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African Journal of Aquatic Science
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Fungi associated with Eichhornia crassipes in South
Africa and their pathogenicity under controlled
conditions
P Ray
a
& MP Hill
a
a
Depart ment of Zoology and Ent omology, Rhodes Universit y, PO Box 94, Grahamst own,
6140, Sout h Af rica
To cite this article: P Ray & MP Hill (2012): Fungi associat ed wit h Eichhornia crassipes in Sout h Af rica and t heir
pat hogenicit y under cont rolled condit ions, Af rican Journal of Aquat ic Science, 37: 3, 323-331
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African Journal of Aquatic Science 2012, 37(3): 323–331
Printed in South Africa — All rights reserved
Copyright © NISC (Pty) Ltd
AFRICAN JOURNAL OF
AQUATIC SCIENCE
ISSN 1608-5914 EISSN 1727-9364
http://dx.doi.org/10.2989/16085914.2012.712912
Fungi associated with Eichhornia crassipes in South Africa and their
pathogenicity under controlled conditions
P Ray* and MP Hill
Downloaded by [P Ray] at 19:00 16 November 2012
Department of Zoology and Entomology, Rhodes University, PO Box 94, Grahamstown 6140, South Africa
* Corresponding author, e-mail: puja.ray@gmail.com
Eichhornia crassipes Mart. Solms-Laubach (Pontederiaceae), water hyacinth, continues to be the world’s worst
aquatic weed. In South Africa, considerable research has been conducted on biological control agents associated
with water hyacinth, with the release of six arthropods and one fungus, but little is known about the occurrence
and impacts of native phytopathogenic fungi. Nation-wide surveys were conducted in 2010 and 2011 on various
aquatic bodies of South Africa to identify the fungal pathogens associated with water hyacinth. Diseased plant
parts were collected and fungi were isolated and identified. Some 250 isolates belonging to more than 25 genera
were collected. Some of these represent new host records, as well as undescribed taxa. Isolates of Acremonium
zonatum (Sawada) Gams, Alternaria eichhorniae Nag Raj and Ponnappa, Bipolaris hawaiiensis (M.B. Ellis) Uchida
and Aragaki, Fusarium Link, Myrothecium roridum Tode ex Fr. and Ulocladium sp., showed the highest pathogenicity and have the potential to be useful in complementing the ongoing biocontrol programme on water hyacinth in
South Africa.
Keywords: biological control, mycoherbicides, plant pathogens, water hyacinth
Introduction
The aggressive aquatic macrophyte water hyacinth, Eichhornia crassipes Mart. Solms-Laubach (Pontederiaceae),
is a free-floating aquatic plant of South American origin.
Currently, the plant is widely distributed throughout the
tropics and subtropics and ranks as one of the most
notorious aquatic weeds worldwide (Holm et al. 1977,
Gopal 1987). It forms dense impenetrable mats of vegetation in freshwater bodies including rivers, lakes and dams,
and irrigation and flood canals, where it impedes water
flow, irrigation and navigation, and increases eutrophication, biodiversity loss and the mosquito vectors of malaria,
encephalitis and filariasis (Terry 1996, Center et al. 2002).
The problems are most severe in developing countries,
where human activities and livelihoods are closely linked to
the freshwater bodies. In South Africa it was first recorded in
the early 1900s. Since then the weed has become invasive
throughout the country, mainly as a result of human activities (Jacot Guillarmod 1979).
Conventional methods of control rely on mechanical/
manual removal and herbicides and are inadequate and
expensive measures to apply on a large scale. Therefore,
biological control has been considered to be a major,
long-term option for water hyacinth. Among the various
biological control agents of water hyacinth, several
phytopathogenic fungi have been successful (Charudattan
2001). There have been various studies on the isolation,
identification and pathogenicity of fungi associated with
the weed in its native range, as well as in several water
hyacinth infested areas of the world (Freeman et al.
1981, Hettiarachchi et al. 1983, Jimenez and Charudattan
1998, Naseema and Balakrishnan 2001, Daddy et al.
2003, Praveena and Naseema 2004, Okunowo et al.
2008, Ray et al. 2008a) but not in South Africa. Some of
the widely reported fungi infecting water hyacinth include
Fusarium equiseti (Corda) Sacc., Corticium sesakii (Shirai)
Matsumoto, Cephalosporium eichhorniae Padwick,
Rhizoctonia solani Kuhn (Nag Raj and Ponnappa 1970,
Freeman and Zettler 1971), Uredo eichhorniae Gonz Frag
and Cif. (Charudattan and Conway 1975, Charudattan et al.
1976), Alternaria eichhorniae Nag Raj and Ponnappa (Nag
Raj and Ponnappa 1970, Shabana et al. 1995a, 1995b),
Bipolaris stenospila (Drechsler) Shoemaker (Charudattan
et al. 1976), Acremonium zonatum (Sawada) Gams (Rintz
1973, Martyn and Freeman 1978), Cercospora piaropi
Tharp. (= C. rodmanii Conway; Conway 1976a, 1976b,
Sanders and Theriot 1980, Charudattan 1984, Freeman
and Charudattan 1984, Martyn 1985, Charudattan 1996,
Tessmann et al. 2001), Myrothecium roridum Tode ex Fr.
(Okunowo et al. 2008).
Pathogens indigenous to a region, and those that cause
endemic diseases, are ideal candidates for development as
non-classical (augmentative or inundative) biological control
agents (Cuda et al. 2008). While there has been considerable research on arthropod biological control agents of
water hyacinth in South Africa (Coetzee et al. 2011), the
role of indigenous fungal pathogens in the control of water
hyacinth has not been studied. The international mycoherbicide programme for water hyacinth control in Africa
(IMPECCA) was established to provide technical assistance
to national programmes across the African continent for the
African Journal of Aquatic Science is co-published by NISC (Pty) Ltd and Taylor & Francis
Ray and Hill
324
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development of a mycoherbicide to control water hyacinth
(Bateman 2001). The programme involved the International
Institute of Tropical Agriculture (IITA), Benin, the University
of Mansoura, Egypt; the Agricultural Research Council–
Plant Protection Research Institute (ARC–PPRI), South
Africa; the Seed Services of the Department of Research
and Specialist Services (DRSS), Zimbabwe; the Danish
Institute of Agricultural Sciences (DIAS), Denmark; and the
CABI Bioscience, UK. The major objective of this collaboration was to survey, collect and isolate fungal pathogens from
water hyacinth and to study their potential as mycoherbicides against the weed. The promising pathogens that were
shortlisted for potential development as mycoherbicides
from around the continent, according to order of preference,
were A. eichhorniae, A. zonatum, C. piaropi, R. solani, A.
alternata and M. roridum (Bateman 2001). However, the
IMPECCA project was terminated without the discovery of
a suitable fungus for mycoherbicide development, and not
much work has yet been undertaken in South Africa.
0
With the growing interest in the use of fungi for weed
control, the aim of this study was to survey for fungal
pathogens on water hyacinth in South Africa and to test their
pathogenicity against the weed under controlled conditions.
Materials and methods
Survey for collection of fungi associated with water
hyacinth
Water hyacinth leaves with disease symptoms, suspected
to be damaged by fungal pathogens, were collected
from various water hyacinth infested sites around South
Africa during 2010 and 2011 (Figure 1, Table 1) covering
wide geographical areas of the country with different
climatic conditions. Plant and leaf specimens with disease
symptoms were wrapped in layers of dry paper towelling
to enable absorbance of their moisture content to prevent
secondary microbial growth, kept in paper bags and sent to
the laboratory for isolation and pathogenicity testing.
400
800 km
AFRICA
South
Africa
Collection sites
24° S
Water hyacinth sites
Johannesburg
26° S
Durban
30° S
ATLANTIC
OCEAN
East London
Cape Town
34° S
Port Elizabeth
INDIAN OCEAN
18° E
24° E
30° E
Figure 1: Areas of South Africa infested by Eichhornia crassipes, and the collection locations of diseased plant parts from which
phytopathogens were isolated
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African Journal of Aquatic Science 2012, 37(3): 323–331
325
Isolation of pathogens
The diseased leaves were washed thoroughly in running
tap water to remove unwanted soil particles. The isolation
of the potential fungal agents was performed by transferring disease marks on the leaves to media plates. About
2 mm2 cross-sectional segments of the leaves and petiole
were cut from the margins of necrotic or chlorotic lesions
and surface-sterilised by sequential immersion in 70%
ethyl alcohol to improve sodium hypochlorite penetration,
10% sodium hypochlorite (v/v) to eliminate contaminating
superficial propagules and, finally, thrice in sterile distilled
water to eliminate traces of the disinfectants used. The
medium for the isolations was rose bengal chloramphenicol
agar (Biolab, Merck, Gauteng, RSA) and potato dextrose
agar (PDA) (Biolab, Merck, Gauteng, RSA) supplemented with chloramphenicol (10% w/v) in Petri dishes,
and incubated under sterile conditions at 27 °C. Isolations
were also attempted on water hyacinth extract dextrose
agar (WhDA) plates (200 g freshwater hyacinth leaves; 15 g
dextrose [Merck Chemicals Pty Ltd, Gauteng, RSA]; 20 g
agar-agar [Biolab, Merck, Gauteng, RSA]; and 1 000 ml
distilled water). The WhDA was prepared as follows:
freshwater hyacinth leaves were washed in running tap
water and then in distilled water. They were chopped into
small fragments and boiled for 20–25 min in 500 ml distilled
water and filtered through cheesecloth for the collection of
extract. Dextrose and agar-agar were added to this extract
and boiled until transparent.
Culturing and sub-culturing
The fungal species isolated from water hyacinth were
purified by streak-plate and sub-culturing techniques
(Agarwal and Hasija 1986). The growing edges of fungal
colonies isolated were transferred to malt yeast extract
agar (MEA) (Biolab, Merck, Gauteng, RSA) plates. Fungi
were transferred serially until pure cultures were obtained.
Cultures that appeared contaminated with other fungus
were sub-cultured and purified.
Table 1: Water hyacinth infested sites surveyed in 2010–2011 for
the collection of fungi
Frequency (%) Number of isolates in a genus 100
Total no. of isolates
Site
Site
no.
1 Goudini Road, Worcester, Western
Cape
2 Kluitjieskraal, Tulbagh, Western
Cape
3 Kubusi River, Stutterheim, Eastern
Cape
4 Lake Nsezi, Empangeni, KwaZuluNatal
5 Muldersdrift, Johannesburg,
Gauteng
6 Nahoon River, East London,
Eastern Cape
7 Nseleni River, Empangeni,
KwaZulu-Natal
8 Princess Vlei, Cape Town, Western
Cape
9 PPRI, Pretoria, Gauteng
10 Rhodes University, Grahamstown,
Eastern Cape
11 Rietondale, Pretoria, Gauteng
12 Swartkops River, Port Elizabeth,
Eastern Cape
13 Tongaat Sugar Estates, Tongaat,
KwaZulu-Natal
14 Wriggleswade Dam, Stutterheim,
Eastern Cape
Latitude
(°S)
33.64420
Longitude
(°E)
19.29980
33.43628
19.17581
32.59257
27.42184
28.73215
31.98987
26.03555
27.84850
32.97392
27.92570
28.74739
31.96890
34.04351
18.48781
25.67732
33.31022
28.28601
26.51847
25.73142
33.99525
28.22393
25.53375
29.27172
31.35584
32.55905
27.49288
Identification and maintenance of isolates
The purified cultures of all the isolates were numbered and
multiplied on PDA plates. The stock cultures of the microorganisms were maintained on PDA slants supplemented
with 10% WhDA and MEA media and stored at 4–70 °C in
refrigerator. The other slants were kept in the BOD incubator
at 27 ± 10 °C and routinely transferred into fresh slants. The
fungi were identified on the basis of their morphological
growth characteristics, sporulation, conidial measurement
and ability to produce pigmentation on growth media, using
various available literature (Gilman 1959, Barnett 1960, Ellis
1971, 1976, Holliday 1993, Domsch et al. 2007).
Frequency of occurrence of fungal isolates
The fungal genera isolated from various water hyacinth
infested sites were counted for their frequency of occurrence
as compared to the other genera. The isolation frequency of
each genus was expressed as the percentage of the total
number of fungal isolates representing a given genus using
the formula:
Pathogenicity
The fungal isolates were tested for their ability to infect
water hyacinth plants in vitro. The pathogenicity trials
were undertaken in two steps. In the first trial a 2 mm disc
of fungus with its medium was cut from actively growing
culture in Petri plate and placed on a piece of water hyacinth
leaf on moist filter paper in a Petri plate and incubated
in a walk-in BOD incubator at 26 ± 2 °C under a photoperiod of 14:10 h (L:D) for for 5–7 days. The leaf discs were
observed for the development of disease symptoms. The
fungi which caused disease on water hyacinth in this first
round went for a second round of assessment, i.e. whole
plant bioassay (Table 2). In the second trial, young water
hyacinth plants were collected from local water bodies and
grown in water tanks in a polyethylene tunnel at Rhodes
University, Grahamstown, South Africa. They were fertilised
with 15-3-12 N:P:K slow-release fertiliser (Multicote 8, Haifa
Chemicals Israel, RSA [Pvt.] Ltd). A commercial iron chelate
(13% Fe) was also added to the water at a concentration of
2 g per 23 litres of water. They were sprayed with insecticide,
Malathion (Kombat [Pty] Ltd), to keep them free from insect
infestation, as and when required. For the pathogenicity test
healthy individual plants were collected from these tanks,
washed thoroughly in running tap water followed by sterile
distilled water. They were wiped with a cotton swab dipped
in 70% alcohol and placed in small tubs containing tap water,
Ray and Hill
326
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Table 2: Fungi isolated from water hyacinth in South Africa
Fungal isolate
Location of isolation *
Acremonium (Cephalosporium) zonatum (Sawada)
Gams
Acremonium sp.
Alternaria alternata (Fr.) Keissler
Alternaria eichhorniae Nag Raj & Ponnappa
12, 13
6, 10, 12
3
1, 3, 7, 10, 12
Alternaria tenuissima (Nees ex Fr.) Wiltshire
Alternaria sp.
Bipolaris hawaiiensis (M.B. Ellis) Uchida & Aragaki
Chaetomium sp.
Chrysosporium merdarium var. roseum W. Gams
Cladosporium sp.
Clonostachys rosea (Link: Fries) Schroers, Samuels,
Seifert & Gams
Colletotrichum sp.
Cylindrocladium sp.
Epicoccum nigrum Link
Eurotium sp.
Exserohilum rostratum (Drechsler) K.J. Leonard & Suggs
Fusarium avenaceum (Fr.) Sacc.
Fusarium equiseti (Corda) Saccardo
Fusarium moniliforme Sheldon
Fusarium oxysporum (Schlecht)
Fusarium solani (Mart.) Sacc.
Fusarium sp.
Gloeosporium sp.
Macrophoma sp.
Myrothecium roridum Tode ex Fr.
10
14
3
11
7
7, 10
7
6
7
1, 7
14
9
9
6
Nigrospora sp.
Periconia sp.
Pestalotia sp.
Phoma sp.
Pythium sp.
Scopulariopsis sp.
Stemphylium sp.
Trichothecium sp.
Ulocladium sp.
4, 7, 13
1
7, 13
7, 10
10
12
2
13
10
10
3, 4, 5, 6, 7, 10, 11
1
1
11
3, 9, 10, 11, 13, 14
4
Countries from which the isolate has
previously been reported
Australia, India, Mexico, Nigeria, Pakistan, Panama,
Uganda, USA, Sudan
Peru
Australia, Bangladesh, Egypt, Ethiopia, India, Mexico
Bangladesh, Egypt, Ghana India, Indonesia, Kenya,
Nigeria, Thailand, USA, Uganda, South Africa, Zimbabwe
Hong Kong
Mexico, Sri Lanka, USA
Mexico
Egypt
**
India, USA
**
**
**
India, Mexico
**
**
**
Ethiopia, India, Sudan
India, Sudan
India, Australia
India, Australia
India, Mexico, Nigeria, Peru, Sri Lanka, Uganda
**
India
Burma, India, Indonesia, Malaysia, Mexico, Nigeria,
Philippines, Sri Lanka, Thailand, Uganda
Mexico, USA
Mexico, USA
India, Mexico
Brazil, India, Peru, USA, Uganda
Ethiopia, India, USA
**
Ethiopia, Mexico, USA
**
Egypt
* See site numbers in Table 1
** Possible new host records
supplemented with fertiliser when required. Before inoculation, some of the leaves from each plant were injured on
their upper surface by making cut marks with a sterile scalpel
blade.
The fungi that were to be tested for their pathogenicity to
water hyacinth were grown on PDA plates and incubated in
a walk-in BOD incubator. The conidial/mycelial suspensions
were prepared from 21-day-old culture in sterile distilled
water. The conidial and mycelial mass was harvested
by flooding the plates with sterile distilled water and then
scraping the mass with a sterilised spatula. To this, Tween
20 (oxysorbic polyxyethylene sorbitan monoleate) was
added as surfactant at the rate of 0.05 ml per 50 ml of spore
suspension. Spore/mycelial suspensions were applied on
water hyacinth until runoff. Control plants were sprayed
with sterile distilled water containing Tween 20. These
plants were then kept in the walk-in incubator at 27 °C and
80% relative humidity and 12 h photoperiod. The plants
were individually enclosed in plastic bags to create a dew
effect that was conducive to fungal growth. The disease
severity was assessed visually every 24 h for a period of
25 days and the intensity of infection was measured using
a score chart framed by Freeman and Charudattan (1984)
and designated as: – (no symptom: healthy plant), + (mild
symptom: plant showing slight symptoms up to 15% of
leaf area), ++ (moderate symptom: plant showing definite
bigger patches of diseased areas from 16% to 59% of leaf
area) and +++ (severe symptom: enlarged lesions covering
60–100% of leaf area).
Results
Samples of diseased water hyacinth leaves collected
from the field had variable diseased lesions including
leaf spots, necrotic flecks, leaf blights, petiole rot, zonate
lesions of various shapes and sizes and dieback symptoms
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African Journal of Aquatic Science 2012, 37(3): 323–331
327
(a)
(b)
(c)
(d)
(e)
(f)
(g)
(h)
(i)
Figure 2: Diseased Eichhornia crassipes leaves affected by various fungal species: (a) Fusarium oxysporum, (b) Acremonium zonatum and F.
solani, (c) Alternaria alternata, (d) and (e) A. zonatum, (f) Alternaria eichhorniae, (g) Bipolaris hawaiiensis, (h) Ulocladium sp., (i) A. eichhorniae
(Figure 2). About 250 fungal isolates belonging to more
than 25 genera were purified from these diseased plant
parts (Table 2). About 150 isolates were eliminated from
further consideration because they were either contaminated, failed to grow, or belonged to the non-pathogenic
and non-sporulating mycelia sterilia group. Several isolates
showing rapid growth rate on PDA plates, mostly those of
Aspergillus, Penicillium and Trichoderma, were excluded
from further consideration after their initial isolation.
Alternaria Nees was the commonest genus, with more than
30 isolates including three species, A. eichhorniae, A. alternata
and A. tenuissima with a frequency of occurrence of 12.4%
(Figure 3). This was followed by Cladosporium Link with 13
isolates (5.2% occurrence frequency) and Acremonium Link
with 12 isolates (4.8% occurrence frequency). There were
10 isolates of Fusarium Link belonging to five species (F.
avenaceum (Fr.) Sacc., F. oxysporum (Schlecht), F. equiseti
(Corda) Saccardo, F. solani (Mart.) Sacc. and F. moniliforme
Sheldon) with a frequency of occurrence of 3.6%.
The pathogens that caused disease to water hyacinth leaf
pieces in the first trial of pathogenicity testing in the Petri
plate bioassay were further subjected to screening for their
pathogenicity and damage to water hyacinth (Table 3) by
whole plant bioassay. The appearance of symptoms on the
leaves started within 3–6 days of application of inoculum
in some of the viable isolates. These included five isolates
of A. eichhorniae, three of A. zonatum and Fusarium
spp. each, and one of Bipolaris hawaiiensis (M.B. Ellis)
Uchida and Aragaki, M. roridum and Ulocladium Preiss sp.
each. They were found to be highly virulent and severely
damaging to the inoculated water hyacinth leaves. Two
isolates of A. eichhorniae caused a rapid rate of infection
and colonisation on the host plant, especially on the
older leaves. In spite of the severe infection, in all cases
new leaves continued to emerge on most of the plants
and several of the plants survived. For example, one of
the isolates of F. oxysporum caused about 90% damage
to water hyacinth by the 30th day of application, yet on
Ray and Hill
328
60
FREQUENCY (%)
50
40
30
20
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Figure 3: Percentage frequency of occurrence of different fungal genera found on water hyacinth in South Africa in 2010–2011
the 45th day new leaves were observed to emerge, thus
compensating for the damage caused by the fungi earlier.
Discussion
These surveys revealed that there is a rich diversity of
fungal pathogens associated with water hyacinth in South
Africa. Several of these mycobiota share a common link
with those recorded in the centre of origin of the weed,
the Amazon River Basin (Evans and Reeder 2001) and
other countries where water hyacinth has been reported
as a major weed. Several of these pathogens possibly
co-evolved with the host plant and were spread to different
parts of the world with the weed itself. However, the
pathogenicity of the pathogens was variable. For example,
highly virulent isolates of A. eichhorniae have been reported
from Egypt (Shabana et al. 1995a, 1995b) and India (Nag
Raj and Ponnappa 1970), while those previously reported
from the USA (Freeman et al. 1974) and South Africa
(Morris et al. 1999) have proved only weakly pathogenic to
water hyacinth. Cercospora piaropi was reported on water
hyacinth in South Africa (Morris 1990) and was reported
to cause a severe decline of plants on a small farm dam
in Mpumalanga province. The fungus was also imported
from Florida, USA, and introduced into South Africa in 1988
(Morris et al. 1999). But, during the present study, it was not
isolated in this country.
Among the pathogens collected, isolates of A.
eichhorniae, A. zonatum, B. hawaiiensis, F. oxysporum,
F. solani, M. roridum, Ulocladium sp. and two isolates
of Fusarium sp. appeared the most potent. Alternaria
eichhorniae has been extensively studied for biocontrol
potential against water hyacinth (Nag Raj and Ponnappa
1970, Shabana 1997, Shabana et al. 2000). It was
observed to be host-specific to water hyacinth (Nag Raj
and Ponnappa, 1970, Shabana et al. 1995a) and capable
of severely damaging and suppressing this weed (Shabana
et al. 1995a, 1995b, 1995c). However, one of the major
obstacles to the use of A. eichhorniae as a mycoherbicide
for water hyacinth is its requirement for at least 10 h of
dew to allow the applied inoculum to germinate and infect
and, to an extent, to colonise the weed (Shabana et al.
1995a). Rintz (1973) undertook extensive studies on A.
zonatum from the biocontrol perspective and showed that
the pathogen did not seem capable of killing water hyacinth,
or of seriously hindering their prolific growth in the USA.
During the present study, isolates of A. zonatum appeared
substantially effective under greenhouse conditions in this
study, but further studies would be needed to determine its
potential as a biological control agent in the field. Several
authors (Ponnappa 1970, Hettiarachchi et al. 1983,
Liyanage and Gunasekera 1989, Okunowo et al. 2010)
have also reported the potential of M. roridum as a biological
control agent. While the isolates of B. hawaiiensis, F.
oxysporum, F. solani and Ulocladium appear to have good
potential, they still need to be evaluated for host specificity,
virulence, and potential under various climatic conditions.
Although these fungal pathogens have been found to be
present on water hyacinth, not much work has been done
to evaluate their biocontrol potential in South Africa. There
are several difficulties associated with the use of fungi in
the biological control of water hyacinth (Zorner et al. 1993,
Boyetchko and Peng 2004); yet, with more studies, the
use of phytopathogenic fungi as biocontrol agents could
be valuable as these pathogens can cause a significant
reduction in water hyacinth biomass, especially following
natural disease outbreaks, after insect attacks, or when
used as inundative bioherbicide agents. They can be
used to manage invasive weeds in natural areas and in
situations where non-chemical alternatives to weed control
African Journal of Aquatic Science 2012, 37(3): 323–331
329
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Table 3: Evaluation of the impact of various fungi on water hyacinth
Isolate
no.
Fungal
isolate
P-37
P-39
P-112
P-117
P-53
P-59
P-77
P-72
P-30
P-50
P-67
P-75
P-90
P-95
P-99
P-18
P-31
P-44
P-71
P-100
P-38
P-47
P-56
P-62
P-46
P-115
P-134
P-113
P-116
P-104
P-23
P-107
P-113
P-62
P-64
P-65
P-4
P-6
P-2
P-12
P-49
P-35
P-43
P-72
Acremonium zonatum
Acremonium zonatum
Acremonium zonatum
Acremonium zonatum
Acremonium sp.
Acremonium sp.
Acremonium sp.
Alternaria alternata
Alternaria eichhorniae
Alternaria eichhorniae
Alternaria eichhorniae
Alternaria eichhorniae
Alternaria eichhorniae
Alternaria eichhorniae
Alternaria eichhorniae
Alternaria tenuissima
Alternaria sp.
Alternaria sp.
Alternaria sp.
Bipolaris hawaiiensis
Cladosporium sp.
Cladosporium sp.
Clonostachys rosea
Colletotrichum sp.
Epicoccum nigrum
Exserohilum rostratum
Fusarium avenaceum
Fusarium oxysporum
Fusarium oxysporum
Fusarium solani
Fusarium sp.
Fusarium sp.
Fusarium sp.
Gloeosporium sp.
Macrophoma sp.
Myrothecium roridum
Nigrospora sp.
Nigrospora sp.
Pestalotia sp.
Phoma sp.
Phoma sp.
Trichothecium sp.
Trichothecium sp.
Ulocladium sp.
Intensity of
infection
on water
hyacinth*
+++
++
+++
+++
–
+
++
++
++
+++
++
+++
+++
+++
+++
–
+
+
+
+++
–
–
+
–
–
+
+
++
+++
+++
–
+++
+++
–
–
+++
+
–
+
+
–
–
–
+++
Disease
symptoms
on leaves
Zonate leaf spots
Leaf spots
Zonate leaf spots
Zonate leaf spots
No symptoms
Leaf spots
Leaf spots
Leaf spot/blight
Leaf spot
Leaf spot/blight
Leaf spots
Leaf spot/blight
Leaf spot/blight
Leaf spot/blight
Leaf spot/blight
No symptoms
Leaf spots
Leaf spots
Leaf spots
Leaf spots
No symptoms
No symptoms
Leaf spot
No symptoms
No symptoms
Leaf spot
Leaf spots
Leaf spots
Leaf spots
Leaf spots
No symptoms
Leaf spot/blight
Leaf spot/blight
No symptoms
No symptoms
Leaf spots
Chlorotic streaking
No symptoms
Leaf spots
Leaf necrosis
No symptoms
No symptoms
No symptoms
Leaf spots
* Intensity of infection (see: Methods: pathogenicity): – = no
symptom, + = mild symptom, ++ = moderate symptom, +++ = severe
symptom
are needed. Although bioherbicides have been used as the
sole option for the management of certain weeds in several
cases (Daniel et al. 1973, Browers 1986, Kenney 1986,
Riddings 1986, Mortensen 1988), for weeds like water
hyacinth, they are likely to need to be supplemented with
other control options such as in combination with different
fungal pathogens (den Breeyen 1998, Ray et al. 2008b), the
release of insect biocontrol agents (Charudattan et al. 1978,
Denoth et al. 2002, Moran 2005, Yamoah et al. 2011) or
used as a major supplement to low doses of conventional
chemical herbicides (Charudattan 1986, Shearer and
Nelson 2002, Nelson and Shearer 2005).
The present study was carried out indoors in controlled
experimental conditions. A follow-up study is needed that
uses the priority pathogens identified in this paper, but in
open, field-type environments.
Acknowledgements — The Working for Water programme and
Rhodes University are gratefully acknowledged for financial
assistance with this project. We are grateful to Angela Bownes
(ARC–PPRI, Hilton), Anthony King (ARC–PPRI, Queenswood),
Julie Coetzee and Philip Weyl (Rhodes University, Grahamstown),
and Roy Jones (Ezemvelo KZN Wildlife, Richards Bay), for helping
collect the diseased plant parts used for this study.
References
Agarwal GP, Hasija SK. 1986. Microorganisms in the laboratory: a
laboratory guide of microbiology, mycology and plant pathology.
Lucknow, India: Print House.
Barnett HL. 1960. Illustrated genera of imperfect fungi (2nd edn).
Minneapolis, Minnesota: Burgess.
Bateman R. 2001. IMPECCA: an international, collaborative
program to investigate the development of a mycoherbicide
for use against water hyacinth in Africa. In: Julien M, Hill MP,
Center TD, Jianqing D (eds), Biological and integrated control
of water hyacinth, Eichhornia crassipes. Proceedings of the
Second Meeting of the Global Working Group for the Biological
and Integrated Control of Water Hyacinth, Beijing, China, 9–12
October 2000. Australian Centre for International Agricultural
Research (ACIAR) Proceedings 102: 57–61.
Boyetchko SM, Peng G. 2004 Challenges and strategies for
development of mycoherbicides. In: Arora DK (ed.), Fungal
biotechnology in agricultural, food and environmental applications.
New York: Marcel Dekker. pp 111–121.
Browers RC. 1986. Commercialisation of CollegoTM – an industrialist’s
view. Weed Science (Supplement 1) 34: 24–25.
Center TD, Hill MP, Cordo H, Julien MH, 2002. Water hyacinth. In: Van
Driesche R (ed.), Biological control of invasive plants in the eastern
United States. Washington, DC: USDA Forest Service. pp 41–64.
Charudattan R. 1984. Role of Cercospora rodmanii and other
pathogens in the biological and integrated controls of water
hyacinth. In: Thyagarajan G (ed.), Proceedings of the International
Conference on Water Hyacinth, Hyderabad, India, 7–11 February,
1983. Nairobi: Nations Environment Programme. pp 834–859.
Charudattan R. 1986. Integrated control of water hyacinth
(Eichhornia crassipes) with pathogen, insects and herbicides.
Weed Science (Supplement 1) 34: 26–30.
Charudattan R. 1996. Pathogens for biological control of water
hyacinth. In: Charudatta R, Labrada R, Center TD, Kelly-Begazo
C (eds), Strategies for water hyacinth control. Rome: Food and
Agriculture Organization. pp 189–196.
Charudattan R. 2001. Biological control of water hyacinth by using
pathogens: opportunities, challenges, and recent developments.
In: Julien MH, Hill MP, Center TD, Jianqing D (eds), Biological
and integrated control of water hyacinth, Eichhornia crassipes.
Proceedings of the Second Meeting of the Global Working Group
for the Biological and Integrated Control of Water Hyacinth,
Eichhornia crassipes, Beijing, China, 9–12 October, 2000.
Australian Centre for International Agricultural Research (ACIAR)
Proceedings 102: 21–28.
Charudattan R, Conway KE. 1975. Comparison of Uredo
eichhorniae, the water hyacynth rust, and Uromyces pontederiae.
Mycologia 67: 653–657.
Charudattan R, Conway KE, Freeman TE. 1976. A blight of water
hyacinth, Eichhornia crassipes caused by Bipolaris stenospila
Downloaded by [P Ray] at 19:00 16 November 2012
330
(Helminthosporium stenospilum). Proceedings of the American
Phytopathological Society 2: 65.
Charudattan R, Perkins BD, Littell RC. 1978. Effects of fungi and
bacteria on the decline of arthropod-damaged water hyacinth
(Eichhornia crassipes) in Florida. Weed Science 26: 101–107.
Coetzee JA, Hill MP, Byrne MJ, Bownes A. 2011. A review of
the biological control programmes on Eichhornia crassipes (C.
Mart.) Solms (Pontederiacaeae), Salvinia molesta D.S.Mitch.
(Salviniaceae), Pistia stratiotes L. (Araceae), Myriophyllum
aquaticum (Vell.) Verdc. (Haloragaceae) and Azolla filiculoides
Lam. (Azollaceae) in South Africa. African Entomology 19:
451–468.
Conway KE. 1976a. Cercospora rodmanii, a new pathogen of water
hyacinth with biocontrol potential. Canadian Journal of Botany
54: 1079–1083.
Conway KE. 1976b. Evaluation of Cercospora rodmanii as a
biological control of water hyacinth. Phytopathology 66: 914–917.
Cuda JP, Charudattan R, Grodowitz MJ, Newman RM, Shearer
JF, Tamayo ML, Villegas B. 2008. Recent advances in biological
control of submersed aquatic weeds. Journal Aquatic Plant
Management 46: 15–32.
Daddy F, Ladu BMB, Beed FD, Birmin-Yauri YA, Owotunse S.
2003. Surveillance of potential pathogenic fungi associated
with water hyacinth in Lake Kainji, Nigeria. Journal of Aquatic
Sciences 18: 125–130.
Daniel JT, Templeton GE, Smith RJ Jr, Fox WT. 1973. Biological
control of northern jointvetch in rice with an endemic fungal
disease. Weed Science 21: 303–307.
den Breeyen A. 1998. Biological control of water hyacinth using
plant pathogens: dual pathogenicity and insect interaction. In:
Hill MP, Julien MH, Center TD (eds), Proceedings of the First
IOBC Global Working Group Meeting for the Biological and
Integrated Control of Water Hyacinth, 16–19 November 1998,
Harare, Zimbabwe. Pretoria: Plant Protection Research Institute.
pp 75–79.
Denoth M, Frid L, Myers JH. 2002. Multiple agents in biological
control: improving the odds? Biological Control 24: 20–30.
Domsch H, Gams W, Anderson TH 2007. Compendium of soil
fungi (2nd edn). Taxonomically revised by W. Gams, Eching:
IHW-Verlag.
Ellis MB. 1971. Dermatiaceous hypomycetes. Kew, Surrey:
Commonwealth Mycological Institute.
Ellis MB. 1976. More dermatiaceous hypomycetes. Kew, Surrey:
Commonwealth Mycological Institute.
Evans HC, Reeder RH. 2001. Fungi associated with Eichhornia
crassipes (water hyacinth) in the upper Amazon Basin and
prospects for their use in biological control. In: Julien MH, Hill MP,
Center TD, Jianqing D (eds), Biological and integrated control of
water hyacinth, Eichhornia crassipes. Proceedings of the Second
Meeting of the Global Working Group for the Biological and
Integrated Control of Water Hyacinth, Eichhornia crassipes, Beijing,
China, 9–12 October, 2000. Australian Centre for International
Agricultural Research (ACIAR) Proceedings 102: 62–70.
Freeman TE, Charudattan R. 1984. Cercospora rodmanii Conway,
a biocontrol agent for water hyacinth. Florida Agriculture
Experiment Station Technical Bulletin 842. Gainesville, Florida:
Institute of Food and Agricultural Science University of Florida.
Freeman TE, Charudattan R, Conway KE, Cullen RE, Martyn RD,
McKinney DE, Olexa MT, Reese DF. 1981. Biological control of
aquatic plants with pathogenic fungi. Technical Report A-81-1.
Vicksburg, Missouri: US Army Engineer Office Waterways
Experiment Station.
Freeman TE, Zettler FW. 1971. Rhizoctonia blight of water hyacinth.
Phytopathology 61: 892.
Freeman TE, Zettler FW, Charudattan R. 1974. Phytopathogens
as biocontrols for aquatic weeds. Pest Articles and News
Summaries (PANS) 20: 181–184.
Ray and Hill
Gilman JC. 1959. A manual of soil fungi (revised 2nd edn). Calcutta:
Oxford and IBH Publishing.
Gopal B. 1987. Water hyacinth. Amsterdam/Oxford/New York/
Tokyo: Elsevier.
Hettiarachchi S, Gunasekera SA, Balasooriya I. 1983. Leaf spot
diseases of water hyacinth in Sri Lanka. Journal of Aquatic Plant
Management 21: 62–65.
Holliday P. 1993. A dictionary of plant pathogens. New Delhi:
Cambridge University Press.
Holm LG, Pluchnet DL, Pancho JV, Herberger JP. 1977. The
world’s worst weeds: distribution and biology. Honolulu: Hawaii
University Press.
Jacot Guillarmod A. 1979. Water weeds in southern Africa. Aquatic
Botany 6: 377–391.
Jimenez MM, Charudattan R. 1998. Survey and evaluation of
Mexican native fungi for potential biocontrol of water hyacinth.
Journal of Aquatic Plant Management 36: 145–148.
Kenney DS. 1986. De Vine – the way it was developed – an
industrialist’s view. Weed Science (Supplement 1) 34: 15–16.
Liyanage NP, Gunasekera SA. 1989. Integration of Myrothecium
roridum and 2,4-D in water hyacinth management. Journal of
Aquatic Plant Management 27: 15–20.
Martyn RD. 1985. Water hyacinth decline in Texas caused by
Cercospora piaropi. Journal of Aquatic Plant Management 23:
29–32.
Martyn RD, Freeman TE. 1978. Evaluation of Acremonium zonatum
as potential biocontrol agent of water hyacinth. Plant Disease
Reporter 62: 604–608.
Moran PJ. 2005. Leaf scarring by the weevils Neochetina eichhorniae
and N. bruchi enhances infection by the fungus Cercospora piaropi
on water hyacinth, Eichhornia crassipes. BioControl 50: 511–524.
Morris MJ. 1990. Cercospora piaropi recorded on the aquatic
weed, Eichhornia crassipes, in South Africa. Phytophylatica 22:
255–256.
Morris MJ, Wood AR, den Breeÿen A. 1999. Plant pathogens and
biological control of weeds in South Africa: a review of projects
and progress during the last decade. African Entomology Memoir
No. 1: 129–137.
Mortensen K. 1988. The potential of an endemic fungus,
Colletotrichum gloeosporioides f. sp. malvae, for biological
control of round-leaved mallow (Malva pusila) and velvetleaf
(Abutilon theophrasti). Weed Science 36: 473–478.
Nag Raj TR, Ponnappa KM. 1970. Blight of water hyacinth by
Alternaria eichhorniae sp. nov. Transactions of the British
Mycological Society 55: 123–130.
Naseema A, Balakrishnan S. 2001. Bioherbicidal potential of fungal
pathogens of water hyacinth. In: Sankaran KV, Murphy ST,
Evans HC (eds), Alien weeds in moist tropical zones: banes and
benefits. Proceedings of the Workshop, Kerala Forest Research
Institute, Peechi, India, 2–4 November 1999. KFRI and CABI
Bioscience, UK. pp 115–121.
Nelson LS, Shearer JF. 2005. 2,4-D and Mycoleptodiscus terrestris
for control of Eurasian watermilfoil. Journal of Aquatic Plant
Management 43: 29–34.
Okunowo WO, Gbenle GO, Osuntoki AA, Adekunle AA. 2008.
Survey, evaluation and molecular characterization of Nigerian
native fungus for potential biocontrol of water hyacinth.
Abstract book, Centennial American Phytopathological Society
conference, 26–31 July 2008, Minneapolis, USA. Phytopathology.
(Supplement) 98: S115.
Okunowo WO, Gbenle GO, Osuntoki AA, Osuntoki AA, Adekunle
AA. 2010. Media studies on Myrothecium roridum Tode: a
potential biocontrol agent for water hyacinth. Journal of Yeast
and Fungal Research 1: 55–61.
Ponnappa KM. 1970. On the pathogenicity of Myrothecium roridum
– Eichhornia crassipes isolate. Hyacinth Control Journal 8: 18–20.
Praveena R, Naseema A. 2004. Fungi occurring on water hyacinth
Downloaded by [P Ray] at 19:00 16 November 2012
African Journal of Aquatic Science 2012, 37(3): 323–331
[Eichhornia crassipes (Mart.) Solms] in Kerala. Journal of
Tropical Agriculture 42: 21–23.
Ray P, Sushilkumar, Pandey AK. 2008a. Survey and selection of
potential pathogens for biological control of water hyacinth. Indian
Journal of Weed Science 40: 75–78.
Ray P, Sushilkumar, Pandey AK. 2008b. Efficacy of pathogens of
water hyacinth (Eichhornia crassipes), singly and in combinations
for its biological control. Journal of Biological Control 22: 173–177.
Riddings WH. 1986. Biological control of stranglervine in citrus – a
researcher’s view. Weed Science 34: 31–32.
Rintz RE. 1973. A zonal leaf spot of water hyacinth caused by
Cephalosporium zonatum. Hyacinth Control Journal 11: 41–44.
Sanders DR, Theriot EA. 1980. Preliminary evaluation of a
Cercospora rodmanii formulation for the biological control of water
hyacinth (Abstract). In: Proceedings of the 33rd Annual Meeting
of the Southern Weed Science Society, 15–17 January 1980, Hot
Springs, Arkansas. p 193.
Shabana YM. 1997. Formulation of Alternaria eichhorniae, a
mycoherbicide for water hyacinth, in invert emulsions averts dew
dependence. Journal of Plant Disease Protection 104: 231–238.
Shabana YM, Charudattan R, Elwakil MA, 1995c. First record of
Alternaria eichhorniae and Alternaria alternata on water hyacinth
in Egypt. Plant Disease 79: 319.
Shabana YM, Charudattan R, Mohamed EA. 1995a. Evaluation
of Alternaria eichhorniae as bioherbicide for water hyacinth
(Eichhornia crassipes) in greenhouse trials. Biological Control 5:
136–144.
331
Shabana YM, Charudattan R, Mohamed EA. 1995b. Identification,
pathogenicity and safety of Alternaria eichhorniae from Egypt
as a bioherbicide agent for water hyacinth. Biological Control 5:
123–135.
Shabana YM, Elwakil MA, Charudattan R. 2000. Effect of media,
light and pH on growth and spore production by Alternaria
eichhorniae, a mycoherbicide agent for water hyacinth. Journal
of Plant Diseases and Protection 107: 617–626.
Shearer JF, Nelson LS. 2002. Integrated use of endothall and a
fungal pathogen for management of the submerged aquatic
macrophyte Hydrilla verticillata. Weed Technology 16: 224–230.
Terry PJ. 1996. The water hyacinth problem in Malawi and foreseen
methods of control. In: Strategies for water hyacinth. FAO Report
of a Panel of Experts Meeting, 11–14 September 1995, Fort
Lauderdale, Florida. pp 57–72.
Tessmann DJ, Charudattan R, Kistler HC, Rosskopf EN. 2001. A
molecular characterization of Cercospora species pathogenic
to water hyacinth and emendation of C. piaropi. Mycologia 93:
323–334.
Yamoah E, Jones EE, Suckling DM, Bourdôt GW, Walter M,
Stewart A. 2011. Using insects as potential vectors of Fusarium
tumidum to control gorse. New Zealand Entomologist 34: 5–11.
Zorner PS, Evans SL, SD Savage. 1993. Perspectives on providing
a realistic technical foundation for the commercialization of
bioherbicides. In: Duke SO, Menn JJ, Plimmer JR (eds), Pest
control with enhanced environmental safety. American Chemical
Society (ACS) Symposium Series 524(6): 79–86.
Received 1 May 2012, accepted 9 July 2012