION
Food is fundamental need of any organism. The increasing population of world
has lead to high demands of food. In 1996 the World Food Summit (WFS) set the
target of ''eradicating hunger in all countries up to 2015". Subsequently, in 2012 UN
announced to ―create a world where no one is hungry‖ (http://www.fao.org/).
However, on the other hand, an infectious disease appears to cause great economic
looses in agricultural production. Several pathogenic bacteria, fungi, viruses and pests
attack important crops worldwide and ultimately the global food production.
Nematodes are small roundworm and there are thousands of species of this pest is
living on the earth. These nematodes rely on different sources for nutrition including
plants, bacteria, fungi, algae, insects etc. and they are one of the components of the
food chain. On the other hand, there are several species which can infect various
plants and animals. Plant-parasitic nematodes feed on plant tissues and cause diseases.
They interfere in agriculture crop production across the globe (Oka et al., 2000). The
root-knot nematodes, Meloidogyne spp. infects a variety of crop plants and it has been
considered the most damaging plant parasitic nematodes (Regaieg et al., 2010). They
damage the root system and thus reduced the growth of crops. Several strategies have
been employed to control these parasitic nematodes (Mukhtar et al., 2014). Among
these, conventional methods included soil sanitation, soil management, organic
amendment, chemical nematicides, use of resistant variety, biological control and
others (Collange et al., 2011). Chemical nematicides are widely used to control the
nematode population due to their instantaneous result although, the excess uses of
nematicides have resulted in the development of resistance in nematode population
and moreover, these chemical are toxic to plant, soil and environment (Anand et al.,
2010). These incidents have directed the attention of agricultural scientists to develop
bioproducts to control the parasitic nematodes. In fact use of biological products to
protect plant form infectious diseases has got momentum in the field of agriculture as
it is an eco friendly and economically feasible approach.
The activities of soil microorganisms shape the soil characteristics as well as
plant health and productivity (Carvalhais et al., 2012). Soil harbours a diverse range of
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fungi and many of them are rivals of nematodes (Singh et al., 2006). Nematode-
trapping fungi form a peculiar trapping structures by which it trap the nematodes and
kill them. These fungi are possible candidate to be used as a biocontrol agent.
Nematophagous fungi nowadays become a research of interest for the scientists all
over the globe to control parasitic nematodes. Many researchers have isolated and
studied their antagonist potential. Species of Arthrobotrys have been found to show
the predatory activity against both plant and animal parasitic nematodes (Carvalho et
al., 2011; Wang et al., 2013). On the other hand, Duddingtonia flagrans has been
extensively studied against animal parasitic nematodes (Nagee et al., 2001; Braga et
al., 2009; Santurio et al., 2011) and it been also reported to produce chlamydospores
on large scale which can undergo gut passage in small ruminants (Ferreira et al., 2011;
Tavela et al., 2013).
As discussed above, increasing population of world has end up in the
chemicalization of agriculture due of high demand of food. Now a day’s pesticides are
extensively used in agriculture to protect crop plants from pathogen. However,
majority of theses pesticides does not seem to be specifically targeting the specific
organism but conjointly have an effect on non target organisms (Johnsen et al., 2001;
Eisenhauer et al., 2009; Cloyd, 2012; Shi et al., 2013). So it may happen that when
one apply nematophagous fungi together with such pesticides, pesticides may interfere
with their performance. Thus, for it to be successfully integrated into field,
nematophagous fungi have to be assess their compatibility with the variety of
fungicides, insecticides and fungicides which are currently widely employed in
agriculture to protect agriculture crop form infectious diseases. Fungi which may
resist such pesticides are often successfully implemented in the field for biocontrol
purpose. In the present work, effect of different fungicides, herbicides and insecticides
on growth of two nematode-trapping fungi i.e. A. conoides and D. flagrans was also
checked under laboratory conditions. The pesticides that we have selected in the
present work are well known and extensively used in agriculture crop protection.
Many researches from India have reported various biological agents to control
root-knot nematodes (Shankar et al., 2011; Sowmya et al., 2012). However, in India
less attention has been paid on biocontrol of plant parasitic nematodes using
nematophagous fungi. In India till date Arthrobotrys oligospora, Dactylaria
brochopaga and Monacrosporium eudermatum have been reported for the control
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Meloidogyne spp. (Singh et al., 2012; Usman and Siddiqui, 2012; Singh et al., 2013a;
Singh et al., 2013b). In the present work, attention has been paid on the control of
Meloidogyne spp. of root-knot nematodes which cause major crop loss. In the present
study, two nematode trapping fungi form agriculture soil of Anand district of Gujarat
was isolated and characterized based on morphology and 18S rDNA gene sequences.
Additionally, their predatory activity against Meloidogyne spp. was investigated.
2.2 MATERIALS AND METHODS
2.2.1 Isolation of nematophagous fungi and its morphological characterization
A total 23 soil samples were collected from the vicinity of tomato roots from various
agricultural fields infected with nematodes in Anand district of Gujarat during
Decembers and January 2011.
2.2.1.1 Harvesting of root-knot nematodes.
Root-knot nematodes used in the present study were collected from infected
roots of tomato plants. Infected roots were rinsed with tap water to remove soil
particles followed by three times washing with sterile water amended with tetracycline
(50µL/mL). After this, roots were chopped into small pieces and dipped into sterile
distilled water containing tetracycline (30μg/mL) for 30 min. Nematodes that were
coming out into the water were collected by centrifugation at 2000rpm for 3 min.
Again nematodes were washed three times with sterilized water containing
tetracycline and collected each time as mentioned above. Throughout the study
whenever nematodes were needed, they were harvested as described above.
2.2.2.2 Isolation of nematode-tapping fungi.
Nematode-trapping fungi were isolated on water agar (Hi-Media, India), pH
7.0 as per the method described by Nagee et al (2001). Briefly, 1g of each soil
samples were sprinkled on 2% water agar plates amended with tetracycline (50μg/mL)
to avoid bacterial contamination. Plates were incubated at 25+1ºC in dark. After 10
days, as different fungi stared growing, 1mL of nematode suspension (~2000-3000
live nematodes) were added to each plate and further incubated. Afterwards plates
were regularly monitored under a light microscope to visualize trapping of nematodes.
Fungi which showed trapping of nematodes were subsequently purified by sub-
culturing on Czapek Dox Agar (CDA) and Corn Meal Agar (CMA). Purified cultures
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were maintained and preserved on 1.7% CMA by inoculating and incubating at
28+1ºC for 6 days. Pattern of trapping structures and morphology of conidiospores
were used to identify the fungi on the basis of morphology according to (Drechsler,
1937; Cooke and Godfrey, 1964). Microscopy and micrometry of fungi were made in
light microscope under magnification 10X and 40X attached with camera (Leica,
MD2500). For further morphological based identification, conidial size was measured
after growing fungi on nine different media described by (Singh et al., 2005). Length
of 10 conidia form each medium was measured and average length was calculated.
Nematode-trapping fungi were partially identified based on their morphology.
2.2.2 Evaluation of predatory activity against living Meloidogyne spp. of
nematodes
Nematode trapping efficiency of isolated fungi was performed in 90 mm
diameter Petri plates containing half strength CMA (0.85%). 8mm diameter agar disk
having mycelial growth form one week old CMA plate previously grow at 28+1ºC
was inoculated upside down on half strength CMA and allowed to grow in dark for 5
days. On 6th
day 200 μL nematode suspension containing ~1000-1100 nematodes
(adult and larvae) were added to each plates and further incubated in dark at 28+1ºC
for 24hr. Plates devoid of fungi were served as control. There were three replicates for
each fungal treatment and control. Over a period of 24hr incubation, plates were
observed in light microscope attached with camera (Leica, MD2500) under 10X and
40X magnifications. After 24 hours, plates were further observed under the light
microscope to evaluate the morphological variation in the fungi in the presence of
living nematodes. After this, plates were flooded with 2 mL normal saline solution to
recover untrapped nematodes. 100 μL of this suspension was deposited on glass slide
6 times in 10mm diameter quarter previously marked and the number of nematodes
was counted under light microscope under 4X magnification. The mean number of
nematodes in these aliquots was used to find the total number of nematodes trapped by
fungi. The results were statistically analyzed by one way ANOVA to compare
parasitism of isolates against a control group.
2.2.3 Optimization of growth conditions
As a fact, environmental factors viz., temperature, nutrient source, pH of soil
as well as other parameters affect the performance of biocontrol agent under the field.
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Optimum growth conditions i.e. suitable medium, pH and temperature was evaluated
for the isolates.
2.2.3.1Effect of media
Effect of nine different media described by (Singh et al., 2005) and other was
assessed to know which medium promotes luxurious growth of our isolates. For all the
media, agar plates were prepared and inoculated centrally with 8mm diameter disc
with mycelial growth from previously grown fungi. Plates were incubated at 28+1ºC
for 5 days. After 5 days, radial diameter of growth was measured and mean from three
plates were calculated and compared.
2.2.3.2 Effect of pH
To investigate which pH supports healthy growth of the isolated nematode-
trapping fungi, isolates were subjected to grow on corn meal agar having different pH
i.e. 4, 5, 6, 7, 8 and 9. The desired pH of medium was adjusted by adding 0.1N NaOH
or 0.1N HCL. Plates were incubated at 28+1ºC for 5 days and effect of pH was
assessed by measuring the radial diameter of growth.
2.2.3.3 Effect of temperature
Both the isolates were further subjected to different temperature conditions for
determination of optimum temperature for growth. Corn Meal Agar (CMA) (pH7) was
prepared, inoculated centrally and incubated at variable temperatures viz., 4+1, 15+1,
25+1, 28+1, 37+1 and 42+1ºC and incubated in the dark for 5 days. Effect of
temperature was evaluated as described above. All the above experiments i.e. effect of
media, pH and temperature were performed in triplicate.
2.2.4 Molecular characterization of fungi
Fungi were identified based on 18S rDNA gene sequencing. For DNA
isolation, 70 mL of 2.4 % potato dextrose medium (PDB) (Hi-media, India) was
prepared in 250 mL Erlenmeyer flask. Medium was inoculated with spore suspension
and allowed to grow on rotary shaker 125rpm at 28+1ºC for 7 days. Mycelia were
harvested by filtering the medium using sterilized Whatman filet paper No.1. After
filtering, approximately 100-150mg of mycelia were crushed to fine powder in
mortar-pestle using liquid nitrogen followed by grinding with glass beads (Sigma) in
lysis buffer containing 0.1M Tris (pH 7.1), 0.3M EDTA (pH 8.0) and 1% SDS. To
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this 70µL of 0.1 % β mercaptoethanol was added per 1 mL of lysis buffer.
Homogenized mycelia were incubated at 70ºC for 1hr. After this the mixture was
centrifuged at 10,000rpm for 10minutes. Supernatant was transferred to new 2mL
microfuge tube and protein was precipitated using equal volume of Tris saturated
phenol (pH 8) and centrifuged at 10,000rpm for 10 minutes. This was followed by
mixing of aqueous phase with an equal volume of phenol-chloroform and again
centrifuged at 10, 000rpm for 10 minutes. Finally DNA was precipitated form the
aqueous phase using double volume of chilled ethanol (75%) and incubated at -20 ºC
for 1hr. Precipitated DNA was palleted by centrifuging at 12,000rpm for 10minutes.
DNA was washed with an absolute alcohol and again collected by centrifuging as
mentioned above. Plates were allowed to air dry and resuspended into 25µL nuclease
free water. Quality of DNA was assessed using gel electrophoresis and quantified
using NanoDrop 1000 spectrophotometer (Thermo scientific, USA). The partial
sequence of 18S rDNA encoding gene was amplified by using the forward 5'
AGGGTTCGATTCCGGAGA 3' and reverse 5' TTGGCAAATGCTTTCGC 3'.
Primers were synthesized from Sigma, India. 25 µL PCR reaction mixture was
prepared as follow.
Table 2.1: PCR reaction mixture used to amplify the 18S rDNA gene sequence.
Reagent Quantity
(µL)
Genomic DNA (50-70ng/µL) 1.0
10X TaqA assay buffer 2.5
Forward primer (10 µM) 1.0
Reverse primer (10 µM) 1.0
dNTPs mix (2.5 mM each) 2.5
Taq DNA polymerase (2.0U/ µL) 0.5
MiliQ water 16.5
Total 25.0
Taq DNA polymerase, 10X TaqA assay buffer and dNTPs were purchased from
Genei, Bangalore, India. PCR conditions was optimized and amplification of the
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target sequence was carried out in thermocycler (Corbett, Korea) with cycling profile
describe below.
Table 2.2: PCR cycling conditions applied to amplify the 18S rDNA gene
sequence.
PCR products were electrophoreses at 100V in submarine system in 1%
agarose gel amended with ethidium bromide dye along with 100 bp DNA marker and
visualized on UV transiluminator. Amplified products were sent to Eurofins
Genomics India Pvt. Ltd., Bangalore for sequencing from both the sides using prime
pair which was used to amplify the target sequence. Sequences were analyzed using
CodonCode aligner v9 (http://www.codoncode.com/aligner/) to make single contig.
Sequences were identified by BLASTn search against NCBI non redundant databse
(http://blast.ncbi.nlm.nih.gov/Blast.cgi) Phylogenetic relationship with 13 different
species of nematophagous fungi having different mechanism to infect/trap nematodes
was conducted by MEGA5.1 (Tamura et al., 2011) using Neighbor-Joining (N-J)
method (Saitou and Nei, 1987). Related nucleotide sequences (Table 2.3) were
downloaded from NCBI nucleotide collection and comparisation was made.
Nucleotide sequences encoding for partial 18S ribosomal small subunit were
deposited in GenBank using Bankit tool
(http://www.ncbi.nlm.nih.gov/WebSub/?tool=genbank).
Temperature °C Time (Minutes) Cycles
94 5 1
94 1
35 58 1
72 1
72 7 1
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Table 2.3: 18S rDNA gene sequences downloaded from NCBI nucleotides
collection for phylogenetic analysis.
Name of fungi Mode of killing
nematodes
Accession No. Reference
Arthrobotrys oligospora
NBAII AO1
Adhesive network JX403728 Direct submission
Orbilia auricolor Adhesive network DQ471001 (Spatafora et al., 2006)
A. musiformis Adhesive network AJ001985 Direct submission
A. robusta Adhesive network AJ001988
Monacrosporium
doedycoides
Adhesive network AJ001994 Direct submission
A. superba Adhesive network EU977561 Direct submission
Dactylella oxyspora Adhesive knob AY902793 (Li et al., 2006)
Gamsyllela gephyropaga Adhesive columns EF445990 (Smith and Jaffee,
2009)
Hirsutella minnesotensis Endoparasitic DQ078757 Direct submission
Pochonia chlamydosporia Egg parasitic EU266591 Direct submission
Verticillium
chlamydosporium
Egg parasitic AJ291806 (Morton et al., 2003)
Paecilomyces lilacinus Egg parasitic FR775537 Direct submission
Nematoctonus concurrens Endoparasitic EF546658 Direct submission
2.2.5 Effect of various pesticides on growth of the isolates
Chemicals
All the experiments were performed in Corn Meal Agar (CMA) 1.7% (Hi-
media, India) pH 7.0. Commercial formulations of different pesticides that are
available in market were used for experiment, only benomyl was purched from Sigma
Aldrich, India.
Fungicides: Tebuconzole (Folicur, 25.9% EC), Mancozeb (85% w/w) and
Carbendazim (Bavistin, 50% WP).
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Herbicides: Glyphosate (Roundup, 41% S.L IPA salt), Methsulfuron methyl (Algrip,
20% WG) and Paraquate trichloride (Grammoxone, 43.8%).
Insecticides: Profenofos (Rocket, 50% EC) and Ethion (Fosmite, 50% EC).
Stock solutions of each pesticide were prepared in sterile distilled water except
benomyl which was dissolved in dimethyl sulfoxide (DMSO) and appropriate volume
was added under aseptic conditions in sterilized CMA so as to achieve the required
final concentrations of active moiety. Fungicides were tested at 10-50 µg/ml
concentrations while herbicides and insecticides were tested at 50, 100, 500 and 1000
µg/ml concentrations. All the plates were inoculated centrally with 8 mm mycelial
disk from CMA plates previously grown at 28+1ºC for 7 days. Control plates contain
only CMA except for benomyl for which, the control pale was amended with DMSO.
Plates were incubated at 28+1ºC for 6 days, and radial growth diameter of fungus
colony in centimeter was measured and % inhibition was calculated by using
following formula
% of growth inhibition = (C-T)/C×100
Where: C= Radial diameter of the fungus growth in control plate.
T= Radial diameter of the fungus growth in treated plate (Johnsen et al., 2001)
.
All the studies were preformed in triplicate and mean values of radial growth diameter
for each concentration were used for calculation.
2.3 RESULTS AND DISCUSSION
2.3.1 Isolation and morphological characterization of nematophagous fungi
Isolation of nematophagous fungi from soil is a little bit complicated due to its
less abundance and over growth of other soil fungi. Water agar is nutritionally poor
medium which permits sluggish growth of organisms so it can be use for isolation of
scarce microorganisms. Two different nematode-trapping fungi were isolated from the
soil samples collected from different fields of Anand district of Gujarat. After 20-25
days of incubation, two different fungi showed trapping of nematodes (Figure 2.1).
Both the fungi were subsequently purified by sub-culturing on CMA/CZP agar plates.
Isolates were designated as RPAN10 and RPAN12. Morphology of conidia and type
of trapping structure forms the basis of identification of nematophagous fungi. Isolate
RPAN12 trap nematodes by means of three dimensional net made up of 3-7 rings
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(Figure 2.2). Initially the trapping device looks like a ring (Figure 2.2a) afterward it
turns in to fully developed net like structure. It produced a mono-septet elongated
conidiospores and the length of conidia range form 19.11-31.61µm with an average
length of 24.13±2.1µm (Table 2.1 and figure 2.4) on erect conidiophores (Figure
2.3c). Conidiophores are unbranched and have up to 18 conidia. These characteristics
are found similar to Arthrobotrys conoides described by (Drechsler, 1937) except for
size of conidia which is reported 30-46 µm and number of conidia on conidiophore
upto 30. Size of conidia is also varying from A. conoides isolate CED (GenBank
accession JN191309) reported from Brazil (Falbo et al., 2013).
Figure 2.1: Nematode trapped on water agar plates by the isolates after
incubation with living nematodes in dark for 20-25 days. (a) & (b) nematodes
trapped into three dimensional networks produced by isolate RPAN12 under 4X
magnification and (c) nematode trapped by ring like structure of isolate RPAN10
under 10X magnification.
Another isolate, RPAN-10 forms ring like trapping structure but not typical
networks likes that of A. conoides (Figure 2.5 a&b). Conidia are simple, single septet
on erect conidiophore (Figure 2.5c) measuring in length range 21.38-53.37µm with an
average length of 33.1±5.71µm (Table 2.1 and figure 2.6). The conidiophore produces
consecutive conidia slightly below and to one side of elder conidia (Figure 2.5d).
These characteristics are analogous to Duddingtonia flagrans reported by (Cooke,
1969) exception in size of conidia. Isolate RPAN10 also produced thick walled
chlamydospores (Figure 2.5c). Based upon these morphological characteristics, isolate
RPANT10 may be D. flagrans.
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Figure 2.2: Morphological variation found in trapping structures of A. conoides
(RPAN12) on corn meal agar under 40X objective. (a) developing trapping
structure and (b), (c), (d), (e) & (f) fully developed adhesive networks.
Figure 2.3: Morphology of conidiospores of A. conoides (RPAN 12) after growing
fungi on CMA at 28ºC for 7 days. (a) & (b) monosepted conidia of A. conoides
under 10X & 40X objective respectively and (c) conidia on conidiophore under
10X objective.
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Figure 2.4: Variation in size of conidia of A. conoides on different media after
incubation at 28°C for 7 days. Upper three from left to right on CDA, CMA &
MM plates and lower three from left to right RM, SDA & YEPSSM.
It is well documented that chlamydospores of D. flagrans can survive gut
passage of rumen and hence can be used as biocontrol agent against animal parasitic
nematodes (Waller et al., 2001; Gomez-Rincon et al., 2006; Assis et al., 2012; Tavela
Ade et al., 2013).
Previously A. oligospora, A. oviformis, A musiformis and D. flagrans have
been isolated from the Gujarat and their predatory activity against animal parasitic
nematodes Haemonchus contortus is reported by (Nagee et al., 2001; Chauhan et al.,
2002b; Sanyal and Mukhopadhyaya, 2003). Here we have first time isolated A.
conoides form the Gujarat region. Moreover, previous studies carried out form the
Gujarat were particularly on parasitism against animal parasitic nematodes, using D.
flagrans and other fungi however, in this study attention have been made on root-knot
using A. conoides and D. flagrans.
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Figure 2.5: Morphological characterization of D. flagrans (RPAN10) after
growing on CZP agar at 28ºC for 7 days. (a) & (b) ring like trapping structure (c)
conidia and chlamydospores under 40X magnification.
Figure 2.6: Variation in size of conidia of D. flagrans on different media after
incubation at 28°C for 7 days. Upper three from left to right on CDA, CMA &
MM plates and lower three from left to right RM, NA & YEPSSM.
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Table 2.4: Variation in size of conidia of A. conoides and D. flagrans on different
media after incubation at 28°C for 6 days.
2.3.2 Predatory activity against living Meloidogyne spp. of root-knot nematodes.
There is an imperative need to develop new technologies for crop protection
against the parasitic nematodes as these tiny creatures are getting resistant against
chemical pesticides and cause great economic loose. Use of biocontrol agents is one of
the prominence methods which could be useful in this prospect. In the present study,
in vitro predatory activity of both the isolates of nematode-trapping fungi was
evaluated against the root-knot nematodes. Predatory activity of both the isolates was
performed against living root-knot nematodes in order to estimate which isolate is
Medium A. Conoides D. flagrans
Length
range (μm)
Average size
conidia (μm)
Length
range (μm)
Average size
conidia (μm)
Corn meal agar 22.27–27.68 25.64±1.24 21.65-47.5 36.1±8.22
Czapek’s agar 21.74–25.32 23.9±1.25 31.17-53.37 40.65±6.53
Sabouraud’s
dextrose agar
21.61–28.25 24.99±2.3 28.88-38.92 29.97±5.37
Potato dextrose agar 20.08–30.8 23.7±3.07 23.26-40.46 33.44±4.77
Richard’s agar 19.11–26.28 23.13±2.04 21.38-38.92 29.97±5.37
Jensen’s agar 23.45–25.31 23.01±1.89 23.57-42.09 31.61±5.64
Yeast extract
peptone soluble
starch agar
20.84–28.46 24.63±2.61 22.28-40.25 33.75±6.39
Martin’s agar 21.78–31.61 25.12±2.66 21.61-30.01 26.16±2.92
Nutrient agar 20.89–27.46 23.01±1.89 28.19-44.8 36.26±6.19
Average 24.13±2.1 33.1±5.71
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better to manage plant parasitic nematodes. After 5 days, ~1000-1100 nematodes were
added to growing fungal plates and predatory efficacy was assessed after 24 hours.
Results showed that very few trapping structures were produced by both the isolates
on half strength CMA after 5 days. The numbers of trapping structures were
drastically increased when nematodes were added to growing fungi. This shows that
in the presence of nematodes, both the isolates got induced. Previously this response
was thought to be because of peptides secreted by nematodes (Pramer and Stoll, 1959;
Pramer and Kuyama, 1963; NORDBRING‐HERTZ, 1973; Nordbring-Hertz, 1977).
However, recently pheromones identified as a ascarocides have been reported by
(Hsueh et al., 2013) which might trigger some signaling pathways in A. oligospora
and ultimately lead to aggressive behavior of this fungus. The response of A. conoides
(RPAN12) was superior compared to D. flagrans (RPAN10). A. conoides started
trapping of nematodes after 5-6 hours of incubation and traps almost 92-95%
nematodes within in 24 hours. Compared to A. conoides, D. flagrans showed very low
trapping efficiency and only 26-29% nematodes were trapped within 24 hours. A
significant difference (P<0.001) in trapping efficiency was observed among control,
A. conoides and D. flagrans (Figure 2.7).
Figure 2.7: Trapping efficacy of isolated nematode-trapping fungi. Bar line
indicate standard devotion of three replicates.
This results make clear that A. conoides is more proficient in trapping of root-
knot nematodes then D. flagrans. The difference in efficiency may be attributed to the
difference in trapping weapons that fungi use to trap nematodes. A. conoides make use
of adhesive network made up of 3-7 rings (Figure 2.2) while D. flagrans produces ring
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like structure but did not form network characteristic like that of A. conoides (Figure
2.5 a&b). Moreover, nematophagous fungi are host specific to some extent. As
previously stated, D. flagrans has been extensively studied against animal parasitic
nematodes rather than plant parasitic nematodes (Vilela et al., 2012; Assis et al., 2013)
whereas A. conoides have been found to be a superior in controlling Meloidogyne
infection. For the first time (Al-Hazmi et al., 1982) reported reduction of Meloidogyne
incognita under microplote and greenhouse conditions. Afterward (Campos and
Campos, 1997; Kalele et al., 2010; Falbo et al., 2013) also said that that A. conoides
can be used for the control of root-knot nematodes. A. conoides is also successful
against animal parasitic nematodes. However, very few researchers studied the effect
of A. conoides on animal parasitic nematodes. Say for example (Maciel et al., 2009)
reported reduction of Ancylostoma spp. by A. conoides. Recently (Paula et al., 2013)
have studied successful trapping of Angiostrongylus cantonensis which causes
eosinophilic meningoencephalitis in humans. The present study reveals the trapping
efficiency of local isolates in plate assay, so further research need to be carried out for
suitable formulation and application methodology for A. conoides so it can serve a
more robust prospective mean to control Meloidogyne spp. infection under the field
applications.
2.3.3 Optimization of the growth conditions
For field application, large scale production of fungi is required. Further,
different abiotic factors especially temperature and soil pH affect the efficiency of
biocontrol agents under field conditions (Hasanzadeh et al., 2012). Here, different
cultural conditions viz., pH, temperature and media for favorable growth of the
isolated nematode-trapping fungi were optimized. Of the 9 different media tested,
Corn Meal Agar (CMA) showed luxurious growth of both the fungi followed by
Martinson’s Medium (MM), Jenson’s Medium (JM), Yeast Extract Peptone Soluble
Starch Medium (EEPSTM) and Remington’s Medium (RM). D. flagrans also grow
well on Czapek Dox Agar (CZP) whereas A. conoides showed moderate growth on
RM as compared to D. flagrans. Both isolates showed slow growth on Sabouraud
Dextrose Agar (SDA), Potato Dextrose Agar (PDA) and Nutrient Agar (NA) (Figure
2.8).
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Figure 2.8: Effect of different media on growth of the isolates. AC-Arhtrobotrys
conoides, DF-Duddingtonia flagrans.
While comparing growth of both isolates, A. conoides was found to be slow
growing compared to D. flagrans. All the media contain different pure as well as
crude carbon and nitrogen sources; still both the fungi were able to grow on all the
media tested. This highlighted the saprophytic nature of nematophagous fungi because
in soil, different carbon and other nutrients are available at varying proportions and
fungi have to manage with this. In case of temperature, both the isolates grew
optimally at 25 and 28ºC. These results are comparable with optimum temperature
reported for A. oligospora (Duponnois et al., 1995; Morgan et al., 1997).
Figure 2.9: Efeect of temperature on Figure 2.10: Effect of pH on growth of
the growth the isolates. of the isolates.
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Stunted growth of both fungi was observed at 42ºC while at 4ºC, isolates were failed
to grow this outcome clearly explained that very high and low temperatures is
inhibitory for both the fungi. A. conoides found to be little bit cold loving than D.
flagrans as it showed enhanced growth at 15 ºC and very poor growth at 37 ºC while
this was reverse in case of D. flagrans (Fig. 2.9). In case of pH, healthy growth of the
isolates was observed between pH 6 to 8. These results are comparable with the
optimum pH reported for A. oligospora (Nagesh et al., 2005) and Pochonia
chlamydosporia (Nagesh et al., 2007). At pH 5 and 9 moderate growth was observed
and no growth was observed at pH 4 (Fig. 2.10). A. conoides showed relatively fair
growth at pH 5 and pH9 than D. flagrans. This result showed that very acidic and
alkaline conditions are not conducive for the growth of both the fungi. These results
are analogous with results of (Hasanzadeh et al., 2012). Thus temperature 25-28 ºC
and pH 6-9 was found to suitable for both the isolates. These parameters should be
kept in mind together with other abiotic and biotic factors at the time of field
application otherwise it may be lead to unanticipated results.
2.3.4 Molecular characterization of the isolates
Nematode-trapping fungi produced different types of trapping devices to catch
and kill nematodes. They make three fundamental types of trapping devices adhesive
knobs, constricting rings and adhesive networks (Rubner, 1996). Previously
nematophagous fungi were identified and classified solely based on the type of
trapping devices and morphology of conidia. However, only morphological base
identification does not lead to concluding results. Knowledge of DNA sequence has
become very essential in any biology study. For the very first time, Ahren and their
colleagues make use of 18S rDNA sequences to study the phylogeny of nematode-
trapping fungi (Ahren et al., 1998). They have concluded that 18S rDNA based
phylogeny is supported with the type of trapping structure produced by fungi.
Afterward (Li et al., 2006) proved the same theory. Yet identification of the
nematophagous fungi using small and large ribosome coding gene sequences is
considered to be more accurate than exclusively morphology based (Yang and Liu,
2006).
A good quality and quantity of genomic DNA (Figure 2.11) was isolated from
both the fungi as described in the methodology.
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Figure 2.11: Genomic DNA isolated form AC- A. conoides and DF- D. flagrans on
1.0 % agarose gel.
Figure 2.12: Amplified 18s rDNA encoding gene partial sequence with 100bp
ladder. AC- A. conoides and DF- D. flagrans.
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Figure 2.13: A Neighbor joining tree showing relationship of our isolates with
other nematophagous fungi.
A 552 and 559bp 18S rDNA ITS gene from both the fungi was amplified
(Figure 2.12) and sequenced. Sequences obtained were further subjected to BLASTn
search against NCBI non-redundant databse. Blast results confirmed that isolates
RPAN-12 is Arthrobotrys conoides and RPAN-10 is Duddingtonia flagrans which
also matches with morphology based data. Sequences were submitted to GenBank
under the accession No. JX979095 and JX979094 for A. conoides and D. flagrans
respectively. The phylogenetic relationship of nematophagous fungi based on 18S
rRNA gene, nuclear loci and elongation factor 1-alpha (EF1-a) sequences is been
reported previously (Ahren et al., 2003; Mayer et al., 2005; Li et al., 2006). Form
these data it can be concluded that, trapping device plays a very important role in
classifying nematophagous fungi. Nematophagous fungi have been classified into
three main group i.e. nematode trapping, endoparasitic and egg/cyst parasitic fungi
based upon the mechanisms they use to trap the nematodes. All these groups form
separate clades when phylogeny of nematophagous fungi is studied on the basis of
18S rDNA gene sequences. Figure 2.13 shows N-J tree, representing the association
of our isolate A. conoides and D. flagrans. Here three different clades with bootstrap
values 98, 51 and 74 is clearly seen, in that nematode-trapping fungi are disconnected
from other nematophagous fungi. A. conoides and D. flagrans both are nematode-
trapping fungi falls in the cluster of nematode-trapping fungi with bootstrap value 98
Chapter 2 2014
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while other fungi which forms either adhesive knob (D. oxyspora) adhesive columns
(G. gephyropaga), endoparasitic or egg parasitic fungi (H. minnesotensis, P.
chlamydosporia, V. chlamydosporium, P. lilacinus and N. concurrens) forms separate
clade (Fig.8). These results are analogous with the previous studies described.
2.3.5 Effect of various pesticides on the growth of isolates
In order to develop a successful biocontrol agent against plants parasitic
nematodes, the antagonist or the predator must sustain the abiotic stresses as well as
should be resistant to diverse pesticides that are readily used by framers to control
other plant pathogens. Further, combination of biological and chemical control agent
also called Integrated Pest Management (IPM) is more effective means to control pest
for sustainable agriculture (Talebi et al., 2008; Unlu et al., 2012). Here in this study,
we also checked compatibility of our isolates with different pesticides that are rapidly
used in agricultural practices.
2.3.6.1 Effect of fungicides
A. conoides and D. flagrans are found to sensitive to all the fungicides tested at
40-50 µg/ml, except D. flagrans which showed a very stunted growth i.e. 92.5%
growth inhibition at 40 µg/mL of mancozeb (Figure 2.14). Among all the fungicides,
carbendazim (Bavistin) found to be more toxic than the others. Carbendazim causes
100% growth inhibition even at 10µg/mL of both the isolates. Isolates were resistant
to benomyl compared to other fungicide tested at 10µg/mL concentration as it causes
only 20 and 18.8% growth inhibition of A. conoides and D. flagrans respectively
(Figure 2.15). Moreover, isolates were partially resistant to tebuconzole at 10 µg/mL
which causes only 32.3 and 26.2% growth inhibition of A. conoides and D. flagrans
respectively (Figure 2.16). On comparing both the fungi, D. flagrans was found to
more resistant to fungicides than A. conoides.
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Figure 2.14: Effect of mancozeb on radial Figure 2.15: Effect of benomyl on
growth of the isolates. radial growth of the isolates.
Figure 2.16: Effect of tebuconazole on radial growth of the isolates.
Various findings have evaluated the adverse effect of fungicides on
economically important fungi. Kubilay and Gokce have reported that, fungicides
(captan, dichlofluanid, iprodione, benomyl, and carbendazim) interfere the
performances of Paecilomyces fumosoroseus (Er and Gökçe, 2004). Khalil et al have
also reported incompatibility of Verticillium lecannii an entomopathogenic fungus
with certain pesticides including mancozeb and benomyl (Khalil et al., 1985). There
are several reports on the adverse effect of fungicides on the growth of various
nematophagous fungi. Grewal and Sohi reported that commonly used pesticides
inhibit the growth of A. conoides and A. oligospora (Grewal and Sohi, 1988).
Chauhan and colleagues have also reported inhibitory effect of carbendazim on A.
musiformis (Chauhan et al., 2002a). Pullen et al reported adverse effect of benomyl
and other fungicides on hyphal growth of Hirsutella rhossoliensis at certain
Chapter 2 2014
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concentrations (Pullen et al., 1990). Furthermore, it has been said that tebuconazole
also affect non target soil microorganisms (Cycon et al.; Bending et al., 2007; Muñoz-
Leoz et al., 2011) and thus influence the soil fertility. In addition, our isolates showed
somewhat similar resistant profile towards mancozeb and carbendazim reported
against A. oligospora (Mohammadi Goltapeh et al., 2010). In this reverence, our
isolates were superior to some extent as isolates showed some degree of resistant to all
the fungicide tested up to 20µg/mL except tebuconazole. Results of the presrnt study
shows that our isolates of nematophagous fungi A. conoides and D. flagrans are
partially resistant to tested fungicides upto 10-20 µg/mL. Thus, once these fungi
applied in the field as a biocontrol agent, such fungicides should be used upto a
precise level otherwise it may interfere with the performance of both the fungi.
2.3.6.2 Effect of Herbicides
Herbicides are chemicals used to inhibit the growth of weeds that grow
unnecessarily with the crops. Both the isolates were resistant to glyphosate and
paraquote dichloride up to 100 µg/mL (Figure 2.17&2.18) whereas, metsulfuryl
methyl was found to toxic even at 50 µg/L (Figure 2.19). A. conoides was most
resistant to glyphosate up to 1000 µg/mL while growth of D. flagrans was inhibited at
500µg/mL concentration (Figure 2.17). Paraquote dichloride a contact herbicide and
Metsulfuryl methyl causes 100% inhibits growth of both the isolates at 500 and 1000
µg/ml respectively (Figure 2.18&2.19). A. conoides was partially resistant to
metsulfuryl methyl at 500 µg/mL showed 85.5% growth inhibition whereas, D.
flagrans failed to grow at this particular concentration (Figure 2.19). In contrast to
fungicides, A. conoides was found to more resistant to herbicides than D. flagrans.
Figure 2.17: Effect of glyphosate on radial Figure 2.18: Effect of paraquate
growth of the isolates. dichloride on radial growth of
the isolates.
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Figure 2.19: Effect of metsulfuryl methyl on radial growth of the isolates.
Poprawaski and Majchorwicz have studied the effect of herbicides on growth
of entomopathogenic fungi and concluded that herbicides interfere with the growth of
fungi (Poprawski and Majchrowicz, 1995). Similar results have been reported by
Paraquat and Abdel-Mallek for cellulose decomposing fungi (Paraquat and Abdel-
Mallek, 1987) and for Phymatotrichum omnivorum by Gunasekaran and Ahuja
(Gunasekaran and Ahuja, 1975). In contrast to these, our isolates of nematophagous
fungi were resistant to all the herbicides upto 100 µg/mL and for some herbicides upto
500 µg/mL. This indicates that these herbicides if applied in the field they may have
not much effect on growth of A. conoides (RPAN12) and D. flagrans (RPAN10).
2.3.6.3 Effect of Insecticides
Among the insecticides, profenofos was found to be more lethal for both the
isolates as 75% growth of both the isolates was diminished at 50 µg/mL concentration
(Figure 2.20). In contrast to this, isolates were resistant to ethion up to 100 µg/mL. A.
conoides was found to more resistant to ethion compare to D. flagrans as at A.
conoides is can survive 500 µg/mL concentration whereas D. flagrans failed to grow
(Figure 2.21). According to Vig et al application of insecticides that they have studied
have no any adverse impact on soil fungi (Vig et al., 2008). In contrast to this, (Asi et
al., 2010) reported significant inhibition of mycelial growth and conidial germination
by various insecticides. Das and Mukherjee have reported some beneficial and
harmful effect of insecticides on soil microorganisms. According to them some
insecticides are detrimental to some organisms whereas the some microorganisms
degrade it and utilize it for the growth (Das and Mukherjee, 2000). These result shows
Chapter 2 2014
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that insecticide ethion can be applied but profenophose can’t be applied together with
these isolates in the field.
Figure 2.20: Effect of profenophose on Figure 2.21: Effect of ethion on
radial growth radial growth of the isolates. growth of the isolates.
The overall results of in vitro study indicate that excluding fungicides and few
insecticides at higher concentration, isolated nematophagous fungi can be successfully
use for the control of nematode in the field as a sustainable biocontrol agent.
However, this in vitro study results do not escort to final conclusion because, these
effects may not necessarily reproduce once applied in the field. The reason is that, in
soil or in field a very complex phenomenon is going on. Further, as said by (Mensin et
al., 2013) different pesticides have different rate of effect on growth of various
nematophagous fungi and at present we have no knowledge regarding the mode of
action of these herbicides and insecticides on fungi.
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2.4 REFERENCES
Ahren, D., Ursing, B.M. and Tunlid, A., 1998. Phylogeny of nematode-trapping fungi
based on 18S rDNA sequences. FEMS Microbiology Letters 158, 179-184.
Al-Hazmi, A., Schmitt, D. and Sasser, J., 1982. Population Dynamics of Meloidogyne
incognita on Corn Grown in Soil in Fested with Arthrobotrys conoides. Journal
of Nematology 14, 44.
Anand, T., Chandrasekaran, A., Kuttalam, S., Senthilraja, G. and Samiyappan, R.,
2010. Integrated control of fruit rot and powdery mildew of chilli using the
biocontrol agent Pseudomonas fluorescens and a chemical fungicide. Biological
Control 52, 1-7.
Asi, M.R., Bashir, M.H., Afzal, M., Ashfaq, M. and Sahi, S.T., 2010. Compatibility of
entomopathogenic fungi, Metarhizium anisopliae and Paecilomyces
fumosoroseus with selective insecticides. Pakistan Journal of Botany 42, 4207-
4214.
Assis, R.C., Luns, F.D., Araujo, J.V., Braga, F.R., Assis, R.L., Marcelino, J.L.,
Freitas, P.C. and Andrade, M.A., 2013. Comparison between the action of
nematode predatory fungi Duddingtonia flagrans and Monacrosporium
thaumasium in the biological control of bovine gastrointestinal nematodiasis in
tropical southeastern Brazil. Veterinary Parasitology 193, 134-140.
Bending, G.D., Rodriguez-Cruz, M.S. and Lincoln, S.D., 2007. Fungicide impacts on
microbial communities in soils with contrasting management histories.
Chemosphere 69, 82-88.
Braga, F.R., Araújo, J.V., Silva, A.R., Araujo, J.M., Carvalho, R.O., Tavela, A.O.,
Campos, A.K. and Carvalho, G.R., 2009. Biological control of horse
cyathostomin (Nematoda: Cyathostominae) using the nematophagous fungus
Duddingtonia flagrans in tropical southeastern Brazil. Veterinary Parasitology
163, 335-340.
Campos, H. and Campos, V., 1997. Effect of timing and forms of application of
Arthrobotrys conoides, Arthrobotrys musiformis, Paecilomyces lilacinus and
Verticillium chlamydosporium in the soil for the control of Meloidogyne exigua
of coffee. Fitopatologia Brasileira 22, 361-365.
Carvalhais, L.C., Dennis, P.G., Tyson, G.W. and Schenk, P.M., 2012. Application of
metatranscriptomics to soil environments. Journal of microbiological methods
91, 246-251.
Carvalho, R.O., Braga, F.R. and Araújo, J.V., 2011. Viability and nematophagous
activity of the freeze-dried fungus Arthrobotrys robusta against Ancylostoma
spp. infective larvae in dogs. Veterinary Parasitology 176, 236-239.
Chauhan, J., Subramanian, R. and Sanyal, P., 2002a. Influence of heavy metals and a
fungicide on growth profiles of nematophagous fungus Arthrobotrys musiformis,
a potential biocontrol agent against animal parasitic nematodes. Indian Journal
of Environment and Toxicology 12, 22-25.
Chauhan, J.B., K., S.P., Mukhopadhyaya, P.N. and Subramanian, R.B., 2002b.
Evaluation of an Indian isolate of Arthrobotrys musiformis, a possible biocontrol
candidate against rounds worms. Journa of Veterinary Parasitology 16, 6.
Cloyd, R.A., 2012. Indirect effects of pesticides on natural enemies. Pesticides—
advances in chemical and botanical pesticides. Intech, Rijeka, Croatia, 127-150.
Chapter 2 2014
Page 58
Collange, B., Navarrete, M., Peyre, G., Mateille, T. and Tchamitchian, M., 2011.
Root-knot nematode Meloidogyne management in vegetable crop production:
The challenge of an agronomic system analysis. Crop Protection 30, 1251-1262.
Cooke, R., 1969. Two nematode-trapping hyphomycetes, Duddingtonia flagrans gen.
et comb. nov. and Monagrosporium mutabilis sp. nov. Transactions of the
British Mycological Society 53, 315-319.
Cooke, R. and Godfrey, B., 1964. A key to the nematode-destroying fungi.
Transactions of the British Mycological Society 47, 61-74.
Cycon, M., Piotrowska-Seget, Z. and Kaczynska, A. Kozdro j, J. 2006.
Microbiological characteristics of a sandy loam soil exposed to tebuconazole
and l-cyhalothrin under laboratory conditions. Ecotoxicology 15, 639-646.
Das, A. and Mukherjee, D., 2000. Soil application of insecticides influences
microorganisms and plant nutrients. Applied Soil Ecology 14, 55-62.
Drechsler, C., 1937. Some hyphomycetes that prey on free-living terricolous
nematodes. Mycologia 29, 447-552.
Duponnois, R., Mateille, T. and Gueye, M., 1995. Biological characteristics and
effects of two strains of Arthrobotrys oligospora from Senegal on Meloidogyne
species parasitizing tomato plants. Biocontrol Science and Technology 5, 517-
526.
Eisenhauer, N., Klier, M., Partsch, S., Sabais, A.C., Scherber, C., Weisser, W.W. and
Scheu, S., 2009. No interactive effects of pesticides and plant diversity on soil
microbial biomass and respiration. Applied Soil Ecology 42, 31-36.
Er, M.K. and Gökçe, A., 2004. Effects of selected pesticides used against glasshouse
tomato pests on colony growth and conidial germination of Paecilomyces
fumosoroseus. Biological Control 31, 398-404.
Falbo, M.K., Soccol, V.T., Sandini, I.E., Vicente, V.A., Robl, D. and Soccol, C.R.,
2013. Isolation and characterization of the nematophagous fungus Arthrobotrys
conoides. Parasitology Research 112, 177-185.
Ferreira, S.R., de Araújo, J.V., Braga, F.R., Araujo, J.M. and Fernandes, F.M., 2011.
In vitro predatory activity of nematophagous fungi Duddingtonia flagrans on
infective larvae of Oesophagostomum spp. after passing through gastrointestinal
tract of pigs. Tropical Animal Health Production 43, 1589-1593.
Gomez-Rincon, C., Uriarte, J. and Valderrabano, J., 2006. Efficiency of Duddingtonia
flagrans against Trichostrongyle infections of sheep on mountain pastures.
Veterinary Parasitology 141, 84-90.
Grewal, P.S. and Sohi, H.S., 1988. A new and cheaper technique for rapid
multiplication of Arthrobotrys conoides and its potential as a bio-nematicide in
mushroom Culture. Current Science 57, 3.
Gunasekaran, M. and Ahuja, A., 1975. Effect of herbicides on mycelial growth of
Phymatotrichum omnivorum. Transactions of the British Mycological Society
64, 324-327.
Hasanzadeh, M., Mohammadifar, M., Sahebany, N. and Etebarian, H.R., 2012. Effect
of cultural condition on biomass production of some Nematophagous fungi as
biological control agent. Egyptian Academic Journal of Biological Sciences 5,
12.
Hsueh, Y.P., Mahanti, P., Schroeder, F.C. and Sternberg, P.W., 2013. Nematode-
trapping fungi eavesdrop on nematode pheromones. Current Biolology 23, 83-
86.
Chapter 2 2014
Page 59
Johnsen, K., Jacobsen, C.S., Torsvik, V. and Sørensen, J., 2001. Pesticide effects on
bacterial diversity in agricultural soils–a review. Biology and Fertility of Soils
33, 443-453.
Kalele, D.N., Affokpon, A., Coosemans, J. and Kimenju, J.W., 2010. Suppression of
root-knot nematodes in tomato and cucumber using biological control agents.
African Journal of Horticultural Science 3, 72-80.
Khalil, S., Shah, M. and Naeem, M., 1985. Laboratory studies on the compatibility of
the entomopathogenic fungus Verticillium lecanii with certain pesticides.
Agriculture, Ecosystems & Environment 13, 329-334.
Li, Y., Jeewon, R., Hyde, K.D., Mo, M.H. and Zhang, K.Q., 2006. Two new species
of nematode-trapping fungi: relationships inferred from morphology, rDNA and
protein gene sequence analyses. Mycological Research 110, 790-800.
Maciel, A., Araújo, J., Campos, A., Lopes, E. and Freitas, L., 2009. Predation of
Ancylostoma spp. dog infective larvae by nematophagous fungi in different
conidial concentrations. Veterinary Parasitology 161, 239-247.
Mensin, S., Soytong, K., McGovern, R.J. and To-anun, C., 2013. Effect of agricultural
pesticides on the growth and sporulation of nematophagous fungi. Journal of
Agricultural Technology 9, 953-961.
Mohammadi Goltapeh, E., Shams-Bakhsh, M. and Pakdaman, B., 2010. Sensitivity of
the nematophagous fungus Arthrobotrys oligospora to fungicides, insecticides
and crop supplements used in the commercial cultivation of Agaricus bisporus.
Journal of Agricultural Science and Technology 10, 383-389.
Morgan, M., Behnke, J.M., Lucas, J. and Peberdy, J.F., 1997. In vitro assessment of
the influence of nutrition, temperature and larval density on trapping of the
infective larvae of Heligmosomoides polygyrus by Arthrobotrys oligospora,
Duddingtonia flagrans and Monacrosporium megalosporum. Parasitology 115,
303-310.
Morton, C.O., Mauchline, T.H., Kerry, B.R. and Hirsch, P.R., 2003. PCR-based DNA
fingerprinting indicates host-related genetic variation in the nematophagous
fungus Pochonia chlamydosporia. Mycological Research 107, 198-205.
Mukhtar, T., Hussain, M.A., Kayani, M.Z. and Aslam, M.N., 2014. Evaluation of
resistance to root-knot nematode (Meloidogyne incognita) in okra cultivars.
Crop Protection 56, 25-30.
Muñoz-Leoz, B., Ruiz-Romera, E., Antigüedad, I. and Garbisu, C., 2011.
Tebuconazole application decreases soil microbial biomass and activity. Soil
Biology and Biochemistry 43, 2176-2183.
Nagee, A., Mukhopadhyaya, P.N., Sanyal, P.K. and Kothari, I.L., 2001. Isolation of
Nematode-trapping fungi with potential for biocontrol of parasitic nematodes in
animal agriculture form ecological niches of Gujarat. Intas polivet 2, 3.
Nagesh, M., Hussaini, S., Chindandaswamy, B., Shubha, M. and Ruby, K., 2007.
Relationship between initial water content of the substrate and mycelial growth
and sporulation of the nematophagous fungi, Paecilomyces lilacinus and
Pochonia chlamydosporia. Nematologia Mediterranea 35, 57.
Nagesh, M., Hussaini, S.S., Chidanandaswamy, B.S. and Biswas, S.R., 2005.
Isolation, in vitro characterization and predaceous activity of an indian isolate of
the fungus, Arthrobotris oligospra on the root-kone nematode, Meloidogyne
incognita. Nematologia Mediterranea 33, 5.
Nordbring-Hertz, B., 1977. Nematode-induced morphogenesis in the predacious
fungus Arthrobotrys oligospora. Nematologica 23, 443-451.
Chapter 2 2014
Page 60
Nordbring‐Hertz, B., 1973. Peptide‐Induced Morphogenesis in the
Nematode‐Trapping Fungus Arthrobotrys oligospora. Physiologia Plantarum 29,
223-233.
Oka, Y., Koltai, H., Bar‐Eyal, M., Mor, M., Sharon, E., Chet, I. and Spiegel, Y., 2000.
New strategies for the control of plant‐parasitic nematodes. Pest Managment
Science 56, 983-988.
Paraquat, I. and Abdel-Mallek, A., 1987. Effect of some herbicides on cellulose-
decomposing fungi in Egyptian soil. Zentralblatt für Mikrobiologie 142, 293-
299.
Paula, A.T.d., Braga, F.R., Carvalho, L.M.d., Lelis, R.T., Mello, I.N.K.d., Tavela,
A.d.O., Soares, F.E.d.F., Junior, A.M., Garcia, J.d.S. and Araújo, J.V.d., 2013.
First report of the activity of predatory fungi on Angiostrongylus cantonensis
(Nematoda: Angiostrongylidae) first-stage larvae. Acta tropica 127, 187-190.
Poprawski, T.J. and Majchrowicz, I., 1995. Effects of herbicides on in vitro vegetative
growth and sporulation of entomopathogenic fungi. Crop Protection 14, 81-87.
Pramer, D. and Kuyama, S., 1963. Symposium on Biochemical Bases of
Morphogenesis in Fungi. Ii. Nemin and the Nematode-Trapping Fungi.
Bacteriology Reviews 27, 282-292.
Pramer, D. and Stoll, N.R., 1959. Nemin: a morphogenic substance causing trap
formation by predaceous fungi. Science 129, 966-967.
Pullen, M., Zehr, E. and Carter Jr, G., 1990. Influences of certain fungicides on
parasitism of the nematode Criconemella xenoplax by the fungus Hirsutella
rhossiliensis. Phytopathology 80, 1142-1146.
Regaieg, H., Ciancio, A., Raouani, N.H., Grasso, G. and Rosso, L., 2010. Effects of
culture filtrates from the nematophagous fungus Verticillium leptobactrum on
viability of the root-knot nematode Meloidogyne incognita. World Journal of
Microbiology and Biotechnology 26, 2285-2289.
Rubner, A., 1996. Revision of predacious hyphomycetes in the Dactylella-
Monacrosporium complex.
Saitou, N. and Nei, M., 1987. The neighbor-joining method: a new method for
reconstructing phylogenetic trees. Molecular Biology and Evolution 4, 406-25.
Santurio, J.M., Zanette, R.A., Da Silva, A.S., Fanfa, V.R., Farret, M.H., Ragagnin, L.,
Hecktheuer, P.A. and Monteiro, S.G., 2011. A suitable model for the utilization
of Duddingtonia flagrans fungus in small-flock-size sheep farms. Experimental
Parasitology 127, 727-731.
Sanyal, P. and Mukhopadhyaya, P., 2003. Influence of faecal dispersal time of
Duddingtonia flagrans chlamydospores on larval translation of ovine
Haemonchus contortus. Indian Journal of Animal Sciences.
Shankar, T., Pavaraj, M., Umamaheswari, K., Prabhu, D. and Baskaran, S., 2011.
Effect of Pseudomonas aeruginosa on the root-knot nematode Meladogyne
incognita infecting tomato, Lycoperiscum esculentum. Academic Journal of
Entomology 4, 114-117.
Shi, Y., Lu, Y., Meng, F., Guo, F. and Zheng, X., 2013. Occurrence of organic
chlorinated pesticides and their ecological effects on soil protozoa in the
agricultural soils of North Western Beijing, China. Ecotoxicology and
environmental safety 92, 123-128.
Singh, R.K., Kumar, N. and Singh, K.P., 2005. Morphological variations in conidia of
Arthrobotrys oligospora on different media. Mycobiology 33, 118-20.
Chapter 2 2014
Page 61
Singh, U., Sahu, A., Sahu, N., Singh, R., Renu, S., Singh, D., Manna, M., Sarma, B.,
Singh, H. and Singh, K., 2013a. Arthrobotrys oligospora‐mediated biological
control of diseases of tomato (Lycopersicon esculentum Mill.) caused by
Meloidogyne incognita and Rhizoctonia solani. Journal of Applied
Microbiology 114, 196-208.
Singh, U.B., Sahu, A., Sahu, N., Singh, R., Prabha, R., Singh, D.P., Sarma, B. and
Manna, M., 2012. Co-inoculation of Dactylaria brochopaga and
Monacrosporium eudermatum affects disease dynamics and biochemical
responses in tomato Lycopersicon esculentum Mill.) to enhance bio-protection
against Meloidogyne incognita. Crop Protection 35, 102-109.
Singh, U.B., Sahu, A., Sahu, N., Singh, R., Singh, D.K., Singh, B.P., Jaiswal, R.,
Singh, D.P., Rai, J. and Manna, M., 2013b. Nematophagous fungi: Catenaria
anguillulae and Dactylaria brochopaga from seed galls as potential biocontrol
agents of Anguina tritici and Meloidogyne graminicola in wheat (Triticum
aestivum L.). Biological Control 67, 475-482.
Singh, U.B., Singh, R. and Srivastava, J., 2006. Prevalence of nematophagous fungi in
the rhizospheric soil in some districts of Uttar Pradesh. Indian Journal of Plant
Pathology 24, 43.
Smith, M.E. and Jaffee, B.A., 2009. PCR primers with enhanced specificity for
nematode-trapping fungi (Orbiliales). Microbial Ecology 58, 117-128.
Sowmya, D., Rao, M., Manoj Kumar, R., Gavaskar, J. and Priti, K., 2012. Bio-
management of meloidogyne incognita and erwinia carotovora in carrot (Daucus
carota l.) using Pseudomonas putida and Paecilomyces lilacinus. Nematology
Mediterr 40, 189-194.
Spatafora, J.W., Sung, G.H., Johnson, D., Hesse, C., O'Rourke, B., Serdani, M.,
Spotts, R., Lutzoni, F., Hofstetter, V., Miadlikowska, J., Reeb, V., Gueidan, C.,
Fraker, E., Lumbsch, T., Lucking, R., Schmitt, I., Hosaka, K., Aptroot, A.,
Roux, C., Miller, A.N., Geiser, D.M., Hafellner, J., Hestmark, G., Arnold, A.E.,
Budel, B., Rauhut, A., Hewitt, D., Untereiner, W.A., Cole, M.S., Scheidegger,
C., Schultz, M., Sipman, H. and Schoch, C.L., 2006. A five-gene phylogeny of
Pezizomycotina. Mycologia 98, 1018-1028.
Talebi, K., Kavousi, A. and Sabahi, Q., 2008. Impacts of pesticides on arthropod
biological control agents. Pest Technology 2, 87-97.
Tamura, K., Peterson, D., Peterson, N., Stecher, G., Nei, M. and Kumar, S., 2011.
MEGA5: molecular evolutionary genetics analysis using maximum likelihood,
evolutionary distance, and maximum parsimony methods. Molecular Biology
and Evolution 28, 2731-2739.
Tavela Ade, O., de Araujo, J.V., Braga, F.R., da Silveira, W.F., Dornelas e Silva,
V.H., Carretta Junior, M., Borges, L.A., Araujo, J.M., Benjamin Ldos, A.,
Carvalho, G.R. and de Paula, A.T., 2013. Coadministration of sodium alginate
pellets containing the fungi Duddingtonia flagrans and Monacrosporium
thaumasium on cyathostomin infective larvae after passing through the
gastrointestinal tract of horses. Research in Veterinary Science 94, 568-752.
Unlu, L., Ogur, E. and Celik, Y., 2012. The importance of integrated pest management
for sustainable agriculture. Journal of Selcuk University Natural and Applied
Science 1, 54-59.
Usman, A. and Siddiqui, M.A., 2012. Effect of some fungal strains for the
management of root-knot nematode (Meloidogyne incognita) on eggplant
(Solanum melongena). Journal of Agricultural Technology 8(1), 213-218.
Chapter 2 2014
Page 62
Vig, K., Singh, D., Agarwal, H., Dhawan, A. and Dureja, P., 2008. Soil
microorganisms in cotton fields sequentially treated with insecticides.
Ecotoxicology and environmental safety 69, 263-276.
Vilela, V.L., Feitosa, T.F., Braga, F.R., de Araujo, J.V., Souto, D.V., Santos, H.E.,
Silva, G.L. and Athayde, A.C., 2012. Biological control of goat gastrointestinal
helminthiasis by Duddingtonia flagrans in a semi-arid region of the northeastern
Brazil. Veterinary Parasitology 188, 127-33.
Waller, P.J., Knox, M.R. and Faedo, M., 2001. The potential of nematophagous fungi
to control the free-living stages of nematode parasites of sheep: feeding and
block studies with Duddingtonia flagrans. Veterinary Parasitology 102, 321-
330.
Wang, J., Wang, R. and Yang, X.-Y., 2013. Efficacy of an Arthrobotrys oligospora N
mutant in nematode-trapping larvae after passage through the digestive tract of
sheep. Veterinary Microbiology 161, 359-361.
Yang, Y. and Liu, X., 2006. new generic approach to the taxonomy of predatory
anamorphic Orbiliaceae (Ascomycotina). Mycotaxon 97, 153-161.