Phenotypic and molecular characterization of Cercospora species pathogenic to waterhyacinth

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Phenotypic and molecular characterization of Cercospora species pathogenic to waterhyacinth
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Tessmann, Dauri Jose, 1962-
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Cercospora   ( lcsh )
Cladistic analysis   ( lcsh )
Water hyacinth -- Diseases and pests   ( lcsh )
Plant Pathology thesis, Ph.D   ( lcsh )
Dissertations, Academic -- Plant Pathology -- UF   ( lcsh )
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Thesis:
Thesis (Ph.D.)--University of Florida, 1999.
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Includes bibliographical references (leaves 114-124).
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by Dauri Jose Tessmann.
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Typescript.
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Vita.

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PHENOTYPIC AND MOLECULAR CHARACTERIZATION OF CERCOSPORA
SPECIES PATHOGENIC TO WATERHYACINTH
















By

DAURI JOSE TESSMANN


A DISSERTATION PRESENTED TO THE GRADUATE SCHOOL
OF THE UNIVERSITY OF FLORIDA IN PARTIAL FULFILLMENT
OF THE REQUIREMENTS FOR THE DEGREE OF
DOCTOR OF PHILOSOPHY

UNIVERSITY OF FLORIDA































To lone, my wife.
To my father and mother.














ACKNOWLEDGMENTS


I thank Dr. Raghavan Charudattan, chairman of my graduate supervisory

committee, for his support, patience, guidance, and encouragement through all stages of

my graduate studies. I thank Dr. H. Corby Kistler for his guidance, counsel, and

encouragement. I thank my committee members, Dr. James W. Kimbrough, Dr. Richard

D. Berger, and Dr. Maria Gallo-Meagher for their support and assistance. My special

thanks to Dr. Gerald Benny for his assistance and advice. I also thank Dr. James F.

Preston III for his advice and assistance and Dr. David J. Mitchell for his counsel. I thank

Jim DeValerio for his assistance and friendship. I extend my thanks to Rodney Pettway,

Ellen Dickstein, Kris Beckham, and Eldon Philman for their help with DNA techniques,

fatty acid analysis, spectral analysis, and greenhouse work. I thank Dr. Erin N. Rosskopf,

Dr. Manjunath Keremane, Dr. Margaret Smither-Kopperl, Dr. Wellington Pereira, and

Dr. Lisias Coelho for their help, support, and friendship. I thank Mr. Jay Harrison for his

advice with statistical analysis. My special thanks to Dr. Jose C. Dianese and his wife

Heloisa Dianese, for having encouraged me to continue my studies and pursue this

degree. I also thank Dr. Robinson A. Pitelli for his support and friendship. I extent my

appreciation to my friends and classmates and laboratory colleagues, Dr. Jugah Kadir, S.

Chandramohan, Camilla Yandoc, Angela Vincent, Dr. Yasser Shabana, Dr. Gabriela

Wyss, and Matt Petterson, in the plant pathology department. I thank the Brazilian

government and its agency CNPq, for my scholarship, the Florida/Brazil Institute for a fee








waiver, and the Universidade Estadual de MaringA for providing me a study leave to

pursue this degree. Finally, I extend my sincere thanks to Dr. George Agrios, chairman of

the plant pathology department, and the official staff of the department.














TABLE OF CONTENTS
page


ACKNOWLEDGMENTS....................................................................................iii

LIST OF TABLES .................................................................................................... vii

LIST OF FIGURES.................................................................. ..............................viii

ABSTRACT.................. .......................................................................... x

CHAPTERS

1 INTRODUCTION........................................................................................ 1

Reproductive Biology, Origin, and Dispersal of Waterhyacinth............................... 2
Biologial Control of Waterhyacinth .................................................. ...................... 5
The Species of Cercospora as Agents for Biological Control.................................. 11
R research R ationale ................................................................................................ 13

2 A MOLECULAR CHARACTERIZATION OF CERCOSPORA SPECIES
FROM WATERHYACINTH...................................................................... 15

Introduction ................................................................................................ 15
Materials and Methods ................. ........................................................... 18
Fungal Isolates..................................... ................. ............... 18
D N A Isolation ................................................................................................. 19
DNA Amplification and Sequencing ...................................... ........................ 20
Southern Blot Analysis of the 3-Tubulin Gene.................................................. 23
Phylogenetic Analysis ....................................................................................... 23
Results ........................................... ....................... ....................... 25
Identification of Isolates.................................................................................. 25
Phylogenetic Analysis ...................................................................................... 29
Discussion ................................................................. .......................... 52

3 PATHOGENIC VARIABILITY AND BIOCHEMICAL
CHARACTERIZATION OF CERCOSPORA SPECIES FROM
W ATERHYACINTH ....... ........................ ........................................................61

Introduction ....................................... ...................... ..... ......... 61








M materials and M ethods................................................................... ....................... 63
Fungal Isolates and Cultural Characteristics................................................... 63
Toxin A analysis ................................................................................................ 64
Pathogenicity Test and Virulence Analysis..................................................... 67
Fatty Acid Analysis ...................... ......................... 69
Results .......................................... .............................................. 71
Cultural Characteristics................................................................................. 71
Virulence and Toxin Analysis........................................................................ 72
Fatty Acid A analysis .............................. ........................................................... 84
Discussion ............................. ...... .................... ........................ 92
Variability of Cultural and Pathogenic Traits ................................................... 92
Fatty Acid Analysis .......................................................................................... 95

4 SUMMARY AND CONCLUSIONS...... .... .......................................................... 97

APPENDICES

A SEQUENCE ALIGNMENTS OF ELONGATION FACTOR-la ...................... 101

B SEQUENCE ALIGNMENTS OF P-TUBULIN GENE..................................... 104

C SEQUENCE ALIGNMENTS OF HISTONE 3 GENE .................................... 108

D SEQUENCE ALIGNMENTS OF rDNA 5.8S GENE AND INTERNAL
TRANSCRIBED SPACER REGIONS............................................................ 111

LIST OF REFERENCES .................... ................. ........................... 114

BIOGRAPHICAL SKETCH......................................................................... 125














LIST OF TABLES


Table page

2-1 Designations, origins, and morphological characteristics of conidia of the
isolates of Cercospora species from waterhyacinth used in this study .............. 26

2-2 Conidial size and morphology of Cercospora piaropi and Cercospora
rodmanii recorded in the literature ........................................ .......................... 27

2-3 Sequencing results for the data sets aligned with CLUSTAL 1.7W and
analyzed using PAUP version 4.0b I.......................................... ...................... 30

2-4 Maximum likelihood comparisons of tree topologies obtained for
elongation factor-let, P-tubulin, and histone 3 sequences...................................47

3-1 Designations and geographic origin of the isolates of Cercospora piaropilC.
rodmanii analyzed in this study ............................................... ....................... 65

3-2 Mycelial growth rates of isolates of Cercospora piaropi/C. rodmanii from
w aterhyacinth, in m m day .................................................................................. 74

3-3 Geographic origin, color of pigments produced in axenic culture,
production of plant pathogenic toxins, and virulence of isolates of
Cercospora piaropi/C. rodmanii ............................................. ....................... 76

3-4 Production of cercosporin by isolates of Cercospora piaropilC. rodmanii
from waterhyacinth, in gtg per g wet mycelium.................................................... 81

3-5 Correlation matrix among some physiological traits of 55 isolates of
Cercospora piaropi/C. rodmanii from several geographical locations .............. 86

3-6 Fatty acid composition of Cercospora spp. isolates from 4-day-old mycelia........ 88

3-7 Fatty acid composition of Cercospora spp. isolates from 5-day-old mycelia........ 89

3-8 Fatty acid composition of Cercospora spp. isolates from 6-day-old mycelia........ 90

3-9 Canonical discriminant analysis of isolates of Cercospora spp. based on
fatty acid profiles with three different harvesting times of mycelia.................... 91














LIST OF FIGURES


Figure page

2-1 Maps of the elongation factor-la, 0-tubulin, and histone-3 genes; and of
ribosomal DNA region (rDNA)............................................... ........................ 22

2-2 Conidia of Cercospora from waterhyacinth with truncate (A) and obconic
(B) bases (x400)............................................... ....................... ...................... 28

2-3 Tree length distribution for elongation factor-la, 1-tubulin, and histone 3
datasets based on 10,000 random trees................ ........................... 31

2-4 Results of the partition-homogeneity test implemented in PAUP*4.0b 1............. 33

2-5 Most-parsimonious tree inferred from elongation factor-I a gene..................... 34

2-6 Maximum likelihood tree inferred from elongation factor-la gene...................... 35

2-7 Neighbor-joining tree on distances derived from sequences of the
elongation factor-I a gene ........................................................ ....................... 36

2-8 Most-parsimonious tree inferred from P-tubulin gene......................................... 38

2-9 Maximum likelihood tree inferred from 0-tubulin gene.................................. .... 39

2-10 Neighbor-joining tree on distances derived from sequences of P-tubulin
gene ....... .................. .... ....................................... ........................... 40

2-11 Sequences of amplified segments of P-tubulin gene for the isolates 2619
and WH83. ............................................................................ 41

2-12 Southern blot analysis of the P-tubulin gene of Cercospora species from
w aterhyacinth ................................................................................................. 43

2-13 One of three equally parsimonious trees inferred from histone 3 gene. ................44

2-14 Maximum likelihood tree inferred from histone 3 gene...................................... 45








2-15 Neighbor-joining tree on distances derived from sequences of the histone 3
gene..................................................... .............................. .......... 46

2-16 Strict consensus of 24 most-parsimonious trees of length 246 based on
parsimony analysis of combined elongation factor-I a, P-tubulin, and
histone 3 datasets ..................... .. .......................... 49

2-17 Maximum likelihood tree inferred from the combined elongation factor-la,
P-tubulin, and histone 3 datasets.............................................. ........................... 50

2-18 Neighbor-joining tree on distances inferred from the combined elongation
factor-la, P-tubulin, and histone 3 datasets ........................ ......................51

3-1 An example of differences in colony characteristics of Cercospora
piaropilC. rodmanii from waterhyacinth after growth for 7 days on PDA............ 73

3-2 Isolates of C. piaropi/C.rodmanii plotted in a descending order by their
virulence in waterhyacinth ..................................................... ......................... 79

3-3 Ultraviolet spectra of crude extracts from the yellow pigment-producer,
isolate WHK (A), and the reddish-purple pigment-producer, isolate BA57
(B), compared to the standards cercosporin (C) and beticolin-1 (D) in ethyl
acetate. ........ ...... .......................................................... 83

3-4 Crude extracts of isolates of Cercospora piaropilC. rodmanii resolved on
thin layer chromatograms under long-ultraviolet light........................................ 85














Abstract of Dissertation Presented to the Graduate School
of the University of Florida in Partial Fulfillment of the
Requirements for the Degree of Doctor of Philosophy

PHENOTYPIC AND MOLECULAR CHARACTERIZATION OF CERCOSPORA
SPECIES PATHOGENIC TO WATERHYACINTH

By

Dauri Jose Tessmann

May 1999

Chairman: R. Charudattan
Major Department: Plant Pathology

Phylogenetic relationships among isolates of Cercospora species pathogenic to

waterhyacinth, collected from several geographic regions of the weed, were examined by

using partial DNA sequences from three protein-coding genes, elongation factor-la, 3-

tubulin, and histone 3. Each gene included at least one intron area. In cladograms from

individual, as well as from combined datasets for 14 isolates, with the maximum

parsimony, maximum likelihood, and neighbor-joining methods, two statistically well-

supported clades were found: a major clade-grouping of isolates from Brazil, Venezuela,

Mexico, Florida (United States), South Africa, and Zambia; and a minor clade-grouping

of isolates from Texas (United States). In the grouping pattern of the phylogenetic trees,

most of the isolates have spread together with the plant host, from its center of origin in

South America. The isolates grouped together by DNA analysis had distinctive

differences in colony color, pigmentation color, and intensity, growth rate, cercosporin








production, and virulence. In addition, the rDNA region containing ITS1, ITS2, and the

5.8S gene were invariant even when compared with C. beticola as the outgroup. In the

discriminant analysis, the fatty acids methyl ester (FAME) profiles had reduced resolution

for differentiating populations and species among isolates of C. piaropi/C. rodmanii and

the level of resolution was influenced by the age of the colonies. The grouping defined

by discriminant analysis of FAME profiles had no relation to the grouping defined by

DNA analysis. Shape and dimensions of conidia were unreliable criteria for taxonomic

differentiation of isolates that composed the two clade-groupings. In addition, the isolate

that typified C. rodmanii did not show differences in DNA sequence in relation to the

other isolates grouped in the major clade, including some that had conidial size and

morphology typical of C. piaropi. Therefore, the separation of these species, besides of

not having strong phenotypic support, did not also have support from the phylogenetic

analysis. Consequently, the description of the species C. piaropi is emended herein to

include C. rodmanii as a synonym.














CHAPTER 1
INTRODUCTION

The use of plant pathogens as agents of biological control has been an effective

alternative to control several important weeds (Adams, 1988; Charudattan, 1988;

Charudattan, 1991; Charudattan and Walker, 1982; TeBeest, 1996; Templeton and

TeBeest, 1979). Many research efforts have been applied during the last two decades to

develop and implement biological control of the aquatic weed, waterhyacinth (Eichhornia

crassipes [Mart.] Solms), with plant pathogens (Charudattan, 1990). These efforts

included the search for highly virulent pathogens in the plant's native and adventive

ranges; evaluation of the potential of these pathogens for biological control, including

studies about their host ranges and life cycles; and development of technology to use

them as bioherbicides.

Waterhyacinth, an aquatic plant indigenous to lowland tropical South America

(Penfound and Earle, 1948; Barrett and Forno, 1982), has during the last hundred years

become one of the most noxious weeds in many tropical and subtropical regions of the

world. The methods used to control this weed include the use of chemical herbicides,

physical removal with mechanical harvesters, and biological control with insects, mites,

and plant pathogens (Harley and Forno, 1990; Charudattan, 1990; Murphy and Barrett,

1990; Gallagher and Haller, 1990; Center, 1996a). Among these control measures,

chemical herbicides and mechanical removal have been the best short-term solutions for

those areas that require immediate removal of populations of this weed. The problem is








that these measures need to be used continuously to keep infested areas under control.

Moreover, the use of mechanical removal has been very expensive, and the continued use

of herbicides on aquatic environments has raised public concern about the harmful side

effects of chemicals on wildlife, agriculture, and human health. Furthermore, many

developing countries can not afford the recurrent expenses with chemical herbicides and

they usually are less prepared than developed countries in technology and regulatory

safeguards to apply chemical herbicides in aquatic environments. For all these reasons,

the development and implementation of biological control has been considered

indispensable for a long-term solution to this weed problem (Labrada et al., 1996).

Reproductive Biology, Origin, and Dispersal of Waterhvacinth

Eichhornia crassipes is the only species in the genus Eichhornia that has become

a noxious, aggressive weed although a few other species may have similar tendencies.

Eichhornia is an unnatural or polyphyletic group (Kohn et al., 1996) that comprises eight

species of freshwater aquatics in the monocotyledonous family Pontederiaceae. This

family includes the North American pickerelweed, Pontederia cordata L. All species

belonging to the genus Eichhoria, with the exception of the exclusively African E. natans

(Beauv.) Solms, are native to the New World tropics (Barrett, 1988).

Studies of evolution and dispersal of Eichhornia species around the world have

been based mostly on polymorphism of their floral organs (Barrett, 1977; Barrett, 1988).

Flowers of E. crassipes can be divided into three sexual types that differ in the length and

position of their reproductive organs, the male stamens and the female pistils. These

types, or floral morphs, are distinguished by their long, medium, or short styles, i.e., the








prolongation of the ovary. Hence, E. crassipes is described as tristylous plant (Barrett,

1977).

From a global perspective, populations of the three style morphs have been found

only in the lowland tropical South America. Outside the native range of this plant, only

the long- and medium-style morphs have been found, with predominance of the medium-

style morph (Barrett, 1977; Barrett and Forno, 1982). The center of origin of E. crassipes

was suggested to be located in the lowland tropical South America, the Amazon basin,

because the greatest diversity of Eichhornia species has been recorded in this region

(Barrett, 1988) and this is the only region were populations of E. crassipes having three

morphs are found (Barrett and Forno, 1982).

Waterhyacinth was introduced into many countries by man for ornamental

purposes because of its beauty: an attractive plant with clusters of violet and yellow

flowers perched atop floating rosettes of bulbous green leaves. This plant is easily spread

because its rosettes of floating leaves are held together only by delicate horizontal stolons

which can easily break apart. In the newly invaded areas, this plant has been a very

effective colonizer because of its free-floating habit, its capacity for rapid vegetative

propagation, and the absence of its natural enemies (Barrett and Forno, 1982; Barrett,

1988). Because growth of waterhyacinth is directly related to the level of available

nutrients in the water in which the plant is growing (Chadwick et al., 1966; Wahlquist,

1972), the explosive growth rate of this weed has been observed mostly in water bodies in

process of eutrophication. Such waters are common particularly in habitats that are

disturbed by human activities (Labrada et al., 1996).








In the 1970s, some misconceptions about the reproductive biology of

waterhyacinth were clarified. Because early investigators did not observe seeds and

seedlings in nature, and based on the generalization that plants that grow exclusively by

vegetative propagation over long periods often lose their ability to reproduce sexually,

many investigators assumed that clones of waterhyacinth were sexually sterile and could

not regenerate from seed. However, later evidence showed the contrary. Barrett (1980a;

1980b) observed that waterhyacinth reproduces sexually both in the native and adventive

ranges of its distribution. The extent of sexual reproduction and its contribution to the

spread of the weed, however, varies greatly in different regions. A number of factors

such as climate, pollinating agents, and factors that affect seed germination have been

considered responsible to limit the efficiency of sexual reproduction. Albeit seed

production may occur in many regions, not all of these regions are favorable for seed

germination and production of seedlings. This may happen because the seeds produced

by waterhyacinth are usually deposited at the bottom of ponds and rivers where they can

remain viable for several years. For germination and seedling development, cyclic

changes in the water level of rivers and ponds must occur, causing periodic drought at

their margins. This is a typical phenomenon observed in the lowlands of tropical South

America. However, in some of the adventive areas, the absence of flooding and drought

cycles may reduce the role of seeds in the spread of this plant.

The problems caused by this plant in areas where it becomes a weed are many.

They include clogged irrigation canals, blocked waterway transport routes, water losses,

reduction in fish population, and destruction of wild life habitats (Pieterse, 1990). The

United States of America was one of the first and foremost countries that faced dramatic








problems caused by this weed and is the country that has accumulated the richest

experience to overcome these problems. Waterhyacinth was first observed in the US

after the Civil War, however the earliest authentic account details its introduction at the

1890 Cotton Centennial Exposition in New Orleans, Louisiana (Penfound and Earle,

1948). Since then and during the following decades, this plant became a very noxious

weed in the Southeast and in California, where it caused a massive negative impact on

human and economic activities in these areas. These problems were reduced only after

the mid-1970s as a result of the introduction and establishment of some the weed's

natural enemies and intensive weed-management programs with herbicides (Center,

1996b; Confrancesco, 1996; Haller, 1996). Other countries that have had long struggles

against this weed include South Africa, Australia, and India.

In recent decades, the problems caused by this weed have increased in magnitude

in many African, Asian, and Latin American countries, mainly in those areas where

human activities have increased (Labrada et al., 1996). In Africa, the rapid spread of

waterhyacinth and the increased problems observed during the last decade in Uganda,

Malawi, Ivory Coast, Benin, and Nigeria have called for urgent international action

(Charudattan et al., 1996). Even though in Brazil waterhyacinth is native to the Amazon

basin, it has been a weed problem but only in water reservoirs located near highly

populated areas (Dr. R. Pitelli, UNESP, Jaboticabal, Brazil, personal communication,

1999).

Biological Control of Waterhvacinth

Biological control strategy for waterhyacinth has been based mostly on arthropods

and plant pathogenic fungi. The most studied and used arthropods for biological control








are the weevils Neochetina bruchi Hustache and N. eichhorniae Warner (Coleoptera:

Curculionidae), the moth Sameodes albiguttalis Warren (Lepidoptera:Pyralidae), and the

mite Orthogalumma terebrantis Wallwork (Acarina:Galumidae). Of these organisms, N.

eichhorniae has been the most important control agent which, along with N. bruchi, has

been successfully introduced from South America to several countries, including USA,

Mexico, South Africa, India, and Australia (Center, 1996a). These insects cause

reduction in the growth of waterhyacinth populations as a consequence of the damage

from feeding on the laminae by adult weevils and the tunneling by larvae at the base of

petioles and into the crown. The potential to increase the damage caused by the

arthropods by the interaction with some fungi and bacteria has been recorded

(Charudattan, 1986; Charudattan et al., 1978). However, no further study with other

insect-plant pathogen associations has been reported. A better understanding of these

interactions can be extremely valuable to increase the efficacy of the biocontrol agents.

Plant pathogens were suggested as possible biocontrol organisms for

waterhyacinth as early as the 1930s. As pointed out by Martyn (1977), the first published

paper concerned with plant pathogens as controls for waterhyacinth was written by

Agharkar and Banerjee (1932) in India. They reported studies with a Fusarium species

pathogenic to waterhyacinth. However, it was not until the 1960s that a serious

consideration was given to pathogens as biological control agents for waterhyacinth. It

was then that scientists at the Commonwealth Institute for Biological Control, Bangalore,

India, discovered many new pathogens and evaluated the biological control potential of

some of them (Nag Raj, 1965; Nag Raj and Ponnappa, 1967; 1970). In 1970, another

program was initiated in Florida to control waterhyacinth with pathogens that led to a








serious effort to collect, identify, and screen pathogens in the United States and in other

countries.

Studies on the biological control ofwaterhyacinth with plant pathogens have been

based only on fungal plant pathogens. However, some bacteria belonging to

Xanthomonas and Erwinia have been reported to be associated with damage caused by

the Neochetina weevils. These bacteria have been associated with a chlorotic halo that

surrounds the weevil's feeding spots (Charudattan, 1996). The most comprehensive list

of fungi reported from waterhyacinth in different parts of the world is presented by R.

Charudattan in Gopal (1987) with more than one hundred different fungi associated with

this plant. However, for most of these associations, etiological studies are incomplete or

non-existing, and only about 10 of them have been considered important for biological

control (Charudattan, 1990; Charudattan et al., 1996).

The first disease recorded on waterhyacinth was a leaf spot caused by the fungus

Cercospora piaropi Tharp. This fungus, described and named by Tharp (1917) based on

a specimen collected in Palestine, Texas, was subsequently recorded in India

(Thirumalachar and Govindu, 1954; Vasudeva, 1963; Nag Rag, 1965), Florida (Freeman

and Charudattan, 1974), South Africa (Morris, 1990), Australia (Galbraith, 1983), Brazil

(Barreto and Evans, 1995), and Mexico (Martinez Jimenez and Charudattan, 1998). The

symptoms of this disease have been described as discrete dark brown leaf spots present

on leaves and petioles. These spots are discrete on young leaves but lead to chlorosis and

necrosis of part or the whole of older leaves and petioles. Even though this fungus did

not appear to cause appreciable damage to the waterhyacinth plants when it was first








noted, severe epidemics of the disease caused by this fungus were reported by Martyn

(1985) in Texas, USA, and Morris (1990) in South Africa.

The second disease reported in waterhyacinth was a rust caused by the fungus

Uredo eichhorniae Fragoso and Ciferri, from the Dominican Republic (Ciferri and

Fragoso, 1927). In addition, a smut, caused by Doassansia eichhorniae Ciferri, was

reported from that country almost at the same time (Ciferri, 1928). Despite the

worldwide distribution of waterhyacinth, there have been no new sightings of D.

eichhorniae from anywhere including the Dominican Republic. Presently, U. eichhorniae

has been recorded only in Argentina (Charudattan and Conway, 1975) and Southern

Brazil (Dr. R. Charudattan, unpublished field records). Uredo eichhorniae has potential

as a biocontrol agent, but its introduction into the United States has not been allowed

because the complete life cycle and consequently possible alternate host(s) are still

unknown (Dr. R. Charudattan, personal communication).

Rhizoctonia solani Kiuhn was recorded on waterhyacinth for the first time in India

(Padwick, 1946), where it caused extensive blotches and streaks, and often killed

individual plants. In the American continent, R. solani was recorded, first from the

anchoring waterhyacinth (E. azurea [Swartz] Kunth) in the Panama Canal Zone (Freeman

and Zettler, 1971) and shown to be pathogenic to E. crassipes (Joyner and Freeman,

1973). It was later found on E. crassipes, in Louisiana, USA (Freeman et al., 1982). The

fungus isolated in Panama was named later as Aquathanathephorus pendulus Tu and

Kimbrough gen. and sp. nov. based on differences on the morphology of the basida

observed through induction of the perfect stage (Tu and Kimbrough, 1978).








The first report of a zonate leaf-spot disease of waterhyacinth was by Padwick,

(1946) in India. He attributed the cause to a new fungus, naming it Cephalosporium

eichhorniae Padwick. However, this species was later considered a synonym of C.

zonatum (Rintz, 1973), and C. zonatum was reclassified to Acremonium zonatum

(Sawada) Gams (Gams, 1971). This fungus has been also recorded on waterhyacinth in

El Salvador, Panama, Florida (Freeman et al., 1973), Louisiana, USA (Rintz, 1973), and

Mexico (Martinez Jimenez and Charudattan, 1998). Rintz (1973) observed that this

fungus attacked several host plants under artificial conditions and did not cause severe

damage on waterhyacinth. Martyn (1977) observed that A. zonatum was able to control

small, young plants rather than large mature plants.

Marasmiellus inoderma (Berk.) Sing. was reported by Nag Raj (1965) as the

causal agent of thread blight on leaves and petioles of waterhyacinth in India. According

to this report, this fungus could spread quickly through plant populations of plants

because it had an abundant aerial mycelial growth. No further studies have been reported

about this pathogen on waterhyacinth.

Myrothecium roridum Tode ex Fries was also described a cause of a disease on

waterhyacinth in India (Ponnappa, 1970). Although this fungus inflicted extensive

damage to waterhycinth by necrotic lesions on leaves, it was not considered for use as a

biological control agent because of its broad host range (Ponnappa, 1970).

Several species of Bipolaris and Helminthosporium (two related genera) have

been reported to attack waterhyacinth in various countries. One species in particular, B.

oryzae (Breda de Haan) Shoemaker (=B. stenospila [Drechs.] Schoemaker) was reported

as the causal agent of severe blighting on shoots of waterhyacinth in the Dominican








Republic (Charudattan et al., 1975). A Dominican isolate of this fungus from

waterhyacinth was also pathogenic to sugarcane, rice, and bermudagrass, and therefore it

is not likely to be safe for use as a bioherbicide (Charudattan, 1996).

Alternaria eichhorniae Naj Raj and Ponnappa has been recorded in India (Nag

Raj and Ponnappa, 1970), Bangladesh (Badur-ud-Din, 1978), Indonesia

(Mangoendihardjo et al., 1977), Thailand (Rakvidhyasastra et al., 1978), and Egypt

(Shabana et al., 1995a). This fungus causes a severe leaf blight on waterhyacinth, has a

narrow host range (Nag Raj and Ponnappa, 1970; Shabana et al., 1995a), and is being

studied for development as a bioherbicide to control waterhyacinth (Shabana, 1996;1997;

Shabana et al.; 1995b; 1997).

A second Cercospora species, C. rodmanii Conway, was described on

waterhyacinth from the Rodman Reservoir, Florida, based on some phenotypic

differences in relation to C. piaropi (Conway, 1976a). Indeed, to circumscribe C.

rodmanii, Conway (1976a) emended the diagnosis of C. piaropi given by Tharp (1917).

Based on Conway's description, C. rodmanii had longer conidia than C. piaropi; conidia

of C. rodmanii had truncate bases compared to the obconic bases observed in C. piaropi;

Cercospora rodmanii was considered more virulent than C. piaropi; the former caused a

general blighting symptom on the foliage compared with more discrete leaf spots caused

by C. piaropi. Moreover, C. rodmanii had the presence of a well-developed stroma at the

base of conidiophores; and the presence of an associated pycnidial state, Asteromella sp.

However, in practice it has been difficult to distinguish between these two species based

on these diagnostic characters (Martyn, 1985; Morris, 1990; Martinez Jimenez and

Charudattan, 1998).











The species of Cercospora as Agents for Biological Control

The isolate of Cercospora from Rodman Reservoir, named C. rodmanii, has been

the most studied pathogen for biological control of waterhyacinth. Even though, severe

epidemics of C. piaropi have been recorded more recently in Texas, USA (Martyn, 1985)

and South Africa (Morris, 1990), early reports considered this pathogen to be incapable of

causing serious damage to waterhyacinth (Nag Rag, 1965; Freeman and Charudattan,

1974).

Cercospora spp. are the most widespread pathogens of waterhyacinth.

Cercospora piaropi and C. rodmanii have been reported collectively, from every

continent in which waterhyacinth has spread (Charudattan, 1996). The host specificity of

Cercospora species on waterhyacinth has been the most important factor to consider

these pathogens as biocontrol candidates (Freeman and Charudattan, 1974; Conway and

Freeman, 1977). Cercospora rodmanii was shown to has a limited host range in a study

that involved 58 species in 22 plant families, and plants that developed symptoms showed

damage only on senescent tissues. The blighted areas showed the presence of both C.

rodmanii and a long-beaked Alteraria sp. (Conway and Freeman, 1977).

In studies of the C. rodmanii-waterhyacinth pathosystem, the infection process

originated from either fungal mycelium or conidia. In both instances the hyphae grew

into the stomata, ramified in the substomatal cavity, and invaded the surrounding tissue.

A stroma developed in the stomatal cavity and a fascicle of conidiophores arose from it

and emerged through the stomata. Primary and secondary conidia were produced on the

conidiophores (Freeman and Charudattan, 1984).








In nature, conidia produced on diseased tissue are spread by wind and serve to

disseminate C. rodmanii from disease foci. The speed and intensity of the epidemic are

directly related to the number of conidia produced, which is related to the amount of

diseased and dead tissue that is available for sporulation. Under natural conditions in

Florida, the sporulation reaches a peak during the fall and early winter (Freeman and

Charudattan, 1984).

Disease stress within a population of waterhyacinth is manifested initially as an

overall chlorotic appearance of the plants. Numerous severely spotted or dead leaves

soon become evident on the plants. As the disease progresses, the entire population of

waterhyacinth turns brownish in appearance. At this stage, the waterhyacinth population

begins to decline (Freeman and Charudattan, 1984).

Biological control of waterhyacinth with C. rodmanii has relied mostly on the

bioherbicidal strategy, but this fungus has been released also as a classical biocontrol

agent in South Africa, Egypt, and Honduras (R. Charudattan, personal communication).

The differences between the classical and the bioherbicidal strategy are that, in the first, a

pathogen is introduced from the geographic origin of the weed into the weed's adventive

range where the control is desired. In the bioherbicidal strategy, native or exotic

pathogens are cultured in vitro on a large scale and applied in fairly high concentration to

the weed (Templeton, 1982; Charudattan, 1985). In the United States, a bioherbicidal

strategy strategy was proposed to control this weed, based on C. rodmanii, which can

provide significant levels of control when used under conditions that limit host growth

rate or in combination with other biotic and abiotic agents (Conway, 1976b; Freeman and

Charudattan 1984; Charudattan, 1986; Charudattan et al., 1985). However, even though








C. rodmanii has shown good potential to control waterhyacinth, the market for aquatic

weed control has been dominated by chemical herbicides that provide fast, economical,

and predictable control. Therefore, the incentive to develop and register C. rodmanii was

discontinued (Charudattan, 1991).

Research Rationale

The differentiation of the species C. piaropi and C. rodmanii, based on the

characters used for their description, has been difficult in practice and has caused

scientific and regulatory (plant quarantine) problems and consequently has prevented to

some extent the development and implementation of biological control of waterhyacinth.

In addition, the species C. piaropi has been recorded in several regions of the world,

including the center of origin of waterhyacinth and its adventive areas, whereas C.

rodmanii has been recorded only in Florida. In view of the fact that these species were

circumscribed based on their morphology and virulence, the possibility arises that, like

many other fungal species, the DNA-based techniques may help to reconcile the two

species. Hence, the objective of this research was to 1) delineate phylogenetically

informative characters based on a collection of isolates of C. piaropi and C. rodmanii

from several geographic origins; 2) characterize the partial DNA sequences of three

evolutionary conserved genes; and 3) assess the validity of the two Cercospora spp.

reported on waterhyacinth. It was hypothesized that this approach will allow the

redefinition of the species based on the phylogenetic species concept and to provide some

insights about the biogeography of these pathogens of waterhyacinth. Another objective

was to determine the extent of variability in some phenotypic traits, including cultural

features, phytotoxins, and virulence among a population of isolates of C. piaropi and C.






14

rodmanii cultured under standard conditions. Finally, an attempt was made to determine

the usefulness of fatty acid methyl ester profiles to discriminate isolates, populations, or

species of Cercospora from waterhyacinth.














CHAPTER 2
A MOLECULAR CHARACTERIZATION OF CERCOSPORA SPECIES FROM
WATERHYACINTH

Introduction

Two Cercospora species have been studied and used as biocontrol agents for the

control of the aquatic weed waterhyacinth (Eichhornia crassipes [Mart.] Solms): C.

piaropi described by Tharp (1917) from a Texas specimen and C. rodmanii described by

Conway (1976a) from a Florida specimen. The species C. rodmanii was described by

Conway (1976a) who examined decaying waterhyacinth plants collected at the Rodman

Reservoir, Florida, where it caused severe epidemics of a Cercospora leaf-spot disease.

He observed that the Rodman isolate had some differences in conidial morphology and in

disease symptomatology in relation to what was described for C. piaropi by Tharp (1917).

Indeed, to circumscribe these two species of Cercospora on waterhyacinth, Conway

(1976a) emended the original description of C. piaropi, given by Tharp (1917), and

differentiated these species based on the observation that conidia of C. rodmanii were

longer than those of C. piaropi; conidia of C. rodmanii had a truncate base while those of

C. piaropi had an obconic base; C. rodmanii was more virulent than C. piaropi and

caused general blighting symptom on the foliage compared to the more discrete leaf spots

caused by C. piaropi; C. rodmanii had the presence of a well-developed stroma at the

base of conidiophores; and the C. rodmanii specimens examined had the presence of an

associated pycnidial state, Asteromella sp. (Conway, 1976a). Based on the Rodman








isolate, a bioherbicidal strategy was proposed to control waterhyacinth with the

expectation that this pathogen can provide significant levels of control when used under

conditions that limit host growth rate or in combination with other biotic and abiotic

agents (Conway, 1976b; Freeman and Charudattan, 1984; Charudattan, 1986;

Charudattan et al., 1985).

Since its original description from Texas, C. piaropi has been recorded also in

India (Thirumalachar and Govindu, 1954; Vasudeva, 1963; Nag Rag; 1965), Florida,

USA (Freeman and Charudattan, 1974), Australia (Galbraith, 1983), South Africa

(Morris, 1990), Brazil (Barreto and Evans, 1995), and Mexico (Martinez Jimenez and

Charudattan, 1998), whereas the species C. rodmanii has been recorded only in Florida.

In fact, it is difficult to distinguish the two species, as observed in the reports of Martyn

(1985) and Morris (1990). The problem is that the variability in size and shape of conidia

sometimes encompasses the description of both species, and the differences in disease

symptoms between these two species are not distinguishable in practice.

The species definition in Cercospora Fres. has been based upon morphological

criteria and host affiliation (Chupp, 1953; Ellis, 1971). However, in addition to the

problems that such criteria cause for species identification, it is still unknown whether

they are appropriate to circumscribe species defined by phylogenetic relationships.

Moreover, even though those Cercospora species with a known teleomorphic stage have

been linked to Mycosphaerella, in the phylum Ascomycota (Sivanesan, 1984), it is still

unknown whether the form-genus Cercospora is in fact a monophyletic group.

Molecular markers have been a valuable source of new diagnostic characters for

studies of fungal taxonomy and evolution (Bruns et al., 1991; Kohn, 1992). In fact, DNA








sequences of conserved genes can be abundant sources of phylogenetically informative

characters, which are appropriate for phylogenetic analysis of fungal taxa and to define

species according to the phylogenetic species concept (O'Donnell and Cigelnik, 1997;

O'Donnell et al., 1998a).

DNA sequence analysis of internal transcribed spacers regions (ITS) of the

ribosomal DNA have been used to distinguish species and groups of species in a number

of taxonomic studies of fungi (Chen et al., 1996; Crawford et al., 1996; Harrington, 1998;

Roy et al., 1998). In addition, in recent studies, protein-coding genes were appropriate for

phylogenetic analysis of fungi at the species level. Such genes may have little or no

variation in their amino acid sequences but their third codon positions and intron regions

appear to have a high rate of nucleotide substitutions. Protein-coding genes have

advantages as molecular markers over ribosomal genes in that they offer a large number

of unlinked sources of phylogenetic information (Geiser et al., 1998a). Indeed, these loci

may show different levels of resolution for the different groups of fungi, and as seen in

Fusarium, the concurrent analyses of more than one independent locus increases the

strength of the conclusions by offering different lines of evidence for a phylogenetic

hypothesis (O'Donnell and Cigelnik, 1997; O'Donnell et al., 1998a; O'Donnell et al.,

1998b).

The objective of this study was to determine the phylogenetic relationships among

isolates of Cercospora species pathogenic to waterhyacinth, from several geographical

origins, based on partial DNA sequences from three protein-coding genes: elongation

fator-1 a, P-tubulin, and histone 3; and the ribosomal DNA regions containing ITS I,

ITS2, and the 5.8S gene. The hypotheses tested were whether C. rodmanii and C. piaropi








are two distinct species according to the phylogenetic concept of species, and whether

genetic divergence, if found, is related to the geographical origin of the isolates.

Materials and Methods

Fungal Isolates

The isolates used in this study were recovered from symptomatic leaves of

waterhyacinth, and monocultures were obtained from hyphal tips. Their designations and

geographic origins are presented in the Table 2-1. These isolates have been preserved in

the fungal collection of the Biological Control of Weeds Laboratory of the Plant

Pathology Department, University of Florida.

For isolate identification, a test tube containing 1 g of autoclaved, wet, seeds of

ryegrass (Lolium multiflorum Lam.) was inoculated with a 5 cm3 mycelial plug removed

from a 7-day-old culture on potato dextrose agar (PDA) plate. The tube was kept at room

temperature with 12 h light. Production of conidia was observed 7 to 14 days after

inoculation. Since the isolates sporulated irregularly, this procedure needed to be

repeated to identify the isolates. A natural substratum, such as ryegrass seed, was

preferred because most of the isolates did not sporulate in axenic cultures even when

grown in several media. Observations and conidial measurements were made with a light

microscope. In addition, the isolates were grown on V-8 agar in petri plates for

observation of cultural attributes, including the production of the typical reddish-purple

pigment, and for determination of the occurrence of the Asteromella conidial state by

flooding the plates with distilled water (Conway, 1976a).








DNA Isolation

Roux bottles containing 90 ml of potato dextrose broth (PDB) (Difco, Detroit,

MI) were inoculated with six 5-mm3 mycelial plugs excised from the margin of an 8-day-

old culture on PDA plate (Difco, Detroit, MI). After 5-6 days of growth, the contents of

the flasks were filtered through sterile cheesecloth, squeezed dry, and rinsed three times

with sterile deionized water. Mycelium was placed into 13-ml plastic tubes, stored for 24

h in a -800C freezer, and lyophilized for 24-48 h. The dry mycelium was then mixed with

liquid nitrogen, ground to a fine powder, and combined with DNA extraction buffer that

consisted of a 1:1:0.4 volume of Nuclei Lysis Buffer (0.3 M sorbitol, 0.1 M Tris, and 20

mM EDTA, at pH 7.5), DNA isolation buffer (0.2 M Tris at pH 7.5, 50 mM EDTA, and

0.2 mM cetyltrimethylammonium bromide), and 0.5% Sarkosyl (Koenig et al., 1997;

Rosskopf, 1997).

Ten milliliters of the extraction buffer were combined with approximately Ig of

ground mycelium in 15-ml tubes, and the tubes were placed in a 65C water bath for 60

minutes. The contents of the tubes were then mixed by inversion, and I ml of the

solution was transferred to a sterile, 1.5-ml microcentrifuge tube. Five hundred

microliters of chloroform:octanol (24:1) solution was added to each tube. The solution

was mixed thoroughly by inversion. The solution was then centrifuged for 10 minutes at

12,000 g in a microcentrifuge, at room temperature. The supernatant was transferred to

sterile 1.5-ml tubes and treated with 5 il of a suspension containing 20 mg RNAse

(Sigma Chemical Company, St.Louis, MO) per ml for 30 minutes at 370C. Following the

RNAse treatment, 5 pl of a suspension of 20 mg Proteinase K (Sigma Chemical

Company, St. Louis, MO) per ml were added and allowed to remain in solution for 20








minutes at 37'C. One volume of ice-cold isopropanol was then added, and the tubes were

shaken until the DNA was visible as a white precipitate. After 60 minutes in a -20C

freezer, the tubes were centrifuged at 10,000 g for 5 minutes and the DNA pellet was

washed with 100 pl of 70% ethanol three times and allowed to dry. Then, the pellet was

suspended in 100 .l of TE buffer (10 mM Tris pH = 7.6; 1 mM EDTA) and samples were

placed at 40C until the DNA was dissolved. The method described here is a modified

version of the DNA extraction method used by Koenig et al. (1997) and Rosskopf (1997).

Samples that were difficult to resuspend were subjected to a LiCI treatment.

Three hundred microliters of ice-cold 4M LiCI solution were added to each tube, and the

tubes were placed on ice for 30 minutes before centrifugation at 12,000 g for 10 minutes

at 4C. The supernatant was transferred to a sterile 1.5-ml tube containing 600 pt of

isopropanol. This solution was mixed, and the tubes were kept at room temperature for

30 minutes. After centrifugation at 12,000 g for 10 minutes at 4C, the supernatant was

discarded and 100 pl of ice-cold 70% ethanol was added to the tubes. After

centrifugation at 12,000 g for 5 minutes, the ethanol was discarded and the DNA was air-

dried in a hood. TE buffer pH = 7.6 (100 ptl) was added to the tubes and they were placed

in a water bath at 650C until the pellet was dissolved. Concentration and purity of DNA

was estimated spectrophotometrically.

DNA Amplification and Sequencing

Approximately 200 ng of template DNA per 100 l reaction mixture was used for

DNA amplification by the polymerase chain reaction (PCR). Primers for amplification of

P-tubulin and histone 3 gene sequences were based on those used by Glass and








Donaldson (1995). The internal transcribed spacer regions (ITS) of the nuclear ribosomal

repeat were analyzed with primers ITS4 and ITS5 (White et al., 1990). Gene maps with

the primer position and sequences are shown in Figure 2-1. For amplification of

elongation factor-la gene sequences, a set of primers was designed with the software

PC/Gene (IntelliGenetics Inc., Mountain View, Ca) based on Aureobasidium pullulans

EF-la gene sequence (GenBank accession no. APU19723). All primers for protein-

coding genes span at least one intron area. The primers were synthesized at either the

University of Florida Interdisciplinary Center for Biotechnology Research

Oligonucleotide Synthesis Laboratory (Gainesville, FL) or by Gibco BRL, Gaithersburg,

MD). PCR was performed using final concentrations of the components in the reaction

mixture as follows: 20 mM Tris-HCI (pH 8.4); 50 mM KC1; 3 mM MgC12; 1 JM of each

primer; 200 pM each of dATP, dCTP, dGTP, and dTTP; and 2.5 U of Taq polymerase

(Gibco-BRL, Gaithersburg, MD) per 100 pl of reaction mixture. A GeneAmp 9600

(Perkin-Elmer Applied Biosystems, Foster City, CA) was used for the amplification. For

amplification of the elongation factor-la, j-tubulin and histone 3 gene sequences, the

cycling conditions used an initial denaturation step of 1 min at 940C, with 35 cycles of

940C for 45 seconds, 620C for 30 seconds, and 720C for 45 seconds. The last cycle

included a 10-minute incubation at 720C and then storage at 40C. For amplification of

the rDNA region, the annealing temperature was dropped to 550C. PCR fragments were

cleaned using PCR Preps Kit (Promega INC., Madison, WI) according to the

manufacturer's instructions. Sequences were obtained with a Perkin-Elmer Applied









Elongation factor- la
EFla


4
EFIb

P-tubulin
Bt2a

c~ III, ___~b


4
Bt2b

Histone 3
H3-la


H3-lb

Nuclear rDNA genes
ITS4

I 18S 5.8 28S
ITSI ITS2 4
ITS5

Bt2a 5' GGTAACCAAATCGGTGCTGCITTC 3'
Bt2b 5' ACCCTCCGTGTAGTGACCCTTGGC 3'
EFIa 5' ATCAACCTCGTCGTTATCGGCCACG 3'
EFIb 5' TCAGACTTCACGTTGTCGAGGACCC 3'
H3-la 5' ACTAAGCAGACCGCCCGCAGG 3'
H3-lb 5' GCGGGCGAGCTGGATGTCCTT 3'
ITS4 5'TCCTCCGCTTATTGATAT 3'
ITS5 5' GGAAGTAAAAGTCGTAACAAGG 3'



Figure 2-1. Maps of the elongation factor-lo, 0-tubulin, and histone-3 genes; and of
ribosomal DNA region (rDNA). Position of primers used for PCR amplification and
sequencing are indicated by arrows. Shaded boxes denote protein-coding sequences
(exons), and cross-hatched boxes denote introns. The P-tubulin and histone 3 maps are
from Neurospora crassa (Glass and Donaldson, 1995), and elongation factor-la map is
from Aureobasidium pullulans (Thomewell et al., 1995). The rDNA region map was
based on White et al. (1990).








Biosystems model 373A or 377 automated DNA sequencer (Perkin-Elmer, Foster City,

California, CA) in the DNA Sequencing Core Laboratory of the University of Florida.

Southern Blot Analysis of the 3-tubulin Gene

The possibility that isolates may have multiple copies of thep-tubulin gene was

tested through Southern blotting analysis. Approximately 5 tg of DNA was digested

with at least 10 units of Pst I, EcoR I, and Hind III restriction enzymes (Gibco-BRL,

Gaithersburg, MD) and incubated for at least 4 h. These restriction enzymes were chosen

based on analysis of the restriction map of the 3-tubulin gene of Colletotrichum

graminicola (Ces.) G.W. Wils. (GenBank accession no. M34492) using the University of

Wisconsin Genetics Computer Group program. Restriction fragments were separated by

electrophoresis in 0.8% agarose in Tris-borate-EDTA (TBE) buffer at pH 7.0. Gels were

run at 50 V for approximately 18 h Lambda phage DNA digested with Hind II was

used for size markers. DNA was detected by staining gels in a solution of ethidium

bromide (0.5 ptg ml') followed by UV trans-illumination. DNA was transferred to Zeta-

probe GT membrane (Bio-Rad Laboratories, Hercules, CA) using the capillary transfer

method. The DNA transfer proceeded for at least 16 h and DNA was immobilized by UV

cross-linking and hybridized to a 32P-labeled DNA probe. The DNA probe was a 380 bp-

PCR product of the p-tubulin gene from the isolate 28-1, amplified with the primers Bt2a

and Bt2b and labeled using the RadPrime DNA Labeling System (Gibco-BRL,

Gaithersburg, MD), according to procedures provided by the manufacturer.

Phylogenetic Analysis

Following alignment with CLUSTAL W1.7 (Thompson et al., 1994), the measure

of the phylogenetic signal for each dataset was estimated by the skewness of the tree-








length distributions (Hillis and Huelsenbeck, 1992; Hillis et al., 1993), implemented in

PAUP version 4.0b1 (Swofford, 1997). Phylogenetic relationships among taxa for

individual and combined datasets were inferred based on the maximum parsimony,

maximum likelihood, and neighbor-joining methods, implemented by PAUP (Swofford,

1997) and MacClade Version 3.01 (Maddison and Maddison, 1992), and performed with

all DNA characters unweighted. All of these methods of phylogenetic inference have

been shown similar accuracy (Hillis et al., 1994). In the maximum parsimony analyses,

trees were obtained using the stepwise addition option in heuristic search with random-

addition sequences, and alignment gaps were treated as missing characters. For neighbor-

joining analysis, a distance matrix was generated using the Jukes-Cantor procedure. For

maximum likelihood analysis, a 2:1 transition/transversion rate was assumed. Clade

stability was estimated with 1000 bootstrap replications (Hillis and Bull, 1993)

implemented in PAUP (Swofford, 1997) and by decay indices (Bremer, 1988) calculated

with TreeRot (Sorenson, 1996). Other measures, including tree length and consistency

and retention indices, were calculated with PAUP.

The partition-homogeneity test (PHT) option in PAUP (Swofford, 1997) was

used to determine whether the elongation factor-la, P-tubulin, and histone 3 datasets

were in conflict, with 1000 replicates. For the PHT, only phylogenetically informative

characters were used. Gaps were considered missing data for all analyses. The Kishino-

Hasegawa test was used to compare alternative constrained and unconstrained tree

topologies using PAUP (Swofford, 1997).








This study used the phylogenetic species concept (Nixon and Wheeler, 1990) as

used by O'Donnell and Cigelnik (1997), which recognizes species as the smallest group

of populations or lineages diagnosable by a unique combination of fixed apomorphies.

Results


Identification of Isolates

Some isolates showed characteristics of conidial size and morphology that

encompassed the description of both Cercospora species described as pathogens of

waterhyacinth, C. piaropi and C. rodmanii. The isolates WH9BR, WHK, BA57, 175-102,

and 2619, from Florida; 62-2, 62-4, and 34 from Brazil; WHV from Venezuela; 2867

from Mexico; and 400 from Zambia (Tables 2-1; 2-2) were in this category. The

presence of conidia with both obconic and truncate bases (Figure 2-2) is not a feature

related with the geographic origin of the isolates. However, some isolates showed only

conidia with obconic base and conidial size concordant with the description of C. piaropi,

such as the isolates WH83 and RR29 from Florida; TX15, TX18, and TX20, from Texas;

10, 28-1 and 18-2, from Brazil; and MX3 from Mexico. The fact that all isolates showed

conidia with obconic bases and some isolates showed conidia with both, obconic and

truncate bases confused the species identification. The isolate WH9BR, which was

collected by Conway and deposited as a type culture of C. rodmanii also showed conidia

with obconic and truncate bases. In general, conidia with truncate bases were longer than

conidia with obconic bases and the range of conidial length for most isolates was more

close to the range described for C. piaropi. In addition, none of these isolates produced

the Asteromella conidial state in V-8 agar cultures flooded with distilled water.








Table 2-1. Designations, origins, and morphological characteristics of conidia of the isolates of Cercospora species from
waterhyacinth used in this study.
Isolate Geographic original Conidial size (um)b Morphology of conidial baseb
WH9BR' Florida, USA 55- (148) -232 x 3.0- (3.8) -4.5 Truncate/obconic
RR29 Florida, USA 44- (132)-196 x 3.0- (4.0) -4.5 Obconic
WH83 Florida, USA 33- (115) -183 x 3.0- (3.4) -4.0 Obconic
WHK Florida, USA 35- (132)-243 x 3.5- (4.0) -5.0 Truncate/obconic
BA57 Florida, USA 31-( 91)-148 x 2.5- (3.5) -4.0 Truncate/obconic
175-102 Florida, USA 48- (127) -205 x 3.0- (3.4) -4.5 Truncate/obconic
2619 Florida, USA 35- ( 92) -253 x 2.5- (3.5) -4.0 Truncate/obconic
TX15 Texas, USA 28- ( 76) -142 x 2.5- (3.0) -3.5 Obconic
TX18 Texas, USA 33- (109)-157 x 2.0- (3.0) -3.5 Obconic
TX20 Texas, USA 35-( 84)-167 x 2.5- (2.8) -3.5 Obconic
62-4 Northeast Brazil 39- (117) -211 x 2.5- (3.4) -4.0 Truncate/obconic
62-2 Northeast Brazil 35- (118)-175 x 2.0- (3.2) -4.0 Truncate/obconic
10 Southeast Brazil 35- (147)-219 x 3.0- (4.0) -4.5 Obconic
28-1 Southwest Brazil 28-( 81)-177 x 2.5- (3.5) -4.5 Obconic
18-2 Southwest Brazil 22-( 69) -158 x 2.0- (2.7) -3.5 Obconic
34 South Brazil 24-(115)-175 x 2.5- (3.0) -3.5 Truncate/obconic
WHV Venezuela 55-(141)-203 x 2.0-(3.0)-3.5 Truncate/obconic
2867 Mexico 44- (139)-225 x 2.5- (3.6) -4.5 Truncate/obconic
MX3 Mexico 42- (108)-163 x 3.0- (3.8)- 4.0 Obconic
114 South Africa NDd ND
400 Zambia 26- (119) -187 x 2.5- (3.7) -4.0 Truncate/obconic
a The isolates RR29, BA57, 175-102, TX15, TX18, and TX20 were collected for this study; the isolates 114 and 400 were obtained from Dr. M.
Morris, Stellenbosch, South Africa; and the other isolates were obtained from the culture collection of the Biological Control of Weeds
Laboratory of the University of Florida. Additional details of geographic origin are available from Dr. R. Charudattan.
b Based on conidia produced by cultures grown on autoclaved ryegrass seeds.
C. rodmanii culture type collected by K. Conway.
d ND = not determined due to lack of sporulation.









Table 2-2. Conidial size and morphology of Cercospora piaropi and Cercospora rodmanii recorded in the literature.

Species Geographic Conidial size (tm) Morphology of
origin conidial base
C. piaropi (Tharp, 1917)a Texas, USA 80-140 x 3 Truncate/obconicb
C. piaropi (Chupp, 1953) USA 25-140 x 2-3.5 Truncate
C. piaropi (Thirumalachar and Govindu, 1954) India 66-120 x 2-4.2 Truncate
C. piaropi (Vasudeva, 1963) India 80-140 x 3 Truncate
C. piaropi (Freeman and Charudattan, 1974) Florida, USA 55-121 x 3.3-4.4 Truncate
C. piaropi (Morris, 1990) South Africa 50- (140) -270 x 1.5- (2.5) -3.5 Truncate/obconic
C. piaropi emendedd by Conway [1976]) -25-( 95)-220 x 2.0- (3.5) -5.0 Obconic
C. rodmanii (Conway, 1976) Florida, USA 66- (172)-374 x 3.0- (4.0) -5.0 Truncate

a Original species description.
b The species diagnostic given by Tharp (1917) describe only truncate conidia; however, Conway (1976a) examined the holotype
and observed mostly obconic conidia.
Original species description.


























pi


Figure 2-2. Conidia of Cercospora from waterhyacinth with truncate (a) and obconic (B)
bases (x400).








The isolate 114 from South Africa did not produce condia in any medium,

substrate, or condition tested. It was identified based on the collector's information.

Phylogenetic Analysis

The aligned DNA sequences of the elongation factor-la, p-tubulin, and histone 3

datasets provided enough phylogentically informative characters to infer relationships

among the isolates of Cercospora sp. from waterhyacinth (Table 2-3; Appendices A, B,

and C).

From 431 characters of elongation factor- Ia dataset, 14 characters were

parsimony-informative and three were variable characters that were parsimony-

uninformative. From the 380 characters of P-tubulin dataset, six characters were

parsimony-informative and 64 variable characters were parsimony-uninformative. From

309 characters of the histone 3 dataset, 17 characters were parsimony-informative and

124 variable characters were parsimony-uninformative. However, the aligned DNA

sequences of the 5.8 rDNA gene and is ITS flanking regions for eight of the isolates were

invariant in this region even when compared with the outgroup, C. beticola (Table 2-3;

Appendix D).

The tree-length distribution analysis, based on 10,000 replicates, showed strongly

significant skewed tree-length distributions for the elongation factor-la, p-tubulin, and

histone 3 datasets (Figure 2-3), with gl values of -2.817, -2.063, and -3.230 (P=0.01),

respectively. This analysis indicated that these datasets were significantly nonrandom

and potentially informative about phylogeny (Hillis et al., 1993; Hillis and Huelsenbeck,

1992). The partition homogeneity test (Farris et al., 1995; Huelsenbeck et al., 1996) was


























Table 2-3. Sequencing results for the data sets aligned with CLUSTAL 1.7W and
analyzed using PAUP version 4.0bl.

Total Variable Parsimony- Parsimony-
Data set characters characters informative uninformative
characters characters
Elongation factor-la 431 17 14 3
P-Tubulin 380 70 6 64
Histone 3 309 141 17 124
5.8S rDNA and ITS regions 586 6 0 6














5000 Elongation factor- la

S4000-

3000 -

E 2000 -
z


'C c'cl N 0r m '


7000-
6000-
5000 -
4000 -
3000 -
2000
1000
n


timMI 1


2500
I-
' 2000
1500
1000
z


P-Tubulin


N 0 CR 0 C '/ ~ ~ N


Histone 3









.n 1. 1


Tree Length (steps)


Figure 2-3. Tree length distribution for elongation factor-la, P-
tubulin, and histone 3 datasets based on 10,000 random trees.


N II1


01 i
C' Cl
fU


' 1- -


0








applied to decide whether these three datasets could be combined for phylogenetic

analysis. With the same 14 isolates from each dataset as ingroups and C. beticola as the

outgroup, this test rejected the null hypothesis of homogeneity in the phylogenetic signal

among datasets (P=0.002). Thus, these datasets should not be combined for phylogenetic

analysis (Figure 2-4). According to this test, the datasets can not be combined whether

the actual sum of the tree length of individual datasets are less than the shortest tree

generated when the dataset are combined. In this study, elongation factor-la, P-tubulin,

and histone 3 had tree lengths of 17, 78, and 20 steps, respectively. Thus, the sum of

these tree lengths (= 115 steps) was lower than the tree length of the shortest tree

generated when these dataset were combined (116 steps).

The most-parsimonious tree inferred from a segment of 431 bp of elongation

factor-la dataset (Figure 2-5), for 16 isolates of C. piaropi/C. rodmanii, showed a strong

grouping (100% bootstrap support, decay index = 8) for the isolates 10, 28-1, and 62-2

from Brazil; 114 from South Africa; 2867 and MX3 (Mexico), WH83, WH9BR, WHK,

BA57, 2619, and 175102 from Florida, USA; 400 from Zambia; and WHV from

Venezuela. In addition, this tree showed a strong grouping (100% bootstrap support,

decay index = 5) for the isolates TX20 and TX 18, from Texas, USA. The grouping of the

isolates 62-2, WH83, WH9BR, WHK, and BA57 inside the major group did not resolve

with strong support (64% bootstrap support and decay index = 1). The phylogenetic tree

inferred through maximum likelihood method showed the same topology as the tree

inferred through parsimony (Figure 2-6). For the neighbor-joining tree, even though it

showed a more branched topology than the other two trees, the only two clades with




















700

600 -

S500-

S400-

S300

S200 P = 0.002


1 0 I ,


110 111 112 113 114 115 116 117 118 119 120
Sum of Tree Lengths


Figure 2-4. Results of the partition-homogeneity test implemented in PAUP*4.0b 1.
The arrow indicates the summed length of 115 steps of the single most
parsimonious tree inferred from the actual elongation factor-la, P-tubulin, and
histone 3 datasets. The vertical bars show the random distribution of the sum of tree
lengths obtained from 1000 random repartitions of the combined datasets.


















10
114
28-1
2867
MX3
62-2
) ( --WH83
100 (1)
8 64 WH9BR

-WHK
BA57
2619
400
WHV
175102
() TX20
5 -- TX18

Cercospora beticola
(outgroup)



Figure 2-5. Most-parsimonious tree inferred from elongation factor- la gene (length = 17,
consistency index = 1.0, retention index = 1.0, rescaled consistency index = 1.0).
Bootstrap replication percentages and decay indices (in parentheses) are indicated above
the nodes. Edge length is indicated below the nodes and branches.

















10
114
28-1
2867
MX3
62-2
WH83
WH9BR
WHK
BA57
2619
400
WHV
175102
TX20
TX18


(outgroup)


Figure 2-6. Maximum likelihood tree inferred from elongation factor-la gene. Bootstrap
replication frequencies are indicated above the nodes.


















10
MX3
62-2
WH9BR
WHK
BA57
WH83
175102
400
114
2867
2619
WHV
28-1
TX20
TX18
Cercospora beticola
(outgroup)


Figure 2-7. Neighbor-joining tree on distances derived from sequences of the elongation
factor-la gene. Bootstrap replication frequencies are indicated above the nodes.








strong support (100% bootstrap) grouped the same isolates as parsimony and maximum

likelihood tree (Figure 2-7). Those clades placed internal to the major clade did not have

strong bootstrap support. Actually, by default PAUP did not show bootstrap values lower

than 50%.

The most-parsimonious tree inferred from a segment of 380 bp of the p-tubulin

dataset (Figure 2-8), for 21 isolates of C. piaropi/C. rodmanii, also showed a major group

containing the isolates MX3 and 2867 from Mexico; 62-4, 28-1, 18-2, 34, 10, and 62-2

from Brazil; WHK, 175102, BA57, WH9BR, WH83, and RR29 from Florida, USA; 114

from South Africa; 400 from Zambia, and WHV from Venezuela. Also, this tree showed

a group containing the isolates TX15, TX18, and TX20, from Texas, USA. The

statistical support for these groupings were 77% bootstrap and decay index = 1 for the

major group, and 71% bootstrap and decay index = 2 for the minor group. According to

this tree, the isolate 2619 from Florida was distinct from all other members. This

happened because the nucleotide sequence of the intron area of the p-tubulin gene in this

isolate accumulated more sequence differences than the other isolates (Figure 2-11).

The maximum likelihood tree inferred from p-tubulin dataset had exactly the

same topology of the maximum parsimony tree (Figure 2-9). This tree showed 71 and

72% bootstrap support for the two major clades, which grouped the same isolates such as

the two major groupings in the parsimony tree. Also the neighbor-joining tree inferred

from this dataset showed a more branched topology; however, only two clades showed

more than 50% bootstrap support (Figure 2-10). The major group, with 71% bootstrap

support, contained the same isolates as the major group in parsimony and maximum

















MX3
62-4
--28-1
WHK
18-2
175102
BA57
(1) WH9BR
77
1 WH83
2867
RR29
34
(2) 10
918 114
--400
--62-2
WHV
(1) TX18
2 1 TX20
TX15
2619
Cercospora beticola
(outgroup)



Figure 2-8. Most-parsimonious tree inferred from (3-tubulin gene (length = 74,
consistency index = 0.973, retention index = 0.818, rescaled consistency index = 0.796).
Bootstrap replication percentages and decay indices (in parentheses) are indicated above
the nodes. Edge length is indicated below the nodes and branches.


















MX3
62-4
28-1
WHK
18-2
175102
BA57
WH9BR
WH83
2867
RR29
34
10
114
400
62-2
WHV
TX18
TX20
TX15
2619
Cercospora beticola
(outgroup)


Figure 2-9. Maximum likelihood tree inferred from P-tubulin gene. Bootstrap replication
percentages are indicated above the nodes.
















MX3
28-1
WHK
175102
400
114
RR29
18-2
10
62-2
WH9BR
WH83
BA57
2867
WHV
62-4
34
TX18
TX20
TX15
2619
Cercospora beticola
(outgroup)


Figure 2-10. Neighbor-joining tree on distances derived from sequences of P-tubulin
gene. Bootstrap replication percentages are indicated above the nodes.















1 GCAGACCATCTCTGGCGAGCACGGTCTCGACAGCAACGGTG ....... 42
I 1 l l l l l l l I I I I I l i I I Il l i i l It I l II
2 GCAGACCATCTCCGGCGAACATGGCCTCGACGGCTCCGGCGTGTATGTGC 51

43 ....... ........ ....... .. ............ .........CTACA 47

52 AGCAGATCGCAATGGATAAATGGAGCAGCGACTGACGTCGTG GGTACA 101

48 ATGGCAGCTCCGAGCTTCAGCTCGAGCGCATGAGCGTTTACTTCAACGAG 97
1 1i1 i i I I I I I I Il l l l l lill i l l l l i I l l l l I l l l l i
102 ATGGCACGTCTGACCTCCAGCTCGAGCGCATGAACGTCTACTTCAACGAG 151

98 GTTCG....... TGGCCCGAACTCCAAACCTTCCGAGATGTCCACAACGC 140
II II II I I I I I IllI I lIII
152 GTACGCCCGCATTGAGCAGAGCACCAAACTGCTCGAACTCGAGCTGACGC 201

141 GTCTCTTGGTTCATACGGACCCACTGACCGCCTCTCCGGCTTCCGGCAA 190
I I I l l l i l i l l l l l
202 G ........................... AACTGCACAGGCTTCCGGCAA 223

191 CAAGTACGTTCCTCGCGCCGTCCTCGTCGATCTTGAGCCCGGTACCATGG 240
1 l i l l l1 I I I I I I I l il i l l l l i lI l lI 1 1 I I 1 1 1 1 1 11i l
221 CAAGTATGTCCCACGTGCCGTCCTCGTCGATTTGGAGCCTGGCACCATGG 273

241 ATGCTGTCCGTGCTGGTCCTTCGGTCAGCTTTTCCGCCCCGACAACTTC 290
I I I I I I I I I I I I I I i I I i l l l i l l l l l l l l l l i lI I I I Il l l
274 ATGCCGTCCGCGCTGGTCCATTCGGCCAGCTTTTCCGCCCAGACAACTTC 323

291 GTTTTCGGCCAGTCTGGTGCTGGCAACAACTGGGCCAAGGGTCACTACAC 340
I I lI l l Il l l I I i I I i l I l l l l ll l l l l l lI ll i I I
324 GTCTTCGGCCAGTCCGGCGCCGGAAACAACTGGGCCAAGGGTCACTAAAC 373




Figure 2-11. Sequences of amplified segments of P-tubulin gene for the isolates 2619
(upper line) and WH83 (lower line). Vertical lines show matching of nucleotides between
aligned sequences. Sequences inside the boxes correspond to exon areas.








likelihood trees. However, in this tree the isolate 2619 grouped with the isolates from

Texas (62% bootstrap support).

The hypothesis that the differences observed in DNA sequences of P-tubulin gene

of isolate 2619 were due to the presence of extra copies of this gene in its genome was

disproved through Southern blot analysis (Figure 2-12). Southern blots showed the

presence of a single copy of p-tubulin gene in all eight isolates screened, including the

isolate 2619. As predicted in the restriction map of 0-tubulin, the DNA probe hybridized

to two DNA fragments digested with Pst I, to one DNA fragment digested with EcoR I,

and to two DNA fragments digested with Hind II.

The Figure 2-13 shows one of the three most-parsimonious trees inferred from a

segment of 309 bp of the histone 3 dataset, for 14 isolates of C. piaropi/C. rodmanii,

which also showed a strong grouping (96% bootstrap support, decay index = 4) for the

isolates WH83, WH9BR, WHK, and 2619 from Florida, USA; 2867 and MX3 from

Mexico; 10, 62-2, and 28-1 from Brazil; 114 from South Africa; WHV from Venezuela;

and 400 from Zambia. In addition, this tree also showed a strong grouping (100%

bootstrap support, decay index = 11) for the isolates TX18 and TX20 from Texas. The

same two well-supported groupings were observed in the maximum likelihood tree

(Figure 2-14) and in the neighbor-joining tree inferred from this dataset (Figure 2-15). In

the neighbor-joining tree only two groupings had strong support (84 and 100% bootstrap).

The statistical comparison between the topologies of the trees inferred from

parsimony analyses showed significant differences among them (Table 2-4). The

elongation factor-la tree differed significantly (P=0.0175) from and the P-tubulin tree

but not from histone 3 tree; however when the P-tubulin tree was constrained by inserting




















Pst I EcoR I Hind III



hiri






i ig ..r
12 3 4 5 6 7 8 1 2 3 4 5 6 7 8 1 2 3 4 5 6 7 8

Figure 2-12. Southern blot analysis of the P-tubulin gene of Cercospora species from
waterhyacinth. Total DNA of isolates 2619 (lane 1), 28-1 (lane 2), WH83 (lane 3),
WH9BR (lane 4), 62-2 (lane 5), WHK (lane 6), 400 (lane 7), and WHV (lane 8) were
digested with Pst I, EcoR I, and Hind III. Digested DNA was probed with a 32P-labeled
380 bp PCR amplification of the P-tubulin gene from the isolate 28-1.
















WH83

WH9BR

114

WHK

MX3


(4) 2867
96
8 10

WHV

62-2

400
28-1
(11)
100 TX20
12 TX18

Cercospora beticola
(outgroup)



Figure 2-13. One of three equally parsimonious trees inferred from histone 3 gene (length
= 147, consistency index = 0.63, retention index = 0.963, rescaled consistency index =
0.956). Bootstrap replication percentages and decay indices (in parentheses) are indicated
above the nodes. Edge length is indicated below the nodes and branches.


















10

WHV
28-1

400

2867
62-2

114
WHK

MX3

WH83
WH9BR

2619

TX18
TX20
Cercospora beticola
(outgroup)


Figure 2-14. Maximum likelihood tree inferred from histone 3 gene. Bootstrap replication
frequencies are indicated above the nodes.

















- WH83
- 2619

- 114

SWHK

SMX3

WH9BR

2867

WHV

62-2

400

10

28-1

TX18

TX20
Cercospora beticola
(outgroup)


Figure 2-15. Neighbor-joining tree on distances derived from sequences of the histone 3
gene. Bootstrap replication percentages are indicated above the nodes.











Table 2-4. Maximum likelihood comparisons of tree topologies obtained for elongation factor-la, P3-tubulin, and
histone 3 sequences a.

Treeb Length In LC Difference SDe Tf Pg
(steps) In Ld difference

EF-la (Fig. 2-5) 17 -698.49004 best
tub2 (Fig. 2-8) 28 -754.20175 55.71171 23.36445 2.3845 0.0175*
Constrained tubh 20 -721.40525 22.91521 19.91336 1.1507 0.2505
H3 (Fig. 2-13) 20 -721.40525 22.91521 19.91336 1.1507 0.2505

a Results were obtained with the Kishino-Hasegawa test implemented with PAUP*4.0b 1.
bMost parsimonious tree from each dataset.
SLog likelihood.
d Difference in log likelihood compared to that of the best tree.
SStandard deviation of log likelihood.
ST-value.
g Probability of getting a more extreme T-value with the two-tailed test under the null hypothesis of no
difference between the two trees.
h Isolate 2619 was placed in the same clade containing the isolates WH83, WHK, WH9BR, 10, 62-2, 28-1,
WHV, 114,400, MX3, and 2867.
* Significant at P-0.05.








the isolate 2619 inside the major clade, it was no longer significantly different.

Therefore, the differences in tree topologies were due to differences in nucleotide

sequences of the intron area of the P-tubulin gene in the isolate 2619 as shown in the

Figure 2-11.

Even though these three datasets should not be combined because they have

heterogeneous phylogenetic signals, the most parsimonious tree inferred from the

combined datasets showed basically the same topology of the trees inferred from

individual datasets. A strict consensus of the 24 most parsimonious trees shown in Figure

2-16, belongs to the same two well-supported groupings of isolates observed in the trees

of individual datasets, with 100% bootstrap support for the minor clade grouping of the

isolates from Texas and 99% bootstrap support for the major clade grouping of the

isolates from the other origins. With combined data, the isolate 2619 was grouped within

the major clade. The topology of the maximum likelihood tree of combined data was the

same as the maximum parsimony tree (Figure 2-17); and the neighbor-joining tree, as in

the individual datasets, showed a more branched topology but with only two well

supported clades, which grouped the same isolates like the other trees (Figure 2-18).

All datasets analyzed separately or combined, through maximum parsimony,

maximum likelihood or neighbor-joining methods, suggested that the isolate WH9BR,

identified as C. rodmanii, was not distinct from the other members of the major clade that

grouped isolates from several geographical locations, including some isolates that showed

conidial size and morphology in concordance with the description of C. piaropi, such as

WH83, RR29, 10, 28-1, 18-2, and MX3. In addition, the characteristics and dimensions
















10

114
28-1

2867

MX3

99 62-2
WH83

WH9BR

WHK

2619

-400
WHV

.oo r TX20
TX18

Cercospora beticola
(outgroup)



Figure 2-16. Strict consensus of 24 most-parsimonious trees of length 246 based on
parsimony analysis of combined elongation factor-la, P-tubulin, and histone 3 data sets.
Bootstrap replication percentages are indicated above the nodes.















-WH83

WH9BR

2867

10

114

98 --WHV

62-2

400

-WHK

MX3

28-1

2619

TX18

L---- TX20

Cercospora beticola
(outgroup)








Figure 2-17. Maximum likelihood tree inferred from the combined elongation factor-la,
P-tubulin, and histone 3 datasets. Bootstrap replication percentages are indicated above
the nodes.


















WH83

WH9BR
WHK

62-2

114

2619

MX3
100
2867

10

400
WHV

28-1

100 TXI8
TX20

Cercospora beticola
(outgroup)


Figure 2-18. Neighbor-joining tree on distances inferred from the combined elongation
factor-la, P-tubulin, and histone 3 datasets. Bootstrap replication percentages are
indicated above the nodes.








of conidia were unreliable criteria for taxonomic differentiation of isolates that composed

the two groupings defined by the phylogenetic analysis.

Discussion

More than 3000 species names have been validated in Cercospora (Pollack,

1987). The circumscription of species in this genus has been based on host affiliation and

conidial morphology of the species (length, width, and morphology of bases and tips) and

conidiophores (length, diameter, geniculation, and fasciculation) (Chupp, 1953; Ellis,

1971). As rationalized by Ellis (1971), "it has been customary for plant pathologists and

mycologists to describe as new any Cercospora found on a host plant for the first time".

Chupp (1953) and Ellis (1971), discussed the reliability of the criteria used for species

circumscription in Cercospora. Some species in this genus have wide host ranges and the

size of conidia and conidiophores can have variations induced by changes in

environmental conditions, especially humidity. These factors complicate the taxonomy

and identification of species belonging to this genus. The study reported here shows that

the criteria used by Conway (1976a) for differentiating the species C. rodmanii from C.

piaropi are not adequate to provide a clear identification of these species.

This study inferred phylogenetic relationships for a population of isolates

belonging to the complex C. piaropi/C. rodmanii and obtained from several geographical

locations. The phylogenetic trees inferred from three independent lines of evidence were

concordant in showing that the population of isolates of C. piaropi/C. rodmanii from

waterhyacinth grouped in two well-supported clades: a major clade, which grouped

isolates collected in the center of origin of waterhyacinth (Brazil) and in several of the

host's adventive areas; and a minor clade, which grouped isolates only from Texas. The








characteristics and dimensions of conidia were unreliable criteria for differentiation of

isolates that composed these two groups, and the isolate WH9BR that is the type culture

of C. rodmanii was not differentiated, based on DNA sequencing data, from the isolates

that had morphological features of C. piaropi. Thus, the differentiation of the species C.

piaropi and C. rodmanii based on phenotypic traits, besides being not clear in some

situations was also not supported by the DNA sequencing data. Based on the finding that

isolates of these species grouped together in a well-supported clade, they should be placed

in the same species according to the phylogenetic species concept (Nixon and Wheeler,

1990; O'Donnell and Cigelnik, 1997).

The differentiation between the species C. piaropi and C. rodmanii is difficult.

Martyn (1985) and Morris (1990) noted that C. piaropi and C. rodmanii appear to be

closely related, both in morphology and disease symptomatology. In addition, in the

emended description of C. piaropi, Conway (1976a) described conidia of C. piaropi as

acicular with obconic bases, but earlier descriptions refer to them as obclavate to acicular

(Thirumalachar and Govindu, 1954), truncate, tapering towards the tip (Tharp, 1917;

Freeman and Charudattan, 1974), or as truncate (Chupp, 1953). Morris (1990) observed

that C. piaropi conidia were mostly acicular with truncate bases, with a few conidia being

obconic. In reference to the size of conidia, Freeman and Charudattan (1984) observed

that frequently both long and short conidia were produced on dead leaf tissue when leaves

infected by C. piaropi were incubated under moist conditions. In this study, the

possibility of having the isolates mixed was ruled out because all cultures were obtained

as monocultures from hyphal tips, and this procedure was done twice.








Therefore, to avoid problems of identification and also problems of

communication among scientists and quarantine officials, I propose a second emendation

to the description of the species C. piaropi and to consider the species C. rodmanii as its

synonym. Cercospora piaropi should remain as the valid name based on the principle of

priority of publication of the International Code of Botanical Nomenclature (Greuter et

al., 1994).

The two groups defined by the phylogenetic analysis as major and minor clade

groupings could be considered to represent two different species of Cercospora,

according to the phylogenetic concept of species (Nixon and Wheeler, 1990; O'Donnell

and Cigelnik, 1997). However, until additional criteria are available to delimit

Cercospora species, these groups must be considered to be taxonomically conspecific.

Wang et al. (1998) also considered conspecific the two groups found in C. zeae-maydis,

identified through AFLP analysis and DNA sequencing of the 5.8S ribosomal DNA and

the ITS regions. Indeed, differently from this study, Wang et al. (1998) found that the

DNA sequences of the 5.8S ribosomal DNA and the ITS regions had eight informative

characters which supported the differentiation of the two groups observed through AFLP

analysis. The authors interpreted these two groups as being sibling species.

The study reported here is a contribution for the taxonomy of species in

Cercospora. However, many questions still need to be answered, such as whether the

Cercospora species defined by host affiliation and morphological criteria correspond to

the distinct lineages identified by cladistic analysis, and whether the form-genus

Cercospora is a monophyletic group. Studies to determinate the correlation between

lineages defined by host affiliation and lineages defined by molecular markers, as well as








their common ancestry, have been presented for other groups of fungi and have been very

insightful. For example, molecular studies identified several clonal lineages with

independent evolutionary origins in Fusarium oxysporum Schlechtend.:Fr.f.sp. cubense

(E.F.Sm) Snyder & Hans, a taxon defined by its pathogenicity to bananas (Koenig et al.,

1997; O'Donnell et al., 1998b). In addition, these pathogens of banana could be as

closely related to pathogens of other hosts. In the anther smut fungus Microbotryum

violaceum (Pers.:Pers.) Deml. and Oberw. correlation was found between haplotypes,

defined by partial DNA sequence of y-tubulin gene and their respective host species of

origin (Garr et al., 1997).

The utility of using elongation factor-la, P-tubulin, and histone 3 genes for

phylogenetic studies at the species level or below has been demonstrated in previous

work. DNA sequences of the P-tubulin gene were used in phylogenetic studies in

Fusarium and in the Gibberellafujikuroi (Sawada) Wollenw. species complex

(O'Donnell et al., 1998a; O'Donnell and Cigelnick, 1997) and in Aspergillus section

Fumigati (Geiser et al., 1998a). DNA sequences of the histone 3 gene were used to

differentiate Fusarium species associated with conifers (Donaldson et al., 1995). More

recently, DNA sequences of elongation factor-1 a were used for phylogenetic studies in

Fusarium oxysporum (Schlecht.) Snyd. and Hans. (O'Donnell et al., 1998b). In this

study, these three loci were shown to be appropriate for phylogenetic inferences in

Cercospora. These loci can provide diagnostic tools needed to investigate species

boundaries and to correlate the host range and biogeography of these fungi.

The question of whether independent datasets should be analyzed separately or

combined and analyzed simultaneously is still controversial. Miyamoto and Fitch (1995)








defend that independent datasets should rarely be combined but should be kept separate

for phylogenetic analysis because their independence increases the significance of

corroboration. According to de Queiroz et al. (1995), assuming that the goal is to

uncover the true phylogeny of the entities in question, arguments to combine data based

on the notion that one should use the "total evidence" available, or that the combined

analysis gives the tree the greatest descriptive and explanatory power, are not compelling.

However, to combine datasets can enhance detection of real phylogenetic groups. On the

other hand, if there is heterogeneity among datasets with respect to some property that

affects estimation of phylogeny, then combining the data can give misleading results. In

this study, even though according to the statistical test the three datasets should not be

combined, the phylogenetic tree inferred from the combined data did not show misleading

results (Figures 2-16; 2-17; 2-18).

Taking in account the limitations in the number of samples analyzed, the

phylogenetic analysis presented in this study provides some insights about a

biogeographic hypothesis for Cercospora on waterhyacinth. The grouping of isolates of

C. piaropi/C. rodmanii from different geographical origins in a same clade, including the

isolates collected in the center of origin of waterhyacinth (Brazil and Venezuela),

suggests that the fungi may have been spread together with the plant host, from its center

of origin. It has been documented that waterhyacinth was dispersed, mostly by man, from

the lowlands of tropical South America to many tropical and subtropical regions of the

world (Barrett, 1977; 1988). In addition, the occurrence of a distinct population of the

pathogen only in a single region outside the center of origin of the host plant (Texas

isolates), suggests that this population may have adapted to waterhyacinth from other








host(s). Thus, a multiple origin of Cercospora pathogens of waterhyacinth cannot be

ruled out.

Biogeographic hypotheses from phylogenetic evidence has not been proposed for

species in Cercospora, but have been proposed for a few other groups of fungi. For the

Gibberellafujikuroi species complex, gene trees inferred from P-tubulin and 28S rDNA

supported a phylogeny consistent with species radiations in South America, Africa, and

Asia (O'Donnell et al., 1998a). These analyses placed the American clade, with 12

species, as a monophyletic sister-group of an African-Asian clade which had 12 and 8

species, respectively. The biogeographic hypothesis proposed in their study reflected

vicariant events associated with the fragmentation of the super-continent Gondwana.

Otherwise, other groups of fungi have a more complex biogeographic history, such as the

genus Lentinula that includes cultivated shiitake mushrooms (Hibbett et al., 1998). Here,

based on phylogenetic analysis of the 5.8S rDNA gene and its ITS flanking regions,

Hibbett et al. (1998) hypothesized that the present distribution of Lentinula in four

continents must result from some combination of vicariance, dispersal, and extinction.

Data from this study suggest a dispersal hypothesis for the Cercospora species

from waterhyacinth following the path of the plant host from its center of origin and in

combination with possible adaptations of native populations of Cercospora in the

adventive areas of the plant host. However, further studies are needed to address this

question, including a greater number of isolates sampled from different areas in the center

of origin and the adventive areas of waterhyacinth.

The large unexpected difference in nucleotide sequences observed in the segment

of P-tubulin gene of the isolate 2619 compared with the other isolates may be due to the








presence of extra copies of this gene in the genome of this isolate, as was observed by

Tsai et al. (1994) in the genome of EpichloE species. These authors interpreted this

finding as being the result of inter-specific hybridization among species belonging to this

genus. However, since the Southern blot analysis of the P-tubulin gene disproved the

hypothesis of extra copies of this gene in the genome of the isolate 2619 (Figure 2-12),

the reasons of such differences in nucleotide sequences are not clear. One possibility is

the occurrence of a history of recombination in this isolate. For instance, based on the

principle that full compatibility among gene genealogies indicates complete asexuality,

and incompatibility indicates mixis, Koufopanou et al. (1997) concluded that the

incompatibility among five protein-coding-gene genealogies inferred from isolates of the

human fungal pathogen Coccidioides immitis Rixford and Gilchrist was an indication of

sexual recombination in the population of this fungus. The same approach was given by

Geiser et al. (1998b) to determine the occurrence of history of recombination in

Aspergillusflavus Link, based on the lack of concordance among gene genealogies from

five protein-coding genes among a population of isolates of this fungus.

This analysis clearly demonstrated the utility of the elongation factor-1 a, 3-

tubulin, and histone 3 genes for phylogenetic analysis of closely related species of fungi.

All three genes harbor considerable phylogenetically informative variation that can

provide diagnostic tools needed to investigate boundaries of fungal species and even

biogeography. Phylogenetic reconstruction based on independent loci, such as the

protein-coding genes used in this study, can be very useful for biological control. It can

provide tools to delimite the species used as biocontrol agents, to track them in the field,

and to understand aspects of their origin and reproductive biology.








I propose a second emendation to the species description of C. piaropi, a species

originally described by Tharp (1917) as follows:

Cercospora piaropi sp. nov.

Spots ovate, grayish-tan centered with purplish-black borders somewhat raised
above, brighter above than below, 1.5-3 x 3-5 mm in diameter, or larger by
confluence; conidiophores epiphyllous, fasciculate but very few in each fascicle,
sparse, bright brown with yellowish apices, denticulate, sometimes branched,
pluriseptate, 100-125 x 3.5-4.5 pim; conidia hyaline, truncate at base, upward
attenuate, pluriseptate at maturity, 80-140 x 3pm.
On living leaves on Piaropus crassipes (Mart.) Britton, Palestine, Texas, Oct.30,
1914, 1. M. Lewis & B. C. Tharp.

Conway (1976a) emended the description given by Tharp (1917) as follows:

Cercospora piaropi Tharp emend. Conway

Leaf spot ovate, dark brown, later with a grayish-tan center with dark brown
borders, 1.5-3 x 3.5 mm, larger by confluence; fruiting amphigenous; stromata
lacking or a few brown cells; conidiophores borne singly or in fascicles of two to
nine, dark brown, multiseptate, not branched, sympodial, 55-200 x 2.5-5 pm; conidia
hyaline, acicular, straight to mildly curved, multiseptate, base obconic, 25-(95)-220 x
2-(3.5)-5 im.
SPECIMEN: Deposited as the National Fungus Collection (BPI).
CULTURE: Deposited at the Florida Division of Plant Industries, Gainesville,
Florida, FTCC719.

The new species, C. rodmanii, was described by Conway (1976a) as follows:

Cercospora rodmanii Conway sp. nov.

Maculae nigrae punctulatae ad rodundas, 1-3 mm latas, phyllum chloriticum et
petiolus chloriticus, extremus phylli mortuus; conidiophora fasciculata, 3-12,
amphigena, brunnea, sympodialia, orientia ex stromate, emergentia per stoma; 84-
(145)-284 x 4-(4.5)-5 lim; conidia hyalina, truncata, acicularia, multiseptata, 66-
(172)-374 x 3-(4)-5 pm.
Pycnidia brunnea, ostiolata, globosa 80-95 x 80-110 jm, sub stomate, deinde
erumpentia; ostiola 30-40 x 25-30 lm; conidia hyalina, in forma bacillorum 2-3.5 x
1-1.5 pum.
HABITAT: In phyllis Eichhornia crassipes (Mart.) Solms.

Leaf spots black, punctate to circular (1-3 mm diam), leaf and petiole chlorotic,
tip of leaf necrotic, conidiophores amphigenous, 3-12 in each fascicle, brown








sympodial, arising from a well developed stroma, emerging through the stroma, 84-
(145)-284 x 4-(4.5)-5 gm; conidia hyaline, truncate at base, acicular, multiseptate,
66-(172)-374 x 3-(4)-5 gm.
ASSOCIATED STATE: Asteromella pynidia dark brown, ostiolate, globose, 80-
95 x 80-110 [tm, substomatal, later erumpent, ostiole 30-40 x 25-30 Rim; condia
hyaline, bacilliform 2-3.5 x 1-1.5 lim.
TYPE SPECIMEN: Deposited at the National Fungus Collection (BPI).
TYPE CULTURE: Florida Division of Plant Industries, Gainesville, Florida,
FTCC 715.
HABITAT: On leaves of waterhyacinth (Eichhornia crassipes [Mart.] Solms).
Collected by K.E. Conway, Rodman Reservoir, Orange Springs, Florida.

Herein I propose a second emend of C. piaropi as follows:

Cercospora piaropi Tharp emend. Tessmann and Charudattan

Leaf spot grayish-tan, dark brown to black, ovate, punctate to circular, 1-3 x 3-5
mrn in diameter, or larger by confluence, leaf and petiole chlorotic, tip of leaf
necrotic; conidiophores amphigenous, stroma lacking, or with few brown cells or
well-developed; conidiophores borne singly or in fascicles of two to twelve, brown
or dark brown, multiseptate, sometimes branched 55-284 x 2.5-5 pm; conidia
hyaline, truncate at base or obconic, acicular, multiseptate, 25-374 x 2-5 Rm.
Associated state: Asteromella pycnidia dark brown, ostiolate, globose, 80-95 x 80-
110 gtm, substomal, later erumpent, ostiole 30-40 x 25-30 im; conidia hyaline,
bacilliform 2-3.5 x 1-1.5 jim.
Specimen: Deposited at the National Fungus Collection (BPI).
Culture: Deposited at ATCC.














CHAPTER 3
PATHOGENIC VARIABILITY AND BIOCHEMICAL CHARACTERIZATION OF
CERCOSPORA SPECIES FROM WATERHYACINTH


Introduction


Waterhyacinth (Eichhoria crassipes [Mart.] Solms), an aquatic plant indigenous

to lowland tropical South America (Perfound and Earle, 1948; Barrett, 1988), was spread

worldwide by man. Its free-floating habit and capacity for rapid vegetative propagation

has enabled it to become one of the most noxious aquatic weeds in many tropical and

sub-tropical regions of the world. Problems caused by this weed include clogged

irrigation canals, blocked waterway transport routes, water losses, and reduction in fish

populations in reservoirs (Pieterse, 1990). The strategies used to control this weed have

included mechanical removal, chemical control with herbicides, and biological control

with pathogens and insects (Charudattan, 1986; 1990). Among the pathogens studied as

biocontrol agents, the species Cercospora piaropi Tharp and C. rodmanii Conway have

been shown to decrease waterhyacinth biomass, and in some instances to cause

substantial decline of waterhyacinth populations (Charudattan et al., 1985; Freeman and

Charudattan, 1984; Martyn, 1985; Morris, 1990).

The species C. piaropi was described by Tharp (1917) in Texas, and almost sixty

years later, Conway (1976a) emended the description of C. piaropi and described a new

species, C. rodmanii. The new species was described based on plant specimens collected

at the Rodman reservoir, Florida, where a severe epidemic of a Cercospora leaf-spot








disease caused the decline of the population of waterhyacinth in that area. This new

species was differentiated from C. piaropi mainly based on conidial morphology, disease

symptomatology, and on C. rodmanii being a more aggressive pathogen than C. piaropi.

This idea was reinforced by the early reports that described C. piaropi as a pathogen not

able to cause serious damage to waterhyacinth (Nag Rag, 1965; Freeman and

Charudattan, 1974) and because the epidemics recorded at the Rodman Reservoir was the

first devastating epidemic of a Cercospora leaf spot on waterhyacinth ever noticed.

However, C. piaropi also was recorded later to cause serious damage to waterhyacinth in

Texas (Martyn, 1985) and in South Africa (Morris, 1990), even though these authors had

difficulties to differentiate this species from C. rodmanii. Indeed, the differentiation

between these two species with Conway's criteria has not been an easy task, as discussed

in Chapter 2. Until the taxonomic status of C. piaropi and C. rodmanii are changed, as

discussed in Chapter 2, this study will refer to these species as the complex C. piaropi/C.

rodmanii complex rather than referring to them as individual species.

The occurrence of pathogenic variability, even though it was presumed to exist,

has not been identified in C. piaropi/C. rodmanii. In addition, isolates of C. piaropi/C.

rodmanii, as many other Cercospora species, show a great variation in their cultural

features, such as mycelial color, pigmentation color and intensity, and growth rate. The

extent these factors are related to pathogenicity or virulence of these isolates is unknown.

Such information would be very important to optimize large-scale inoculum production,

which is an important step in the bioherbicide strategy to control this weed (Charudattan

et al., 1985; Charudattan, 1986). In Cercospora spp., two colored secondary metabolites

have been identified as phytopathogenic toxins: a red-purple compound, called








cercosporin, which was identified in several species in this genus (Assante et al., 1977;

Jenns et al., 1989; Lynch and Geoghegan, 1977; Upchurch et al., 1991), and a yellow

compound, identified only in Cercospora beticola, which corresponds to a group of six

toxins, named beticolins and formerly known as Cercospora beticola toxin (CBT) (Milat

and Blein, 1995).

The objective of this study was to determine the extent of variation in virulence

and in some physiological characteristics, including the production of phytopathogenic

toxins, among a population of C. piaropi/C. rodmanii isolates collected in several

geographic locations. An additional objective was to determine the extent of variation of

fatty acid methyl ester profiles (FAME), with the purpose to find useful biochemical

markers to differentiate isolates, populations, and species among a collection of isolates

of C. piaropi/C. rodmanii. FAME profiles have been used as biochemical characters to

address taxonomic issues in fungi at species and sub-species levels (Augustyn et al.,

1990; Bentivenga and Morton, 1996; Berger et al., 1991; Graham et al., 1995; Johnk and

Jones, 1993; Malfeito-Ferreira et. al. 1989; Martinez et al., 1991; Nemec et al., 1997; da

Silva et al., 1998; Stahl and Klug, 1996; Viljoen and Kock, 1989). The knowledge of

pathogen variability can provide valuable information for biological control programs to

select the most effective strains, to infer the stability of these strains, and to define the

boundaries of species and populations.

Materials and Methods

Fungal Isolates and Cultural Characteristics

The origin, designation, and the name of the collectors of the isolates used in this

study are listed in Table 3-1. All isolates were obtained from symptomatic leaves of








waterhyacinth and monocultures were obtained from mycelial tips. These isolates have

been preserved in the fungal collection of the Biological Control of Weeds Laboratory of

the Plant Pathology Department, University of Florida.

Fungal cultural characteristics were studied on plates containing 10 ml of PDA.

Three replicates for each isolate were inoculated with 4-mm-diameter mycelial plugs

removed from the margin of 7-day-old colonies growing in 9-cm-diameter plates

containing 20 ml of PDA. Inoculated plates were incubated in a growth chamber at a

temperature of 2420C with 12 h light, and radial growth was measured 8 days later. The

mean diameter was calculated as the average of two diagonal measurements of colony

diameter on each plate. Colony morphology, pigment diffusion, and mycelial

pigmentation were also recorded at 8 and 14 days after inoculation. The experiment had a

completely randomized design with three replicates for each isolate and was repeated

once.

Toxin Analysis

Toxin analysis was based on the protocol developed by Milat and Blein (1995).

Isolates were cultured in 50-ml test tubes containing 15 ml of V8 broth medium (200 ml

V8 juice, 3 g CaCO3, and 800 ml of tap water; sterilized). Each tube was inoculated with

eight, 4-mm-diameter, mycelial plugs removed from the margin of 7-day-old colonies

growing in 9-cm-diameter plates containing 20 ml of PDA. After incubation for 14 days

at 2520C under constant light, the mycelium was separated from the broth by filtration

using a double-layered cheese cloth, and blended for about 5 sec in a Waring blender in

ethyl acetate (20 ml/g of wet mycelium). The crude extract was separated from mycelial

debris and resolved by thin-layer chromatography using pre-coated TLC plates of silica









Table 3-1. Designations and geographic origin of the isolates of Cercospora piaropilC.
rodmanii analyzed in this study.


Designation"
WH83*
WH9R
WH9BR*
WHK*
2619*
SGS25
SGS32
SGS35
SGS49
J2
J6
175-99
175-101
175-102
175-104
175-108
LJ37
TX2
TX15
TX18*
TX20*
RR4
RR22
RR24
RR27
RR29
RR31
BA55
BA57
BA59
BC31
2702
2703
2704
65-2
67-1
69-1
70-12
61-5
62-2
62-1
62-4


Geographic origin
Gainesville, FL, USA
Rodman Reservoir, FL, USA
Rodman Reservoir, FL, USA
Kissimee, FL, USA
Suwannee River, FL, USA
Marion Co., FL, USA
Marion Co., FL, USA
Marion Co., FL, USA
Marion Co., FL, USA
Marion Co., FL, USA
Marion Co., FL, USA
Sarasota Co., FL, USA
Sarasota Co., FL, USA
Sarasota Co., FL, USA
Sarasota Co., FL, USA
Lee Co., FL, USA
Leon Co., FL, USA
Lake Conroe, TX, USA
Lake Conroe, TX, USA
Lake Conroe, TX, USA
Lake Conroe, TX, USA
Rodman Reservoir, FL, USA
Rodman Reservoir, FL, USA
Rodman Reservoir, FL, USA
Rodman Reservoir, FL, USA
Rodman Reservoir, FL, USA
Rodman Reservoir, FL, USA
Gainesville, FL, USA
Gainesville, FL, USA
Gainesville, FL, USA
Vero Beach, FL, USA
Minas Gerais, Brazil
Minas Gerais, Brazil
Minas Gerais, Brazil
Alagoas, Brazil
Alagoas, Brazil
Alagoas, Brazil
Alagoas, Brazil
Pernambuco, Brazil
Pernambuco, Brazil
Pernambuco, Brazil
Pernambuco, Brazil


Collector
R. Charudattan
K. Conway
K. Conway
R. Charudattan
R. Charudattan
J. DeValerio
J. DeValerio
J. DeValerio
J. DeValerio
J. DeValerio
J. DeValerio
D. Tessmann
D. Tessmann
D. Tessmann
D. Tessmann
D. Tessmann
D. Tessmann
D. Tessmann
D. Tessmann
D. Tessmann
D. Tessmann
D. Tessmann
D. Tessmann
D. Tessmann
D. Tessmann
D. Tessmann
D. Tessmann
D. Tessmann
D. Tessmann
D. Tessmann
J. DeValerio
R. Charudattan
R. Charudattan
R. Charudattan
R. Charudattan
R. Charudattan
R. Charudattan
R. Charudattan
R. Charudattan
R. Charudattan
R. Charudattan
R. Charudattan








Table 3-1--continued
Designation Geographic origin Collector Date
46-4 Pernambuco, Brazil R. Charudattan 1997
49-1 Pernambuco, Brazil R. Charudattan 1997
49-2 Pernambuco, Brazil R. Charudattan 1997
16-1 Corumba, Mato G. do Sul, Brazil R. Charudattan 1995
18-2 Corumba, Mato G. do Sul, Brazil R. Charudattan 1995
28-1* Rio Verde, Mato G. do Sul, Brazil R. Charudattan 1995
10* Sao Paulo City, SP, Brazil R. Charudattan 1996
34 Rio Grande do Sul, Brazil R. Charudattan 1997
2867* Mexico R. Charudattan 1994
2943 Mexico R. Charudattan 1994
MX3* Mexico R. Charudattan 1997
WHV* Venezuela R. Charudattan 1982
114* South Africa M. Morris ?
279 South Africa M. Morris ?
400* Zambia M. Morris ?

a Designation corresponds to notation of isolates used in this study. Isolates labeled with
an asterisk (*) were included in the fatty acid analysis.








gel without fluorescent indicator (E. Merck, Darmstadt, Germany). A total of 5 pl of

crude extract was spotted in the TLC plate for each isolate. Chromatograms were

developed using chloroform/methanol/water (80:20:2, v/v) as elution mixture. The

standards used were cercosporin from Sigma Chemical Co.(St. Louis, MO), and

beticolin-1, obtained from Dr. L. Milat (INRA, Dijon Cedex, France). In addition, the

ultraviolet spectrum of the crude extract was determined in a spectrophotometer and

cercosporin was quantified in each sample through reading of its absorbance in

spectrophotometer at 473 nm. The amount of cercosporin was calculated using the

formula: absorbancece at 473 nm/31,455)534] gg, where 31,455 equaled the molar

extinction coefficient at 473 nm and 534 equaled the molecular weight of cercosporin.

This formula contained a corrected extinction coefficient with ethyl acetate as solvent,

since in the original formula, used by Jenns et al. (1989) and Velicheti and Sinclair

(1994), 5 N KOH was used as solvent. This correction was needed because the

maximum absorption of cercosporin in 5 N KOH was 480 nm and in ethyl acetate was

473 nm. Each sample corresponded to an individually growing isolate. The experiment

had three replicates and was performed twice in a randomized block design.

Pathogenicity Test and Virulence Analysis

Waterhyacinth plants for virulence tests were vegetatively propagated from plants

collected from Lake Alice, located on the campus of the University of Florida. The plants

were propagated in pots with tap water supplemented (0.05% w/v) with chelated iron

(Keel-Iron, NaFe EDTA 5%; Chase & Company, Sanford, FL). For the pathogenicity

tests and screening for virulence, daughter ramets (=offsets or clones) of waterhyacinth,

with 2 leaves, and with approximately the same root mass, were placed in pots containing








125 ml of tap water with the supplement described above. Each pot had one plant and the

volume of the liquid in the pots was maintained at the initial level by daily additions of

water.

Due to irregular, or lack of, sporulation in culture, the inoculum was prepared

from mycelium grown in still liquid cultures. Isolates were cultured in 250-ml flasks

containing 50 ml of V8 broth medium (200 ml V8 juice, 3 g CaCO3, and 800 ml of tap

water; sterilized). Each tube was inoculated with eight 4-mm-diameter mycelial plugs

removed from the margin of 7-day-old colonies grown in 9-cm-diameter plates containing

20 ml of PDA. After inoculation for 14 days at 2530C with 12 h light, the mycelium

was separated from the broth by filtration using a double-layered cheese cloth, and

blended for about 5 sec in a Waring blender at a concentration of 80 mg mycelium per ml

of water. The suspension was amended with 0.5% Metamucil (psyllium mucilloid;

Procter & Gamble, Cincinnati, OH) and 0.05% Silwet L-77 (polyalkyleneoxide modified

heptamethyltrisiloxane, 0.02% v/v, OSI Corp., Loveland Ind., Inc., Greeley, CO).

A bioassay was developed which consisted of immersing waterhyacinth leaves,

attached to plants, in a suspension of inoculum. Control leaves were immersed in a

solution containing only the amendments. This procedure was used after other

inoculation procedures, such as foliar spraying and droplet deposition on leaves, were

evaluated and found to be unsuitable. The inoculated and control plants were placed in a

dew chamber in darkness, at 2520C for 12 h, and then held in a greenhouse for 2 wk.

This experiment was conducted in a quarantine greenhouse. Disease severity was

assessed 7 and 14 days after inoculation. Disease was rated using a rating scale, where: 0

= no symptoms; I = less than 1% of the lamina surface with spots; 2 = greater than 1%








and less than 10% of the lamina surface with spots; 3 = greater than 10% and less than

25% of the lamina surface with spots; 4 = greater than 25% and less than 50% of the

lamina surface with spots; 5 = greater than 50% and less than 75% of the lamina surface

with spots; 6 = greater than 75% of the lamina surface with spots; and 7 = dead lamina.

The treatments were arranged in a randomized block design with four replications

(pots) per isolate, each with one 2- to 3-leaved plant per pot. Statistical analyses of

pathogenic variability, growth rate, and cercosporin production were done with the GLM

and CORR procedures of SAS (SAS Institute, Cary, NC).

Fatty Acids Analysis

Fourteen isolates of C. piaropi/C. rodmanii, representing different geographical

locations, and three other species of Cercospora (outgroups) were included in this study

(Table 3-1). All fungi were grown in 80 ml of modified TSB (trypticase soy broth with

10 g dextrose per liter; BBL Microbiology Systems, Becton Dickinson and Co.,

Cockeysville, MD) in 250-ml flasks in a slow-shake culture (130 rpm) for 4, 5, and 6

days at 242C in dark before harvest and fatty acid extraction. Four, 4-mm diameter

plugs removed from the periphery of 7-day-old cultures in PDA were used as initial

inoculum. Cultures were harvested using a side-arm Erlenmeyer flask fitted with a

Buchner funnel and filter attached to a vacuum pump. A nylon-type filter (polypropylene,

mesh opening of 105 .m) was used to prevent the sample from becoming contaminated

with paper fibers. After harvest, the fungal mycelium was transferred to 15-ml

polypropylene tubes and kept in a freezer at -700C until lyophilization.








Fifty-gram (dry weight) of samples of fungal tissue were placed in clean screw-

cap test tubes (13 by 100 mm; with Teflon cap liners), 2 ml of a saponification reagent

(45g sodium hydroxide in I liter of 50% methanol) was added and the mixture was

homogenized with a vortex mixer for 10 sec. The homogenate was then saponified at

1000C in a water bath for 5 min, homogenized in a vortex mixer for 10 sec, and kept in a

water bath for 25 min at 100C, then cooled in a room-temperature water bath. To

methylate the liberated fatty acids, 2 ml of 54% 6 N HC1 in methanol was added to each

tube. Sub-samples were then placed in an 800C water bath for 10 min and immediately

cooled to room temperature. To extract fatty acid methyl esters from the aqueous phase,

1.25 ml of 50% hexane-50% methyl tert-butyl ether was added to each tube, and the tubes

were rotated end-over-end for 10 min. Next, the aqueous phase (bottom of tube)

contained fungal debris was removed with a Pasteur pipette, and 3.0 ml of 1.2 NaOH in

H20 was added to each tube; the tube was then rotated end-over-end for 5 min. Finally,

the organic phase (top of tube) contained the fatty acid methyl esters was removed from

the tubes and placed in a crimp-top gas chromatography vial.

Fatty acid extracts were analyzed by gas-liquid chromatography with the

Microbial Identification System (MIS) developed by Microbial I.D., Inc. (Newark, DE).

This system was designed to analyze the fatty acid composition of unknown bacteria and

to identify them by matching the fatty acid composition of an unknown organism to one

of the fatty acid profiles in its computer database of known organisms. The MIS consists

of a chromatographic unit chromatographh, integrator, and autosampler) coupled to a

computer system. The gas-liquid chromatograph is equipped with a 25 m by 0.2 mm

phenyl-methyl-silicone-fused capillary column (Hewlett-Packard, Wilmington, DE) and a








flame ionization detector. Data from chromatographic analysis are sent to the computer

system, where fatty acids are identified on the basis of their retention times relative to

known standards, and quantified relative to other fatty acids in the sample on the basis of

peak width and area data. The system is calibrated with known fatty acid standards when

it is started and after every 10th sample. Results of each sample analysis are printed out in

a fatty acid composition report and also stored on a hard disk within the computer.

Samples were run through the gas chromatography column for 38 min, long enough for

fatty acids up to 28 carbons long to pass through.

Fatty acids profiles of individual isolates were based on analysis of the cellular

fatty acid content of three independently grown cultures in each of the three harvest days.

The value for each fatty acid in a given profile was the mean from all analyzed cultures of

that isolate. Concentrations of each fatty acid was expressed as a percentage of the total

fatty acid content. To determine if the fatty acid compositions of the isolates were

statistically different, discriminant analysis, implemented by SAS (SAS Institute, Cary,

NC), was used to test the hypothesis that isolate x(fal, fa2, fa3...fa) = y(fal, fa2, fa3...fan)

= ...= z(fal, fa2, fa3... fa), where fa is fatty acid (Stahl and Klug, 1996).

Results

Cultural Characteristics

The isolates compared in this study formed colonies on PDA with colors that

ranged from pale- to dark gray, and pale pink to pink mycelium. The colonies produced

reddish purple or yellow compounds, with different intensities. In addition, some isolates

showed dark gray mycelia and these did not produce pigments (Figure 3-1). Reddish

purple pigment was produced by isolates with all the mycelial colors described above,








while only isolates with-pale pink mycelium produced yellow pigment. The growth rate

among the isolates ranged less than one fold, from 2.17-2.34 to 4.00-4.29 mm per day

(Table 3-2).

Virulence and Toxin Analysis

A total of 55 isolates of C. piaropi/C. rodmanii were used in the virulence

testing. Significant differences (P<0.05) were observed in the ability of the isolates to

cause disease (i.e., difference in virulence) among the isolates (Table 3-3). The most

virulent isolates necrosed almost 75% of the leaf area in 14 days after inoculation. Most

isolates showed intermediate levels of virulence, and some isolates were nonpathogenic

to waterhyacinth. There was no relation between the degree of virulence and the

geographical origin of the isolates. Highly virulent and nonpathogenic isolates were

found for collections from inside and outside South America, the geographical center of

origin of waterhyacinth.

The ranking of the isolates for level of virulence was conserved between the

experiments, with some variation due to uncontrolled factors (Figure 3-2). The disease

grades of the third experiment were lower than the grades of the first and second

experiment. This variation is due to influence of external temperature, since the third

experiment was run at a cooler temperature (May) compared to the first and second

experiments (June/July).

All highly virulent isolates produced purple pigments while the yellow-pigment

producers were weak to moderate in virulence. The nonpigment-producers were

nonpathogenic (Table 3-3).












































Figure 3-1. An example of differences in colony characteristics of Cercospora piaropilC.
rodmanii from waterhyacinth after growth for 7 days on PDA.








Table 3-2. Mycelial growth rates of isolates of Cercospora piaropilC. rodmanii from
waterhyacinth, in mm day .


Isolate
65-2
18-2
1-75-99
16-1
114
70-12
TX2
34
LJ37
279
400
MX3
WH9BR
SGS35
WH83
62-1
175101
WHV
TX15
2704
RR4
WHK
SGS32
WH9R
175102
2943
28-1
67-1
2703
10
BA59
2867
69-1
175108
49-1
J6
J2
49-2
2702
61-5
RR24
62-4


1st Experiment
4.29 az
4.09 ab
3.92 abc
3.75 abcd
3.59 bcde
3.59 bcde
3.46 cdef
3.38 cdef
3.37 cdef
3.33 cdefg
3.29 defgh
3.10 efghi
3.00 efghij
3.00 efghij
2.92 fghijk
2.92 fghijk
2.88 fghijkl
2.88 fghijkl
2.75 ghijklm
2.75 ghijklm
2.75 ghijklm
2.71 hijklmn
2.71 hijklmn
2.71 hijklmn
2.71 hijklmn
2.71 hijklmn
2.67 ijklmno
2.67 ijklmno
2.67 ijklmno
2.63 ijklmno
2.59 ijklmno
2.58 ijklmno
2.55 ijklmno
2.55 ijklmno
2.54 ijklmno
2.54 ijklmno
2.50 ijklmno
2.50 ijklmno
2.41 jklmno
2.38 klmno
2.38 klmno
2.34 klmno


2n Experiment
4.00 a
3.92 ab
3.87 ab
3.54 abc
3.30 cde
3.25 cdef
3.50 bcd
3.25 cdef
3.29 cde
3.54 abc
2.71 gn
3.12 cg
2.92 efghij
2.96 efghi
3.00 defgh
2.92 efghij
2.96 efghi
2.87 efghijk
2.75 ghijklm
2.37 Imnoppqrs
2.79 fghijkl
2.79 efghijk
2.75 ghijklm
2.67 ghijklmno
2.63 hijklmno
2.42 klmnopqr
2.54 hijklmnop
2.63 hijklmno
2.79 fghijkl
2.71 ghijklmn
2.63 hijklmno
2.34 Imnopqrs
2.38 Imnopqrs
2.58 hijklmno
2.46 jklmnopqr
2.50 ijklmnopq
2.46 jklmnopqr
2.38 klmnoprs
2.29 mnopqrs
2.17 opqrs
2.33 Imnopqrs
2.17 opqrs








Table 3-2--continued.

Isolate Ist Experiment 2nd Experiment

BC31 2.30 Imno 2.25 nopqrs
TX18 2.30 Imno 2.17 opqrs
SGS49 2.30 Imno 2.29 mnopqrs
2619 2.25 mno 2.17 opqrs
175104 2.25 mno 2.25 nopqrs
BA57 2.25 mno 2.21 opqrs
RR22 2.25 mno 2.08 pqrs
RR29 2.21 mno 2.17 opqrs
TX20 2.21 mno 1.92 s
BA55 2.21 mno 2.17 opqrs
RR31 2.17 mno 2.08 pqrs
62-2 2.13 no 1.96 rs
SGS25 2.08 o 2.04 qrs
RR27 2.08 o 2.04 qrs


z Values are the means of three replicates of each isolate. Means followed by the same
letter in each column do not differ according to Tukey's Honestly Significant Difference
procedure (P=0.05).








Table 3-3. Geographic origin, color of pigments produced in axenic culture, production of plant pathogenic toxins, and virulence of
isolates of Cercospora piaropi/C. rodmanii.

Isolate Origin Pigment colorw Cercosporinx Diseasey


BA57
BA59
2867
2619
WH9R
62-2
2703
2702
WHV
175-101
RR31
SGS49
2943
RR29
67-1
62-4
RR27
BA55
175-108
175-102
RR24
175-104
RR22
WH83
BC31


Florida, USA
Florida, USA
Mexico
Florida, USA
Florida, USA
Northeast Brazil
Southeast Brazil
Southeast Brazil
Venezuela
Florida, USA
Florida, USA
Florida, USA
Mexico
Florida, USA
Northeast Brazil
Northeast Brazil
Florida, USA
Florida, USA
Florida, USA
Florida, USA
Florida, USA
Florida, USA
Florida, USA
Florida, USA
Florida, USA


Reddish purple
Reddish purple
Reddish purple
Reddish purple
Reddish purple
Reddish purple
Reddish purple
Reddish purple
Reddish purple
Reddish purple
Reddish purple
Reddish purple
Reddish purple
Reddish purple
Reddish purple
Reddish purple
Reddish purple
Reddish purple
Reddish purple
Reddish purple
Reddish purple
Reddish purple
Reddish purple
Reddish purple
Reddish purple


az
ab
abc
abcd
abcde
abcde
abcde
abcde
abcdef
abcdef
abcdef
abcdef
abcdef
abcdefg
abcdefgh
bcdefghi
cdefghi
cdefghij
defghijk
defghijk
defghijk
efghijkl
fghijklm
ghijklm
ghijklmn









Table 3-3--continued
Isolate


Origin


10
SGS25
61-5
WHK
RR4
WH9BR
28-1
49-2
J2
2704
69-1
62-1
400
70-12
TX18
SGS35
TX20
J6
LJ37
65-2
279
49-1
TX2
TX15
SGS32
114
34


Southeast Brazil
Florida, USA
Northeast Brazil
Florida, USA
Florida, USA
Florida, USA
Southwest Brazil
Northeast Brazil
Florida, USA
Southeast Brazil
Northeast Brazil
Northeast Brazil
Zambia
Northeast Brazil
Texas, USA
Florida, USA
Texas, USA
Florida, USA
Florida, USA
Northeast Brazil
South Africa
Northeast Brazil
Texas, USA
Texas, USA
Florida, USA
South Africa
South Brazil


Cercosporin


Pigment color
Reddish purple
Yellow
Reddish purple
yellow
Reddish purple
Reddish purple
Reddish purple
Reddish purple
Yellow
Reddish purple
Yellow
Reddish purple
Reddish purple
Reddish purple
Reddish purple
Reddish purple
Reddish purple
Yellow
Reddish purple
Reddish purple
Reddish purple
Reddish purple
Reddish purple
Np
Np
Reddish purple
Np


Disease
hijklmn
ijklmno
jklmnop
klmnopq
Imnopq
mnopqr
mnopqr
mnopqr
mnopqrs
nopqrst
nopqrst
opqrstuv
pqrstuv
pqrstuv
qrstuv
qrstuv
rstuvw
stuvw
tuvw
uvw
vw
vw
vw
w
w
w
w








Table 3-3--continued
Isolate Origin Pigment color Cercosporin Disease
16-1 Southwest Brazil Np 0.00 w
18-2 Southwest Brazil Np 0.00 w
175-99 Florida, USA Np 0.00 w
Control -- 0.00 w

CV =29.15%
NT = not tested.
Np = non-pigmented.
w Observed in 10-day-old cultures grown on PDA plates at 251 C and 12 h light.
x Production of Cercospora toxins in cultures, detected in thin-layer chromatograms, where plus sign (+) means presence of
cercosporin and minus (-) sign means cercosporin was not detected. The toxin beticolin-l was not detected in any sample. The
procedures for culture, extraction, and detection by TLC are explained under Material and Methods. This experiment was done
twice.
Y Based on disease reaction on waterhyacinth leaves 14 days after inoculation, where: 0, no symptoms; 1, less than 1% of the lamina
surface with spots; 2, greater than 1 and less than 10% of the lamina surface with spots; 3, greater than 10 and less than 25% of the
lamina surface with spots; 4, greater than 25 and less than 50% of the lamina surface with spots; 5, greater than 50 and less than 75%
of the lamina surface with spots; 6, greater than 75% of the lamina surface with spots; and 7, dead lamina. Data were combined from
three experiments conducted in a quarantine greenhouse.
z Means followed by the same letter do not differ according to Tukey's Honestly Significant Difference procedure (P=0.05).












7 B* Experiment 1
SExperiment 2
6 -I s A Experiment 3
-e n --- Average
5 -A ** m *0


A AAAA A A A A





A #A A
1 AA A UM A

-- -





Isolates

Figure 3-2. Isolates of C. piaropi/C.rodmanii plotted in a descending order by their virulence in waterhyacinth. The experiments 1, 2,
and 3 were performed in the periods of 11 to 25/June/1997, 17 to 31/July/1997, and 27/April to 1 l/May/1998, respectively.








The toxin cercosporin was present in the crude extract of most isolates analyzed

(Table 3-3); however toxin beticolin-1 was not detected in the samples analyzed. In one

isolate (114) cercosporin was detected but this isolate was nonpathogenic. This isolate, in

culture, showed a reddish purple pigment of very low intensity. Cercosporin was not

detected by the TLC method in nonpigmented isolates. However, it was detected in low

levels by spectrophotometric analysis (Table 3-4).

The isolates showed significant differences (P=0.05) in cercosporin production,

according to the quantification through absorption at 473 nm with the aid of a

spectrophotometer (Table 3-4). Even though the two experiments were run at the same

conditions, some variation was observed among them. The differences in cercosporin

production among the isolates ranged up to 17 fold in the first experiment and 12 fold in

the second.

Figure 3-3 shows the ultraviolet spectra of crude extracts for typical yellow and

reddish purple isolates, compared with the spectra of the standards for cercosporin and

beticolin-1 in ethyl acetate. The absorption spectrum of the crude extract of the yellow

pigment-producer had a maximum peak at 473 nm (Figure 3-3 A). However, it also

showed two extra peaks at 447 and 504 nm compared to the spectrum of the reddish

purple isolate (Figure 3-3 B), which was very close to the spectrum of pure cercosporin

(Figure 3-3 C). Beticolin-1 was expected to be detected through the UV spectrum,

having the maximum absorption at 435 nm (Figure 3-3 D); however, its presence was not

clear in the samples analyzed with a spectrophotometer. The isolates that were the

strongest cercosporin producers were those with more intense reddish purple

pigmentation in culture.








Table 3-4. Production of cercosporin by isolates of Cercospora piaropilC. rodmanii from
waterhyacinth, in .tg per g wet mycelium.

Isolate 1" Experiment 2nd Experiment
BA57 0.919 az 0.813 ab
175101 0.895 ab 0.747 abc
62-2 0.887 ab 0.945 a
2619 0.862 bc 0.626 bcde
62-4 0.750 bcd 0.759 ba
2702 0.737 bcde 0.664 bcd
WH9R 0.727 bcde 0.658 bcd
RR29 0.682 bcdef 0.659 bcd
SGS25 0.671 bcdefg 0.617 bcde
2703 0.668 bcdefgh 0.542 defg
61-5 0.620 cdeefghi 0.520 defgh
RR31 0.592 defghij 0.557 cdef
RR27 0.592 defghij 0.527 defgh
175102 0.566 defghij 0.296 ijklmo
BA55 0.499 efghijkl 0.333 hijklm
175108 0.456 fghijklm 0.284 ijklmno
SGS49 0.449 fghijklm 0.389 fghijk
2704 0.444 fghijklm 0.345 ghijklmn
TX20 0.431 fghijklmn 0.197 klmnop
175104 0.422 ghijklmn 0.413 fghijk
BA59 0.420 hijklmno 0.363 fghijkl
WHV 0.419 hijklmno 0.510 defg
MX3 0.394 ijklmnop 0.449 efghi
46-4 0.389 ijklmnop 0.217 jklmnop
RR22 0.385 ijklmnop 0.387 fghijk
10 0.359 jklmnopq 0.274 ijklmnop
67-1 0.359 jklmnopq 0.537 defg
BC31 0.358 jklmnopq 0.251 ijklmnop
WHK 0.357 jklmnopq 0.249 ijklmnop
2867 0.353 jklmnopq 0.278 ijklmnop
28-1 0.351 jklmnopq 0.246 jklmnop
69-1 0.335 klmnopq 0.283 ijklmno
WH9BR 0.332 klmnopqr 0.291 ijklmno
WH83 0.322 klmnopqr 0.258 ijklmnop
TX18 0.286 Imnopqrs 0.265 ijklmnop
2943 0.283 Imnopqrs 0.300 ijklmno
RR24 0.245 mnopqrs 0.242 jklmnop
SGS35 0.242 mnopqrs 0.205 klmnop
J6 0.239 mnopqrs 0.260 ijklmnop
62-1 0.234 mnopqrs 0.167 Imnop
70-12 0.228 mnopqrs 0.183 Imnop
49-1 0.222 mnopqrs 0.159 mnop








Table 3-4--continued.

Isolate 1st Experiment 2nd Experiment
J2 0.211 mnopqrs 0.249 ijklmnop
RR4 0.206 mnopqrs 0.166 Imnop
LJ37 0.188 nopqrs 0.129 op
65-2 0.186 nopqrs 0.192 klmnop
49-2 0.171 opqrs 0.222 jklmnop
114 0.170 opqrs 0.137 nop
279 0.168 pqrs 0.121 op
400 0.162 pqrs 0.131 op
34 0.153 pqrs 0.125 op
TX2 0.115 qrs 0.179 Imnop
SGS32 0.084 rs 0.118 op
TX15 0.062s 0.109 op
16-1 0.055s 0.11l op
18-2 0.054s 0.105 op
17599 0.054s 0.081 p

z Values are the means of three replicates of each isolate. Means followed by the same
letter in each column do not differ according to Tukey's Honestly Significant Difference
procedure (P=0.05).




























S0.4


P 1.6 C

1.2
0.8
0.4

o0

1.6 D
1.2-
0.8
0.4
0 -- <^ I --- --- --I --- ^ = = -
240 280 320 360 400 440 480 520 560 600

Wavelength (nm)


Figure 3-3. Ultraviolet spectra of crude extracts from the yellow pigment-producer,
isolate WHK (A), and the reddish-purple pigment-producer, isolate BA57 (B), compared
to the standards cercosporin (C) and beticolin-1 (D) in ethyl acetate.








Based on the presence of yellow metabolites in some cultures of isolates of C.

piaropi/C.rodmanii, it was hypothesized that the yellow color could be related to the

toxins beticolins, formerly called the Cercospora beticola toxin (CBT). However, the

analyses based on thin-layer chromatograms, resolved under long-UV light and having a

standard beticolin-l, did not show the presence of beticolins in the samples, but only

cercosporin with an Rf value of 0.61 (e.g., isolate WHK in the Figure 3-4 A). According

to Milat and Blein (1995), the six beticolin toxins would be expected to have R values

below that of cercosporin, as shown by the standard for beticolin-1, which had an R f

value of 0.46. The Figure 3-4 B shows the thin-layer chromatogram of a crude extract

from an isolate that produced a typical reddish-purple pigment having a band

corresponding to cercosporin.

The virulence of the isolates was positively correlated with their ability to produce

cercosporin, which ranged from 72 to 89% (P<0.0001; Table 3-5) and negatively

correlated with mycelial growth rate, which ranged from -55 to -61% (P<0.0001).

Cercosporin production and mycelial growth were negatively correlated (-50 to -62%;

P<0.003) as was the correlation between mycelial growth and virulence (-55 to -0.66%;

P<0.001).

Fatty Acid Analysis
The isolates of C. piaropi/C.rodmanii analyzed contained the fatty acids palmitic

acid (16:0), oleic acid (18:1w9c), stearic acid (18:0), and the unresolved mixtures, named

sum feature 4 (16: lo7cl5iso20H) and sum feature 6 (18:2co6,9c/18:0 anteiso). In

addition, two of the outgroup species also contained the fatty acid myristic acid (C14:0).

The mean fatty acid composition with standard deviation for each isolate from three



















A B
Ref.




0.61

0.46







1 2 3 4 1 2 3



Figure 3-4. Crude extracts of isolates of Cercospora piaropilC. rodmanii resolved on thin
layer chromatograms under long-ultraviolet light. A, lanes 1 to 4, standard cercosporin,
standard beticolin-1, nonpigmented isolate 17599, and the typical yellow pigment-
producer isolate WHK; B, lanes 1 to 3, standard cercosporin, standard beticolin-1, and
the typical purple pigment-producer isolate BA57.








Table 3-5. Correlation matrix among some physiological traits of 55 isolates of Cercospora piaropi/C. rodmanii from several
geographical locations.

Virulence Virulence Virulence Cercosporin Cercosporin Mycelial Mycelial
production production growth growth
1st 2nd 3rd t 2nd 1st 2nd
experiment experiment experiment experiment experiment experiment experiment

Virulence
It experiment

Virulence 0.93
2nd experiment (0.0001)

Virulence 0.92 0.91
3" experiment (0.0001) (0.0001)

Cercosporin production 0.78 0.72 0.89
1st experiment (0.0001) (0.0001) (0.0001)

Cercosporin production 0.80 0.73 0.85 0.87
2"d experiment (0.0001) (0.0001) (0.0001) (0.0001)

Mycelial growth -0.60 -0.61 -0.66 -0.62 -0.54
1i" experiment (0.0001) (0.0001) (0.0001) (0.0001) (0.0001)

Mycelial growth -0.55 -0.55 -0.61 -0.57 -0.50 0.81
2 experiment (0.0001) (0.0001) (0.0001) (0.0001) (0.0003) (0.0001)








different culture ages are presented in the Tables 3-6, 3-7, and 3-8. The isolates of C.

piaropilC. rodmanii did not differ in the number and kind of fatty acids present but

differed in the relative concentration of each type. The relative concentration of an

individual fatty acid ranged from less than 1% of the total fatty acid content to over 50%.

The results of canonical discriminant analysis of the fatty acid profiles showed an

accentuated effect of the age of the cultures in the resolution of taxa (Table 3-9). FAME

profiles from 4-day-old mycelium differentiated more of the isolates of C. piaropi/C.

rodmanii than the profiles from 5- and 6-day-old mycelia. The differentiation of isolates

of C. piaropi/C. rodmanii using 4-day-old mycelia did not relate with geographical origin

of the isolates. In addition, some of the species of Cercospora used as the outgroup did

not differentiate from the isolates of C. piaropi/C. rodmanii. Basically, the isolates of C.

piaropi/C. rodmanii could not be differentiated based on fatty acid profiles from the 5-

and 6-day-old mycelia; with the exception of the isolate 2619 at the 6-day-old mycelium

comparison.

The FAME profile of the isolate WH9BR, which was collected and identified by

Conway (1976a), who described it as C. rodmanii, was significantly different (P=0.05)

from only one isolate of C. piaropi/C. rodmanii when 6-day-old mycelia were used. It

was not significantly different from any isolates of C. piaropi/C. rodmanii when 5-day-

old mycelia were used. In addition, the FAME profile of isolate WH9BR was not

significantly different from other two outgroup Cercospora species when 5- and 6-day

old mycelia were used.








Table 3-6. Fatty acid composition of Cercospora spp. isolates from 4-day-old mycelia.

Isolate / species % Of total fatty acid content (mean SD)
14:0 Sum feature 4a 16:0 Sum feature 6b 18:1 0)9c 18:0

WH83 0.00 0.00 0.79 0.01 19.36 + 3.11 49.00 + 6.10 24.66 + 1.83 6.06 0.96
WHK 0.00 + 0.00 0.74 + 0.05 24.92 2.03 39.54 2.38 27.41 0.23 7.21 0.88
WH9BR 0.00 + 0.00 0.97 0.13 26.00 5.54 39.58 8.59 26.72 1.98 6.60 1.16
2619 0.00 0.00 1.00 + 0.20 22.77 1.38 45.39 2.43 25.19 + 0.52 5.40 0.41
TX18 0.00 +0.00 0.83 0.08 19.33 + 2.16 47.65 3.08 28.42 0.89 3.78 0.32
TX20 0.00 0.00 1.03 0.12 18.79 1.33 51.92 1.43 24.63 0.99 3.62 0.67
2867 0.00 0.00 1.06 0.22 18.39 2.44 48.07 1.24 27.35 1.40 4.96 + 0.43
MX3 0.00 + 0.00 0.86 0.14 16.29 2.28 53.59 3.43 24.18 1.38 4.60 0.63
62-2 0.00 + 0.00 1.35 + 0.28 23.08 5.90 42.56 + 7.05 26.60 + 0.51 6.02 0.55
10 0.00 0.00 0.94 0.19 20.89 2.86 45.42 4.17 25.64 1.56 6.80 0.84
28-1 0.00 0.00 0.79 0.05 16.46 0.99 53.46 1.48 24.85 + 0.51 4.03 + 0.19
400 0.00 0.00 0.96 0.08 16.97 + 0.46 54.63 + 0.62 23.28 0.34 3.58 0.39
114 0.00 0.00 0.83 + 0.09 15.00 + 0.49 53.51 + 1.29 25.75 1.34 4.38 0.64
WHV 0.00 0.00 1.15 0.11 15.63 0.27 50.61 1.22 29.31 1.05 3.31 0.01
C. beticola 0.24 0.03 0.87 0.11 25.43 + 0.49 35.24 0.36 30.26 0.57 7.71 0.69
C. oenotherae 0.35 0.04 0.97 0.09 25.59 0.26 40.89 + 0.71 25.69 0.51 6.63 0.67
Cercospora sp.c 0.00 0.00 0.56 0.02 25.41 1.73 27.74 2.40 37.61 1.38 8.60 + 0.74
C. gossypina 0.00 0.00 0.61 +0.08 19.24 1.38 51.78 2.62 21.33 1.17 6.91 0.25

a Sum feature 4 is an unresolved mixture of 16:1 ow7c15 iso 20H;
b Sum feature 6 is an unresolved mixture of 18:2 co6, 9c/18:0 anteiso.
c From Jasminum sp.








Table 3-7. Fatty acid composition of Cercospora spp. isolates from 5-day-old mycelia.

Isolate / species % Of total fatty acid content (mean SD)
14:0 Sum feature 4a 16:0 Sum feature 6b 18:1 o9c 18:0

WH83 0.00 + 0.00 0.80 0.03 18.84 1.44 49.03 + 2.36 25.84 2.26 5.18 2.03
WHK 0.00 0.00 0.83 0.10 23.72 0.73 41.06 1.81 27.57 0.61 6.83 0.48
WH9BR 0.00 0.00 1.01 0.12 22.16 3.38 43.93 6.38 26.76 2.30 6.15 0.81
2619 0.00 0.00 1.14 0.10 23.28 2.41 43.71 3.61 26.28 1.27 5.48 0.29
TX18 0.00 0.00 0.79 0.05 19.00 0.68 48.41 1.73 28.17 1.02 3.49 0.29
TX20 0.00 0.00 0.98 0.06 18.58 0.18 52.78 0.38 24.17 0.43 3.49 0.21
2867 0.00 0.00 1.04 0.10 19.11 2.56 48.47 + 3.48 26.75 0.06 4.64 0.87
MX3 0.00 t 0.00 0.90 0.03 18.81 3.39 48.85 7.79 25.58 3.14 5.65 1.65
62-2 0.00 0.00 1.19 0.10 20.67 0.95 44.65 :0.50 27.16 0.57 5.73 0.32
10 0.00 +0.00 1.08 0.38 20.23 1.90 44.71 +t2.97 26.95 +0.49 6.52 0.38
28-1 0.00 t0.00 0.85 +0.04 15.73 + 0.64 54.41 + 1.58 24.84 0.84 3.95 0.12
400 0.00 + 0.00 1.04 0.08 16.16 0.83 52.81 2.36 23.24 1.27 3.54 0.21
114 0.00 0.00 0.88 0.12 16.34 1.41 53.22 1.85 24.98 0.59 4.21 0.52
WHV 0.00 0.00 1.18 0.08 15.29 0.42 50.77 0.69 29.55 1.13 3.22 0.10
C. beticola 0.24 0.02 0.86 + 0.01 26.10 + 0.95 34.10 + 0.79 30.40 0.57 8.13 1.13
C. oenotherae 0.36 +0.09 1.07 +0.07 25.36 7.37 39.11 i 12.63 28.80 4.10 5.13 1.30
Cercospora sp.C 0.00 0.00 0.94 0.45 25.69 2.33 32.06 6.85 34.36 6.22 7.14 i 1.23
C. gossypina 0.00 +0.00 0.58 0.03 20.71 1.03 48.06 2.55 23.13 1.04 7.19 0.20

a Sum feature 4 is an unresolved mixture of 16:1 w7c 15 iso 20H;
b Sum feature 6 is an unresolved mixture of 18:2 06, 9c/18:0 anteiso.
c From Jasminum sp.




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