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Molecular analysis of genetic diversity and variability in Colletotrichum gloeosporioides

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Molecular analysis of genetic diversity and variability in Colletotrichum gloeosporioides
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Liyanage, Hemachandra D., 1958-
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viii, 148 leaves : ill.,photos ; 29 cm.

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Colletotrichum ( jstor )
DNA ( jstor )
Electrophoresis ( jstor )
Enzymes ( jstor )
Fungi ( jstor )
Gels ( jstor )
Genes ( jstor )
Mycology ( jstor )
Ribosomal DNA ( jstor )
Yeasts ( jstor )
Citrus -- Diseases and pests ( lcsh )
Colletotrichum gloeosporioides -- Analysis ( lcsh )
Fungal diseases of plants ( lcsh )
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Thesis:
Thesis (Ph. D.)--University of Florida, 1992.
Bibliography:
Includes bibliographical references (leaves 131-147).
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Typescript.
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Vita.
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by Hemachandra D. Liyanage.

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MOLECULAR ANALYSIS OF GENETIC DIVERSITY AND VARIABILITY IN
COLLETOTRICHUM GLOEOSPORIOIDES














BY

HEMACHANDRA D. LIYANAGE


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

1992


UF FL:Nf L!ES


































Dedicated
to
Mother















ACKNOWLEDGEMENTS


I sincerely thank Dr. Corby Kistler for offering me

this opportunity to work in his lab, for all the advice,

guidance, constructive criticism and help throughout the

research study and my career. I am very thankful to Dr. R.

T. McMillan, Jr., for providing financial arrangements and

for being helpful in many ways. I gratefully acknowledge

the willing assistance and advice given by Dr. Frank Martin

and Dr. Ron Sonoda. I thank my committee members, Dr. Daryl

Pring, Dr. James Kimbrough, and Dr. Curtis Hannah for their

assistance and guidance throughout this study.

A special word of thanks to my loving wife, Thamara,

who spent her time and effort helping me throughout the

period of study and in all my difficult times.


iii
















TABLE OF CONTENTS

Page

ACKNOWLEDGEMENTS................................... iii

ABSTRACT................. ............................ vii

CHAPTERS

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

2 OBSERVED VARIABILITY IN COLLETOTRICHUM
GLOEOSPORIOIDES CAUSING POST BLOOM FRUIT
DROP IN CITRUS ................................. 7

Introduction.................................... 7
Materials and Methods......................... 9
Strains of Colletotrichum gloeosporioides. 9
Pathogenicity............................. 10
Benomyl Tolerance......................... 10
Results.......................................... 11
Colletotrichum gloeosporioides Strains
from Citrus are Morphologically
Variable................................. 11
Colletotrichum gloeosporioides Strains
have Different Nuclear Numbers
in their Spores......................... 13
Both Type 1 and Type 2 Strains are
Pathogenic to Tahiti Lime Flowers....... 15
Type 1 and Type 2 Strains Differ in their
Tolerance to Benomyl.................... 15
Discussion....................................... 16

3 DNA POLYMORPHISMS FOUND AT MANY GENETIC LOCI
EXAMINED IN COLLETOTRICHUM GLOEOSPORIOIDES... 25

Introduction................... ....... ...... 25
Ribosomal DNA in Fungi.................... 25
Ribosomal DNA is Polymorphic in Many
Fungi................................... .27
Fungal Cutinase Genes and Cutinase
Isozymes................................ 31
Restriction Fragment Length Polymorphisms
(RFLP) in Fungi. .......... .. ........... 36










Page


Materials and Methods.......................... 39
Strains of Colletotrichum gloeosporioides. 39
DNA Extraction ........................... 40
DNA Cloning and Restriction Enzyme Mapping 41
Enzyme Assays and Electrophoresis of
Cutinase .............................. 42
Probes Containing Cutinase Gene Sequences. 43
Detection of Restriction Fragment Length
Polymorphisms............ .... .............. 44
Results.......................................... 45
Ribosomal DNA is Polymorphic in
Colletotrichum gloeosporioides........... 45
Ribosomal RNA Genes....................... 47
Diverse Cutinases and Cutinase Genes are
Found in Type 1 and Type 2 Strains of
Colletotrichum gloeosporioides.......... 49
Subgroups of Colletotrichum gloeosporioides
have Distinct RFLP Patterns Detected by
Many Genetic Markers................... 53
Discussion...................................... 73

4 VARIABILITY OF MOLECULAR KARYOTYPES AND
CHROMOSOMOL DNAS IN COLLETOTRICHUM
GLOEOSPORIOIDES.............................. 81

Introduction ...................... ..... .... 81
Pulsed Field Gel Electrophoresis.......... 81
Molecular Karyotypes of Fungi.............. 82
Materials and Methods......................... 85
Strains of Colletotrichum gloeosporioides. 85
Preparation of Protoplast Plugs............ 85
Electrophoresis and Southern Analysis..... 86
Results........................ ................ 87
Discussion............................... ...... 97

5 GENERAL DISCUSSION AND CONCLUSIONS............. 99

APPENDICES

A STRAINS OF COLLETOTRICHUM GLOEOSPORIOIDES..... 102

B ANALYSIS OF VARIANCE TABLES.................... 104

C PROCEDURES FOR DNA LABELLING AND SOUTHERN
HYBRIDIZATION...... ......................... 110

D CALCULATED MB SIZES FOR CHROMOSOMAL DNAS IN
COLLETOTRICHUM GLOEOSPORIOIDES............... 112











pace

LITERATURE CITED.................................... 131

BIOGRAPHICAL SKETCH................................. 148















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

MOLECULAR ANALYSIS OF GENETIC DIVERSITY AND VARIABILITY IN
COLLETOTRICHUM GLOEOSPORIOIDES

By

Hemachandra D. Liyanage

August 1992

Chairman: Dr. R. T. McMillan, Jr.
Cochairman: Dr. Corby H. Kistler
Major Department: Plant Pathology

Results from this study suggest two distinct genetic

subpopulations of Colletotrichum gloeosporioides from citrus

based on DNA variation, cultural morphology, and growth.

Type 1 strains are slow growing, morphologically stable,

benomyl tolerant, and contain a single form of ribosomal DNA

(rDNA) as detected by common HindIII, PstI, SphI and SstI

fragments hybridizing to cloned Neurospora crassa rDNA. Type

2 strains are faster growing, morphologically less stable,

benomyl sensitive, and have rDNA distinct from type 1

strains. The rDNA from type 1 and type 2 strains were cloned

and mapped for 10 restriction enzyme sites and genes coding

for large subunit, small subunit and 5.8S rRNA. A subclone

constructed from the non-transcribed spacer region of type 1

rDNA clone hybridizes only to rDNA from type 1 strains. DNA

polymorphisms detected by heterologous hybridization with

vii









cloned N. crassa genes for glutamate dehydrogenase,

anthranilate synthetase, histidinol dehydrogenase, and S-

tubulin corresponded to type 1 or type 2 strains. All

strains liberate free fatty acid from [3H]-labelled cutin

and hydrolyze cutin model substrates. Serine esterases from

extracellular fluids of cutin-grown C. gloeosporioides

strains were detected by labelling proteins separated by

sodium dodesyl sulfate polyacrylamide gel electrophoresis

with 3H-diisofluorophosphate. The two major esterases from

type 1 strains have molecular weights of 26 and 20

kilodaltons (kd) whereas the type 2 esterases were 24 and 22

kd. A DNA probe containing the cloned cutinase gene from C.

gloeosporioides hybridized strongly to DNA from type 2

strains but poorly to type 1 strains. Distinct cutinase

genes may be present in the two types of C. gloeosporioides

strains from citrus. Chromosome-sized DNAs separated by

pulsed-field gel electrophoresis corresponded to type 1 or

type 2 strains. Type 1 strains had five large chromosomal

DNAs 7.6, 7.0, 4.7, 3.7, and 3.3 (or 2.8) million base pairs

(Mb) in size and one or two smaller chromosomes (1.6 to 0.63

Mb). Type 2 strains had three large chromosomal DNAs (7.8,

4.7, and 3.7 Mb) and two to four smaller chromosomal DNAs

(0.52-0.28 Mb).


viii















CHAPTER 1
INTRODUCTION


Historically, the study of plant diseases dates back to

Theophrastus (371-287 B.C.) who first described disease

conditions of plants, mostly cereal rusts, in Historia

plantarum and De causis plantarum (Ainsworth 1981).

Afterwards, studies of causal agents of plant diseases and

control measures played a major role in human survival. The

period from the mid-eighteenth to the mid-nineteenth century

was marked by the accumulation of experimental evidence for

the pathogenicity of fungi to plants. Almost all groups of

fungi include some plant pathogenic species but the greatest

number of plant pathogens is to be found among the imperfect

fungi (Ainsworth 1971).

The most important technique for the identification of

plant pathogenic fungi has always been macroscopic and

microscopic morphological examination. Morphology always

took precedence over other considerations in describing

genera and species of plant pathogenic fungi (Ainsworth

1981). The genus Colletotrichum Corda was established in

1831 and was characterized by having setose acervuli

containing hyaline, curved fusiform conidia (Baxter et al.

1985). However, there was always confusion in describing









2

fungi to this genus due to similar morphological characters

of Vermicularia Tode and Gloeosporium Desm. & Mont. (Dickson

1925; Duke 1928; Arx 1957; Baxter et al. 1985). Duke (1928)

suggested that type species of Vermicularia and

Colletotrichum represented the same fungus. Species in the

genus Gloeosporium probably represent the same fungi as in

the genus Colletotrichum because the Gloeosporium species,

which supposedly lack setae, were found to produce them on

certain substrates (Baker et al. 1940). Arx (1957) accepted

Colletotrichum and Vermicularia as separate genera while

rejecting the more heterogenous genus Gloeosporium.

The species concept of Colletotrichum gloeosporioides

(Penz.) is still uncertain (Van Der Aa et al. 1990).

Colletotrichum gloeosporioides was first described in 1882

by Penzig as Vermicularia gloeosporioides, and in 1887 it

was renamed Colletotrichum gloeosporioides (Burger 1921).

The presence of this fungus in the United States was first

observed in 1886 in Florida and was reported by Underwood

(1891). Arx (1957) recognized eleven species in the genus

Colletotrichum, and the name C. gloeosporioides with nearly

600 synonyms was maintained to designate the variable

anamorph of Glomerella cingulata (Stonem.) Spauld. & Schr.

He recognized nine forms within the species C.

gloeosporioides. Arx (1970, 1987) introduced the concept of

host forms of C. gloeosporioides but did not accept these

forms as species or intraspecific taxa with certainty.










3

Sutton (1980) considered C. gloeosporioides a group species

showing excessively wide variation.

The conidia of C. gloeosporioides are straight, obtuse

at the apex, 9-24 x 3-4.5 pm and appressoria are 6-20 x 4-12

jm, clavate or irregular, sometimes becoming complex (Sutton

1980).

Colletotrichum gloeosporioides is a ubiquitous fungus

and often causes a variety of diseases commonly known as

anthracnose on fruits, leaves and stems of a wide range of

host species. The host range of this fungus is so wide that

nearly 200 susceptible host species were listed under C.

gloeosporioides in the Index of Plant Diseases in Florida

(Alfieri et al. 1984). Many tropical fruit crops are

attacked by this fungus in the field and in post-harvest

condition (Nolla 1926; Simmonds 1965; Brown 1975). Citrus is

one of the major fruit crops attacked by this fungus, and

the diseases of citrus caused by C. gloeosporioides have

been known since 1886 when it was first isolated from citrus

plants in Florida, U.S.A. (Underwood 1891). Rolfs (1904 and

1905) described a group of citrus diseases (wither tip, leaf

spot, lemon spot, canker, and anthracnose) caused by C.

gloeosporioides. A more recently described citrus disease,

post bloom fruit drop, PFD (Fagan 1979; Sonoda and Pelosi

1988; McMillan and Timmer 1989) is caused by the same

species. The name PFD was suggested by Fagan (1979) to

distinguish this disease condition of citrus characterized









4

by premature fruit drop or blossom blight from normal

physiological thinning of fruits. The symptoms first appear

as small, brown spots on flower buds or light pink water-

soaked spots on open petals. These spots may enlarge and

rapidly cover the petals within 24 h. Afterwards, the petals

become brown and desiccated. Eventually, young fruitlets

become discolored, and they abscise, leaving the calyxes

behind as persistent buttons (Figure 1). The disease is

economically important in regions where citrus is grown

(Denham and Waller 1981; Fagan 1984a). In Florida the

disease has been reported from all commercially grown citrus

(Sonoda and Pelosi 1988; McMillan and Timmer 1989).

The causal agent of PFD of citrus was identified as C.

gloeosporioides (Fagan 1979; Sonoda and Pelosi 1988;

McMillan and Timmer 1989). Colletotrichum gloeosporioides

isolated from citrus diseases were reported to be variable

in morphology and pathogenicity (Burger 1921). Morphological

variability also has been observed in the strains of this

fungus causing PFD (Denham and Waller 1981; Sonoda and

Pelosi 1988). Three strains C. gloeosporioides varying in

morphology and pathogenicity were reported to be associated

with diseased plants by Fagan (1980). Because of the

inconsistency of the morphological characteristics, it is

uncertain whether the strains or forms of C. gloeosporioides

recognized by morphological criteria alone are really

different at genetic and molecular level. The morphological









5

changes could be caused by environmental effects, genetic

differences or both. Therefore, study of this group species

at the molecular level to understand the genetic and

molecular differences among the strains is important.

The present study was undertaken to investigate the

morphological variability and potential genetic variation of

C. gloeosporioides at the molecular level. The objectives of

this study are:

1. To examine the basis for morphological variability

of C. gloeosporioides causing PFD disease of citrus.

2. To investigate genetic variation of C.

gloeosporioides at the molecular level.

3. To examine variations in chromosome-size DNA and to

describe the molecular karyotypes.

Morphological and growth diversity arising from single

spore cultures of different C. gloeosporioides strains is

examined in Chapter 2. Molecular investigations using

genetic markers were carried out to study the variation

within citrus strains of C. gloeosporioides, and they are

reported in Chapter 3. Chapter 4 describes the chromosomal

variation of C. gloeosporioides strains and the molecular

karyotypes of the fungus. The results obtained in these

studies are reviewed comprehensively in Chapter 5, and a

concept of genetically distinct subpopulations of C.

gloeosporioides is proposed.











































Figure 1 Symptoms of post bloom fruit drop disease on sweet
orange (Citrus sinensis var. Valencia) caused by
Colletotrichum gloeosporioides.















CHAPTER 2
OBSERVED VARIABILITY IN COLLETOTRICHUM GLOEOSPORIOIDES
CAUSING POST BLOOM FRUIT DROP IN CITRUS


Introduction


More than six hundred synonyms for the fungal species

Colletotrichum gloeosporioides have been published (Arx

1957). This reflects the considerable amount of diversity

and variability observed for the fungus by different

investigators throughout the world. Sutton (1980) could not

give a standard morphological description of C.

gloeosporioides. He considered the different forms of this

fungus to be within a group species.

The association of C. gloeosporioides with citrus dates

back to 1886-1891 (Underwood 1891). The fungus has been

found to cause various diseases on this crop for the past

century (see Chapter 1). The most thorough studies on the

morphological variation of C. gloeosporioides were done by

Burger (1921). The fungus he studied was the causal agent of

bloom drops and leaf spots in citrus. Cultural

characteristics such as mycelial color, growth and

sporulation enabled him to classify C. gloeosporioides

strains into five groups. However, some strains did not fit

into any of the morphological classes due to inconsistency









8

of mycelial and sporulation characteristics in continuous

culture.

Mycelial sectors distinguished by growth and color

differences within single spore cultures of strains are

another type of variability observed in C. gloeosporioides.

Burger (1921) observed black and white mycelial sectors in

single spore cultures of the fungus. When single spored,

these black and white sectors were able to maintain their

identity in continuous culture.

Burger (1921), after studying cultural characteristics,

spore dimensions, and sectoring, concluded that C.

gloeosporioides is constantly giving off new types under

natural conditions as well as in artificial cultures. He

further suggested that these variabilities of C.

gloeosporioides may have arisen from environmental effects

as well as from high frequency mutations.

Morphological and pathogenic variability in C.

gloeosporioides causing PFD of citrus has been reported by

Fagan (1979,1980) and Denham and Waller (1981). Three

different forms of C. gloeosporioides were recognized by

Fagan (1980). Two forms, cgm with gray to dark gray mycelium

and cgc with light gray mycelium, were isolated from

senescent leaves and were nonpathogenic to citrus flowers.

The pathogenic form, cgp, had off-white to pink mycelium and

was isolated from floral parts of citrus. Fagan (1980)

concluded that at least two strains of C. gloeosporioides









9

causing PFD occurred in Belize. These strains corresponded

to morphological groups of C. gloeosporioides described by

Burger (1921).

The objective of this study is to examine the

morphological and phenotypic diversity of C. gloeosporioides

causing PFD of Tahiti lime (Citrus aurantifolia Swingle) and

Sweet orange (Citrus sinensis Osbek).

Materials and Methods


Strains of Colletotrichum gloeosporioides


Strain number, host, place and year of isolation are

tabulated in Appendix A. Isolation of C. gloeosporioides

from host plants was carried out as follows. Host plant

tissues were surface sterilized in 1% sodium hypochlorite

(Clorox Co., Oakland, CA) for 30-60 s, rinsed 3 times with

sterilized water and plated on potato dextrose agar (PDA,

Difco laboratories, Detroit, MI) plates. Edges from growing

mycelia were isolated and maintained in the laboratory as

strains. Strains were grown in 20% (w/v) V-8 juice (Campbell

Soup Co., Camden, NJ) for 7 d at 250 rpm on a Lab-Line orbit

shaker (Lab-Line Instruments Inc., Melrose Park, IL). Spores

were collected by centrifugation at 7000 x g for 5 min and

washed 2 times with sterilized water before storing in 50%

glycerol (in water) at -800C. To obtain single spore

cultures, spores were spread on PDA plates; 14-16 h later,

germinating spores were isolated under a dissecting









10

microscope (25x10 magnification) and plated on PDA plates.

Morphology of colony growth, mycelial color and sectoring

were examined in PDA culture and still liquid culture,

potato dextrose broth (PDB Difco laboratories, Detroit, MI).

To examine the nuclear number, spores were stained with

1% aniline blue (Sigma Chemical Co., St. Louis, MO) in 50%

glycerin in water (Tu and Kimbrough 1973). To stain nuclei,

a drop of spores in water was placed on a microscopic slide,

and a drop of stain was added. The slide was then heated

over a flame for 5-10 seconds. Approximately 1000 spores

were examined for each strain.


Pathogenicity


All the strains were tested for their ability to infect

flowers of Tahiti lime under natural conditions in the field

as well as in the laboratory. Strains were grown in 20% V-8

juice for 7 days and inoculum containing 107 spores ml'1

water were prepared. Tahiti Lime flowers were sprayed using

a hand sprayer to wetness with inoculum or water, and

symptom development was observed for 3 days. Each treatment

contained 10-15 flowers. The control was sprayed with water.


Benomvl Tolerance


The growth of C. gloeosporioides strains was examined

in PDA medium containing 0, 2 and 10 pg benomyl (methyl-

(butyl carbamoyl)-2-benzimidazolecarbamate, Sigma Chemical









11

Co., St. Louis, MO) ml-1. Radial growth of the mycelial

colony was measured every 24 h for a 10 day period. Growth

rates in mm h-' were estimated by the slopes obtained with

linear regression analysis of the growth curve. A comparison

of slopes was made using analysis of variance (Appendix B).

Each treatment was replicated 5 times, and the experiment

was repeated once with 2 replicates.


Results


Colletotrichum gloeosporioides Strains from Citrus are
Morpholocicallv Variable


The C. gloeosporioides strains examined can be grouped

into two major categories based on morphology and growth

characteristics. Type 1 strains (H-l, H-3, H-9, H-21, H-22,

H-25B, H-36, IMB-3, LP-1, Maran, OCO, and TUR-1) produce

morphologically stable and relatively slow-growing mycelial

colonies in PDB. The colonies are orange-colored and have

appressed mycelia with abundant sporodochia (Figure 2.1).

Type 2 strains (H-4, H-ll, H-12, H-23, H-24, H-46, H-47, H-

48, 180269 and 226802) grow faster and produce mostly gray,

fluffy mycelial colonies (Figure 2.1). The type 1 strains

grow at a significantly slower rate from 0.008 to 0.10 mm

h'1; type 2 strains grow significantly faster from 0.12 to

0.15 mm/ h"1 as calculated by slopes of linear regression

data (Table 2.1). The strain types also differ in culture












Table 2.1 Effect of benomyl concentration on the estimated
radial growth rates in mm h'1 of Colletotrichum
gloeosporioides type 1 and type 2 strains.


Strain Benomyl concentration jg/ml
0 2 10


Type 1

H-1

H-3

H-9

H-25B

H-3 6

IMB-3

LP-1

Maran

OCO

Type 2

H-4

H-1l

H-12

H-46

H-47

H-48

180269

226802


0.10

0.10

0.10

0.041

0.095

0.008

0.10

0.041

0.095


0.141

0.125

0.121

0.133

0.133

0.145

0.133

0.150


0.033

0.033

0.045

0.041

0.037

0.008

0.050

0.029

0.037



0.00

0.00

0.00

0.00

0.00

0.00

0.00

0.00


0.029

0.033

0.041

0.041

0.033

0.004

0.041

0.033

0.033



0.00

0.00

0.00

0.00

0.00

0.00

0.00

0.00









13

stability as determined by their ability to produce sectors

of different color, morphology and growth habit. To

quantitate these levels of instability, 100 conidia were

isolated from three type 1 strains and two type 2 strains

and tested for morphological stability. One hundred single

spore cultures from strains H-l, H-3, and H-25B (type 1)

grown in PDB were found to produce identical colonies. One

hundred single spore cultures from strain H-12 and H-48

(type 2) produced 100% sectoring colonies. These colonies

varied in colony color from dark gray to gray, white, and

orange with different growth rates (Figures 2.2, 2.3, and

2.4). Sporodochia production was scattered or inhibited but

could be stimulated by mycelial injury (Figure 2.5). One

hundred injury-induced spores from an H-48 gray mycelial

sector produced 50 sectoring colonies, 24 dark gray with no

sporodochia and 26 orange colonies with scattered

sporodochia production.


Colletotrichum gloeosporioides Strains have Different
Nuclear Numbers in their Spores


The nuclear number observed by aniline blue staining

varied from 1 to 3 per single spore (Table 2.2). All spores

examined from all isolates were single called. All the

isolates examined contained spores with more than one

nucleus as identified by dark stained objects distinguished

from the lightly stained cytoplasm under the high












Table 2.2 Percentage of spores carrying different numbers of
nuclei in Colletotrichum gloeosporioides strains.


Percentage of spores*
Strain Number of nuclei
1 2 3


H-l

H-3

H-4

H-9

H-12

H-25B

H-4 6

H-4 8

180269

226802

LP-1

Maran


92.6

97.8

94.3

96.4

98.8

96.9

93.4

92.8

92.9

94.7

99.6

98.6


6.8

2.2

5.4

3.4

1.2

2.9

6.0

6.6

6.6

5.2

0.4

1.4


0.6

<0.1

0.3

0.2

<0.1

0.2

0.6

0.6

0.5

0.1

<0.1

<0.1


*=Calculated from 1000 spores










15

magnification (46x10) of a light microscope. Spores

containing a single nucleus varied from 92.6 to 99.6% in the

15 isolates studied. The maximum number of nuclei observed

within a single spore was 3, and the percentage of spores

containing three nuclei varied from <0.1-0.6%. The

percentage of spores containing two nuclei varied from 0.4-

6.8%.


Both Type 1 and Type 2 Strains are Pathoqenic to Tahiti Lime
Flowers


Brown lesions developed in flowers individually

inoculated with all strains of the pathogen 24 h after

spraying. The petals were blighted completely at 36 h and

had dropped at 48 h. Flowers sprayed with water alone were

not blighted after 72 h. The fungal strains reisolated from

infected tissues were found to be morphologically like the

original strains. The relative virulence of strains was not

measured in this study.


Type 1 and Type 2 Strains Differ in their Tolerance to
Benomvl


All type 2 strains were completely inhibited by 2 or 10

Ag ml" benomyl in PDA, but type 1 strains were more

tolerant. Average growth rates for individual type 1 and

type 2 strains are listed in Table 2.1. Analysis of variance

showed that benomyl concentration had a significant effect

on type 1 strains. There was a significant interaction









16

between strains and concentration indicating that growth

rate of each strain may respond differently to different

concentrations of benomyl (Appendix B).



Discussion


Grouping of Colletotrichum gloeosporioides strains

based on morphological and physiological observations was

first attempted by Burger (1921). However, morphologically

based groups of C. gloeosporioides strains have been

inconsistently described in this and subsequent studies

(Burger 1921; Arx 1957; Sutton 1980). Fagan (1980), Denham

and Waller (1981), and Sonoda and Pelosi (1988) reported

morphological variations associated with this fungus

isolated from Sweet orange cultivars. The type 1 strains in

this study show similarities in morphology, growth and

sporodochia production to strains designated cgp by Fagan's

(1980) description and correspond to the orange colored,

slow-growing colonies described by Sonoda and Pelosi (1988).

The more variable type 2 strains show similarities to the

cgm and cgc strains of Fagan and correspond to faster

growing colonies described by Sonoda and Pelosi.

Both type 1 and type 2 strains were isolated from sweet

orange (C. sinensis) as well as Tahiti lime (Appendix A).

Both types were pathogenic to Tahiti lime flowers as










































Figure 2.1. Morphology of type 1 (left) and type 2 (right)
strains of Colletotrichum gloeosporioides grown
in potato dextrose broth.












































Figure 2.2 Fluffy and appressed mycelial sectors
produced by a single spore culture of
Colletotrichum gloeosporioides type 2 strain in
potato dextrose agar.












































Figure 2.3. Dark gray, light gray and orange mycelial
sectors produced by a single spore culture of a
Colletotrichum gloeosporioides type 2 strain in
potato dextrose broth.












































Figure 2.4. Multi-colored mycelial sectors produced by a
single spore culture of a Colletotrichum
gloeosporioides type 2 strain in potato dextrose
broth.











































Figure 2.5. Mycelial injury can induce type 2 (non-
sporodochia-forming) type 2 strains to form
sporodochia.









22

indicated by inoculation tests, confirming previous results

(Sonoda and Pelosi 1988). Variation in the ability of type 1

and type 2 strains to cause disease on Tahiti lime flowers

was not observed. However, only a single high inoculum

concentration was used for pathogenicity testing.

Differences in virulence perhaps could be found if a

dilution series of inoculum were used in assessing the

disease-causing potential of the strains. Sonoda and Pelosi

(1988) and Agostini et al (1992) suggested that slowly

growing, orange-colored strains (type 1 strains) were the

actual causal agent of PFD because only they could be

consistently isolated from diseased petals in the field

while the gray (type 2) strains were isolated primarily from

leaves, stems and fruit. This observation has been confirmed

by other workers (Agostini et al. 1992; Gantotti and Davis,

personal communication). Clearly, further pathogenicity

testing and field sampling are needed to confirm whether one

or both types of the pathogen are important in PFD

epidemics.

The genetic and molecular basis of the morphological

diversity caused by sectoring of C. gloeosporioides is yet

to be elucidated. One of the probable genetic explanations

for the observed morphological diversity and sectoring of

type 2 strains may be heterokaryosis followed by

parasexuality or nuclear sorting out. Heterokaryosis and

parasexuality have been found to contribute to the variation









23

of C. gloeosporioides as reviewed by Baxter et al. (1985).

The strains studied carried only <7% spores with 2-3 nuclei.

Therefore, heterokaryosis or nuclear sorting out may not be

a cause of 100% sectoring observed in type 2 strains.

Multinuclear spores have been reported in C. gloeosporioides

as well as many other fungi (Panaccione et al. 1989; Shirane

et al. 1989; TeBeest et al. 1989). The multinuclear

condition may arise from division of the nucleus without

division of the spore (Churchill 1982). Hence, it may

represent homokaryotic condition.

An interesting phenomenon observed between type 1 and

type 2 strains is the differential sensitivity to benomyl

(methyl-2-benzimidazole carbamate: active ingredient in the

commonly used fungicide benlate). Type 2 strains were

completely inhibited by the levels of benomyl tested while

type 1 strains were tolerant although their growth rates

were significantly reduced (Table 2.2). The benomyl

tolerance of type 1 strains may have practical consequences

to the control of this disease. Current control measures

include spraying benomyl to control PFD (Fagan 1984b). If

type 1 strains are the primary causal agent of PFD as

previously suggested (Sonoda and Pelosi 1988; Agostini et

al. 1992; Gantotti and Davis, personal communication),

spraying in the field may only partially inhibit the

virulent pathogen while completely eliminating the less









24

virulent form. While slowing the epidemic in the short run,

this practice may have the long-term effect of selecting for

the most virulent form of the fungus.
















CHAPTER 3
DNA POLYMORPHISMS FOUND AT MANY GENETIC LOCI EXAMINED
IN COLLETOTRICHUM GLOEOSPORIOIDES


Introduction


Ribosomal DNA in Fungi


The nuclear ribosomal RNA (rRNA) genes of eukaryotes

are clustered in tandemly repeating units known as ribosomal

DNA (rDNA) unit repeats. In fungi as in many other

eukaryotes, each rDNA unit repeat consists of coding regions

for small subunit, SSU (17-18S), 5.8S, and large subunit,

LSU (25-26S) rRNA and intervening internal and external

transcribed and non-transcribed spacer (NTS) regions

(Fedoroff 1979; Chambers et al. 1986). Each rDNA unit repeat

codes for a 35S rRNA precursor which gives rise to SSU and

LSU rRNAs. In Neurospora crassa and Saccharomyces cerevisiae

a 35-37S rRNA precursor cleaves into a 17-18S rRNA of the

small ribosomal subunit (37S), and the 5.8S and 25S rRNAS of

large ribosomal subunit (60S) required for building 80S

ribosomes (Russell et al. 1976; Bell et al. 1977; Planta et

al. 1980).

The number of times rRNA genes are repeated varies

depending on the species of the organism. It was estimated









26

that there are about 185-225 copies of rDNA unit repeats in

Neurospora crassa (Krumlauf and Marzluf 1980; Rodland and

Russell 1982), 140 copies in yeast, Saccharomyces cerevisiae

(Schweizer et al. 1969; Rubin and Sulston 1973), 59 copies

in Rhizoctonia solani (Thanatephorus praticola AG-4)

(Vilgalys and Gonzalez 1990) and 60-90 copies in Coprinus

cinereus (Cassidy et al. 1984) per haploid genome.

Another rRNA gene recognized in fungi is 5S rRNA gene.

The 5S rRNA gene may be present within the rDNA unit repeat

or dispersed elsewhere in the genome. The 5S rDNA sequences

are located within the same rDNA unit repeat in S.

cerevisiae (Bell et al. 1977), Mucor racemosus (Cihlar and

Sypherd 1980), S. rose and S. carlsbergensis (Verbeet et

al. 1983), C. cinereus (Cassidy et al. 1984), Schizophyllum

commune (Buckner et al. 1988), R. solani (Vilgalys and

Gonzalez 1990), the slime mold, Dictyostelium discoideum

(Maizels 1976), and water mold, Achlya ambisexualis, (Rozek

and Timberlake 1979). It is located elsewhere in the genome

in N. crassa (Free et al. 1979; Selker et al. 1981),

Schizosaccharomyces pombe (Tabata 1981), Aspergillus

nidulans (Borsuk et al. 1982), yeasts, Yarrowia lipolytica,

(Van Heerikhuizen et al. 1985), and Cochliobolus

heterostrophus (Garber et al. 1988).

In all known cases the 5S rRNA is transcribed

independently as a primary transcript separate from the 35-

37S rRNA precursor transcript (Udem and Warner 1972;









27

Miyazaki 1974). When the 5S rRNA gene is within the same

unit repeat the 5S rRNA gene could be located in the same

strand, transcribed in the same direction as the other rRNA

genes or in the opposite strand, and transcribed in an

antiparallel manner (Aarstad and Oyen 1975). In C. cinereus

the 5S rRNA gene is transcribed in the same direction as the

rest of the rRNA genes (Cassidy et al. 1984).


Ribosomal DNA is Polymorphic in Many Fungi


Ribosomal DNA is a unique genetic marker which can be

used in the study of relatedness among organisms. Generally

the number of rDNA unit repeats is maintained from

generation to generation of an organism. The meiotic

recombination is suppressed within the rDNA array. In N.

crassa (Russell et al. 1988) and in C. cinerus (Cassidy et

al. 1984) the rDNA was shown to be inherited in a simple

stable Mendelian fashion exhibiting an approximately 1:1

ratio of the two parental rDNA types. No meiotic

recombinants were detected among the progeny indicating that

non-sister chromatid crossing over was highly suppressed in

the rDNA region of these organisms. However, Butler and

Metzenberg (1989 and 1990) demonstrated that N. crassa rDNA

can undergo unequal sister chromatid exchange and that the

number of rDNA unit repeats does not segregate in a simple

Mendelian fashion. Their observations suggested that









28
although the same rDNA RFLP can be inherited, the number of

unit repeats can be different from either of the parents.

Within a given species, the members of the rRNA gene

family are reasonably homogeneous in sequence, as are their

associated spacer sequences, despite frequent length and

restriction site differences among the latter. Yet there are

interspecific differences in sequence and these appear to be

much more pronounced for spacers than for genes. Smith

(1973) suggested that the differences between genes and

spacers might be in the rate at which they accumulate

mutations. Chromosomes containing mutations deleterious to

gene function would be eliminated by natural selection while

neutral spacer mutations would be retained in the

population. Hence, spacers change more rapidly than genes

simply by retaining a larger fraction of mutation (Smith

1973).

Length heterogeneity and restriction site polymorphisms

in rDNA has been commonly observed in many fungi. These

polymorphisms were common among strains of S. cerevisiae

(Petes and Botstein 1977), N. crassa (Russell et al. 1984),

S. commune (Specht et al. 1984), C. cinerus (Wu et al.

1983), and Y. lipolytica (Clare et al. 1986). Both

restriction site and length polymorphisms have been also

observed among biological species of Armillaria (Anderson et

al. 1989). Polymorphisms of rDNA in many fungi are located

within the NTS region of the rDNA unit repeat (Cassidy et









29

al. 1984; Van Heerikhuizen et al. 1985; Rogers et al. 1989).

Chambers et al. (1986) compared the 8.4 kb rDNA unit repeat

of N. intermedia and N. sitophila with the 8.7 kb long rDNA

unit repeat of N. crassa and found that the 300 bp

difference was within the NTS region. Verbeet et al. (1983),

comparing S. rose and S. carlsbergensis by heteroduplex

analysis, concluded that the NTS regions are largely non-

homologous in sequence whereas the transcribed regions are

essentially homologous. Russell et al. (1984) studied the

organization of the rDNA unit repeat in the strains of N.

crassa, N. tetrasperma, N. sitophila, N. intermedia, and N.

discreta and found that the size of the unit repeat has been

highly conserved among the strains of Neurospora. However, a

restriction enzyme site polymorphism in the NTS region was

found between the strains. This restriction site

polymorphism was strain-specific and not species-specific.

Restriction enzyme mapping of rDNA in yeast, Kluyveromyces

species has shown a length variation, and the variability

was found to reside in the NTS region (Lachance 1989).

Martin (1990) reported the presence of restriction site and

length polymorphisms within single oospore isolates of the

Oomycete genus Pythium, and the differences were found

within the NTS region and 3' end of the 26S coding region.

The NTS region has also been useful to study the

phylogenetic relatedness among fungal species and other

organisms (Verma and Dutta 1987).









30

The transcribed intergenic spacer (ITS) has also been

shown to be variable in fungi. In S. commune location of

strain-dependent length polymorphisms resided in the ITS

region between 18S and 5.8S cistrons (Buckner et al. 1988).

Chambers et al. (1986) compared the sequences of ITS regions

for N. crassa and S. carlsbergensis, and found that there is

a general lack of homology between the internal transcribed

spacer regions between 5.8S and 26S rRNA genes of these two

species. Buchko and Klassen (1990), using PCR technique to

amplify the ITS region, demonstrated length heterogeneity in

strains of Pythium ultimum.

The locations of rDNA polymorphisms were not confined

to the ITS regions. Polymorphisms within the coding regions

of rRNA genes due to addition or deletion of restriction

enzyme sites were found in fungi (Chambers et al. 1986).

Another cause of rDNA polymorphism in eukaryotes is the

presence of introns in the coding regions. In fruit fly,

Drosophila melanogastor, the presence of an intron in the

coding region of the 28S rRNA gene has given rise to

polymorphism (Glover and Hogness 1977) of rDNA in this

organism. In fungi there are no conclusive reports for the

presence of introns in rRNA genes. However, Buckner et al.

(1988) examining the strain-dependent rDNA length

polymorphism in S. commune suggested the possibility of

having an intron in the coding region of the 18S rRNA gene.









31

Deletions of large fragments of rDNA may also occur in

organisms as reported by Malezka and Clark-Walker (1989). A

deletion of a 300 kb chromosomal fragment containing 35-40

rRNA cistrons has given rise to a new petite positive strain

of Kluyveromyces lactis.

One of the objectives of this study is to investigate

the variation of rDNA among the strains of C.

gloeosporioides causing PFD.


Fungal Cutinase Genes and Cutinase Isozvmes


Plant pathogenic fungi penetrate their hosts through

the cuticle of epidermal cells or through cutinized cells

below natural apertures. Penetration may take place by

mechanical pressure (Brown and Harvey 1927; Brown 1936;

Pristou and Gallegly 1954; Chakravarty 1957; Wood 1960;

Meredith 1964; Bonnen and Hammerschmidt 1989b), or by

enzymatic degradation of the cuticle (De Bary 1887; Miyoshi

1895; Linskens et al. 1965; Akai et al. 1968; Kunoh and Akai

1969; Shayakh et al. 1977; Kolattukudy 1985) or by both

(Ellingboe 1968; Shishiyama et al. 1970; Nicholson et al.

1972).

The plant cuticular barrier is composed of a

biopolymer, cutin and associated waxes providing a

protective covering against pathogen invasions and hazardous

effects of environment (Martin and Juniper 1970). The

structure of cuticle varies from one plant to another, and











it is influenced by genetic background as well as

environmental factors (Martin and Batt 1958; Martin 1964).

Almost all parts of the plant, surfaces of epidermal cells

of aerial plant parts, substomatal areas, mesophyll and

palisade cells (Martin and Juniper 1970; Sitholey 1971),

flower parts, seed coat (Kolattukudy et al. 1974) fruit

(Espelie et al. 1980), roots and tubers (Kolattukudy and

Agrawal 1974; Kolattukudy et al. 1975) contain a cuticular

layer.

The biopolymer, cutin is composed of C16 and C18 hydroxy

and hydroxy epoxy fatty acids (Van den Ende and Linskens

1974; Espelie et al. 1980; Kolattukudy 1980, 1981). The

composition of the cutin polymer and the proportions of C16

and C,1 fatty acid monomers may vary depending on plant

species or varieties, organs of the same plant, or on growth

conditions (Espelie et al. 1979; Kolattukudy 1980).

The enzyme, cutinase can facilitate the hydrolysis of

cutin into its components (Baker and Bateman 1978; Dickman

et al. 1982). These hydrolysis products of cutin are also

potent inducers of the cutinase gene of the penetrating

fungus (Woloshuk and Kolattukudy 1986). A small amount of

cutinase is constitutively expressed in the fungal spore

which senses the contact with the plant cuticle via the

unique cuticle monomers generated by this small amount of

cutinase. Consequently, these monomers trigger the

expression of the cutinase gene/ genes needed for the









33

production of cutinases which eventually degrade the cuticle

(Kbller et al. 1982; Kolattukudy 1985; Woloshuk and

Kolattukudy 1986; Podila et al. 1988; Kolattukudy et al.

1989).

Many plant pathogenic fungi examined have shown

production of different levels of cutinase isozymes (Purdy

and Kolattukudy 1975a; Lin and Kolattukudy 1980; Kolattukudy

et al. 1981). Direct observational, enzymological and

histochemical evidences have suggested that cutinase is

essential for the penetration of the plant by the fungal

pathogens. Specific antibodies prepared against cutinase

from Nectria haematococca (Fusarium solani f. sp. pisi,

Shaykh et al. 1977) and/ or diisopropylfluorophosphate

(DFP), a potent inhibitor of serine esterases, can prevent

infection of the host by this fungus indicating that

cutinase plays an essential role in the infection process

(Kolattukudy 1979). Dickman and Patil (1986) obtained

cutinase-deficient mutants of C. gloeosporioides, the causal

agent of papaya anthracnose, and found that they were

nonpathogenic to the intact papaya fruit. However, these

cutinase-deficient mutants produced normal lesions when

papaya surfaces were artificially wounded or treated with

purified cutinase enzyme. Dickman et al. (1989) were able to

introduce a Fusarium cutinase gene into a wound pathogen,

Mycosphaerella species through genetic transformation. These

transformants acquired the capacity to infect intact papaya









34

fruits, and the infection by them was prevented by the

treatment of antibodies against Fusarium cutinase.

Cutinolytic enzymes have been purified and

characterized from various plant pathogens (Kolattukudy

1980, 1985; K6ller 1991) including Colletotrichum

gloeosporioides. The single enzyme produced by a strain of

C. gloeosporioides isolated from papaya fruit had a

molecular weight of 24 kd (Dickman et al. 1982) which is

very similar in size to other fungal cutinases (Kolattukudy

1980, 1985; K6ller 1991).

There is considerable heterogeneity of molecular,

immunological and enzymological properties and primary

sequences of the cutinase enzymes (Kolattukudy 1985;

Ettinger et al. 1987; Trail and Kl11er 1990). Sequence

comparison of the cutinase genes cloned from C.

gloeosporioides and N. haematococca revealed considerable

dissimilarity. Even though both cutinase genes shared

homologous regions critical for activity and structural

integrity, only 43% of the amino acids were directly

conserved (Ettinger et al. 1987). Profound differences in

cutinase appear to exist even among Colletotrichum species

(Kolattukudy 1987). A cDNA clone of the cutinase gene from

C. capsici hybridized to genomic DNA from C. graminicola and

C. gloeosporioides, but not with C. orbiculare (syn. C.

lagenarium) or C. coccodes DNA. Though not extensively

investigated, the phenomenon of cutinase diversity is also









35

reflected in enzyme kinetics and activity. For example,

cutinolytic activity of esterases purified from N.

haematococca (Purdy and Kolattukudy 1975b) and F. roseum

culmorum (Soliday and Kolattukudy 1976) was highest at

alkaline conditions (pH 10), whereas an optimum of pH 6.5

was determined for the enzyme derived from Venturia

inaequalis (Kller and Parker 1989). Baker and Bateman

(1978) assayed sixteen plant pathogenic fungi, Botrytis

cinera, B. squamosa, Cladosporium cucumerinum, C.

graminicola, N. haematococca, F. roseum, Gloeocercospora

sorghi, Helminthosporium carbonum, H. maydis (race T),

Pythium aphanidermatum, P. arrhenomanes, P. ultimum, R.

solani, Stemphylum loti, and Sclerotium rolfsii and found

that they can produce various levels of cutinase isozymes

with acidic or alkaline pH optimas. Evidence has been

presented that these differences in enzymatic properties may

allow for the tissue specificity of pathogens. Trail and

K511er (1990) reported an acidic pH optimum for the leaf

pathogen, Cochliobolus heterostrophus, pH 6.5 and an

alkaline pH optimum for the stem pathogen, R. solani, pH

8.5. The leaf and stem pathogen, Alternaria brassicola,

produced two cutinases, one with acidic and the other with

alkaline pH optima, pH 7.0 and 9.0 respectively. Differences

also have been reported for the cutinases produced by N.

haematococca and C. gloeosporioides. Only the enzyme from

the latter accepted palmitate as a substrate and the









36

specific esterase activity with both p-nitrophenol butyrate

and polymeric cutin was reported to be substantially lower

(Dickman et al. 1982).

The ability of a pathogen to produce cutinase can be

used to measure the infecting capacity of the fungal

pathogen (Dickman et al. 1982; K6ller et al. 1982). Thus the

regulation of expression of the cutinase gene could be

highly relevant to pathogenesis. Therefore, the cutinase

gene may be a good genetic marker to examine polymorphisms

among populations of fungi with differing specificities and

capabilities of causing plant disease.

One of the goals in this study was to investigate if

differences in morphologically defined type 1 and type 2

strains (see Chapter 2) are also reflected in the cutinase

isozymes and genetic organization of cutinase gene or genes.


Restriction Fragment Length Polymorphisms (RFLP) in Fungi


When fungal nuclear DNA is digested with a restriction

enzyme an enormous number of fragments generally result. In

order to study the restriction fragment pattern of DNA from

a specific chromosomal locus these fragments are size

fractionated by gel-electrophoresis, and individual

fragments are identified by Southern hybridization to

labelled probes (Southern 1975; Bernatzky 1988). Each

restriction fragment that hybridizes to a given probe

constitute a discrete chromosomal locus. Alleles can be









37

differentiated by the variation in restriction sites.

Restriction fragment length polymorphisms result from

specific differences in DNA sequence such as single base

pair substitution, additions, deletions, or chromosomal

changes (inversions and translocations) that alter the

fragment size obtained by restriction enzyme digestion.

First demonstrated by Grodzicker et al. (1974) for mapping

temperature-sensitive mutations in adenovirus, RFLP analysis

has contributed significantly in genetic analysis of many

organisms.

Genetic studies of plant pathogenic fungi have been

difficult due to lack of easily assayed genetic markers.

Restriction fragment length polymorphism has become a

popular tool for studying genetics of fungi because RFLP

markers are precise, codominant, selectively neutral, easy

to assay, and provide an unlimited number of genetic markers

(Michelmore and Hulbert 1987). Restriction fragment length

polymorphism could provide sufficient markers for the

development of detailed linkage maps for the plant

pathogenic fungi. It is also useful in studying genetic

variation, genomic organization and population genetics of

fungi. Combined with pulsed field gel electrophoresis RFLP

analysis provides a powerful tool to monitor genetic changes

throughout the genome (Michelmore and Hulbert 1987).

Analysis of RFLP markers that flank genetic loci such as

virulence genes can provide information on the genetic basis









38

of any changes in phenotype. Closely linked RFLPs can be

used as tags for important traits. With RFLP markers it is

possible to create a molecular fingerprint of specific

individuals in a population. Hence, RFLPs provide a tool for

studying asexually reproducing populations of fungi. Engels

(1981) and Hudson (1982) presented mathematical models for

the genetic determination of variation among individuals in

a population using RFLP.

Use of RFLPs to measure genetic relatedness among

strains and closely related species of plant pathogenic

fungi is still in its beginning. Genetic variability of

several plant pathogenic fungi, Armillaria mellea (Anderson

et al. 1987), Sclerotinia species (Kohn et al. 1988),

Septoria tritici (McDonald and Martinez 1990) and

Aspergillus species (Someren et al. 1991) has been studied

using RFLP genetic markers. In C. gloeosporioides two

population subgroups were recognized by distinct RFLP

patterns detected by human minisatelite probes for

hypervariable regions within the genome (Braithwaite and

Manners 1989). Linkage maps have been developed for the

lettuce downey mildew fungus, Bremia lactucae using RFLPs as

genetic markers (Hulbert and Michelmore 1988; Hulbert et al.

1988). Hulbert et al. (1988) also reported the linkage of an

avirulance gene and a RFLP locus and suggested the

possibility of cloning the avirulance gene by chromosome

walking.









39

Castle et al. (1987) distinguished the commercial

mushroom, Agaricus brunnescens, from A. bitorquis using

distinct RFLP patterns. These RFLP patterns were used in the

identification of homokaryotic, heterokaryotic and hybrid

strains of this fungus (Castle et al. 1987) Summerbell et

al. (1989) followed the segregation of RFLPs in wild

collected and artificially synthesized heterokaryotic

strains of A. brunnescens to investigate meiosis and the

meiotic recombination in this fungus.

The objective of this study is to determine if type 1

and type 2 Colletotrichum gloeosporioides are genetically

distinguishable. This will be done by examining rDNA

polymorphisms, the diversity of cutinase enzymes at the

isozyme and molecular level, and other molecular markers to

examine RFLPs in type 1 and type 2 strains of C.

gloeosporioides causing PFD of Tahiti lime and sweet orange.


Materials and Methods


Strains of Colletotrichum qloeosporioides


Strains of C. gloeosporioides, host and geographic

location are listed in the appendix A. Each strain is a

single spore culture grown. The place, year, and the host

tissue of isolation are mentioned in appendix A.











DNA Extraction


Fungal mycelium was grown in potato dextrose broth

(PDB) for seven days, harvested, frozen at -80 oC overnight

and lyophilized to complete dryness. The mycelium was ground

into a powder in liquid nitrogen using a mortar and pestle.

The mycelium powder was mixed with extraction buffer (100 mM

Tris pH 8.0, 50 mM EDTA, 100 mM NaC1, 10 mM 8-

mercaptoethanol, 1% SDS, in H20) to make a slurry and

incubated at 65 OC for 30 min. One half volume of 5 M

potassium acetate (60 ml of 5 M potassium acetate, 11.5 ml

glacial acetic acid and 28.5 ml H20) was added to samples

and incubated on ice for 30 min. The supernatant was

collected by centrifugation at 12000 x g for 15 min and was

treated with 30-50 Ag/ml of DNase free RNase (30 min at 37

OC, Sigma Chemical Co., St. Louis, MO). After RNase

treatment 200-250 gg/ml Proteinase K (Sigma Chemical Co.,

St. Louis, MO) was added and incubated for an additional 20

min. Samples were purified by phenol:isoamyl alcohol:

chloroform (25:1:24 by volume) extraction and DNA was

precipitated by addition of a two-fold volume of absolute

ethanol. DNA pellets were dissolved in 100 Al of TE (10 mM

Tris pH 8.0, 1 mM EDTA) and further purified by

precipitation with 0.7 volume of PEG/NaCl ((20% PEG 8000 in

2.5 M NaC1, Sigma Chemical Co., St. Louis, MO ) for 20-30

min on ice. Precipitated DNAs were resuspended in TE and

stored at -20 OC.









41

DNA Cloning and Restriction Enzyme Mapping


The rDNA of C. gloeosporioides was identified by

heterologous hybridization with N. crassa rDNA unit repeat

(pMF2, Free et al. 1979). Total DNA from C. gloeosporioides

strains H-25B and H-48 was digested with restriction enzyme

PstI and fractionated on a 0.7% agarose (FMC BioProducts,

Rockland, ME) gel. The piece of the gel containing the 7 to

10 kb (H-25B) or 6-10 kb (H-48) DNA range was cut out and

the DNA eluted by the freeze squeeze method (Thuring et al.

1975). The DNA was ligated to PstI-cut pUC119 (Sambrook et

al. 1989; Yanisch-Perron et al. 1985) and transformed into

Escherichia coli strain ER1647 (E. coli K-12 mcrB', Ref.

Raleigh et al. 1989; Woodcock et al. 1989) or DH5-a

(Sambrook et al. 1989). Ligation, preparation of competent

cells and transformation was carried out according to

Sambrook et al. (1989). Clones hybridizing to pMF2 were

identified and restriction mapped. For constructing

restriction maps, single and double restriction enzyme

digestions of two presumptive rDNA clones, called pCGR1 (8.4

kb from strain H-25B), pCGR2 (6.8 kb from strain H-48), were

size fractionated in 1% agarose gels. Regions homologous to

the LSU and SSU rRNA of N. crassa were mapped by Southern

hybridization to heterologous probes subcloned from plasmid

pMF2 (Free et al. 1979; Martin 1990). The probe specific to

LSU was a 1.7 kb XbaI + BamHI fragment comprising all but









42

150 bp of the 5' end of the 17S coding region. A 2.9 kb

EcoRI fragment containing all but approximately 700 bp of

the 3' end of the 26S coding region in addition to 200 bp of

transcribed spacer sequences adjacent to 5' end was used as

the probe specific to LSU. These subclones were provided by

Dr. F. N. Martin, Plant Pathology Department, University of

Florida. A probe to detect the 5.8 rRNA gene was prepared by

polymerase chain reaction using primer flanking the gene.

The primers 5' TCCGTAGGTGAACCTGCGC 3' and 5'

GCTGCGTTCTTCATCGATGC 3' amplify a 290 bp fragment which

includes the transcribed spacer of the 3' end of the SSU

rRNA gene and the entire 5.8S gene (White et al. 1990).


Enzyme Assays and Electrophoresis of Cutinase


Colletotrichum gloeosporioides cultures were grown in a

mineral medium (Hankin and Kolattukudy 1968) amended with

tritiated cutin as the sole carbon source. Esterase

activities of strains were measured using p-nitrophenyl

butyrate (PNB) and p-nitrophenyl palmitate (PNP) as model

substrates (Kller and Kolattukudy 1982; K6ller and Parker

1989; Purdy and Kolattukudy 1975a). Assays for cutinase

activity, sodium dodecyl sulfate polyacrylamide gel

electrophoresis (SDS-PAGE) and detection of active serine

esterases by tritium-labelled diisopropyl fluorophosphate

(3H-DFP) were as previously described (K611er and Parker

1989; Trail and Kller 1990). These experiments were









43
conducted by Dr. Wolfram K6ller at the New York State

Agricultural Experiment Station, Geneva.


Probes Containing Cutinase Gene Sequences


Oligonucleotide primers, 5' TGCCCCAAGGTCATCTACATC 3'and

5'GAAGTTGGAGGCCAGGTCGGC 3' were synthesized to amplify a 220

bp fragment (intron and flanking sequences) of C.

gloeosporioides cutinase gene by polymerase chain reaction

(PCR). The PCR reaction mixture was prepared in a total of

100 jl containing 100 pM of each primer, 1.25 mM each of

dATP, dTTP, dCTP and dGTP, 2 gg of template DNA, and 10 Al

of reaction buffer (50mM KC1, 10 mM Tris-HCl pH 8.3, 1.5mM

MgCl2, 0.01% gelatin). The PCR mixture was denatured by

boiling 10 min and chilled on ice before adding 2 units of

Taq DNA Polymerase (Promega Inc. Madison, WI). The PCR

temperature cycles were programmed as following in a Coy

Temp Cycler II (Coy Corp., Grass Lake, MI). Denaturation

temperature was 94 OC, annealing temperature was 37 oC and

primer extension was at 72 OC. The first cycle was run 6 min

at 940C, 2 min at 37 oC and 3 min at 720C and the subsequent

30 cycles were run at 1, 2, and 3 min time intervals,

respectively, at these temperatures. Primer extension time

for the final cycle was 10 min. The fragments amplified by

PCR were labelled with 32P dCTP or digoxigenin (dig) dUTP

(Boehringer Mannheim Corp. Indianapolis, IN; see Appendix

C). Both a 220 and a 260 bp DNA fragment amplified by PCR









44

from strain H-48 hybridized to a genomic clone containing,

the cutinase gene, a 2.2 kb SphI DNA fragment, from C.

gloeosporioides (Ettinger et al. 1987). This clone was

provided by Dr. M. B. Dickman, Department of Plant

Pathology, University of Nebraska, Lincoln, NE.

High stringency Southern hybridization (Southern 1975)

using 32P-labelled probes was carried out according to

methods described by Sambrook et al. (1989). Hybridization

and washing of blots were carried out at 68 OC. First and

second washes were with 2X SSC, 0.1% SDS and 0.1X SSC, 0.1%

SDS respectively. The conditions of low stringency Southern

hybridization were as follows. DNA hybridization was at

65 OC and first and second washes were with 2X SSC at 55 oC.

Autoradiography was performed with Kodak X-OMAT AR5 film

(Eastman Kodak Co., Rochester, NY) and Dupont Hi-Plus

intensifying screens at -80 OC.


Detection of Restriction Fragment Length Polymorphisms


Plasmids containing N. crassa genes for anthranilate

synthetase (pNC2, Schechtman and Yanofsky 1983), glutamate

dehydrogenase (pJR2, Kinsey and Rambosek 1984), histidinol

dehydrogenase (pNH60, Legerton and Yanofsky 1985), and B-

tubulin (pSV50, Vollmer and Yanofsky 1986) were used to

detect DNA polymorphisms. Clones of N. crassa genes were

obtained from the Fungal Genetics Stock Center (Department

of Microbiology, University of Kansas Medical Center, Kansas









45

City, KS). Southern hybridization of 32P-labelled probes

were carried out according to methods described by Sambrook

et al. (1989) and Appendix C.


Results


Ribosomal DNA is Polymorphic in Colletotrichum
gloeosporioides


Southern hybridization of 32P labelled pMF2 to PstI-

digested total blots from C. gloeosporioides strains

detected polymorphic forms of rDNA (Figure 3.1 and 3.2). The

type 1 strains (see Chapter 2 and Appendix A) contained only

a single form (8.4 kb PstI fragment) of rDNA (Figures 3.1

and 3.2). Although the size of the rDNA fragment in type 1

strains H-36 and OCO appears to be slightly higher than 8.4

kb in Figure 3.1, other restriction enzyme digestion tests

concluded that it is 8.4 kb in size (compare figures 3.5,

3.6, and 3.7). The 8.4 kb PstI rDNA fragment was cloned from

Type 1 strain, H-25B and will be referred to as type 1 rDNA.

The restriction fragments obtained by digesting with 10

restriction enzymes and the restriction map of cloned type 1

rDNA unit (pCGR1) for these enzymes are illustrated in Table

3.1 and Figure 3.3 respectively. The restriction map of the

cloned rDNA unit was compared with the total rDNA

restriction fragments of the strain H-25B for 7 enzymes and

was identical (Figure 3.4). The map of cloned rDNA unit from

H-25B was tested against all type 1 strains for three









46

restriction enzymes, HindIII, SphI and SstI (Figures 3.5,

3.6, and 3.7 respectively). All the type 1 strains fell into

an identical group and the hybridization fragments for the

three enzymes agreed with restriction map of the cloned rDNA

fragment from Strain H-25B. Several subcloned fragments from

the NTS region were tested to determine if they can

specifically hybridize to type 1 strains. A 0.4 kb KpnI-PstI

subclone (pCGR1N) from the 3' end of NTS region (Figure 3.3)

was found to hybridize only to type 1 rDNA. The same total

DNA blot in Figure 3.1 was reprobed with the type 1-specific

subclone after removing the previous probe and only type 1

strains show hybridization to the subclone (compare Figures

3.1 and 3.8).

Ribosomal DNA among type 2 strains (see chapter 2) was

polymorphic for PstI (Figures 3.1 and 3.2), SphI (Figure

3.6), and SstI (Figure 3.7). However, HindIII digested DNA

shows a similar pattern of rDNA polymorphism among all type

2 strains (Figure 3.5). Two hybridizing PstI fragments were

detected in strains H-48, 180269, 226802 (8.4 and 6.8 kb),

H-11 (5.0 and 3.4 kb), and H-47 (8.4 and 7.8 kb) by 32P

labelled pMF2 (Figures 3.1 and 3.2). However, the

hybridization intensity of the 8.4 kb band in strains 180269

and 226802 was very low and almost undetectable compared to

6.8 kb hybridizing band (Figure 3.1). All other type 2

strains had only one 6.8 kb hybridizing band. The 6.8 kb

PstI fragment was cloned from strain H-48, and the









47

restriction map was identical when compared with the total

rDNA restriction fragments of the strain H-48 for 7 enzymes

(Figure 3.4). The cloned 6.8 kb PstI fragment from type 2

strain H-48 hybridizing to pMF2 will be referred to as pCGR2

or type 2 rDNA. The restriction fragments obtained by

digestion of clones of type 1 and type 2 rDNA unit with

various restriction enzymes and restriction enzyme maps are

listed in Table 3.1. The length of type 1 rDNA differs from

type 2 by the size of the NTS region of the unit (Figure

3.3). The restriction map of pCGR2 is distinct from that of

type 1 rDNA and the NTS is 1.6 kb shorter.


Ribosomal RNA Genes


In addition to the length heterogeneity, type 1 and

type 2 rDNA units differ by having restriction site

polymorphisms and addition and deletion of restriction sites

within coding regions for rRNA as well as intergenic

regions. Restriction sites for SmaI and SstI within the SSU

coding region and a EcoRI site within the 5.8 S coding

region were found in type 1 rDNA. For the 10 restriction

enzyme sites examined none was detected within the SSU rRNA

and 5.8S RNA coding regions of type 2 rDNA. The coding

region for the LSU lies within BamHI and EcoRI sites for

both type 1 and type 2 rDNA, and were detected as 3.1 and

3.0 kb hybridizing fragments respectively. Restriction site












TABLE 3.1 Restriction fragments obtained by complete
digestion of the three ribosomal DNA clones


No. Restriction Fragment size (kb)
enzyme pCGR1 pCGR2


PstI
HindIII


SphI


EcoRI


BamHI

KpnI




HincII


XbaI

SmaI


SstI


Clal


4.6


8.4
4.1
3.2
1.1
6.0
1.4
1.0
2.7
2.5
2.4
0.8
5.5
2.9
6.2
0.6
0.6
0.6
0.4
3.3
2.8
1.4
0.7
0.2
7.8
0.6
2.2
2.2
2.0
1.0
1.0

2.0
1.8
N


6.8
3.2
3.1
0.5
4.2
2.5
0.1
3.2
2.6
1.0

4.0
2.8
6.4
0.4


3.0
1.6
1.0
0.6
0.6
6.3
0.5
4.3
1.6
0.9


4.6


1.8
0.4
N


N = No restriction site detected









49

polymorphisms for enzymes HindIII, EcoRI, SstI, HincII, and

SmaI were detected within the coding region for LSU rRNA in

the type 1 and type 2 rDNA forms. The NTS region of type 1

rDNA has 4 KpnI sites with three present at equal distance

of 0.6 kb, whereas within the NTS region of type 2 rDNA

there is only a single KpnI site. The additional 1.6 kb NTS

region fragment in type 1 rDNA contains two KpnI sites, each

0.6 kb apart and a 0.4 kb PstI/KpnI type 1 rDNA specific

fragment.


Diverse Cutinases and Cutinase Genes are Found in Type 1 and
Type 2 Strains of Colletotrichum gloeosporioides


Cutinase production by fungal mycelium can be induced

by cutin monomers (Lin and Kolattukudy 1978). Similarly, all

Tahiti lime and Sweet orange strains of C. gloeosporioides

excreted esterases under these inductive conditions when

cutin was used as the sole carbon source. Although, both

model cutin substrates, p-nitrophenol-butyrate and -

palmitate, were hydrolyzed, the ratio of these two

activities was remarkably different (Table 3.2). Cutinolytic

activity was identified for all isolates and was

consistently higher at pH 6.0 than at the alkaline pH of

9.5. Extracellular proteins were labelled with 3H-DFP and

used as active site probe for serine esterase. Two esterases

in the molecular weight range common to many known fungal

cutinases (17kd-32kd, Kolattukudy 1980; Tanabe et al 1988;










50

TABLE 3.2 Extracellular enzyme activities of Colletotrichum
gloeosporioides



Strain PNBase PNPase Cutinase PNB/PNP PNB/
mg/ml mg/ml kBq/h/mg Cutinase pH 6
pH 6 pH9.5 ratio




H-1 4731 255 18.7 2.0 9.4 18.6 254

IMB-3 6475 351 25.4 2.8 9.1 18.5 255

H-3 3678 302 23.9 4.5 5.3 12.2 154

H-4 8784 752 57.5 8.0 7.2 11.7 153

LP-1 5844 499 28.6 8.3 3.4 11.7 204

H-46 477 50 5.9 1.5 3.9 9.6 81

H-12 4692 554 40.0 1.3 30.8 8.5 117

H-25B 3627 488 27.5 5.5 5.0 7.4 132

Maran 3575 484 23.1 10.3 2.2 7.4 155

H-9 2193 329 18.8 4.2 4.4 6.7 117

H-48 4099 662 27.0 6.1 4.4 6.2 152

180269 4668 771 28.1 4.9 5.7 6.1 166

226802 4695 782 49.3 10.7 4.6 6.0 95

H-36 1863 369 23.7 5.8 4.1 5.0 78

Control 1456 332 12.8 7.9 1.6 4.4 144


PNB=p-nitrophenyl butyrate
PNP=p-nitrophenyl palmitate
Control consisted of all treatment without a fungal strain.










51
Trail and Kller 1990) were present for all strains (Figure

3.9). The molecular weight of these proteins differed among

strains and was correlated to C. gloeosporioides RFLP-types.

All strains of type 1 contained bands of 24 and 21 kd,

whereas all strains of type 2 contained 26 and 19 kd bands.

An additional esterase with a molecular weight of about 70

kd, which was not reported for the papaya isolate of C.

gloeosporioides (Dickman et al. 1982), was present

throughout the set of isolates. The high molecular weight

esterase was slightly larger for type 1 strains. The

relative contribution of this high molecular weight esterase

to the total esterase and cutinase activities remains

unknown. The enzyme might be similar to the 60 kd alkaline

cutinolytic esterase isolated from C. lagenarium (Bonnen and

Hammerschmidt 1989a) or the 54 kd non-specific esterase of

N. haematococca (Purdy and Kolattukudy 1975a).

A genomic clone containing the cutinase gene from a

papaya strain of C. gloeosporioides (Ettinger et al 1987)

was 32P-labelled and used to probe SphI-digested DNA of C.

gloeosporioides from citrus. This clone contained a 2.2 kb

SphI fragment which included the cutinase gene (189 bp exon-

52 bp intron-486 bp exon) and 5' and 3' flanking sequences

(Ettinger et al 1987). The probe hybridized to a 2.2 kb SphI

fragment only in type 2 strains (Figure 3.10). DNA from type

1 strains showed no detectable level of hybridization at the

high level of stringency (see materials and methods) used









52

for Southern hybridization. Although the cutinase gene

sequence shows no polymorphism among type 2 strains for the

restriction enzyme SphI, a restriction fragment length

polymorphism can be detected among type 2 strains for

HindIII (Figure 3.11). A 9.0 kb HindIII fragment hybridized

to the probe in all type 2 strains except 226802 which show

hybridization to a 8.0 kb fragment. Type 1 strains did not

show any detectable level of hybridization to the probe at

this level of stringency used for Southern hybridization

(see Appendix C). The long exposure (>1 month) of this blot

resulted in appearance of 4.8, 5.4, 6.6, 7.4, and 9.0 kb

HindIII hybridizing fragments of low level homology (Figure

3.12). Non-radioactive hybridization (Genius, Boehringer

Mannheim Corp. Indianapolis, IN) under low stringency

conditions shows weak hybridization of the probe to a 7.4 kb

HindIII fragment from type 1 strains (Figure 3.13).

Polymerase chain reaction amplification using

oligonucleotide primers flanking an intron sequence in the

cutinase gene resulted in a single 220 bp fragment when H-3

(type 1 strain) or H-12 (type 2 strain) DNA was used as a

template. However, strains LP-1 (type 1) and H-48 (type 2)

produced two amplified fragments, 220 and 260 bp (Figure

3.14). The 220 and 260 bp fragment amplified by PCR from

strain LP-1, when used as probes for Southern hybridization,

also hybridized to numerous HindIII fragments. Approximately

5-10 restriction fragments, ranging in size from 0.5 to >10









53

kb were identified (Figure 3.15 and 3.16 respectively).

Restriction fragments from all type 1 strains were almost

entirely identical. Hybridizing fragments from the type 2

strains showed dissimilar patterns. Hybridization of these

two probes to total DNA resulted in distinct DNA fingerprint

for type 1 strains.


Subgroups of Colletotrichum gloeosporioides have Distinct
RFLP Patterns Detected by Many Genetic Markers


Four clones of N. crassa genes were used as

heterologous probes to identify additional genetic loci in

HindIII-digested DNA from C. gloeosporioides strains. The

probe pSV50, containing the gene for B-tubulin, hybridized

to a 3.2 kb fragment in type 1 strains but a 5.0 kb fragment

in type 2 strains (Figure 3.17). The probe, pJR2, containing

the gene for glutamate dehydrogenase, hybridized to a 3.2 kb

fragment only in type 2 strains, but only diffuse

hybridization was observed in type 1 strains (Figure 3.18).

The probe, pNH60, containing the gene for histidinol

dehydrogenase, hybridized to both a 3.8 and a 4.3 kb

fragment in type 1 strains but hybridized to 3.3 and 4.8 kb

fragments in type 2 strains (Figure 3.19).


























'00
CO N CO kO
I I CO I U
Sa H a0

OR0DcJ


Figure 3.1


Polymorphic forms of ribosomal DNA unit in C.
gloeosporioides strains. Total DNA was digested
with PstI and Southern hybridized with 32P
labelled pMF2 (N. crassa rDNA unit repeat).
Lanes H-12, 226802, H-36, and OCO shows slower
migration of DNA than expected. Lane H-4 DNA is
degraded. Numbers at the left indicate the size
of restriction fragments in kilobases (kb).


I I


go rg
Iv In f
a4 a


24


r
























kb l




5-0
3-4


Figure 3.2


kb TTT


Various restriction fragments contain ribosomal
DNA in type 2 strains of C. gloeosporioides.
Total DNA was digested with PstI and Southern
hybridized with P labelled pMF2 (N. crassa rDNA
unit repeat). Numbers at the left indicate the
size of the major restriction fragments in
kilobases (kb).


- *












1 8 3 109 3 475 2 7 10 9


88U 5.8 8


L8U


104 572 7 9 97 74 2 1


II II


8U 5.8 8
SaU 6.8 8


Figure 3.3


I I I II I CR2
LSU


Restriction enzyme maps for cloned rDNA units
(pCGR1 from Colletotrichum gloeosporioides type 1
strain, H-25B, and pCGR2 from type 2 strain, H-
48). Regions hybridizing to large subunit rRNA
(LSU), small subunit rRNA (SSU) from N.crassa and
the PCR amplified 5.8S rRNA gene from C.
gloeosporioides are indicated by solid boxes.
Restriction enzyme sites are as follows. PstI
(1), HindIII (2), SphI (3), EcoRI (4), BamHI (5),
KpnI (6), HincII (7), XbaI (8), SmaI (9), and
SstI (10).


1 3108


l III I i iI I I II_ I I I I I pCR1


7 4 9 46 9 8 26 671



























8 4




H H H
8b 4 ) Qu ^QQ0 (*Q

25 ~~cc>;q~iq
84 A ~ k *


I -T


Figure 3.4


e4 H

'UJ.J0 i"
--
C- -
r^ BQ-:
& rr 3
\n''q~
H H1- fg
(0 *U 4J 00


I Q(Q0*


The rDNA of C. gloeosporioides strains H-25B
(first 10 lanes from left) and H-48 (next 10
lanes) digested with various restriction enzymes
and detected by Southern hybridization using 32P
labelled pMF2 (N. crassa rDNA unit repeat) as a
probe. The numbers at the left indicate the size
of major restriction fragments in kilobases (kb).


n




















I v S I I I & I I I f0 I I 0 N I U







4-3 .......
4-1
3 -1 1. .E:...


"" ... ,,E ..
::!!


Figure 3.5


Ribosomal DNA polymorphism in C. gloeosporioides
strains. Total DNA was digested with HindIII
and Southern hybridized with 32P labelled pMF2
(N. crassa rDNA unit repeat). H-4 DNA is
degraded. The numbers at the left indicate the
size of restriction fragments in kilobases (kb).

























kb CJ IN



7.o0


2-5


1*4 o-


~I r 'c
Iou I~c


Figure 3.6


Ribosomal DNA polymorphism in C. gloeosporioides
strains. Total DNA was digested with SphI and
Southern hybridized with P labelled pMF2
(N. crassa rDNA unit repeat). The numbers at the
left indicate the size of major restriction
fragments in kilobases (kb).


\oh
~0
CO
0..Io


409M 4


MODO



















c) me 00
9 0 02 k m o 0uco\
I -'e If to I I O C-41 4U I
kb C-4 g 0a- Mcu--





6-4



2-0


Figure 3.7


Ribosomal DNA polymorphism in C. gloeosporioides
strains. Total DNA was digested with SstI and
Southern hybridized with P labelled pMF2
(N. crassa rDNA unit repeat). The numbers at the
left indicate the size of major restriction
fragments in kilobases (kb).























I w* I
kcb


04

Iin


OR
$40
EU


%0
~0
00
cEO


8-4
























Figure 3.8 A 0.4 kb PstI/KpnI fragment (pCGR1N) from the
non-transcribed spacer region of cloned rDNA
unit from C. gloeosporioides isolate H-25B
hybridizes only to the 8.4 kilobase (kb) rDNA
form in type 1 strains. Total DNA was digested
with PstI and Southern hybridized with 2P
labelled pCGR1N.


























kd i~:I10


M R
f-4 C4 1.4
rI iaa


N00
0 %D'
OD C4O
T-1 C


21 .- ,W
19 d


Figure 3.9


Fluorography of 3HDFP-treated proteins after SDS-
polyacrylamide gel electrophoresis of
extracellular fluid from C. gloeosporioides
cultures grown on cutin as the sole carbon
source. Numbers at the left indicate the
molecular weight in kilodaltons (kd).





















(A N
rlr~~M E W 0QU
I$ 04 W0 0 A g
kb ( H N 0- U
r((u30


- MO


Figure 3.10 Cutinase gene in C. gloeosporioides type 2
strains. Total DNA was digested with SphI and
hybridized to a 32P labelled probe containing a
cloned cutinase gene. Numbers at the left
indicate the size of restriction fragment in
kilobases (kb).




















0)
I 1-4 00C CO C4

kb 0 I 1 1 Iu I 1 0


Mq
0
G %0
%cD MO
C4 I
M = 0


- "


Figure 3.11 Restriction fragments hybridize to the cloned
cutinase gene only within type 2 strains. Total
DNA was digested with HindIII and hybridized to
a 32P labelled probe containing a cloned
cutinase gene sequence from C. gloeosporioides.
H-4 DNA is degraded. Numbers at the left
indicate the size of restriction fragments in
kilobases (kb).

























kb I Z 1 I1f 1 OC4 IU


Figure 3.12


Hybridization of total DNA from the indicated
strains to a cloned cutinase gene sequence. The
DNA was digested with HindIII and hybridized to
a 32P labelled probe containing a cloned
cutinase gene sequence from C. gloeosporioides.
The blot was overexposed by placing it next to
X-ray film for more than 4 weeks. H-4 DNA is
partially degraded. Numbers at the left indicate
the size of major restriction fragments in
kilobases (kb).





















IM N


kb #0 1P 1 0N 4 1


)-0
f-4 jr


Figure 3.13


Presumptive cutinase genes in C. gloeosporioides
type 1 and type 2 strains. Total DNA was
digested with HindIII and Southern hybridized to
a digoxigenin labelled probe containing a cloned
cutinase gene sequence from C. gloeosporioides.
H-4 DNA is degraded. Numbers at the left
indicate the size of restriction fragments in
kilobases (kb).























1-4
0


bp U










260
220


Figure 3.14


PCR Amplified fragments using oligonucleotide
primers flanking the intron in the
C. gloeosporioides cutinase gene. Lanes contain
DNA from PCR reactions using for the template
the cloned cutinase gene (CG), or total DNA from
the strains indicated. The control reaction lane
contained no template DNA. Numbers at the left
indicate the size of DNA restriction fragments
in basepairs (bp).










68











a\ C





10















Figure 3.15 RFLP patterns in total DNA from the indicated C.
gloeosporioides strains detected by Southern
hybridization to a 220 bp fragment amplified by
1 0 4 k- WO N























PCR. The total DNAs were digested with HindIII
and the probe was labelled with digoxigenin
dUTP. H-4 DNA was partially degraded. Numbers at
the left indicate the size of major restriction
fragments in kilobases (kb).
fragments in kilobases (kb).





























kb
~eH~ l~cfT 0 vf


CONDo %
v 0 4O
1 ON1U
r-4 C4 C


.a- L


Figure 3.16 RFLP patterns in total DNA from the indicated C.
gloeosporioides strains detected by Southern
hybridization to a 260 bp fragment amplified by
PCR. The total DNAs were digested with HindIII
and the probe was labelled with digoxigenin
dUTP. H-4 DNA was partially degraded. Numbers at
the left indicate the size of major restriction
fragments in kilobases (kb).


. t




























N (A
%0 C(VO M t in cv %o r-i
OMWO\t (A $4 C*4 V-1 V I V M(r CO -4
kb UO MlNN e 04 = =1 v =1=1 =


II I


Figure 3.17


RFLP patterns in total DNA from the indicated C.
gloeosporioides strains detected by Southern
hybridization to the 32P labelled 8-tubulin gene
from N. crassa plasmidd pSV50). The total DNAs
were digested with HindIIl. Numbers at the left
indicate the size of restriction fragments in
kilobases (kb).


*M 0





















%0 C4 In gIo
ri ^- "' "~


Figure 3.18


RFLP patterns in total DNA from the indicated C.
gloeosporioides strains detected by Southern
hybridization to the 32P labelled glutamate
dehydrogenase gene from N. crassa plasmidd
pJR2). The total DNAs were digested with
HindIII. Numbers at the left indicate the size
of the major restriction fragment in kilobases
(kb).


I
-4C
kb I1
r-


en N
'Do
.ugVOD
l Oon
I rON


000


i~i~i~i~i~i~i~i~i~i~i~i~i~i~i~i~i~i~i~i~

















%VI 4 0 oI 0
kb i Z i i P i i i to i i co c4 i u
=H==A===Z r-4 N_.a= 0_
Gi n iaiti


33 1
!+!: o+ \,


Figure 3.19


RFLP patterns in total DNA from the indicated C.
gloeosporioides strains detected by Southern
hybridization to the 32P labelled histidinol
dehydrogenase gene from N. crassa plasmidd
pNH60). The total DNAs were digested with
HindIII. Numbers at the left indicate the size
of the major restriction fragments in kilobases
(kb).


*9









73

Discussion


In Chapter 2, two types of Colletotrichum

gloeosporioides strains were described based on

morphological and growth characteristics. However,

morphological features vary with culture conditions and

time. Morphological variability always has been the case

with C. gloeosporioides (Burger 1921; Arx 1957). Despite

confusion associated with morphological inconsistency, at

the molecular level we see distinct differences between type

1 and type 2 strains.

The rDNA unit proved to be a molecular marker which can

be used to detect type 1 strains of C. gloeosporioides

(Figure 3.8). Based on the Southern hybridization results,

morphologically stable type 1 strains contain a single form

of rDNA. The restriction map of the cloned rDNA unit (Figure

3.3, pCGR1) from type 1 strain, H-25B was identical when

compared with the restriction map of the genomic rDNA

(compare Figures 3.3 and 3.4). The three enzyme sites

HindIII, SphI, and SstI examined for all type 1 strains

agree with the map of the rDNA clone from strain H-25B

indicating that the form of rDNA present in type 1 strains

has been cloned and mapped (compare figures 3.3, 3.4, 3.5,

3.6 and 3.7). The size of the type 1 rDNA unit, 8.4 kb in C.

gloeosporioides, is within the range of observed rDNA unit

repeat sizes for filamentous fungi such as N. crassa, 9.23

kb (Free et al. 1979), Aspergillus nidulans, 7.8 kb (Borsuk









74

et al. 1982), Schizophyllum commune 9.2-9.6 kb (Specht et

al. 1984), and Thanatephorus praticola, 8.8 kb (Vilgalys and

Gonzalez 1990). The given order of rRNA genes, 5' SSU-5.8S-

LSU 3' (Figure 3.3) within the rDNA unit repeat is similar

in all the fungi examined (Free et al. 1979; Cihlar and

Sypherd 1980; Borsuk et al. 1982; Cassidy et al. 1984;

Buckner et al 1988; Garber et al. 1988; Vilgalys and

Gonzalez 1990). The rDNA unit repeats in C. gloeosporioides

may also code for large, 35-37S, precursor rRNAs which give

rise to SSU, 5.8S, and LSU rRNAs required for the building

of 80S ribosomes (Russell et al. 1976; Bell et al. 1977;

Planta et al. 1980). The search for a specific fragment of

rDNA which can detect only type 1 rDNA was successful. The

sub-clone pCGR1N containing a 0.4 kb PstI-KpnI from the NTS

region was strain specific (compare Figures 3.1 and 3.8).

Therefore, type 1 strains can be defined as having a single

homogeneous form of rDNA as detected by a 8.4 kb Pst-1

fragment hybridizing to pMF2 and pCGR1N.

Morphologically variable type 2 strains (see Chapter 2)

were also diverse at DNA levels having different forms of

rDNA (Figures 3.2). The PstI-SphI-and SstI digested total

DNA blots, show RFLPs for rDNA within type 2 strains

(Figures 3.1, 3.2, 3.6, and 3.7). Only the HindIII-digested

blot shows a similar pattern of rDNA bands for type 2

strains (Figure 3.5). These results suggest that rDNA among

type two strains is heterogeneous. In addition, the specific









75

detection of only type 1 rDNA by pCGR1N suggest that

although some type 2 strains contain a rDNA form similar in

size to type 1 rDNA, the sequences may be different at least

at the NTS region. Several subclones from the NTS region of

type 2 rDNA hybridized to both type 1 and type 2 rDNA.

Therefore, a type 2 strain specific rDNA marker was not

found. Restriction site polymorphisms and length

heterogeneity in the rDNA unit repeat have been commonly

observed in many fungi. These polymorphisms were found

within the NTS region (Verbeet et al. 1983; Cassidy et al.

1984; Russell et al. 1984; Van Heerikhuizen et al 1985;

Chambers et al. 1986; Lachance 1989; Rogers et al. 1989),

ITS region (Chambers et al. 1986; Buckner et al. 1988) or

within coding regions (Chambers et al. 1986; Martin 1990).

In the fruit fly Drosophila melanogastor the presence of

introns has given rise to polymorphic forms of rDNA (Glover

and Hogness 1977). In fungi evidence for the presence of

introns in the rDNA coding regions is inconclusive (Buckner

et al. 1988).

Type 1 and type 2 C. gloeosporioides strains,

distinguished by rDNA polymorphisms were different both in

cutinase isozymes and molecular organization of relevant DNA

sequences. The slightly acidic pH optimum (pH 6.0) of

cutinolytic activity (Table 3.2) of these strains is a

reflection of their pathogenicity to aerial plant parts

(flowers or leaves) as observed for other aerial plant









76

pathogens such as V. inaequalis (KBller and Parker 1989),

Botrytis cinera (Salinas et al. 1986) and Cochliobolus

heterostrophus (Trail and K6ller 1990). This pH preference

is also congruent with the hypothesis that pathogens with

this type of cutinase are specialized for infecting aerial

plant surfaces rather than stem bases and roots (Trail and

Kal1er 1990).

All cutinases are serine esterases, and therefore they

can be detected by 3H-DFP which phosphorylates and inhibits

specific serine esterases (Kller and Kolattukudy 1982;

Kolattukudy 1985; K6ller and Parker 1989). The two 3H-DFP

labelled bands (Figure 3.9) which are within the range of

molecular weights of fungal cutinases (Kolattukudy 1980;

Tanabe et al. 1988; Trail and K11er 1990) were not

previously seen in Colletotrichum species including papaya

isolate of C. gloeosporioides (Dickman et al. 1982;

Kolattukudy 1985). Although molecular weight and culture

conditions prior to electrophoresis suggest that cutinase

enzymes are present, DFP binding is serine esterase but not

cutinase specific (K6ller and Kolattukudy 1982; Kolattukudy

1985; K1ller 1991). These bands may not represent two

different primary gene products but may be the result of

post-translational modification (K6ller 1991; Soliday et al.

1984). Some cutinase enzymes may undergo a proteolytic nick

and appear as two fragments after reduction of a disulfide

bridge and electrophoresis under denaturing conditions









77

(KSller 1991; Lin and Kolattukudy 1980; Purdy and

Kolattukudy 1975b; Soliday and Kolattukudy 1976). Conclusive

demonstration that these two bands are actually two

cutinases awaits further experiments such as purification

and characterization of the catalytic activity. Other fungal

pathogens, N. haematococca (Purdy and Kolattukudy 1975a,

1975b), A. brassicola, and R.solani (Trail and K11er 1990)

are known to produce at least two distinct cutinases as

detected by distinct isozyme bands and enzyme catalysis.

For type 2 strains, only one DNA restriction fragment

hybridizes to the cutinase gene probe but two cutinase

isozymes may be present. One possibility is that these may

be two forms of cutinases encoded by distinctly different

genes in the same organism. Only one form of cutinase has

been described biochemically from the papaya strain of C.

gloeosporioides (Dickman et al. 1982). The poor

hybridization of type 1 strains to the cutinase clone

despite the fact that they have abundant cutinase activity

suggests considerable evolutionary diversification of

cutinase gene sequences. Since the cDNA clone for cutinase

from C. capsici hybridizes readily with the total DNA from

C. graminicola and the C. gloeosporioides strain from

papaya, these species may be more related to type 2 than

type 1.


Another line of evidence for distinct genetic forms of









78

C. gloeosporioides from citrus comes from PCR amplification

of a cutinase gene sequence. The published DNA sequences for

cutinase genes from C. gloeosporioides and C. capsici

(Ettinger et al. 1987) show both conserved and diversified

regions of the gene. Two regions of identical sequence flank

a short stretch of DNA which includes the 52 bp intron of

the C. gloeosporioides cutinase and the 57 bp intron of the

C. capsici cutinase. The primer sequences are conserved in

cutinase genes of C. gloeosporioides, and C. capsici and

respectively should amplify a 220 bp or 222 bp fragment when

used as primers for PCR (Ettinger et al. 1987). Indeed, one

or two amplified fragments of about 220-260 bp were obtained

from the DNA of the four strains tested. When used as a

probe for Southern hybridization these fragments were

anticipated to hybridize to conserved elements of the

cutinase gene and divergent sequences expected within the

intron. However, these sequences hybridize to multiple

restriction fragments producing DNA finger prints that

correspond to RFLP types.

Hybridization to these probes must not be specific for

cutinase sequences. The exact nucleotide sequences of the

two amplified fragments were not determined. Therefore, it

is necessary to determine the nucleotide sequence of the 220

and 260 bp amplified fragments before reaching any

conclusions. However, a computer search of the GenBank

database (Release 70.0, December 15, 1991) indicated no









79

significant sequence similarity between the 220 bp targeted

sequence of the cutinase gene from C. gloeosporioides

(Ettinger et al 1987) and genes other than for C. capsici

and C. gloeosporioides cutinase. Hybridization may be to

sequences common to other introns such as putative splice

junction sequences identified previously (Ettinger et al.

1987). Hybridization to the repetitive sequences may be

detected using the 220 bp probe and not the entire cutinase

clone because the intron sequence represents <1% of the

clone but 25% of the PCR fragment.

Cutinase is not the only extracellular enzyme produced

by fungi in the process of plant infection. There are many

others such as cellulolytic and pectinolytic enzymes

produced by fungi in culture and in diseased leaves (Oke

1989; Prusky et al. 1989). The type 1 and type 2 strains

also produce different isozymes of pectinesterase that

correlate to type 1 and type 2 (personal communication

Gantotti and Davis, Homestead, FL).

DNA polymorphisms detected by hybridization to 3 of 4

"housekeeping" genes from N. crassa also correspond directly

to type 1 and type 2 strains. To detect DNA polymorphisms

correlated to type 1 and type 2 strains by Southern

hybridization using these molecular markers it was not

necessary to search for specific restriction enzymes or to

test multiple loci. The only enzyme used to digest total

DNA, HindIII was sufficient to provide RFLPs capable of









80

separating type 1 from type 2 strains. However, exceptions

to the strict correlation between polymorphism and strain

type was seen for hybridization to pNC2. In this case

variation was seen among type 1 strains. On the whole all

the molecular markers suggest that the two types are

different at the molecular level indicating they are indeed

genetically distinct populations of C. gloeosporioides.















CHAPTER 4
VARIABILITY OF MOLECULAR KARYOTYPES AND CHROMOSOMAL DNAS IN
COLLETOTRICHUM GLOEOSPORIOIDES


Introduction


Pulsed Field Gel Electrophoresis


Macromolecules such as nucleic acids and proteins can

be separated on the basis of size, charge or conformation by

gel-electrophoresis. Schwartz et al. (1982) made use of the

relaxation properties (Klotz and Zimm 1972) of large DNA

molecules for their separation in agarose gels by using two

alternating electric fields known as pulsed field gel

electrophoresis (PFGE). A major advance of the pulsed field

gel electrophoresis was achieved by Chu et al. (1986). They

applied the principles of electrostatics to calculate the

voltages needed to generate homogeneous electric fields

using multiple electrodes arranged around a closed contour.

In this system, contour clamped homogeneous electric field

(CHEF) gel electrophoresis, twenty four electrodes were

arranged in a hexagonal contour which offers reorientation

angles of 60 or 120.











Molecular Karvotypes of Fungi


Fungal chromosomes are too small to be observed readily

by conventional cytological methods using light microscopy.

However, electrophoretic karyotyping and molecular analysis

of chromosome-size DNA have become the new methods for

studying genomic structure of various organisms including

filamentous fungi. Many studies have conclusively

demonstrated that DNAs resolved by PFGE corresponds to

chromosomes (Carle and Olson 1985; Orbach et al. 1988; Brody

and Carbon 1989; Kayser and Wostemeyer 1991). However, the

number of bands need not be equal to the number of

chromosomes (Horton and Raper 1991). Electrophoretic

analysis of chromosomes provide a very convenient and rapid

way of assigning genes to chromosomes and for monitoring

entire genomes for any chromosomal rearrangements.

Pulsed field gel electrophoresis has been used to

separate chromosome-size DNA and analyze molecular

karyotypes of many fungi such as S. cerevisiae (Schwartz and

Cantor 1984; Chu et al. 1986), Candida albicans (Snell and

Wilkins 1986), Schizosaccharomyces pombe (Smith et al 1987;

Vollrath and Davis 1987), Neurospora crassa (Orbach et al.

1988), Ustilago maydis (Kinscherf and Leong 1988), Candida

stellatoidea (Kwon-Chung 1988, 1989: Wickes et al. 1991),

Aspergillus nidulans (Brody and Carbon 1989), C.

gloeosporioides (Masel et al. 1990), Ustilago hordei,

Tilletia caries, T. controversy (McCluskey et al. 1990),









83

Schizophyllum commune (Horton and Raper 1991), Absidia

glauca (Kayser and Wostemeyer 1991), Septoria tritici

(McDonald and Martinez 1991), Nectria haematococca (Miao et

al. 1991), Acremonium species (Smith et al. 1991; Walz and

Kuck 1991), Leptosphaeria maculans (Taylor et al. 1991), and

Fusarium oxysporum (Momol and Kistler 1992).

Pulsed field gel electrophoresis of chromosomal DNA

combined with Southern analysis using linkage group-specific

probes were key methods in defining molecular karyotypes of

N. crassa (Orbach et al. 1988) and Aspergillus nidulans

(Brody and Carbon 1989). Molecular karyotyping of N. crassa

by Orbach et al. (1988) confirmed the seven linkage groups

previously defined by genetic analysis (Perkins et al.

1982). The genome size of A. nidulans was estimated by Brody

and Carbon (1989) to be approximately 31 Mb with six

chromosome-sized DNA bands. Kayser and Wostemeyer (1991)

reported differences in electrophoretic karyotypes for

mating types of the Zygomucete Absidia glauca.

Molecular karyotypes of many plant pathogenic fungi

examined to date have been variable. Kinscherf and Leong

(1988) analyzed the molecular karyotype of U. maydis and

demonstrated that considerable chromosomal length

heterogeneity exists in this fungus. DNA hybridization

analysis suggested that stable large scale inter-chromosomal

exchange has given rise to novel chromosomes in one of the

strains. Taylor et al. (1991) using TAFE demonstrated that









84

the karyotypes of highly virulent and weakly virulant

strains of Leptosphaeria maculans (black leg of crucifers)

were polymorphic in both chromosome number and size. Highly

variable karyotypes for N. haematococca with unique

karyotypes for each strain were reported by Miao et al.

(1991). Deletions of large amounts of DNA from chromosomes

have given rise to karyotype variation as well as a

decreased frequency of the pisatin demethylase gene in N.

haematococca. Masel et al. (1990) suggested that chromosomal

rearrangements may play a role in generating variability of

karyotype of C. gloeosporioides. Distinct electrophoretic

karyotypes were reported for strains from two types of C.

gloeosporioides causing different anthracnose diseases in

Stylosanthus species in Australia. The strains showed

extensive chromosomal polymorphisms for both length and

number in the mini-chromosomes (molecules less than 2

million base pairs (Mb) in length) within each type.

The present study was undertaken to investigate the

variation of molecular karyotypes and chromosomal DNAs in

two types of C. gloeosporioides (see Chapter 2 and 3)

causing post bloom fruit drop of Tahiti lime and Sweet

orange.









85

Materials and Methods


Strains of Colletotrichum qloeosporioides


Strains used were obtained from several different areas

of Florida, Mexico, and from the Commonwealth Institute of

Mycology, England. They were isolated from diseased lime or

orange tissues. Details of host, place of collection and

date are tabulated in the appendix A.


Preparation of Protoplast Plugs


The strains of C. gloeosporioides were grown for 7 days

in 50 ml of 20% (w/v) V-8 juice (Campbell Co., Camden, NJ)

in at 250 rpm in Erlinmayer flasks on a Lab-Line orbit

shaker (Lab-Line Instruments Inc., Melrose Park, IL) at

ambient temperature (21-23 oC), and conidia were collected

by centrifugation at 7000 x g for 5 min. About 109 spores

per ml were resuspended in 50 ml potato dextrose broth (PDB)

and incubated at room temperature (23 to 25 OC) for 16-24 h

at 200 rpm. When over 90% of spores were germinated, the

germlings were pelletted by centrifuging at 7000 x g for 5

min. Protoplasts were made by adding germlings to a 10 ml

solution containing NovoZym 234 (Novo Industries, Bagsvaerd,

Denmark) a complex mixture of wall-degrading enzymes. The

NovoZym solution was prepared by mixing 1.5 ml of 1 M

sorbitol, 50 mM sodium citrate containing 0.2 g of NovoZym

234 with 8.5 ml of 1.4 M MgSo4, and 50 mM Sodium Citrate pH










86

5.8. Germlings were incubated in this solution with gentle

rocking on a Bellco rocker (Bellco Biotechnology, Vineland

NJ) at 4 rpm for 3 to 6 h at ambient temperature for 3 to 6

hours until most cells were protoplasts. The protoplasts

were filtered through 4 layers of cheese cloth in order to

remove cell debris and undigested germlings. The filtrate

was centrifuged at 3000 rpm for 25 min at room temperature.

Protoplasts were removed from the top and washed three times

with 1 M sorbitol-50 mM EDTA pH 8.0. Protoplast inserts for

PFGE were made as described by method 1 of Orbach et al.

(1988).


Electrophoresis and Southern Analysis


A commercially available apparatus (BioRad CHEF DRII,

Richmond, CA) using different pulse time combinations was

employed in order to separate chromosome-sized DNAs.

Electrophoresis was done with 0.6% FastLane agarose (FMC

BioProducts, Rockland, ME) gels in 0.25X Tris Borate EDTA

(TBE) buffer (Sambrook et al. 1989) at 4 OC with rapid

circulation of the buffer. The gels were run at 40 volts for

6-10 days. Pulse times were "ramped" for various times

ranging from 10 to 180 min. For the separation of smaller

chromosome sized DNA 1% SeaKem Agarose (FMC BioProducts,

Rockland, ME) in 0.5X TBE buffer was used. These gels were

run at 200 V for 24 h with pulse times of 30-60 s or 50-90s.









87

Southern hybridization (Appendix C) experiments with

32P labelled ribosomal DNA (pMF2), B-tubulin gene (pSV50)

and cutinase gene (see chapter 3) were carried out to assign

these sequences to chromosome-size DNAs separated in CHEF-

gels.

Results


Chromosome-size DNAs (henceforth called chromosomes)

from type 1 and type 2 strains of C. gloeosporioides were

separated by PFGE using Saccharomyces cerevisiae and

Schizosaccharomyces pombe chromosome size DNA as standards

(BioRad Laboratories, Richmond, CA). The sizes in Table 4.1

represent the average size calculated for each chromosome

size-DNA band from independent CHEF-gels. Calculated sizes

for individual chromosome size DNA bands and relevant

figures are compiled in the Appendix D. Type 1 strains have

chromosomes distinguishable from type 2 strains (Table 4.1).

The chromosomes of C. gloeosporioides isolated from

Stylosanthes have been classified by Masel et al. (1990)

into larger, similar-sized chromosomes (>2 Mb) and smaller

variable-sized elements called "minichromosomes" (<2 Mb). A

similar arrangement was noted for strains isolated from

Tahiti lime and Sweet orange. Type 1 strains possess 5

chromosomes (Figures 4.1 and 4.2) and an additional 1 or 2

minichromosomes (Figure 4.3 and 4.5). Type 2 strains possess

3 chromosomes (Figures 4.1 and 4.2) in addition to 2 to 4










88

Table 4.1 Estimated megabase sizes for chromosome-size DNA
from Colletotrichum gloeosporioides type 1 and
type 2 strains


Strain Estimated size" (Mb)
I II III IV V VI VII


Type-1 strains

H-1

H-3

H-9

H-25B

H-36

LP-1

Maran

IMB-3

OCO

Type-2 strains

H-4

H-12

H-46

H-48

180269

226802


7.6

7.6

7.6

7.6

7.6

7.6

7.6

7.6

7.6


7.8

7.8

7.8

7.8

7.8

7.8


7.0

7.0

7.0

7.0

7.0

7.0

7.0

7.0

7.0


4.7

4.7

4.7

4.7

4.7

4.7


4.7

4.7

4.7

4.7

4.7

4.7

4.7

4.7

4.7


3.7

3.7

3.7

3.7

3.7

3.7


3.7

3.7

3.7

3.7

3.7

3.7

3.7

3.7

3.7


0.42

0.46

0.52

0.46

0.43

0.44


3.3

3.3

3.3

3.3

3.3

3.3

2.8

3.3

2.8


0.38

0.38

0.47

0.43

0.41

0.42


1.1

1.1

1.1

1.1

0.77

1.6



1.1


0.42

0.40

0.39

0.39


0.63

0.63

0.63

0.63

0.63

0.63

0.63

0.63

0.65


0.27





0.37


Schizosaccharomyces pombe and Saccharomyces cerevisiae size
standard were used for the calculation of Mb sizes. Megabase
sizes greater than 5.6 were estimated by extending the
calibration curve and therefore may be considered
approximate sizes.
- = not detected in any of the gels.










89

minichromosomes (Figures 4.4 and 4.5) depending on the

strain. Within each type, strains show variations in

chromosome number and size. However type 2 strains show more

total variation in chromosome and minichromosome size

(Figures 4.4 and 4.5).

A Southern blot separating larger chromosome-size DNAs

was hybridized with a 32P labelled ribosomal DNA probe. The

rDNA is associated with the 4.7 Mb chromosome in type 1

strains and with the 7.8 Mb chromosome in type 2 strains

(Figure 4.6). The cutinase gene can be assigned to the 4.7

Mb chromosome-size DNA only in type-2 strains (Figure 4.7).

Homologous regions were not detected in chromosomes of type

1 strains. The B-tubulin gene hybridizes to both the 7.0 and

7.6 Mb chromosome doublet in type-1 strains and the 7.8 Mb

chromosome in type-2 strains (results are not shown due to

weak signals on X-ray film).
























0% N
%00
1O 00 N 1 1 13
MbUO4 0 1 0" .1
Mb'W T- CMTT


5-7
4.6
3-5
2"2










Figure 4.1 Chromosome-sized DNAs from C. gloeosporioides
compared to those for yeast (S. cerevisiae) and
fission yeast (S. pombe). DNAs were separated on
agarose gels using CHEF electrophoresis. Gels
were stained with ethidium bromide (0.5 gg/ml)
and fluorescence photographed with UV
transillumination. DNAs from the indicated
strains were run in 0.25X TBE, 0.6% agarose for
168 h at 40 V. Pulse switching times were ramped
from 40-70 min. Numbers at the left indicate size
standards in megabases (Mb).
























Ica 0 %D
cm Ln o % 0

Mba 04 1 1 0 $

5.7
4 6~ltP


Figure 4.2


Chromosome-sized DNAs from C. gloeosporioides
compared to those for fission yeast (S. pombe).
DNAs were separated on agarose gels using CHEF
electrophoresis. Gels were stained with ethidium
bromide (0.5 Ag/ml) and fluorescence photographed
with UV transillumination. DNAs from the
indicated strains were run in 0.25X TBE, 0.6%
agarose for 168 h at 40 V. Pulse switching times
were ramped from 20-60 min. Numbers at the left
indicate size standards in megabases (Mb).

















n3 0 e
I -I L 0 Uo
Sn oZ ,- I ( 4 0
li : X


Figure 4.3


Chromosome-sized DNAs from C. gloeosporioides
compared to those for yeast (S. cerevisiae). DNAs
were separated on agarose gels using CHEF
electrophoresis. Gels were stained with ethidium
bromide (0.5 jg/ml) and fluorescence photographed
with UV transillumination. DNAs from the
indicated strains were run in 0.5X TBE, 1.0%
agarose for 24 h at 200 V. Pulse switching times
were ramped from 60-90 s. Numbers at the left
indicate size standards in kilobases (kb).


kb 0






1125





630



245




Full Text
MOLECULAR ANALYSIS OF GENETIC DIVERSITY AND VARIABILITY IN
COLLETOTRICHUM GLOEOSPORIOIDES
BY
HEMACHANDRA D. LIYANAGE
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
1992
UNIVERSITY 0? FLORIDA UIRARIES

Dedicated
to
Mother

ACKNOWLEDGEMENTS
I sincerely thank Dr. Corby Kistler for offering me
this opportunity to work in his lab, for all the advice,
guidance, constructive criticism and help throughout the
research study and my career. I am very thankful to Dr. R.
T. McMillan, Jr., for providing financial arrangements and
for being helpful in many ways. I gratefully acknowledge
the willing assistance and advice given by Dr. Frank Martin
and Dr. Ron Sonoda. I thank my committee members, Dr. Daryl
Pring, Dr. James Kimbrough, and Dr. Curtis Hannah for their
assistance and guidance throughout this study.
A special word of thanks to my loving wife, Thamara,
who spent her time and effort helping me throughout the
period of study and in all my difficult times.
iii

TABLE OF CONTENTS
Page
ACKNOWLEDGEMENTS iii
ABSTRACT vii
CHAPTERS
1 INTRODUCTION 1
2 OBSERVED VARIABILITY IN COLLETOTRICHUM
GLOEOSPORIOIDES CAUSING POST BLOOM FRUIT
DROP IN CITRUS 7
Introduction 7
Materials and Methods 9
Strains of Colletotrichum gloeosporioides. 9
Pathogenicity 10
Benomyl Tolerance 10
Results 11
Colletotrichum gloeosporioides Strains
from Citrus are Morphologically
Variable 11
Colletotrichum gloeosporioides Strains
have Different Nuclear Numbers
in their Spores 13
Both Type 1 and Type 2 Strains are
Pathogenic to Tahiti Lime Flowers 15
Type 1 and Type 2 Strains Differ in their
Tolerance to Benomyl 15
Discussion 16
3 DNA POLYMORPHISMS FOUND AT MANY GENETIC LOCI
EXAMINED IN COLLETOTRICHUM GLOEOSPORIOIDES... 25
Introduction 25
Ribosomal DNA in Fungi 25
Ribosomal DNA is Polymorphic in Many
Fungi 27
Fungal Cutinase Genes and Cutinase
Isozymes 31
Restriction Fragment Length Polymorphisms
(RFLP) in Fungi 3 6
iv

Page
Materials and Methods 39
Strains of Colletotrichum gloeosporioides. 39
DNA Extraction 4 0
DNA Cloning and Restriction Enzyme Mapping 41
Enzyme Assays and Electrophoresis of
Cutinase 42
Probes Containing Cutinase Gene Sequences. 43
Detection of Restriction Fragment Length
Polymorphisms 44
Results 45
Ribosomal DNA is Polymorphic in
Colletotrichum gloeosporioides 4 5
Ribosomal RNA Genes 47
Diverse Cutinases and Cutinase Genes are
Found in Type 1 and Type 2 Strains of
Colletotrichum gloeosporioides 4 9
Subgroups of Colletotrichum gloeosporioides
have Distinct RFLP Patterns Detected by
Many Genetic Markers 53
Discussion 73
4 VARIABILITY OF MOLECULAR KARYOTYPES AND
CHROMOSOMOL DNAS IN COLLETOTRICHUM
GLOEOSPORIOIDES 81
Introduction 81
Pulsed Field Gel Electrophoresis 81
Molecular Karyotypes of Fungi 82
Materials and Methods 85
Strains of Colletotrichum gloeosporioides. 85
Preparation of Protoplast Plugs 85
Electrophoresis and Southern Analysis 8 6
Results 87
Discussion 97
5 GENERAL DISCUSSION AND CONCLUSIONS 99
APPENDICES
A STRAINS OF COLLETOTRICHUM GLOEOSPORIOIDES 102
B ANALYSIS OF VARIANCE TABLES 104
C PROCEDURES FOR DNA LABELLING AND SOUTHERN
HYBRIDIZATION 110
D CALCULATED MB SIZES FOR CHROMOSOMAL DNAS IN
COLLETOTRICHUM GLOEOSPORIOIDES 112
v

page
LITERATURE CITED 131
BIOGRAPHICAL SKETCH 143
vi

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
MOLECULAR ANALYSIS OF GENETIC DIVERSITY AND VARIABILITY IN
COLLETOTRICHUM GLOEOSPORIOIDES
By
Hemachandra D. Liyanage
August 1992
Chairman: Dr. R. T. McMillan, Jr.
Cochairman: Dr. Corby H. Kistler
Major Department: Plant Pathology
Results from this study suggest two distinct genetic
subpopulations of Colletotrichum gloeosporioides from citrus
based on DNA variation, cultural morphology, and growth.
Type 1 strains are slow growing, morphologically stable,
benomyl tolerant, and contain a single form of ribosomal DNA
(rDNA) as detected by common Hindlll, PstI, SphI and SstI
fragments hybridizing to cloned Neurospora crassa rDNA. Type
2 strains are faster growing, morphologically less stable,
benomyl sensitive, and have rDNA distinct from type 1
strains. The rDNA from type 1 and type 2 strains were cloned
and mapped for 10 restriction enzyme sites and genes coding
for large subunit, small subunit and 5.8S rRNA. A subclone
constructed from the non-transcribed spacer region of type 1
rDNA clone hybridizes only to rDNA from type 1 strains. DNA
polymorphisms detected by heterologous hybridization with
vii

cloned N. crassa genes for glutamate dehydrogenase,
anthranilate synthetase, histidinol dehydrogenase, and /?-
tubulin corresponded to type 1 or type 2 strains. All
strains liberate free fatty acid from [3H]-labelled cutin
and hydrolyze cutin model substrates. Serine esterases from
extracellular fluids of cutin-grown C. gloeosporioides
strains were detected by labelling proteins separated by
sodium dodesyl sulfate polyacrylamide gel electrophoresis
with 3H-diisofluorophosphate. The two major esterases from
type 1 strains have molecular weights of 26 and 20
kilodaltons (kd) whereas the type 2 esterases were 24 and 22
kd. A DNA probe containing the cloned cutinase gene from C.
gloeosporioides hybridized strongly to DNA from type 2
strains but poorly to type 1 strains. Distinct cutinase
genes may be present in the two types of C. gloeosporioides
strains from citrus. Chromosome-sized DNAs separated by
pulsed-field gel electrophoresis corresponded to type 1 or
type 2 strains. Type 1 strains had five large chromosomal
DNAs 7.6, 7.0, 4.7, 3.7, and 3.3 (or 2.8) million base pairs
(Mb) in size and one or two smaller chromosomes (1.6 to 0.63
Mb). Type 2 strains had three large chromosomal DNAs (7.8,
4.7, and 3.7 Mb) and two to four smaller chromosomal DNAs
(0.52-0.28 Mb).
viii

CHAPTER 1
INTRODUCTION
Historically, the study of plant diseases dates back to
Theophrastus (371-287 B.C.) who first described disease
conditions of plants, mostly cereal rusts, in Historia
plantarían and De causis plantarum (Ainsworth 1981) .
Afterwards, studies of causal agents of plant diseases and
control measures played a major role in human survival. The
period from the mid-eighteenth to the mid-nineteenth century
was marked by the accumulation of experimental evidence for
the pathogenicity of fungi to plants. Almost all groups of
fungi include some plant pathogenic species but the greatest
number of plant pathogens is to be found among the imperfect
fungi (Ainsworth 1971) .
The most important technique for the identification of
plant pathogenic fungi has always been macroscopic and
microscopic morphological examination. Morphology always
took precedence over other considerations in describing
genera and species of plant pathogenic fungi (Ainsworth
1981). The genus Colletotrichum Corda was established in
1831 and was characterized by having setose acervuli
containing hyaline, curved fusiform conidia (Baxter et al.
1985). However, there was always confusion in describing
1

2
fungi to this genus due to similar morphological characters
of Vermicularia Tode and Gloeosporium Desm. & Mont. (Dickson
1925; Duke 1928; Arx 1957; Baxter et al. 1985). Duke (1928)
suggested that type species of Vermicularia and
Colletotrichum represented the same fungus. Species in the
genus Gloeosporium probably represent the same fungi as in
the genus Colletotrichum because the Gloeosporium species,
which supposedly lack setae, were found to produce them on
certain substrates (Baker et al. 1940). Arx (1957) accepted
Colletotrichum and Vermicularia as separate genera while
rejecting the more heterogenous genus Gloeosporium.
The species concept of Colletotrichum gloeosporioides
(Penz.) is still uncertain (Van Der Aa et al. 1990).
Colletotrichum gloeosporioides was first described in 1882
by Penzig as Vermicularia gloeosporioides, and in 1887 it
was renamed Colletotrichum gloeosporioides (Burger 1921).
The presence of this fungus in the United States was first
observed in 1886 in Florida and was reported by Underwood
(1891). Arx (1957) recognized eleven species in the genus
Colletotrichum, and the name C. gloeosporioides with nearly
600 synonyms was maintained to designate the variable
anamorph of Glomerella cingulata (Stonem.) Spauld. & Schr.
He recognized nine forms within the species C.
gloeosporioides. Arx (1970, 1987) introduced the concept of
host forms of C. gloeosporioides but did not accept these
forms as species or intraspecific taxa with certainty.

3
Sutton (1980) considered C. gloeosporioides a group species
showing excessively wide variation.
The conidia of C. gloeosporioides are straight, obtuse
at the apex, 9-24 x 3-4.5 pm and appressoria are 6-20 x 4-12
pm, clavate or irregular, sometimes becoming complex (Sutton
1980).
Colletotrichum gloeosporioides is a ubiquitous fungus
and often causes a variety of diseases commonly known as
anthracnose on fruits, leaves and stems of a wide range of
host species. The host range of this fungus is so wide that
nearly 200 susceptible host species were listed under C.
gloeosporioides in the Index of Plant Diseases in Florida
(Alfieri et al. 1984). Many tropical fruit crops are
attacked by this fungus in the field and in post-harvest
condition (Nolla 1926; Simmonds 1965; Brown 1975). Citrus is
one of the major fruit crops attacked by this fungus, and
the diseases of citrus caused by C. gloeosporioides have
been known since 1886 when it was first isolated from citrus
plants in Florida, U.S.A. (Underwood 1891). Rolfs (1904 and
1905) described a group of citrus diseases (wither tip, leaf
spot, lemon spot, canker, and anthracnose) caused by C.
gloeosporioides. A more recently described citrus disease,
post bloom fruit drop, PFD (Fagan 1979; Sonoda and Pelosi
1988; McMillan and Timmer 1989) is caused by the same
species. The name PFD was suggested by Fagan (1979) to
distinguish this disease condition of citrus characterized

4
by premature fruit drop or blossom blight from normal
physiological thinning of fruits. The symptoms first appear
as small, brown spots on flower buds or light pink water-
soaked spots on open petals. These spots may enlarge and
rapidly cover the petals within 24 h. Afterwards, the petals
become brown and desiccated. Eventually, young fruitlets
become discolored, and they abscise, leaving the calyxes
behind as persistent buttons (Figure 1). The disease is
economically important in regions where citrus is grown
(Denham and Waller 1981; Fagan 1984a). In Florida the
disease has been reported from all commercially grown citrus
(Sonoda and Pelosi 1988; McMillan and Timmer 1989).
The causal agent of PFD of citrus was identified as C.
gloeosporioides (Fagan 1979; Sonoda and Pelosi 1988;
McMillan and Timmer 1989). Colletotrichum gloeosporioides
isolated from citrus diseases were reported to be variable
in morphology and pathogenicity (Burger 1921) . Morphological
variability also has been observed in the strains of this
fungus causing PFD (Denham and Waller 1981; Sonoda and
Pelosi 1988). Three strains C. gloeosporioides varying in
morphology and pathogenicity were reported to be associated
with diseased plants by Fagan (1980). Because of the
inconsistency of the morphological characteristics, it is
uncertain whether the strains or forms of C. gloeosporioides
recognized by morphological criteria alone are really
different at genetic and molecular level. The morphological

5
changes could be caused by environmental effects, genetic
differences or both. Therefore, study of this group species
at the molecular level to understand the genetic and
molecular differences among the strains is important.
The present study was undertaken to investigate the
morphological variability and potential genetic variation of
C. gloeosporioides at the molecular level. The objectives of
this study are:
1. To examine the basis for morphological variability
of C. gloeosporioides causing PFD disease of citrus.
2. To investigate genetic variation of C.
gloeosporioides at the molecular level.
3. To examine variations in chromosome-size DNA and to
describe the molecular karyotypes.
Morphological and growth diversity arising from single
spore cultures of different C. gloeosporioides strains is
examined in Chapter 2. Molecular investigations using
genetic markers were carried out to study the variation
within citrus strains of C. gloeosporioides, and they are
reported in Chapter 3. Chapter 4 describes the chromosomal
variation of C. gloeosporioides strains and the molecular
karyotypes of the fungus. The results obtained in these
studies are reviewed comprehensively in Chapter 5, and a
concept of genetically distinct subpopulations of C.
gloeosporioides is proposed.

6
Figure 1 Symptoms of post bloom fruit drop disease on sweet
orange (Citrus sinensis var. Valencia) caused by
Colletotrichum gloeosporioides.

CHAPTER 2
OBSERVED VARIABILITY IN COLLETOTRICHUM GLOEOSPORIOIDES
CAUSING POST BLOOM FRUIT DROP IN CITRUS
Introduction
More than six hundred synonyms for the fungal species
Colletotrichum gloeosporioides have been published (Arx
1957). This reflects the considerable amount of diversity
and variability observed for the fungus by different
investigators throughout the world. Sutton (1980) could not
give a standard morphological description of C.
gloeosporioides. He considered the different forms of this
fungus to be within a group species.
The association of C. gloeosporioides with citrus dates
back to 1886-1891 (Underwood 1891). The fungus has been
found to cause various diseases on this crop for the past
century (see Chapter 1). The most thorough studies on the
morphological variation of C. gloeosporioides were done by
Burger (1921). The fungus he studied was the causal agent of
bloom drops and leaf spots in citrus. Cultural
characteristics such as mycelial color, growth and
sporulation enabled him to classify C. gloeosporioides
strains into five groups. However, some strains did not fit
into any of the morphological classes due to inconsistency
7

8
of mycelial and sporulation characteristics in continuous
culture.
Mycelial sectors distinguished by growth and color
differences within single spore cultures of strains are
another type of variability observed in C. gloeosporioides.
Burger (1921) observed black and white mycelial sectors in
single spore cultures of the fungus. When single spored,
these black and white sectors were able to maintain their
identity in continuous culture.
Burger (1921), after studying cultural characteristics,
spore dimensions, and sectoring, concluded that C.
gloeosporioides is constantly giving off new types under
natural conditions as well as in artificial cultures. He
further suggested that these variabilities of C.
gloeosporioides may have arisen from environmental effects
as well as from high frequency mutations.
Morphological and pathogenic variability in C.
gloeosporioides causing PFD of citrus has been reported by
Fagan (1979,1980) and Denham and Waller (1981). Three
different forms of C. gloeosporioides were recognized by
Fagan (1980). Two forms, cgm with gray to dark gray mycelium
and cgc with light gray mycelium, were isolated from
senescent leaves and were nonpathogenic to citrus flowers.
The pathogenic form, cgp, had off-white to pink mycelium and
was isolated from floral parts of citrus. Fagan (1980)
concluded that at least two strains of C. gloeosporioides

9
causing PFD occurred in Belize. These strains corresponded
to morphological groups of C. gloeosporioides described by
Burger (1921).
The objective of this study is to examine the
morphological and phenotypic diversity of C. gloeosporioides
causing PFD of Tahiti lime (Citrus aurantifolia Swingle) and
Sweet orange (Citrus sinensis Osbek).
Materials and Methods
Strains of Colletotrichum gloeosporioides
Strain number, host, place and year of isolation are
tabulated in Appendix A. Isolation of C. gloeosporioides
from host plants was carried out as follows. Host plant
tissues were surface sterilized in 1% sodium hypochlorite
(Clorox Co., Oakland, CA) for 30-60 s, rinsed 3 times with
sterilized water and plated on potato dextrose agar (PDA,
Difco laboratories, Detroit, MI) plates. Edges from growing
mycelia were isolated and maintained in the laboratory as
strains. Strains were grown in 20% (w/v) V-8 juice (Campbell
Soup Co., Camden, NJ) for 7 d at 250 rpm on a Lab-Line orbit
shaker (Lab-Line Instruments Inc., Melrose Park, IL). Spores
were collected by centrifugation at 7000 x g for 5 min and
washed 2 times with sterilized water before storing in 50%
glycerol (in water) at -80°C. To obtain single spore
cultures, spores were spread on PDA plates; 14-16 h later,
germinating spores were isolated under a dissecting

10
microscope (25x10 magnification) and plated on PDA plates.
Morphology of colony growth, mycelial color and sectoring
were examined in PDA culture and still liquid culture,
potato dextrose broth (PDB Difco laboratories, Detroit, MI).
To examine the nuclear number, spores were stained with
1% aniline blue (Sigma Chemical Co., St. Louis, MO) in 50%
glycerin in water (Tu and Kimbrough 1973). To stain nuclei,
a drop of spores in water was placed on a microscopic slide,
and a drop of stain was added. The slide was then heated
over a flame for 5-10 seconds. Approximately 1000 spores
were examined for each strain.
Pathogenicity
All the strains were tested for their ability to infect
flowers of Tahiti lime under natural conditions in the field
as well as in the laboratory. Strains were grown in 20% V-8
juice for 7 days and inoculum containing 107 spores ml'1
water were prepared. Tahiti Lime flowers were sprayed using
a hand sprayer to wetness with inoculum or water, and
symptom development was observed for 3 days. Each treatment
contained 10-15 flowers. The control was sprayed with water.
Benomvl Tolerance
The growth of C. gloeosporioides strains was examined
in PDA medium containing 0, 2 and 10 pg benomyl (methyl-
(butyl carbamoyl)-2-benzimidazolecarbamate, Sigma Chemical

11
Co., St. Louis, MO) ml'1. Radial growth of the mycelial
colony was measured every 24 h for a 10 day period. Growth
rates in mm h'1 were estimated by the slopes obtained with
linear regression analysis of the growth curve. A comparison
of slopes was made using analysis of variance (Appendix B).
Each treatment was replicated 5 times, and the experiment
was repeated once with 2 replicates.
Results
Colletotrichum crloeosporioides Strains from Citrus are
Morphologically Variable
The C. gloeosporioides strains examined can be grouped
into two major categories based on morphology and growth
characteristics. Type 1 strains (H-l, H-3, H-9, H-21, H-22,
H-25B, H-36, IMB-3, LP-1, Maran, 0C0, and TUR-1) produce
morphologically stable and relatively slow-growing mycelial
colonies in PDB. The colonies are orange-colored and have
appressed mycelia with abundant sporodochia (Figure 2.1).
Type 2 strains (H-4, H-ll, H-12, H-23, H-24, H-46, H-47, H-
48, 180269 and 226802) grow faster and produce mostly gray,
fluffy mycelial colonies (Figure 2.1). The type 1 strains
grow at a significantly slower rate from 0.008 to 0.10 mm
h'1; type 2 strains grow significantly faster from 0.12 to
0.15 mm/ h’1 as calculated by slopes of linear regression
data (Table 2.1). The strain types also differ in culture

12
Table 2.1 Effect of benomyl concentration on the estimated
radial growth rates in mm h‘1 of Colletotrichum
gloeosporioides type 1 and type 2 strains.
Strain
Benomyl
0
concentration /¿g/ml
2
10
Type 1
H-l
0.10
0.033
0.029
H-3
0.10
0.033
0.033
H-9
0.10
0.045
0.041
H-25B
0.041
0.041
0.041
H-3 6
0.095
0.037
0.033
IMB-3
0.008
0.008
0.004
LP-1
0.10
0.050
0.041
Maran
0.041
0.029
0.033
OCO
0.095
0.037
0.033
Type 2
H-4
0.141
0.00
0.00
H-ll
0.125
0.00
0.00
H-12
0.121
0.00
0.00
H-46
0.133
0.00
0.00
H-47
0.133
0.00
0.00
H-48
0.145
0.00
0.00
180269
0.133
0.00
0.00
226802
0.150
0.00
0.00

13
stability as determined by their ability to produce sectors
of different color, morphology and growth habit. To
quantitate these levels of instability, 100 conidia were
isolated from three type 1 strains and two type 2 strains
and tested for morphological stability. One hundred single
spore cultures from strains H-l, H-3, and H-25B (type 1)
grown in PDB were found to produce identical colonies. One
hundred single spore cultures from strain H-12 and H-48
(type 2) produced 100% sectoring colonies. These colonies
varied in colony color from dark gray to gray, white, and
orange with different growth rates (Figures 2.2, 2.3, and
2.4). Sporodochia production was scattered or inhibited but
could be stimulated by mycelial injury (Figure 2.5). One
hundred injury-induced spores from an H-48 gray mycelial
sector produced 50 sectoring colonies, 24 dark gray with no
sporodochia and 26 orange colonies with scattered
sporodochia production.
Colletotrichum aloeosporioides Strains have Different
Nuclear Numbers in their Spores
The nuclear number observed by aniline blue staining
varied from 1 to 3 per single spore (Table 2.2). All spores
examined from all isolates were single celled. All the
isolates examined contained spores with more than one
nucleus as identified by dark stained objects distinguished
from the lightly stained cytoplasm under the high

14
Table 2.2 Percentage of spores carrying different numbers of
nuclei in Colletotrichum gloeosporioides strains.
Strain
Percentage of spores*
Number of nuclei
1 2
3
H-l
92.6
6.8
0.6
H-3
97.8
2.2
<0.1
H-4
94.3
5.4
0.3
H-9
96.4
3.4
0.2
H-12
98.8
1.2
<0.1
H-25B
96.9
2.9
0.2
H-4 6
93.4
6.0
0.6
H-4 8
92.8
6.6
0.6
180269
92.9
6.6
0.5
226802
94.7
5.2
0.1
LP-1
99.6
0.4
<0.1
Maran
98.6
1.4
<0.1
*=Calculated from 1000 spores

15
magnification (46x10) of a light microscope. Spores
containing a single nucleus varied from 92.6 to 99.6% in the
15 isolates studied. The maximum number of nuclei observed
within a single spore was 3, and the percentage of spores
containing three nuclei varied from <0.1-0.6%. The
percentage of spores containing two nuclei varied from 0.4-
6.8%.
Both Type 1 and Type 2 Strains are Pathogenic to Tahiti Lime
Flowers
Brown lesions developed in flowers individually
inoculated with all strains of the pathogen 24 h after
spraying. The petals were blighted completely at 36 h and
had dropped at 48 h. Flowers sprayed with water alone were
not blighted after 72 h. The fungal strains reisolated from
infected tissues were found to be morphologically like the
original strains. The relative virulence of strains was not
measured in this study.
Type 1 and Type 2 Strains Differ in their Tolerance to
Benomvl
All type 2 strains were completely inhibited by 2 or 10
Hq ml'1 benomyl in PDA, but type 1 strains were more
tolerant. Average growth rates for individual type 1 and
type 2 strains are listed in Table 2.1. Analysis of variance
showed that benomyl concentration had a significant effect
on type 1 strains. There was a significant interaction

16
between strains and concentration indicating that growth
rate of each strain may respond differently to different
concentrations of benomyl (Appendix B).
Discussion
Grouping of Colletotrichum gloeosporioides strains
based on morphological and physiological observations was
first attempted by Burger (1921). However, morphologically
based groups of C. gloeosporioides strains have been
inconsistently described in this and subsequent studies
(Burger 1921; Arx 1957; Sutton 1980). Fagan (1980), Denham
and Waller (1981), and Sonoda and Pelosi (1988) reported
morphological variations associated with this fungus
isolated from Sweet orange cultivars. The type 1 strains in
this study show similarities in morphology, growth and
sporodochia production to strains designated cgp by Fagan's
(1980) description and correspond to the orange colored,
slow-growing colonies described by Sonoda and Pelosi (1988).
The more variable type 2 strains show similarities to the
cgm and cgc strains of Fagan and correspond to faster
growing colonies described by Sonoda and Pelosi.
Both type 1 and type 2 strains were isolated from sweet
orange (C. sinensis) as well as Tahiti lime (Appendix A).
Both types were pathogenic to Tahiti lime flowers as

17
Figure 2.1. Morphology of type 1 (left) and type 2 (right)
strains of Colletotrichum gloeosporioides grown
in potato dextrose broth.

18
Figure 2.2 Fluffy and appressed mycelial sectors
produced by a single spore culture of
Colletotrichum gloeosporioides type 2 strain in
potato dextrose agar.

19
Figure 2.3. Dark gray, light gray and orange mycelial
sectors produced by a single spore culture of a
Colletotrichum gloeosporioides type 2 strain in
potato dextrose broth.

20
Figure 2.4. Multi-colored mycelial sectors produced by a
single spore culture of a Colletotrichum
gloeosporioides type 2 strain in potato dextrose
broth.

21
Figure 2.5. Mycelial injury can induce type 2 (non-
sporodochia-forming) type 2 strains to form
sporodochia.

22
indicated by inoculation tests, confirming previous results
(Sonoda and Pelosi 1988). Variation in the ability of type 1
and type 2 strains to cause disease on Tahiti lime flowers
was not observed. However, only a single high inoculum
concentration was used for pathogenicity testing.
Differences in virulence perhaps could be found if a
dilution series of inoculum were used in assessing the
disease-causing potential of the strains. Sonoda and Pelosi
(1988) and Agostini et al (1992) suggested that slowly
growing, orange-colored strains (type 1 strains) were the
actual causal agent of PFD because only they could be
consistently isolated from diseased petals in the field
while the gray (type 2) strains were isolated primarily from
leaves, stems and fruit. This observation has been confirmed
by other workers (Agostini et al. 1992; Gantotti and Davis,
personal communication). Clearly, further pathogenicity
testing and field sampling are needed to confirm whether one
or both types of the pathogen are important in PFD
epidemics.
The genetic and molecular basis of the morphological
diversity caused by sectoring of C. gloeosporioides is yet
to be elucidated. One of the probable genetic explanations
for the observed morphological diversity and sectoring of
type 2 strains may be heterokaryosis followed by
parasexuality or nuclear sorting out. Heterokaryosis and
parasexuality have been found to contribute to the variation

23
of C. gloeosporioides as reviewed by Baxter et al. (1985).
The strains studied carried only <7% spores with 2-3 nuclei.
Therefore, heterokaryosis or nuclear sorting out may not be
a cause of 100% sectoring observed in type 2 strains.
Multinuclear spores have been reported in C. gloeosporioides
as well as many other fungi (Panaccione et al. 1989; Shirane
et al. 1989; TeBeest et al. 1989). The multinuclear
condition may arise from division of the nucleus without
division of the spore (Churchill 1982). Hence, it may
represent homokaryotic condition.
An interesting phenomenon observed between type 1 and
type 2 strains is the differential sensitivity to benomyl
(methyl-2-benzimidazole carbamate: active ingredient in the
commonly used fungicide benlate). Type 2 strains were
completely inhibited by the levels of benomyl tested while
type 1 strains were tolerant although their growth rates
were significantly reduced (Table 2.2). The benomyl
tolerance of type 1 strains may have practical consequences
to the control of this disease. Current control measures
include spraying benomyl to control PFD (Fagan 1984b). If
type 1 strains are the primary causal agent of PFD as
previously suggested (Sonoda and Pelosi 1988; Agostini et
al. 1992; Gantotti and Davis, personal communication),
spraying in the field may only partially inhibit the
virulent pathogen while completely eliminating the less

24
virulent form. While slowing the epidemic in the short run,
this practice may have the long-term effect of selecting for
the most virulent form of the fungus.

CHAPTER 3
DNA POLYMORPHISMS FOUND AT MANY GENETIC LOCI EXAMINED
IN COLLETOTRICHUM GLOEOSPORIOIDES
Introduction
Ribosomal DNA in Funai
The nuclear ribosomal RNA (rRNA) genes of eukaryotes
are clustered in tandemly repeating units known as ribosomal
DNA (rDNA) unit repeats. In fungi as in many other
eukaryotes, each rDNA unit repeat consists of coding regions
for small subunit, SSU (17-18S), 5.8S, and large subunit,
LSU (25-26S) rRNA and intervening internal and external
transcribed and non-transcribed spacer (NTS) regions
(Fedoroff 1979; Chambers et al. 1986). Each rDNA unit repeat
codes for a 35S rRNA precursor which gives rise to SSU and
LSU rRNAs. In Neurospora crassa and Saccharomyces cerevisiae
a 35-37S rRNA precursor cleaves into a 17-18S rRNA of the
small ribosomal subunit (37S), and the 5.8S and 25S rRNAS of
large ribosomal subunit (60S) required for building 80S
ribosomes (Russell et al. 1976; Bell et al. 1977; Planta et
al. 1980).
The number of times rRNA genes are repeated varies
depending on the species of the organism. It was estimated
25

26
that there are about 185-225 copies of rDNA unit repeats in
Neurospora crassa (Krumlauf and Marzluf 1980; Rodland and
Russell 1982), 140 copies in yeast, Saccharomyces cerevisiae
(Schweizer et al. 1969; Rubin and Sulston 1973), 59 copies
in Rhizoctonia solani (Thanatephorus praticola AG-4)
(Vilgalys and Gonzalez 1990) and 60-90 copies in Coprinus
cinereus (Cassidy et al. 1984) per haploid genome.
Another rRNA gene recognized in fungi is 5S rRNA gene.
The 5S rRNA gene may be present within the rDNA unit repeat
or dispersed elsewhere in the genome. The 5S rDNA sequences
are located within the same rDNA unit repeat in S.
cerevisiae (Bell et al. 1977), Mucor racemosus (Cihlar and
Sypherd 1980), S. rosei and S. carlsbergensis (Verbeet et
al. 1983), C. cinereus (Cassidy et al. 1984), Schizophyllum
commune (Buckner et al. 1988), R. solani (Vilgalys and
Gonzalez 1990), the slime mold, Dictyostelium discoideum
(Maizels 1976), and water mold, Achlya ambisexualis, (Rozek
and Timberlake 1979). It is located elsewhere in the genome
in N. crassa (Free et al. 1979; Selker et al. 1981),
Schizosaecharomyces pombe (Tabata 1981), Aspergillus
nidulans (Borsuk et al. 1982), yeasts, Yarrowia lipolytica,
(Van Heerikhuizen et al. 1985), and Cochliobolus
heterostrophus (Garber et al. 1988).
In all known cases the 5S rRNA is transcribed
independently as a primary transcript separate from the 35-
37S rRNA precursor transcript (Udem and Warner 1972;

27
Miyazaki 1974) . When the 5S rRNA gene is within the same
unit repeat the 5S rRNA gene could be located in the same
strand, transcribed in the same direction as the other rRNA
genes or in the opposite strand, and transcribed in an
antiparallel manner (Aarstad and Oyen 1975). In C. cinereus
the 5S rRNA gene is transcribed in the same direction as the
rest of the rRNA genes (Cassidy et al. 1984).
Ribosomal DNA is Polymorphic in Many Fungi
Ribosomal DNA is a unique genetic marker which can be
used in the study of relatedness among organisms. Generally
the number of rDNA unit repeats is maintained from
generation to generation of an organism. The meiotic
recombination is suppressed within the rDNA array. In N.
crassa (Russell et al. 1988) and in C. cinerus (Cassidy et
al. 1984) the rDNA was shown to be inherited in a simple
stable Mendelian fashion exhibiting an approximately 1:1
ratio of the two parental rDNA types. No meiotic
recombinants were detected among the progeny indicating that
non-sister chromatid crossing over was highly suppressed in
the rDNA region of these organisms. However, Butler and
Metzenberg (1989 and 1990) demonstrated that N. crassa rDNA
can undergo unequal sister chromatid exchange and that the
number of rDNA unit repeats does not segregate in a simple
Mendelian fashion. Their observations suggested that

28
although the same rDNA RFLP can be inherited, the number of
unit repeats can be different from either of the parents.
Within a given species, the members of the rRNA gene
family are reasonably homogeneous in sequence, as are their
associated spacer sequences, despite frequent length and
restriction site differences among the latter. Yet there are
interspecific differences in sequence and these appear to be
much more pronounced for spacers than for genes. Smith
(1973) suggested that the differences between genes and
spacers might be in the rate at which they accumulate
mutations. Chromosomes containing mutations deleterious to
gene function would be eliminated by natural selection while
neutral spacer mutations would be retained in the
population. Hence, spacers change more rapidly than genes
simply by retaining a larger fraction of mutation (Smith
1973) .
Length heterogeneity and restriction site polymorphisms
in rDNA has been commonly observed in many fungi. These
polymorphisms were common among strains of S. cerevisiae
(Petes and Botstein 1977), N. crassa (Russell et al. 1984),
S. commune (Specht et al. 1984), C. cinerus (Wu et al.
1983), and Y. lipolytica (Clare et al. 1986). Both
restriction site and length polymorphisms have been also
observed among biological species of Armillaria (Anderson et
al. 1989). Polymorphisms of rDNA in many fungi are located
within the NTS region of the rDNA unit repeat (Cassidy et

29
al. 1984; Van Heerikhuizen et al. 1985; Rogers et al. 1989).
Chambers et al. (1986) compared the 8.4 kb rDNA unit repeat
of N. intermedia and N. sitophila with the 8.7 kb long rDNA
unit repeat of N. crassa and found that the 300 bp
difference was within the NTS region. Verbeet et al. (1983),
comparing S. rosei and S. carlsbergensis by heteroduplex
analysis, concluded that the NTS regions are largely non-
homologous in sequence whereas the transcribed regions are
essentially homologous. Russell et al. (1984) studied the
organization of the rDNA unit repeat in the strains of N.
crassa, N. tetrasperma, N. sitophila, N. intermedia, and N.
discreta and found that the size of the unit repeat has been
highly conserved among the strains of Neurospora. However, a
restriction enzyme site polymorphism in the NTS region was
found between the strains. This restriction site
polymorphism was strain-specific and not species-specific.
Restriction enzyme mapping of rDNA in yeast, Kluyveromyces
species has shown a length variation, and the variability
was found to reside in the NTS region (Lachance 1989).
Martin (1990) reported the presence of restriction site and
length polymorphisms within single oospore isolates of the
Oomycete genus Pythium, and the differences were found
within the NTS region and 3' end of the 26S coding region.
The NTS region has also been useful to study the
phylogenetic relatedness among fungal species and other
organisms (Verma and Dutta 1987).

30
The transcribed intergenic spacer (ITS) has also been
shown to be variable in fungi. In S. commune location of
strain-dependent length polymorphisms resided in the ITS
region between 18S and 5.8S cistrons (Buckner et al. 1988).
Chambers et al. (1986) compared the sequences of ITS regions
for N. crassa and S. carlsbergensis, and found that there is
a general lack of homology between the internal transcribed
spacer regions between 5.8S and 26S rRNA genes of these two
species. Buchko and Klassen (1990), using PCR technique to
amplify the ITS region, demonstrated length heterogeneity in
strains of Pythium ultimum.
The locations of rDNA polymorphisms were not confined
to the ITS regions. Polymorphisms within the coding regions
of rRNA genes due to addition or deletion of restriction
enzyme sites were found in fungi (Chambers et al. 1986) .
Another cause of rDNA polymorphism in eukaryotes is the
presence of introns in the coding regions. In fruit fly,
Drosophila melanogastor, the presence of an intron in the
coding region of the 28S rRNA gene has given rise to
polymorphism (Glover and Hogness 1977) of rDNA in this
organism. In fungi there are no conclusive reports for the
presence of introns in rRNA genes. However, Buckner et al.
(1988) examining the strain-dependent rDNA length
polymorphism in S. commune suggested the possibility of
having an intron in the coding region of the 18S rRNA gene.

31
Deletions of large fragments of rDNA may also occur in
organisms as reported by Malezka and Clark-Walker (1989). A
deletion of a 300 kb chromosomal fragment containing 35-40
rRNA cistrons has given rise to a new petite positive strain
of Kluyveromyces lactis.
One of the objectives of this study is to investigate
the variation of rDNA among the strains of C.
gloeosporioides causing PFD.
Fungal Cutinase Genes and Cutinase Isozymes
Plant pathogenic fungi penetrate their hosts through
the cuticle of epidermal cells or through cutinized cells
below natural apertures. Penetration may take place by
mechanical pressure (Brown and Harvey 1927; Brown 1936;
Pristou and Gallegly 1954; Chakravarty 1957; Wood 1960;
Meredith 1964; Bonnen and Hammerschmidt 1989b), or by
enzymatic degradation of the cuticle (De Bary 1887; Miyoshi
1895; Linskens et al. 1965; Akai et al. 1968; Kunoh and Akai
1969; Shayakh et al. 1977; Kolattukudy 1985) or by both
(Ellingboe 1968; Shishiyama et al. 1970; Nicholson et al.
1972) .
The plant cuticular barrier is composed of a
biopolymer, cutin and associated waxes providing a
protective covering against pathogen invasions and hazardous
effects of environment (Martin and Juniper 1970). The
structure of cuticle varies from one plant to another, and

32
it is influenced by genetic background as well as
environmental factors (Martin and Batt 1958; Martin 1964).
Almost all parts of the plant, surfaces of epidermal cells
of aerial plant parts, substomatal areas, mesophyll and
palisade cells (Martin and Juniper 1970; Sitholey 1971),
flower parts, seed coat (Kolattukudy et al. 1974) fruit
(Espelie et al. 1980), roots and tubers (Kolattukudy and
Agrawal 1974; Kolattukudy et al. 1975) contain a cuticular
layer.
The biopolymer, cutin is composed of C16 and C18 hydroxy
and hydroxy epoxy fatty acids (Van den Ende and Linskens
1974; Espelie et al. 1980; Kolattukudy 1980, 1981). The
composition of the cutin polymer and the proportions of C16
and C18 fatty acid monomers may vary depending on plant
species or varieties, organs of the same plant, or on growth
conditions (Espelie et al. 1979; Kolattukudy 1980).
The enzyme, cutinase can facilitate the hydrolysis of
cutin into its components (Baker and Bateman 1978; Dickman
et al. 1982). These hydrolysis products of cutin are also
potent inducers of the cutinase gene of the penetrating
fungus (Woloshuk and Kolattukudy 1986) . A small amount of
cutinase is constitutively expressed in the fungal spore
which senses the contact with the plant cuticle via the
unique cuticle monomers generated by this small amount of
cutinase. Consequently, these monomers trigger the
expression of the cutinase gene/ genes needed for the

33
production of cutinases which eventually degrade the cuticle
(Roller et al. 1982; Kolattukudy 1985; Woloshuk and
Kolattukudy 1986; Podila et al. 1988; Kolattukudy et al.
1989) .
Many plant pathogenic fungi examined have shown
production of different levels of cutinase isozymes (Purdy
and Kolattukudy 1975a; Lin and Kolattukudy 1980; Kolattukudy
et al. 1981). Direct observational, enzymological and
histochemical evidences have suggested that cutinase is
essential for the penetration of the plant by the fungal
pathogens. Specific antibodies prepared against cutinase
from Nectria haematococca (Fusarium solani f. sp. pisi,
Shaykh et al. 1977) and/ or diisopropylfluorophosphate
(DFP), a potent inhibitor of serine esterases, can prevent
infection of the host by this fungus indicating that
cutinase plays an essential role in the infection process
(Kolattukudy 1979). Dickman and Patil (1986) obtained
cutinase-deficient mutants of C. gloeosporioides, the causal
agent of papaya anthracnose, and found that they were
nonpathogenic to the intact papaya fruit. However, these
cutinase-deficient mutants produced normal lesions when
papaya surfaces were artificially wounded or treated with
purified cutinase enzyme. Dickman et al. (1989) were able to
introduce a Fusarium cutinase gene into a wound pathogen,
Mycosphaerella species through genetic transformation. These
transformants acquired the capacity to infect intact papaya

34
fruits, and the infection by them was prevented by the
treatment of antibodies against Fusarium cutinase.
Cutinolytic enzymes have been purified and
characterized from various plant pathogens (Kolattukudy
1980, 1985; Roller 1991) including Colletotrichum
gloeosporioides. The single enzyme produced by a strain of
C. gloeosporioides isolated from papaya fruit had a
molecular weight of 24 kd (Dickman et al. 1982) which is
very similar in size to other fungal cutinases (Kolattukudy
1980, 1985; Roller 1991).
There is considerable heterogeneity of molecular,
immunological and enzymological properties and primary
sequences of the cutinase enzymes (Kolattukudy 1985;
Ettinger et al. 1987; Trail and Roller 1990). Sequence
comparison of the cutinase genes cloned from C.
gloeosporioides and N. haematococca revealed considerable
dissimilarity. Even though both cutinase genes shared
homologous regions critical for activity and structural
integrity, only 43% of the amino acids were directly
conserved (Ettinger et al. 1987). Profound differences in
cutinase appear to exist even among Colletotrichum species
(Kolattukudy 1987). A cDNA clone of the cutinase gene from
C. capsici hybridized to genomic DNA from C. graminicola and
C. gloeosporioides, but not with C. orbiculare (syn. C.
lagenarium) or C. coccodes DNA. Though not extensively
investigated, the phenomenon of cutinase diversity is also

35
reflected in enzyme kinetics and activity. For example,
cutinolytic activity of esterases purified from N.
haematococca (Purdy and Kolattukudy 1975b) and F. roseum
culmorum (Soliday and Kolattukudy 1976) was highest at
alkaline conditions (pH 10), whereas an optimum of pH 6.5
was determined for the enzyme derived from Venturia
inaequalis (Roller and Parker 1989). Baker and Bateman
(1978) assayed sixteen plant pathogenic fungi, Botrytis
ciñera, B. squamosa, Cladosporium cucumerinum, C.
graminicola, N. haematococca, F. roseum, Gloeoceroospora
sorghi, Helminthosporium carbonum, H. maydis (race T),
Pythium aphanidermatum, P. arrhenomanes, P. ultimum, R.
solani, Stemphylum loti, and Sclerotium rolfsii and found
that they can produce various levels of cutinase isozymes
with acidic or alkaline pH óptimas. Evidence has been
presented that these differences in enzymatic properties may
allow for the tissue specificity of pathogens. Trail and
Roller (1990) reported an acidic pH optimum for the leaf
pathogen, Cochliobolus heterostrophus, pH 6.5 and an
alkaline pH optimum for the stem pathogen, R. solani, pH
8.5. The leaf and stem pathogen, Alternaría brassicola,
produced two cutinases, one with acidic and the other with
alkaline pH optima, pH 7.0 and 9.0 respectively. Differences
also have been reported for the cutinases produced by N.
haematococca and C. gloeosporioides. Only the enzyme from
the latter accepted palmitate as a substrate and the

36
specific esterase activity with both p-nitrophenol butyrate
and polymeric cutin was reported to be substantially lower
(Dickman et al. 1982) .
The ability of a pathogen to produce cutinase can be
used to measure the infecting capacity of the fungal
pathogen (Dickman et al. 1982; Roller et al. 1982). Thus the
regulation of expression of the cutinase gene could be
highly relevant to pathogenesis. Therefore, the cutinase
gene may be a good genetic marker to examine polymorphisms
among populations of fungi with differing specificities and
capabilities of causing plant disease.
One of the goals in this study was to investigate if
differences in morphologically defined type 1 and type 2
strains (see Chapter 2) are also reflected in the cutinase
isozymes and genetic organization of cutinase gene or genes.
Restriction Fragment Length Polymorphisms (RFLP) in Fungi
When fungal nuclear DNA is digested with a restriction
enzyme an enormous number of fragments generally result. In
order to study the restriction fragment pattern of DNA from
a specific chromosomal locus these fragments are size
fractionated by gel-electrophoresis, and individual
fragments are identified by Southern hybridization to
labelled probes (Southern 1975; Bernatzky 1988). Each
restriction fragment that hybridizes to a given probe
constitute a discrete chromosomal locus. Alleles can be

37
differentiated by the variation in restriction sites.
Restriction fragment length polymorphisms result from
specific differences in DNA seguence such as single base
pair substitution, additions, deletions, or chromosomal
changes (inversions and translocations) that alter the
fragment size obtained by restriction enzyme digestion.
First demonstrated by Grodzicker et al. (1974) for mapping
temperature-sensitive mutations in adenovirus, RFLP analysis
has contributed significantly in genetic analysis of many
organisms.
Genetic studies of plant pathogenic fungi have been
difficult due to lack of easily assayed genetic markers.
Restriction fragment length polymorphism has become a
popular tool for studying genetics of fungi because RFLP
markers are precise, codominant, selectively neutral, easy
to assay, and provide an unlimited number of genetic markers
(Michelmore and Hulbert 1987). Restriction fragment length
polymorphism could provide sufficient markers for the
development of detailed linkage maps for the plant
pathogenic fungi. It is also useful in studying genetic
variation, genomic organization and population genetics of
fungi. Combined with pulsed field gel electrophoresis RFLP
analysis provides a powerful tool to monitor genetic changes
throughout the genome (Michelmore and Hulbert 1987) .
Analysis of RFLP markers that flank genetic loci such as
virulence genes can provide information on the genetic basis

38
of any changes in phenotype. Closely linked RFLPs can be
used as tags for important traits. With RFLP markers it is
possible to create a molecular fingerprint of specific
individuals in a population. Hence, RFLPs provide a tool for
studying asexually reproducing populations of fungi. Engels
(1981) and Hudson (1982) presented mathematical models for
the genetic determination of variation among individuals in
a population using RFLP.
Use of RFLPs to measure genetic relatedness among
strains and closely related species of plant pathogenic
fungi is still in its beginning. Genetic variability of
several plant pathogenic fungi, Armillaria mellea (Anderson
et al. 1987), Sclerotinia species (Kohn et al. 1988),
Septoria tritici (McDonald and Martinez 1990) and
Aspergillus species (Someren et al. 1991) has been studied
using RFLP genetic markers. In C. gloeosporioides two
population subgroups were recognized by distinct RFLP
patterns detected by human minisatelite probes for
hypervariable regions within the genome (Braithwaite and
Manners 1989). Linkage maps have been developed for the
lettuce downey mildew fungus, Bremia lactucae using RFLPs as
genetic markers (Hulbert and Michelmore 1988; Hulbert et al.
1988). Hulbert et al. (1988) also reported the linkage of an
avirulance gene and a RFLP locus and suggested the
possibility of cloning the avirulance gene by chromosome
walking.

39
Castle et al. (1987) distinguished the commercial
mushroom, Agaricus brunnescens, from A. bitorquis using
distinct RFLP patterns. These RFLP patterns were used in the
identification of homokaryotic, heterokaryotic and hybrid
strains of this fungus (Castle et al. 1987) . Summerbell et
al. (1989) followed the segregation of RFLPs in wild
collected and artificially synthesized heterokaryotic
strains of A. brunnescens to investigate meiosis and the
meiotic recombination in this fungus.
The objective of this study is to determine if type 1
and type 2 Colletotrichum gloeosporioides are genetically
distinguishable. This will be done by examining rDNA
polymorphisms, the diversity of cutinase enzymes at the
isozyme and molecular level, and other molecular markers to
examine RFLPs in type 1 and type 2 strains of C.
gloeosporioides causing PFD of Tahiti lime and sweet orange.
Materials and Methods
Strains of Colletotrichum gloeosporioides
Strains of C. gloeosporioides, host and geographic
location are listed in the appendix A. Each strain is a
single spore culture grown. The place, year, and the host
tissue of isolation are mentioned in appendix A.

40
DNA Extraction
Fungal mycelium was grown in potato dextrose broth
(PDB) for seven days, harvested, frozen at -80 °C overnight
and lyophilized to complete dryness. The mycelium was ground
into a powder in liquid nitrogen using a mortar and pestle.
The mycelium powder was mixed with extraction buffer (100 mM
Tris pH 8.0, 50 mM EDTA, 100 mM NaCl, 10 mM B-
mercaptoethanol, 1% SDS, in H20) to make a slurry and
incubated at 65 °C for 3 0 min. One half volume of 5 M
potassium acetate (60 ml of 5 M potassium acetate, 11.5 ml
glacial acetic acid and 28.5 ml H20) was added to samples
and incubated on ice for 30 min. The supernatant was
collected by centrifugation at 12000 x g for 15 min and was
treated with 30-50 /¿g/ml of DNase free RNase (30 min at 37
°C, Sigma Chemical Co., St. Louis, MO). After RNase
treatment 200-250 /ug/ml Proteinase K (Sigma Chemical Co.,
St. Louis, MO) was added and incubated for an additional 20
min. Samples were purified by phenol:isoamyl alcohol:
chloroform (25:1:24 by volume) extraction and DNA was
precipitated by addition of a two-fold volume of absolute
ethanol. DNA pellets were dissolved in 100 ¿a 1 of TE (10 mM
Tris pH 8.0, 1 mM EDTA) and further purified by
precipitation with 0.7 volume of PEG/NaCl ((20% PEG 8000 in
2.5 M NaCl, Sigma Chemical Co., St. Louis, MO ) for 20-30
min on ice. Precipitated DNAs were resuspended in TE and
stored at -20 °C.

41
DNA Cloning and Restriction Enzyme Mapping
The rDNA of C. gloeosporioides was identified by
heterologous hybridization with N. crassa rDNA unit repeat
(pMF2, Free et al. 1979). Total DNA from C. gloeosporioides
strains H-25B and H-48 was digested with restriction enzyme
Pst I and fractionated on a 0.7% agarose (FMC BioProducts,
Rockland, ME) gel. The piece of the gel containing the 7 to
10 kb (H-25B) or 6-10 kb (H-48) DNA range was cut out and
the DNA eluted by the freeze squeeze method (Thuring et al.
1975). The DNA was ligated to Pstl-cut pUC119 (Sambrook et
al. 1989; Yanisch-Perron et al. 1985) and transformed into
Escherichia coli strain ER1647 (E. coli K-12 mcrB', Ref.
Raleigh et al. 1989; Woodcock et al. 1989) or DH5-a
(Sambrook et al. 1989). Ligation, preparation of competent
cells and transformation was carried out according to
Sambrook et al. (1989). Clones hybridizing to pMF2 were
identified and restriction mapped. For constructing
restriction maps, single and double restriction enzyme
digestions of two presumptive rDNA clones, called pCGRl (8.4
kb from strain H-25B), pCGR2 (6.8 kb from strain H-48), were
size fractionated in 1% agarose gels. Regions homologous to
the LSU and SSU rRNA of N. crassa were mapped by Southern
hybridization to heterologous probes subcloned from plasmid
pMF2 (Free et al. 1979; Martin 1990). The probe specific to
LSU was a 1.7 kb Xbal + BamHl fragment comprising all but

42
150 bp of the 5' end of the 17S coding region. A 2.9 kb
EcoRI fragment containing all but approximately 700 bp of
the 3' end of the 26S coding region in addition to 200 bp of
transcribed spacer sequences adjacent to 5' end was used as
the probe specific to LSU. These subclones were provided by
Dr. F. N. Martin, Plant Pathology Department, University of
Florida. A probe to detect the 5.8 rRNA gene was prepared by
polymerase chain reaction using primer flanking the gene.
The primers 5' TCCGTAGGTGAACCTGCGC 3' and 5'
GCTGCGTTCTTCATCGATGC 3' amplify a 290 bp fragment which
includes the transcribed spacer of the 3' end of the SSU
rRNA gene and the entire 5.8S gene (White et al. 1990).
Enzyme Assays and Electrophoresis of Cutinase
Colletotrichum gloeosporioides cultures were grown in a
mineral medium (Hankin and Kolattukudy 1968) amended with
tritiated cutin as the sole carbon source. Esterase
activities of strains were measured using p-nitrophenyl
butyrate (PNB) and p-nitrophenyl palmitate (PNP) as model
substrates (Roller and Kolattukudy 1982; Roller and Parker
1989; Purdy and Kolattukudy 1975a). Assays for cutinase
activity, sodium dodecyl sulfate polyacrylamide gel
electrophoresis (SDS-PAGE) and detection of active serine
esterases by tritium-labelled diisopropyl fluorophosphate
(3H-DFP) were as previously described (Roller and Parker
1989; Trail and Roller 1990). These experiments were

43
conducted by Dr. Wolfram Roller at the New York State
Agricultural Experiment Station, Geneva.
Probes Containing Cutinase Gene Sequences
Oligonucleotide primers, 5' TGCCCCAAGGTCATCTACATC 3'and
5'GAAGTTGGAGGCCAGGTCGGC 3' were synthesized to amplify a 220
bp fragment (intron and flanking sequences) of C.
gloeosporioides cutinase gene by polymerase chain reaction
(PCR). The PCR reaction mixture was prepared in a total of
100 pi containing 100 pM of each primer, 1.25 mM each of
dATP, dTTP, dCTP and dGTP, 2 pq of template DNA, and 10 ill
of reaction buffer (50mM KCl, 10 mM Tris-HCl pH 8.3, 1.5mM
MgCl2, 0.01% gelatin). The PCR mixture was denatured by
boiling 10 min and chilled on ice before adding 2 units of
Taq DNA Polymerase (Promega Inc. Madison, WI). The PCR
temperature cycles were programmed as following in a Coy
Temp Cycler II (Coy Corp., Grass Lake, MI). Denaturation
temperature was 94 °C, annealing temperature was 37 °C and
primer extension was at 72 °C. The first cycle was run 6 min
at 94°C, 2 min at 37 °C and 3 min at 72°C and the subsequent
30 cycles were run at 1, 2, and 3 min time intervals,
respectively, at these temperatures. Primer extension time
for the final cycle was 10 min. The fragments amplified by
PCR were labelled with 32P dCTP or digoxigenin (dig) dUTP
(Boehringer Mannheim Corp. Indianapolis, IN; see Appendix
C). Both a 220 and a 260 bp DNA fragment amplified by PCR

44
from strain H-48 hybridized to a genomic clone containing,
the cutinase gene, a 2.2 kb SphI DNA fragment, from C.
gloeosporioides (Ettinger et al. 1987). This clone was
provided by Dr. M. B. Dickman, Department of Plant
Pathology, University of Nebraska, Lincoln, NE.
High stringency Southern hybridization (Southern 1975)
using 32P-labelled probes was carried out according to
methods described by Sambrook et al. (1989). Hybridization
and washing of blots were carried out at 68 °C. First and
second washes were with 2X SSC, 0.1% SDS and 0.IX SSC, 0.1%
SDS respectively. The conditions of low stringency Southern
hybridization were as follows. DNA hybridization was at
65 °C and first and second washes were with 2X SSC at 55 °C.
Autoradiography was performed with Kodak X-OMAT AR5 film
(Eastman Kodak Co., Rochester, NY) and Dupont Hi-Plus
intensifying screens at -80 °C.
Detection of Restriction Fragment Length Polymorphisms
Plasmids containing N. crassa genes for anthranilate
synthetase (pNC2, Schechtman and Yanofsky 1983), glutamate
dehydrogenase (pJR2, Kinsey and Rambosek 1984), histidinol
dehydrogenase (pNH60, Legerton and Yanofsky 198 5) , and B-
tubulin (pSV50, Vollmer and Yanofsky 1986) were used to
detect DNA polymorphisms. Clones of N. crassa genes were
obtained from the Fungal Genetics Stock Center (Department
of Microbiology, University of Kansas Medical Center, Kansas

45
City, KS). Southern hybridization of 32P-labelled probes
were carried out according to methods described by Sambrook
et al. (1989) and Appendix C.
Results
Ribosomal DNA is Polymorphic in Colletotrichum
aloeosporioides
Southern hybridization of 32P labelled pMF2 to Pstl-
digested total blots from C. gloeosporioides strains
detected polymorphic forms of rDNA (Figure 3.1 and 3.2). The
type 1 strains (see Chapter 2 and Appendix A) contained only
a single form (8.4 kb PstI fragment) of rDNA (Figures 3.1
and 3.2). Although the size of the rDNA fragment in type 1
strains H-36 and 0C0 appears to be slightly higher than 8.4
kb in Figure 3.1, other restriction enzyme digestion tests
concluded that it is 8.4 kb in size (compare figures 3.5,
3.6, and 3.7). The 8.4 kb Pst I rDNA fragment was cloned from
Type 1 strain, H-25B and will be referred to as type 1 rDNA.
The restriction fragments obtained by digesting with 10
restriction enzymes and the restriction map of cloned type 1
rDNA unit (pCGRl) for these enzymes are illustrated in Table
3.1 and Figure 3.3 respectively. The restriction map of the
cloned rDNA unit was compared with the total rDNA
restriction fragments of the strain H-25B for 7 enzymes and
was identical (Figure 3.4). The map of cloned rDNA unit from
H-25B was tested against all type 1 strains for three

46
restriction enzymes, HindIII, SphI and SstI (Figures 3.5,
3.6, and 3.7 respectively). All the type 1 strains fell into
an identical group and the hybridization fragments for the
three enzymes agreed with restriction map of the cloned rDNA
fragment from Strain H-25B. Several subcloned fragments from
the NTS region were tested to determine if they can
specifically hybridize to type 1 strains. A 0.4 kb KpnI-PstI
subclone (pCGRIN) from the 3' end of NTS region (Figure 3.3)
was found to hybridize only to type 1 rDNA. The same total
DNA blot in Figure 3.1 was reprobed with the type 1-specific
subclone after removing the previous probe and only type 1
strains show hybridization to the subclone (compare Figures
3.1 and 3.8).
Ribosomal DNA among type 2 strains (see chapter 2) was
polymorphic for Pst I (Figures 3.1 and 3.2), SphI (Figure
3.6), and SstI (Figure 3.7). However, HindIII digested DNA
shows a similar pattern of rDNA polymorphism among all type
2 strains (Figure 3.5). Two hybridizing PstI fragments were
detected in strains H-48, 180269, 226802 (8.4 and 6.8 kb),
H-ll (5.0 and 3.4 kb), and H-47 (8.4 and 7.8 kb) by 32P
labelled pMF2 (Figures 3.1 and 3.2). However, the
hybridization intensity of the 8.4 kb band in strains 180269
and 226802 was very low and almost undetectable compared to
6.8 kb hybridizing band (Figure 3.1). All other type 2
strains had only one 6.8 kb hybridizing band. The 6.8 kb
Pst I fragment was cloned from strain H-48, and the

47
restriction map was identical when compared with the total
rDNA restriction fragments of the strain H-48 for 7 enzymes
(Figure 3.4). The cloned 6.8 kb PstI fragment from type 2
strain H-48 hybridizing to pMF2 will be referred to as pCGR2
or type 2 rDNA. The restriction fragments obtained by
digestion of clones of type 1 and type 2 rDNA unit with
various restriction enzymes and restriction enzyme maps are
listed in Table 3.1. The length of type 1 rDNA differs from
type 2 by the size of the NTS region of the unit (Figure
3.3). The restriction map of pCGR2 is distinct from that of
type 1 rDNA and the NTS is 1.6 kb shorter.
Ribosomal RNA Genes
In addition to the length heterogeneity, type 1 and
type 2 rDNA units differ by having restriction site
polymorphisms and addition and deletion of restriction sites
within coding regions for rRNA as well as intergenic
regions. Restriction sites for Smal and SstI within the SSU
coding region and a EcoRI site within the 5.8 S coding
region were found in type l rDNA. For the 10 restriction
enzyme sites examined none was detected within the SSU rRNA
and 5.8S RNA coding regions of type 2 rDNA. The coding
region for the LSU lies within BamHI and EcoRI sites for
both type 1 and type 2 rDNA, and were detected as 3.1 and
3.0 kb hybridizing fragments respectively. Restriction site

48
TABLE 3.1 Restriction fragments obtained by complete
digestion of the three ribosomal DNA clones
No.
Restriction
enzyme
Fragment
pCGRl
size (kb)
pCGR2
1
RstI
8.4
6.8
2
HindIII
4.1
3.2
3.2
3.1
1.1
0.5
3
SphI
6.0
4.2
1.4
2.5
1.0
0.1
4
EcoRI
2.7
3.2
2.5
2.6
2.4
1.0
0.8
5
BamHI
5.5
4.0
2.9
2.8
6
Kpnl
6.2
6.4
0.6
0.4
0.6
0.6
0.4
7
Hindi
3.3
3.0
2.8
1.6
1.4
1.0
0.7
0.6
0.2
0.6
8
Xbal
7.8
6.3
0.6
0.5
9
Smal
2.2
4.3
2.2
1.6
2.0
0.9
1.0
1.0
10
SstI
4.6
4.6
2.0
1.8
1.8
0.4
11
Clal
N
N
N = No restriction site detected

49
polymorphisms for enzymes tfindlll, EcoRI, Sstl, Hindi, and
Smal were detected within the coding region for LSU rRNA in
the type 1 and type 2 rDNA forms. The NTS region of type 1
rDNA has 4 Kpnl sites with three present at egual distance
of 0.6 kb, whereas within the NTS region of type 2 rDNA
there is only a single Kpnl site. The additional 1.6 kb NTS
region fragment in type 1 rDNA contains two Kpnl sites, each
0.6 kb apart and a 0.4 kb Pstl/Kpnl type 1 rDNA specific
fragment.
Diverse Cutinases and Cutinase Genes are Found in Type 1 and
Type 2 Strains of Colletotrichum gloeosoorioides
Cutinase production by fungal mycelium can be induced
by cutin monomers (Lin and Kolattukudy 1978). Similarly, all
Tahiti lime and Sweet orange strains of C. gloeosporioides
excreted esterases under these inductive conditions when
cutin was used as the sole carbon source. Although, both
model cutin substrates, p-nitrophenol-butyrate and -
palmitate, were hydrolyzed, the ratio of these two
activities was remarkably different (Table 3.2). Cutinolytic
activity was identified for all isolates and was
consistently higher at pH 6.0 than at the alkaline pH of
9.5. Extracellular proteins were labelled with 3H-DFP and
used as active site probe for serine esterase. Two esterases
in the molecular weight range common to many known fungal
cutinases (17kd-32kd, Kolattukudy 1980; Tanabe et al 1988;

50
TABLE 3.2 Extracellular enzyme activities of Colletotrichum
gloeosporioides
Strain
PNBase
mg/ml
PNPase
mg/ml
Cutinase PNB/PNP
kBq/h/mg
pH 6 pH9.5 ratio
PNB/
Cutinase pH 6
H-l
4731
255
18.7
2.0
9.4
18.6
254
IMB-3
6475
351
25.4
2.8
9.1
18.5
255
H-3
3678
302
23.9
4.5
5.3
12.2
154
H-4
8784
752
57.5
8.0
7.2
11.7
153
LP-1
5844
499
28.6
8.3
3.4
11.7
204
H-4 6
477
50
5.9
1.5
3.9
9.6
81
H-12
4692
554
40.0
1.3
30.8
8.5
117
H-25B
3627
488
27.5
5.5
5.0
7.4
132
Maran
3575
484
23.1
10.3
2.2
7.4
155
H-9
2193
329
18.8
4.2
4.4
6.7
117
H-48
4099
662
27.0
6.1
4.4
6.2
152
180269
4668
771
28.1
4.9
5.7
6.1
166
226802
4695
782
49.3
10.7
4.6
6.0
95
H-3 6
1863
369
23.7
5.8
4.1
5.0
78
Control
1456
332
12.8
7.9
1.6
4.4
144
PNB=p-nitrophenyl butyrate
PNP=p-nitrophenyl palmitate
Control consisted of all treatment without a fungal strain.

51
Trail and Koller 1990) were present for all strains (Figure
3.9). The molecular weight of these proteins differed among
strains and was correlated to C. gloeosporioides RFLP-types.
All strains of type 1 contained bands of 24 and 21 kd,
whereas all strains of type 2 contained 26 and 19 kd bands.
An additional esterase with a molecular weight of about 70
kd, which was not reported for the papaya isolate of C.
gloeosporioides (Dickman et al. 1982), was present
throughout the set of isolates. The high molecular weight
esterase was slightly larger for type 1 strains. The
relative contribution of this high molecular weight esterase
to the total esterase and cutinase activities remains
unknown. The enzyme might be similar to the 60 kd alkaline
cutinolytic esterase isolated from C. lagenarium (Bonnen and
Hammerschmidt 1989a) or the 54 kd non-specific esterase of
N. haematococca (Purdy and Kolattukudy 1975a).
A genomic clone containing the cutinase gene from a
papaya strain of C. gloeosporioides (Ettinger et al 1987)
was 32P-labelled and used to probe Sphl-digested DNA of C.
gloeosporioides from citrus. This clone contained a 2.2 kb
Sphl fragment which included the cutinase gene (189 bp exon-
52 bp intron-486 bp exon) and 5' and 3' flanking sequences
(Ettinger et al 1987). The probe hybridized to a 2.2 kb Sphl
fragment only in type 2 strains (Figure 3.10). DNA from type
1 strains showed no detectable level of hybridization at the
high level of stringency (see materials and methods) used

52
for Southern hybridization. Although the cutinase gene
sequence shows no polymorphism among type 2 strains for the
restriction enzyme SphI, a restriction fragment length
polymorphism can be detected among type 2 strains for
Hindlll (Figure 3.11). A 9.0 kb Hindlll fragment hybridized
to the probe in all type 2 strains except 226802 which show
hybridization to a 8.0 kb fragment. Type 1 strains did not
show any detectable level of hybridization to the probe at
this level of stringency used for Southern hybridization
(see Appendix C). The long exposure (>1 month) of this blot
resulted in appearance of 4.8, 5.4, 6.6, 7.4, and 9.0 kb
Hindlll hybridizing fragments of low level homology (Figure
3.12). Non-radioactive hybridization (Genius, Boehringer
Mannheim Corp. Indianapolis, IN) under low stringency
conditions shows weak hybridization of the probe to a 7.4 kb
Hindlll fragment from type 1 strains (Figure 3.13).
Polymerase chain reaction amplification using
oligonucleotide primers flanking an intron sequence in the
cutinase gene resulted in a single 220 bp fragment when H-3
(type 1 strain) or H-12 (type 2 strain) DNA was used as a
template. However, strains LP-1 (type 1) and H-48 (type 2)
produced two amplified fragments, 220 and 260 bp (Figure
3.14). The 220 and 260 bp fragment amplified by PCR from
strain LP-1, when used as probes for Southern hybridization,
also hybridized to numerous Hindlll fragments. Approximately
5-10 restriction fragments, ranging in size from 0.5 to >10

53
kb were identified (Figure 3.15 and 3.16 respectively).
Restriction fragments from all type 1 strains were almost
entirely identical. Hybridizing fragments from the type 2
strains showed dissimilar patterns. Hybridization of these
two probes to total DNA resulted in distinct DNA fingerprint
for type 1 strains.
Subgroups of Colletotrichum aloeosporioides have Distinct
RFLP Patterns Detected by Many Genetic Markers
Four clones of N. crassa genes were used as
heterologous probes to identify additional genetic loci in
HindiII-digested DNA from C. gloeosporioides strains. The
probe pSV50, containing the gene for fi-tubulin, hybridized
to a 3.2 kb fragment in type 1 strains but a 5.0 kb fragment
in type 2 strains (Figure 3.17). The probe, pJR2, containing
the gene for glutamate dehydrogenase, hybridized to a 3.2 kb
fragment only in type 2 strains, but only diffuse
hybridization was observed in type 1 strains (Figure 3.18).
The probe, pNH60, containing the gene for histidinol
dehydrogenase, hybridized to both a 3.8 and a 4.3 kb
fragment in type 1 strains but hybridized to 3.3 and 4.8 kb
fragments in type 2 strains (Figure 3.19).

54
n
i
H Q n I S I l
X H K E
I
a. i
â–º4 X
CTl
(N
CQ
E
U3
O
in
(0
co
(N
CO
VO
CM
u
cn
O
vo
ro
o
1
IÜ
l
l
CO
IN
1
u
X
a
X
X
rH
X
o
Figure 3.1 Polymorphic forms of ribosomal DNA unit in C.
gloeosporioides strains. Total DNA was digested
with Pst I and Southern hybridized with 32P
labelled pMF2 (N. crassa rDNA unit repeat).
Lanes H-12, 226802, H-36, and OCO shows slower
migration of DNA than expected. Lane H-4 DNA is
degraded. Numbers at the left indicate the size
of restriction fragments in kilobases (kb).

55
kb
co r* yo
-*r - I I I
as x x
8-4 «
6-8
Figure 3.2 Various restriction fragments contain ribosomal
DNA in type 2 strains of C. gloeosporioides.
Total DNA was digested with PstI and Southern
hybridized with labelled pMF2 (N. crassa rDNA
unit repeat). Numbers at the left indicate the
size of the major restriction fragments in
kilobases (kb).

56
1 8 3 109 3 475 27 10 9 749 4696 26 671
K IK
pCGR1
SSU 5.8 S LSU
1 3108
104 3572 7 9 97 74 261
o m
N
N
pCGR2
SSU 5.8 S LSU
1 kb
Figure 3.3 Restriction enzyme maps for cloned rDNA units
(pCGRl from Colletotrichum gloeosporioides type 1
strain, H-25B, and pCGR2 from type 2 strain, H-
48). Regions hybridizing to large subunit rRNA
(LSU), small subunit rRNA (SSU) from N.crassa and
the PCR amplified 5.8S rRNA gene from C.
gloeosporioides are indicated by solid boxes.
Restriction enzyme sites are as follows. PstI
(1), Hindlll (2), SphI (3), EcoRI (4), BamHI (5),
Kpnl (6) , Hindi (7) , Xbal (8) , Smal (9) , and
SstI (10).

57
kb
H
-u
w
H N
H H
H H H
H H H
H ¡B N
t 6 TJ
HÃœH
TJ g tJ
q <0 q
q <0 q
H
•H CQ
H
•h cq *H
H
tc
H
a:
H H H
\HN
H
H H
NH \
â– oho;
qx: o
a 0
H H
(a -u
(0 X5 W
a, S; to tq Bq x to
H Qi
J3 0
a 0
(0
&: to bq oq
H
<0
*
Figure 3.4 The rDNA of C. gloeosporioides strains H-25B
(first 10 lanes from left) and H-48 (next 10
lanes) digested with various restriction enzymes
and detected by Southern hybridization using 32P
labelled pMF2 (N. crassa rDNA unit repeat) as a
probe. The numbers at the left indicate the size
of major restriction fragments in kilobases (kb).

58
o\
CM
CQ
E
VO
O
l
H
VO
CM
to
(0
at
CM
CO
VO
1—1
CQ
CO
|
rH
CM
u
Ov
•o>
O
VO
CO
O
1
£
1
1
04
1
1
1
|
1
CO
CM
1
U
sc
H
SC
sc
i-3
SC
X
SC
£
SC
SC
iH
CM
sc
O
Figure 3.5 Ribosomal DNA polymorphism in C. gloeosporioides
strains. Total DNA was digested with HindIII
and Southern hybridized with 32P labelled pMF2
(N. crassa rDNA unit repeat). H-4 DNA is
degraded. The numbers at the left indicate the
size of restriction fragments in kilobases (kb).

59
Figure 3
o
kb g
cm
cv
o
VO
CQ
n
VO
CO
CM
CO
<0
in
CM
VO
H
i
m
vo
O
CTl
U
CM
rH
1
n
CQ
rH
1
CM
oo
1
1
(0
1
1
1
04
1
1
£
1
32
CM
1-1
X
K
£
as
X
X
â–º4
X
a;
H
£
.6 Ribosomal DNA polymorphism in C. gloeosporioides
strains. Total DNA was digested with SphI and
Southern hybridized with labelled pMF2
(N. crassa rDNA unit repeat). The numbers at the
left indicate the size of major restriction
fragments in kilobases (kb).

60
kb
n aa e
I rl v IN in d
H (D M 'í I N1 rl IN ^
I E I I CU I I l (0
X H 53 g >-3 XSKS
0\ (N
vO O
CO OIN’OIOnOHN'
I I CO CM I U I I
BKhmKOSSK
Figure 3.7 Ribosomal DNA polymorphism in C. gloeosporioides
strains. Total DNA was digested with SstI and
Southern hybridized with labelled pMF2
(N. crassa rDNA unit repeat). The numbers at the
left indicate the size of major restriction
fragments in kilobases (kb).

61
kb
OV
CM
r>
CQ
e
VO
O
1
rH
VO
CM
in
<0
00
CM
00
VO
(0
m
1
rH
(N
u

O
VO
m
o
|
£
1
1
cu
1
1
1
(0
1
1
CO
CM
1
u
X
H
PS
X
.J
X
X
X
E
X
X
rH
CM
X
o
Figure 3.8 A 0.4 kb Pstl/Kpnl fragment (pCGRIN) from the
non-transcribed spacer region of cloned rDNA
unit from C. gloeosporioides isolate H-25B
hybridizes only to the 8.4 kilobase (kb) rDNA
form in type 1 strains. Total DNA was digested
with PstI and Southern hybridized with 32P
labelled pCGRIN.

62
O'
(N
n
CQ
e
VO
O
1
rH
VO
(N
in
(0
CO
(N
CO
VO
H
CQ
n
1
Tf
rH
CM
ov
O
VO
m
o
1
S3
1
1
cu
1
1
l
(3
1
l
CO
CN
1
u
X
H
SC
s
33
33
SC
s
33
X
rH
52
o
Figure 3.9 Fluorography of 3HDFP-treated proteins after SDS-
polyacrylamide gel electrophoresis of
extracellular fluid from C. gloeosporioides
cultures grown on cutin as the sole carbon
source. Numbers at the left indicate the
molecular weight in kilodaltons (kd).

63
kb
n
1
rH
CQ
n
*T
1
S
l
1
IX
H
SS
as
CTl
(N
VO
ca
e
VO
O
CN
in
<0
CO
fM
CO
vo
â– 'T
rH
CM
u
a\
• O
vo
r~i
o
1
1
i
to
i
1
CO
CM
1
u
as
X
X
s
as
as
rH
fM
as
o
2-2
Figure 3.10 Cutinase gene in C. gloeosporioides type 2
strains. Total DNA was digested with SphI and
hybridized to a 32P labelled probe containing a
cloned cutinase gene. Numbers at the left
indicate the size of restriction fragment in
kilobases (kb) .

64
CTl CM
o
CQ
e
VO
o
1
iH
vo
CM
in
10
CO
CM
CO
VO
t-H
CQ
rr
1
f-i
CM
CT»
**
O
VO
m
O
1
se
l
1
PU
1
1
i
(0
1
1
CO
CM
1
o
S3
H
X
X
â–º4
X
S3
S3
s
S3
S3
iH
CM
S3
o
90
8 0
Figure 3.11 Restriction fragments hybridize to the cloned
cutinase gene only within type 2 strains. Total
DNA was digested with Hindlll and hybridized to
a 32P labelled probe containing a cloned
cutinase gene sequence from C. gloeosporioides.
H-4 DNA is degraded. Numbers at the left
indicate the size of restriction fragments in
kilobases (kb).

65
kb
90
70
4-8
ov
CM
n
CQ
e
vo
O
1
f—i
VO
cm
in
<0
CO
CM
CO
VO
H
CQ
m
■«f
1
pH
CM
u
cn
'T
O
vo
n
o
1
2
1
1
cu
1
1
1
n)
l
1
CO
CM
l
o
X
H
X
X
â–ºJ
X
X
X
X
as
X
pH
CM
X
o
Figure 3.12 Hybridization of total DNA from the indicated
strains to a cloned cutinase gene sequence. The
DNA was digested with Hindlll and hybridized to
a 32P labelled probe containing a cloned
cutinase gene sequence from C. gloeosporioides.
The blot was overexposed by placing it next to
X-ray film for more than 4 weeks. H-4 DNA is
partially degraded. Numbers at the left indicate
the size of major restriction fragments in
kilobases (kb).

66
CD CN
n CQ E *0 0
I H VO CM If) (0 CO CM CO vO
pH 03 M TJ1 I Vh I S I I CU I I I (0 I I CON I CJ
KDSHsa JsssasssHNXo
Figure 3.13 Presumptive cutinase genes in C. gloeosporioides
type 1 and type 2 strains. Total DNA was
digested with Hindlll and Southern hybridized to
a digoxigenin labelled probe containing a cloned
cutinase gene sequence from C. gloeosporioides.
H-4 DNA is degraded. Numbers at the left
indicate the size of restriction fragments in
kilobases (kb).

67
Figure 3.14 PCR Amplified fragments using oligonucleotide
primers flanking the intron in the
C. gloeosporioides cutinase gene. Lanes contain
DNA from PCR reactions using for the template
the cloned cutinase gene (CG), or total DNA from
the strains indicated. The control reaction lane
contained no template DNA. Numbers at the left
indicate the size of DNA restriction fragments
in basepairs (bp).

68
kb
10
4-8
2-2
cn
n
X
£
vO
O
1
rH
vO
CN
ID
<0
CO
CM
00
vo
1—1
CD
m
1
'T
iH
CM
OV
rr
O
VO
CO
1
2
1
1
04
1
1
1
(0
1
1
CO
CM
l
X
H
X
X
X
X
s
X
2
X
rH
04
=
Figure 3.15 RFLP patterns in total DNA from the indicated C.
gloeosporioides strains detected by Southern
hybridization to a 220 bp fragment amplified by
PCR. The total DNAs were digested with Hindlll
and the probe was labelled with digoxigenin
dUTP. H-4 DNA was partially degraded. Numbers at
the left indicate the size of major restriction
fragments in kilobases (kb).

69
kb
m
I
H 03 n
I S I
X M B
CTl
(N
03
E
VO
O
H
VO
CM
If)
(0
CO
(N
CO
VO
1
rH
M
c\
o
VO
m
o
1
CU
1
1
|
n)
1
1
CO
CM
i
u
X
tJ
X
X
X
X
X
X
<—1
OJ
X
o
T'
\
i
9 0 j. v>
4 8 ZZZZ'
3 7 *"""
>-
- <*
Figure 3.16 RFLP patterns in total DNA from the indicated C.
gloeosporioides strains detected by Southern
hybridization to a 260 bp fragment amplified by
PCR. The total DNAs were digested with HindIII
and the probe was labelled with digoxigenin
dUTP. H-4 DNA was partially degraded. Numbers at
the left indicate the size of major restriction
fragments in kilobases (kb).

70
kb
CMCT»
ovo E CO n
VO CO 04 GO 10 IT) OJ VO r-( |
Onvoo^m hNHv i ^ n u h
U I CMOO I I 10 I I I CU I I X I
OSiNHKSgKXKJSSHS
Figure 3.17 RFLP patterns in total DNA from the indicated C.
gloeosporioides strains detected by Southern
hybridization to the 32P labelled B-tubulin gene
from N. crassa (plasmid pSV50). The total DNAs
were digested with HindIII. Numbers at the left
indicate the size of restriction fragments in
kilobases (kb).

71
kb
m
I
h ffl n
I X I
X H DC
r
i
cu
CQ
vo cm in
'f H N
I
X DC X X £
0\
in
oo CM
o
I CO
X H
CM
O
CO l£>
io n O
CM I U
CM X O
3-2
Figure 3.18 RFLP patterns in total DNA from the indicated C.
gloeosporioides strains detected by Southern
hybridization to the 32P labelled glutamate
dehydrogenase gene from N. crassa (plasmid
pJR2). The total DNAs were digested with
Hindlll. Numbers at the left indicate the size
of the major restriction fragment in kilobases
(kb) .

72
cn cm
m CQ E vo o
l h vo (m in ra to n co vo
h m rMf i h N j-icn^ovonO
kb isiicuiiimiicocNiu
ShXS^SSSSSXhinSO
t ••• •«
Figure 3.19 RFLP patterns in total DNA from the indicated C.
gloeosporioides strains detected by Southern
hybridization to the 32P labelled histidinol
dehydrogenase gene from N. crassa (plasmid
pNH60). The total DNAs were digested with
Hindlll. Numbers at the left indicate the size
of the major restriction fragments in kilobases
(kb) .

73
Discussion
In Chapter 2, two types of Colletotrichum
gloeosporioides strains were described based on
morphological and growth characteristics. However,
morphological features vary with culture conditions and
time. Morphological variability always has been the case
with C. gloeosporioides (Burger 1921; Arx 1957). Despite
confusion associated with morphological inconsistency, at
the molecular level we see distinct differences between type
1 and type 2 strains.
The rDNA unit proved to be a molecular marker which can
be used to detect type 1 strains of C. gloeosporioides
(Figure 3.8). Based on the Southern hybridization results,
morphologically stable type 1 strains contain a single form
of rDNA. The restriction map of the cloned rDNA unit (Figure
3.3, pCGRl) from type 1 strain, H-25B was identical when
compared with the restriction map of the genomic rDNA
(compare Figures 3.3 and 3.4). The three enzyme sites
Hindlll, Sphl, and SstI examined for all type 1 strains
agree with the map of the rDNA clone from strain H-25B
indicating that the form of rDNA present in type 1 strains
has been cloned and mapped (compare figures 3.3, 3.4, 3.5,
3.6 and 3.7). The size of the type 1 rDNA unit, 8.4 kb in C.
gloeosporioides, is within the range of observed rDNA unit
repeat sizes for filamentous fungi such as N. crassa, 9.23
kb (Free et al. 1979), Aspergillus nidulans, 7.8 kb (Borsuk

74
et al. 1982), Schizophyllum commune 9.2-9.6 kb (Specht et
al. 1984), and Thanatephorus praticola, 8.8 kb (Vilgalys and
Gonzalez 1990). The given order of rRNA genes, 5' SSU-5.8S-
LSU 3' (Figure 3.3) within the rDNA unit repeat is similar
in all the fungi examined (Free et al. 1979; Cihlar and
Sypherd 1980; Borsuk et al. 1982; Cassidy et al. 1984;
Buckner et al 1988; Garber et al. 1988; Vilgalys and
Gonzalez 1990). The rDNA unit repeats in C. gloeosporioides
may also code for large, 35-37S, precursor rRNAs which give
rise to SSU, 5.8S, and LSU rRNAs required for the building
of 80S ribosomes (Russell et al. 1976; Bell et al. 1977;
Planta et al. 1980). The search for a specific fragment of
rDNA which can detect only type 1 rDNA was successful. The
sub-clone pCGRIN containing a 0.4 kb Pstl-Kpnl from the NTS
region was strain specific (compare Figures 3.1 and 3.8).
Therefore, type 1 strains can be defined as having a single
homogeneous form of rDNA as detected by a 8.4 kb Pst-1
fragment hybridizing to pMF2 and pCGRIN.
Morphologically variable type 2 strains (see Chapter 2)
were also diverse at DNA levels having different forms of
rDNA (Figures 3.2). The Pstl-Sphl-and SstI digested total
DNA blots, show RFLPs for rDNA within type 2 strains
(Figures 3.1, 3.2, 3.6, and 3.7). Only the HindIll-digested
blot shows a similar pattern of rDNA bands for type 2
strains (Figure 3.5). These results suggest that rDNA among
type two strains is heterogeneous. In addition, the specific

75
detection of only type 1 rDNA by pCGRIN suggest that
although some type 2 strains contain a rDNA form similar in
size to type 1 rDNA, the sequences may be different at least
at the NTS region. Several subclones from the NTS region of
type 2 rDNA hybridized to both type 1 and type 2 rDNA.
Therefore, a type 2 strain specific rDNA marker was not
found. Restriction site polymorphisms and length
heterogeneity in the rDNA unit repeat have been commonly
observed in many fungi. These polymorphisms were found
within the NTS region (Verbeet et al. 1983; Cassidy et al.
1984; Russell et al. 1984; Van Heerikhuizen et al 1985;
Chambers et al. 1986; Lachance 1989; Rogers et al. 1989),
ITS region (Chambers et al. 1986; Buckner et al. 1988) or
within coding regions (Chambers et al. 1986; Martin 1990).
In the fruit fly Drosophila melanogastor the presence of
introns has given rise to polymorphic forms of rDNA (Glover
and Hogness 1977). In fungi evidence for the presence of
introns in the rDNA coding regions is inconclusive (Buckner
et al. 1988).
Type 1 and type 2 C. gloeosporioides strains,
distinguished by rDNA polymorphisms were different both in
cutinase isozymes and molecular organization of relevant DNA
sequences. The slightly acidic pH optimum (pH 6.0) of
cutinolytic activity (Table 3.2) of these strains is a
reflection of their pathogenicity to aerial plant parts
(flowers or leaves) as observed for other aerial plant

76
pathogens such as V. inaequalis (Roller and Parker 1989),
Botrytis ciñera (Salinas et al. 1986) and Cochliobolus
heterostrophus (Trail and Roller 1990). This pH preference
is also congruent with the hypothesis that pathogens with
this type of cutinase are specialized for infecting aerial
plant surfaces rather than stem bases and roots (Trail and
Roller 1990).
All cutinases are serine esterases, and therefore they
can be detected by 3H-DFP which phosphorylates and inhibits
specific serine esterases (Roller and Rolattukudy 1982;
Rolattukudy 1985; Roller and Parker 1989). The two 3H-DFP
labelled bands (Figure 3.9) which are within the range of
molecular weights of fungal cutinases (Rolattukudy 1980;
Tanabe et al. 1988; Trail and Roller 1990) were not
previously seen in Colletotrichum species including papaya
isolate of C. gloeosporioides (Dickman et al. 1982;
Rolattukudy 1985). Although molecular weight and culture
conditions prior to electrophoresis suggest that cutinase
enzymes are present, DFP binding is serine esterase but not
cutinase specific (Roller and Rolattukudy 1982; Rolattukudy
1985; Roller 1991). These bands may not represent two
different primary gene products but may be the result of
post-translational modification (Roller 1991; Soliday et al.
1984). Some cutinase enzymes may undergo a proteolytic nick
and appear as two fragments after reduction of a disulfide
bridge and electrophoresis under denaturing conditions

77
(Roller 1991; Lin and Kolattukudy 1980; Purdy and
Kolattukudy 1975b; Soliday and Kolattukudy 1976) . Conclusive
demonstration that these two bands are actually two
cutinases awaits further experiments such as purification
and characterization of the catalytic activity. Other fungal
pathogens, N. haematococca (Purdy and Kolattukudy 1975a,
1975b), A. brassicola, and R.solani (Trail and Roller 1990)
are known to produce at least two distinct cutinases as
detected by distinct isozyme bands and enzyme catalysis.
For type 2 strains, only one DNA restriction fragment
hybridizes to the cutinase gene probe but two cutinase
isozymes may be present. One possibility is that these may
be two forms of cutinases encoded by distinctly different
genes in the same organism. Only one form of cutinase has
been described biochemically from the papaya strain of C.
gloeosporioides (Dickman et al. 1982) . The poor
hybridization of type 1 strains to the cutinase clone
despite the fact that they have abundant cutinase activity
suggests considerable evolutionary diversification of
cutinase gene sequences. Since the cDNA clone for cutinase
from C. capsici hybridizes readily with the total DNA from
C. graminicola and the C. gloeosporioides strain from
papaya, these species may be more related to type 2 than
type 1.
Another line of evidence for distinct genetic forms of

78
C. gloeosporioides from citrus comes from PCR amplification
of a cutinase gene sequence. The published DNA sequences for
cutinase genes from C. gloeosporioides and C. capsici
(Ettinger et al. 1987) show both conserved and diversified
regions of the gene. Two regions of identical sequence flank
a short stretch of DNA which includes the 52 bp intron of
the C. gloeosporioides cutinase and the 57 bp intron of the
C. capsici cutinase. The primer sequences are conserved in
cutinase genes of C. gloeosporioides, and C. capsici and
respectively should amplify a 220 bp or 222 bp fragment when
used as primers for PCR (Ettinger et al. 1987) . Indeed, one
or two amplified fragments of about 220-260 bp were obtained
from the DNA of the four strains tested. When used as a
probe for Southern hybridization these fragments were
anticipated to hybridize to conserved elements of the
cutinase gene and divergent sequences expected within the
intron. However, these sequences hybridize to multiple
restriction fragments producing DNA finger prints that
correspond to RFLP types.
Hybridization to these probes must not be specific for
cutinase sequences. The exact nucleotide sequences of the
two amplified fragments were not determined. Therefore, it
is necessary to determine the nucleotide sequence of the 220
and 260 bp amplified fragments before reaching any
conclusions. However, a computer search of the GenBank
database (Release 70.0, December 15, 1991) indicated no

79
significant sequence similarity between the 220 bp targeted
sequence of the cutinase gene from C. gloeosporioides
(Ettinger et al 1987) and genes other than for C. capsici
and C. gloeosporioides cutinase. Hybridization may be to
sequences common to other introns such as putative splice
junction sequences identified previously (Ettinger et al.
1987). Hybridization to the repetitive sequences may be
detected using the 220 bp probe and not the entire cutinase
clone because the intron sequence represents <1% of the
clone but 25% of the PCR fragment.
Cutinase is not the only extracellular enzyme produced
by fungi in the process of plant infection. There are many
others such as cellulolytic and pectinolytic enzymes
produced by fungi in culture and in diseased leaves (Oke
1989; Prusky et al. 1989). The type 1 and type 2 strains
also produce different isozymes of pectinesterase that
correlate to type 1 and type 2 (personal communication
Gantotti and Davis, Homestead, FL).
DNA polymorphisms detected by hybridization to 3 of 4
"housekeeping" genes from N. crassa also correspond directly
to type 1 and type 2 strains. To detect DNA polymorphisms
correlated to type 1 and type 2 strains by Southern
hybridization using these molecular markers it was not
necessary to search for specific restriction enzymes or to
test multiple loci. The only enzyme used to digest total
DNA, Hindlll was sufficient to provide RFLPs capable of

80
separating type 1 from type 2 strains. However, exceptions
to the strict correlation between polymorphism and strain
type was seen for hybridization to pNC2. In this case
variation was seen among type 1 strains. On the whole all
the molecular markers suggest that the two types are
different at the molecular level indicating they are indeed
genetically distinct populations of C. gloeosporioides.

CHAPTER 4
VARIABILITY OF MOLECULAR KARYOTYPES AND CHROMOSOMAL DNAS IN
COLLETOTRICHUM GLOEOSPORIOIDES
Introduction
Pulsed Field Gel Electrophoresis
Macromolecules such as nucleic acids and proteins can
be separated on the basis of size, charge or conformation by
gel-electrophoresis. Schwartz et al. (1982) made use of the
relaxation properties (Klotz and Zimm 1972) of large DNA
molecules for their separation in agarose gels by using two
alternating electric fields known as pulsed field gel
electrophoresis (PFGE). A major advance of the pulsed field
gel electrophoresis was achieved by Chu et al. (1986). They
applied the principles of electrostatics to calculate the
voltages needed to generate homogeneous electric fields
using multiple electrodes arranged around a closed contour.
In this system, contour clamped homogeneous electric field
(CHEF) gel electrophoresis, twenty four electrodes were
arranged in a hexagonal contour which offers reorientation
angles of 60 or 120.
81

82
Molecular Karyotypes of Fungi
Fungal chromosomes are too small to be observed readily
by conventional cytological methods using light microscopy.
However, electrophoretic karyotyping and molecular analysis
of chromosome-size DNA have become the new methods for
studying genomic structure of various organisms including
filamentous fungi. Many studies have conclusively
demonstrated that DNAs resolved by PFGE corresponds to
chromosomes (Carle and Olson 1985; Orbach et al. 1988; Brody
and Carbon 1989; Kayser and Wostemeyer 1991). However, the
number of bands need not be equal to the number of
chromosomes (Horton and Raper 1991). Electrophoretic
analysis of chromosomes provide a very convenient and rapid
way of assigning genes to chromosomes and for monitoring
entire genomes for any chromosomal rearrangements.
Pulsed field gel electrophoresis has been used to
separate chromosome-size DNA and analyze molecular
karyotypes of many fungi such as S. cerevisiae (Schwartz and
Cantor 1984; Chu et al. 1986), Candida albicans (Snell and
Wilkins 1986), Schizosaccharomyces pombe (Smith et al 1987;
Vollrath and Davis 1987), Neurospora crassa (Orbach et al.
1988), Ustilago maydis (Kinscherf and Leong 1988), Candida
stellatoidea (Kwon-Chung 1988, 1989: Wickes et al. 1991),
Aspergillus nidulans (Brody and Carbon 1989), C.
gloeosporioides (Masel et al. 1990), Ustilago hordei,
Tilletia caries, T. controversa (McCluskey et al. 1990),

83
Schizophyllum commune (Horton and Raper 1991), Absidia
glauca (Kayser and Wostemeyer 1991), Septoria tritici
(McDonald and Martinez 1991), Nectria haematococca (Miao et
al. 1991), Acremonium species (Smith et al. 1991; Walz and
Kuck 1991), Leptosphaeria maculans (Taylor et al. 1991), and
Fusarium oxysporum (Momol and Kistler 1992) .
Pulsed field gel electrophoresis of chromosomal DNA
combined with Southern analysis using linkage group-specific
probes were key methods in defining molecular karyotypes of
N. crassa (Orbach et al. 1988) and Aspergillus nidulans
(Brody and Carbon 1989). Molecular karyotyping of N. crassa
by Orbach et al. (1988) confirmed the seven linkage groups
previously defined by genetic analysis (Perkins et al.
1982). The genome size of A. nidulans was estimated by Brody
and Carbon (1989) to be approximately 31 Mb with six
chromosome-sized DNA bands. Kayser and Wostemeyer (1991)
reported differences in electrophoretic karyotypes for
mating types of the Zygomucete Absidia glauca.
Molecular karyotypes of many plant pathogenic fungi
examined to date have been variable. Kinscherf and Leong
(1988) analyzed the molecular karyotype of 17. maydis and
demonstrated that considerable chromosomal length
heterogeneity exists in this fungus. DNA hybridization
analysis suggested that stable large scale inter-chromosomal
exchange has given rise to novel chromosomes in one of the
strains. Taylor et al. (1991) using TAFE demonstrated that

84
the karyotypes of highly virulent and weakly virulant
strains of Leptosphaeria maculans (black leg of crucifers)
were polymorphic in both chromosome number and size. Highly
variable karyotypes for N. haematococca with unique
karyotypes for each strain were reported by Miao et al.
(1991). Deletions of large amounts of DNA from chromosomes
have given rise to karyotype variation as well as a
decreased frequency of the pisatin demethylase gene in N.
haematococca. Masel et al. (1990) suggested that chromosomal
rearrangements may play a role in generating variability of
karyotype of C. gloeosporioides. Distinct electrophoretic
karyotypes were reported for strains from two types of C.
gloeosporioides causing different anthracnose diseases in
Stylosanthus species in Australia. The strains showed
extensive chromosomal polymorphisms for both length and
number in the mini-chromosomes (molecules less than 2
million base pairs (Mb) in length) within each type.
The present study was undertaken to investigate the
variation of molecular karyotypes and chromosomal DNAs in
two types of C. gloeosporioides (see Chapter 2 and 3)
causing post bloom fruit drop of Tahiti lime and Sweet
orange.

85
Materials and Methods
Strains of Colletotrichum crloeosporioides
Strains used were obtained from several different areas
of Florida, Mexico, and from the Commonwealth Institute of
Mycology, England. They were isolated from diseased lime or
orange tissues. Details of host, place of collection and
date are tabulated in the appendix A.
Preparation of Protoplast Plugs
The strains of C. gloeosporioides were grown for 7 days
in 50 ml of 20% (w/v) V-8 juice (Campbell Co., Camden, NJ)
in at 250 rpm in Erlinmayer flasks on a Lab-Line orbit
shaker (Lab-Line Instruments Inc., Melrose Park, IL) at
ambient temperature (21-23 °C) , and conidia were collected
by centrifugation at 7000 x g for 5 min. About 109 spores
per ml were resuspended in 50 ml potato dextrose broth (PDB)
and incubated at room temperature (23 to 25 °C) for 16-24 h
at 200 rpm. When over 90% of spores were germinated, the
germlings were pelletted by centrifuging at 7000 x g for 5
min. Protoplasts were made by adding germlings to a 10 ml
solution containing NovoZym 234 (Novo Industries, Bagsvaerd,
Denmark) a complex mixture of wall-degrading enzymes. The
NovoZym solution was prepared by mixing 1.5 ml of 1 M
sorbitol, 50 mM sodium citrate containing 0.2 g of NovoZym
234 with 8.5 ml of 1.4 M MgSo4, and 50 mM Sodium Citrate pH

86
5.8. Germlings were incubated in this solution with gentle
rocking on a Bélico rocker (Bélico Biotechnology, Vineland
NJ) at 4 rpm for 3 to 6 h at ambient temperature for 3 to 6
hours until most cells were protoplasts. The protoplasts
were filtered through 4 layers of cheese cloth in order to
remove cell debris and undigested germlings. The filtrate
was centrifuged at 3000 rpm for 25 min at room temperature.
Protoplasts were removed from the top and washed three times
with 1 M sorbitol-50 mM EDTA pH 8.0. Protoplast inserts for
PFGE were made as described by method 1 of Orbach et al.
(1988).
Electrophoresis and Southern Analysis
A commercially available apparatus (BioRad CHEF DRII,
Richmond, CA) using different pulse time combinations was
employed in order to separate chromosome-sized DNAs.
Electrophoresis was done with 0.6% FastLane agarose (FMC
BioProducts, Rockland, ME) gels in 0.25X Tris Borate EDTA
(TBE) buffer (Sambrook et al. 1989) at 4 °C with rapid
circulation of the buffer. The gels were run at 40 volts for
6-10 days. Pulse times were "ramped" for various times
ranging from 10 to 180 min. For the separation of smaller
chromosome sized DNA 1% SeaKem Agarose (FMC BioProducts,
Rockland, ME) in 0.5X TBE buffer was used. These gels were
run at 200 V for 24 h with pulse times of 30-60 s or 50-90s.

87
Southern hybridization (Appendix C) experiments with
32P labelled ribosomal DNA (pMF2), B-tubulin gene (pSV50)
and cutinase gene (see chapter 3) were carried out to assign
these sequences to chromosome-size DNAs separated in CHEF-
gels.
Results
Chromosome-size DNAs (henceforth called chromosomes)
from type 1 and type 2 strains of C. gloeosporioides were
separated by PFGE using Saccharomyces cerevisiae and
Schizosaccharomyces pombe chromosome size DNA as standards
(BioRad Laboratories, Richmond, CA). The sizes in Table 4.1
represent the average size calculated for each chromosome
size-DNA band from independent CHEF-gels. Calculated sizes
for individual chromosome size DNA bands and relevant
figures are compiled in the Appendix D. Type 1 strains have
chromosomes distinguishable from type 2 strains (Table 4.1).
The chromosomes of C. gloeosporioides isolated from
Stylosanthes have been classified by Masel et al. (1990)
into larger, similar-sized chromosomes (>2 Mb) and smaller
variable-sized elements called "minichromosomes" (<2 Mb). A
similar arrangement was noted for strains isolated from
Tahiti lime and Sweet orange. Type 1 strains possess 5
chromosomes (Figures 4.1 and 4.2) and an additional 1 or 2
minichromosomes (Figure 4.3 and 4.5). Type 2 strains possess
3 chromosomes (Figures 4.1 and 4.2) in addition to 2 to 4

88
Table 4.1 Estimated megabase sizes for chromosome-size DNA
from Colletotrichum gloeosporioides type 1 and
type 2 strains
Strain
I
Estimated
II III
size*
IV
(Mb)
V
VI
VII
Type-1
strains
H-l
7.6
7.0
4.7
3.7
3.3
1.1
0.63
H-3
7.6
7.0
4.7
3.7
3.3
1.1
0.63
H-9
7.6
7.0
4.7
3.7
3.3
1.1
0.63
H-25B
7.6
7.0
4.7
3.7
3.3
1.1
0.63
H-36
7.6
7.0
4.7
3.7
3.3
0.77
0.63
LP-1
7.6
7.0
4.7
3.7
3.3
1.6
0.63
Maran
7.6
7.0
4.7
3.7
2.8
-
0.63
IMB-3
7.6
7.0
4.7
3.7
3.3
1.1
0.63
0C0
7.6
7.0
4.7
3.7
2.8
-
0.65
Type-2
strains
H-4
7.8
4.7
3.7
0.42
0.38
-
-
H-12
7.8
4.7
3.7
0.46
0.38
-
-
H-46
7.8
4.7
3.7
0.52
0.47
0.42
0.27
H-4 8
7.8
4.7
3.7
0.46
0.43
0.40
-
180269
7.8
4.7
3.7
0.43
0.41
0.39
-
226802
7.8
4.7
3.7
0.44
0.42
0.39
0.37
Schizosaccharomyces pombe and Saccharomyces cerevisiae size
standard were used for the calculation of Mb sizes. Megabase
sizes greater than 5.6 were estimated by extending the
calibration curve and therefore may be considered
approximate sizes.
- = not detected in any of the gels.

89
minichromosomes (Figures 4.4 and 4.5) depending on the
strain. Within each type, strains show variations in
chromosome number and size. However type 2 strains show more
total variation in chromosome and minichromosome size
(Figures 4.4 and 4.5).
A Southern blot separating larger chromosome-size DNAs
was hybridized with a 32P labelled ribosomal DNA probe. The
rDNA is associated with the 4.7 Mb chromosome in type 1
strains and with the 7.8 Mb chromosome in type 2 strains
(Figure 4.6). The cutinase gene can be assigned to the 4.7
Mb chromosome-size DNA only in type-2 strains (Figure 4.7).
Homologous regions were not detected in chromosomes of type
1 strains. The fi-tubulin gene hybridizes to both the 7.0 and
7.6 Mb chromosome doublet in type-1 strains and the 7.8 Mb
chromosome in type-2 strains (results are not shown due to
weak signals on X-ray film).

90
Figure 4.1 Chromosome-sized DNAs from C. gloeosporioides
compared to those for yeast (S. cerevisiae) and
fission yeast (S. pombe). DNAs were separated on
agarose gels using CHEF electrophoresis. Gels
were stained with ethidium bromide (0.5 /¿g/ml)
and fluorescence photographed with UV
transillumination. DNAs from the indicated
strains were run in 0.25X TBE, 0.6% agarose for
168 h at 40 V. Pulse switching times were ramped
from 40-70 min. Numbers at the left indicate size
standards in megabases (Mb).

91
cn
CN
03
E
VO
o
CM
ID
(0
VO
co
CN
00
CM
1
u
cn
o
VO
O
|
(T3
l
i
i
co
(N
u
35
35
s
33
X
33
H
o
Figure 4.2 Chromosome-sized DNAs from C. gloeosporioides
compared to those for fission yeast (S. pombe).
DNAs were separated on agarose gels using CHEF
electrophoresis. Gels were stained with ethidium
bromide (0.5 /¿g/ml) and fluorescence photographed
with UV transillumination. DNAs from the
indicated strains were run in 0.25X TBE, 0.6%
agarose for 168 h at 40 V. Pulse switching times
were ramped from 20-60 min. Numbers at the left
indicate size standards in megabases (Mb).

92
n ca e
i h in A
Figure 4.3 Chromosome-sized DNAs from C. gloeosporioides
compared to those for yeast (S. cerevisiae). DNAs
were separated on agarose gels using CHEF
electrophoresis. Gels were stained with ethidium
bromide (0.5 /xg/ml) and fluorescence photographed
with UV transillumination. DNAs from the
indicated strains were run in 0.5X TBE, 1.0%
agarose for 24 h at 200 V. Pulse switching times
were ramped from 60-90 s. Numbers at the left
indicate size standards in kilobases (kb).

93
CM
OV
O
VO
e
CQ
VO
CO
CN
CO
(0
in
CN
VO
r-t
n
VO
O
OV
u
CN
iH
1
m
u
1
CM
CO
1
i
(0
i
1
1
cu
1
1
w
as
CN
r-i
as
as
s
X
X
as
X
as
Figure 4.4 Chromosome-sized DNAs from C. gloeosporioides
compared to those for yeast (S. cerevisiae). DNAs
were separated on agarose gels using CHEF
electrophoresis. Gels were stained with ethidium
bromide (0.5 /¿g/ml) and fluorescence photographed
with UV transillumination. DNAs from the
indicated strains were run in 0.5X TBE, 1.0%
agarose for 24 h at 200 V. Pulse switching times
were ramped from 30-60 s. Numbers at the left
indicate size standards in kilobases (kb).

94
vo o CQ CD
Figure 4.5 Chromosome-sized DNAs from C. gloeosporioides
compared to those for yeast (S. cerevisiae). DNAs
were separated on agarose gels using CHEF
electrophoresis. Gels were stained with ethidium
bromide (0.5 /xg/ml) and fluorescence photographed
with UV transillumination. DNAs from the
indicated strains were run in 0.5X TBE, 1.0%
agarose for 24 h at 200 V. Pulse switching times
were 60 s (15 h) and 90 s (9 h). Numbers at the
left indicate size standards in kilobases (kb).

95
VO [". CO CN
rr rH
Mb 0* 0 1 1 1 1
,T,U w w
(N CV
CQ CQ O VO
in in co oj
n o> m cm vo o
I I I IN CO
Figure 4.6 Southern blot of chromosome-sized DNAs from C.
gloeosporioides strains, yeast (S. cerevisiae)
and fission yeast (S. pombe) hybridized to a
32P labelled clone containing the ribosomal DNA
repeat from N. crassa. Hybridization is to a 4.7
or 7.8 Mb for type 1 or type 2 strains
respectively. Gel running conditions were 0.25X
TBE, 0.6% agarose for 264 h at 40 V. Pulse
switching times were ramped from 10-120 min.
Numbers at the left indicate size in megabases
(Mb) .

96
a> cm
vo o
VO
r'
CO
CM
CO
r-i
f—1
(N
CO
O
VO
o\
1
rH
m
rH
1
rH
1
1
l
CO
OJ
1
cu
1
1
1
i
i
CU
1
1
X
as
X
CM
PS
â–º4
X
X
as
X
X
hJ
X
X
IJ I
Figure 4.7 Southern blot of chromosome-sized DNAs from C.
gloeosporioides strains, hybridized to a 32P
labelled clone containing the cutinase gene.
Hybridization is to a 4.7 Mb for type 2 strains.
Gel running conditions were 0.25X TBE, 0.6%
agarose for 240 h at 40 V. Pulse switching times
were 40 min (120 h) and 100 min (120 h).

97
Discussion
In previous studies, the DNAs resolved in agarose gels
by PFGE corresponded to chromosomes (Carle and Olson 1985;
Orbach et al 1988; Brody and Carbon 1989; Kayser and
Waltsmeyer 1991). The number of bands visible in an ethidium
bromide stained gel may not always be equal to the number of
chromosomes. Sometimes different chromosomes of same size
may co-migrate resulting in a single band (Brody and Carbon
1989; Horton and Raper 1991). In general the variation in
the number and migration lengths of these bands may
represent differences in size and number of chromosomes in
the organism. Chromosomal variations produced by
translocations, recombinations and losses in fungi may be
easily studied by electrophoretic karyotyping.
Pulsed field gel electrophoresis has allowed detection
of chromosomal variation within species of phytopathogenic
fungi (Kinscherf and Leong 1988; Kistler and Miao 1992;
Masel et al. 1990). Similarly, variation in chromosome-sized
DNA in C. gloeosporioides from Tahiti lime and Sweet orange
has been observed. Two distinct electrophoretic patterns for
type 1 and type 2 can be described (Table 4.1). These
patterns show similarities to those described by Masel et al
(1990) for type-A and type-B strains of C. gloeosporioides
from Stylosanthus. Similar to type-A, the type 1 strains
which have 5 large chromosomes, whereas type-B is similar to
type 2 strains by having 3 large chromosomes.

98
Minichromosomes are variable and within each type there are
differences in number and size. Type 2 strains have a
greater diversity in size and number of minichromosomes. The
differences in the size of the larger chromosomes between
types is best illustrated by the differently sized molecules
hybridizing to pMF2 and cutinase gene. The smaller
chromosomal DNA observed in these strains may be similar to
B chromosomes (Jones and Rees 1982) . In other fungi, B
chromosomes have been shown to contain unique pathogenicity
or virulence genes (Miao et al. 1991) or simply duplicated
sequences from larger chromosomes (Rikkerink et al. 1990).

CHAPTER 5
GENERAL DISCUSSION AND CONCLUSIONS
The objectives of these studies were to examine
variability of Colletotrichum gloeosporioides from Tahiti
lime and Sweet orange at the morphological, molecular, and
chromosomal levels.
Morphological variability of C. gloeosporioides is
reported in Chapter 2. These results are comparable to
morphological observations made by Burger (1921), Fagan
(1980), Denham and Waller (1981), and Sonoda and Pelosi
(1988). A reasonable level of morphological and growth
differences exist between type 1 and type 2 strains. By
observing mycelial color, growth rate, and sectoring in
potato dextrose agar and potato dextrose broth, one may be
able to easily recognize type-1 and type-2 strains at
morphological level.
The two sub-populations recognized in Chapter 2 show
their phenotypic and genetic distinctiveness as reported in
Chapter 2 and 3. The differential sensitivity to benomyl of
type 1 and type 2 strains has an important practical aspect.
The present use of benomyl as a fungicide to control PFD
needs to be reconsidered in light of benomyl tolerance of
virulent type-1 strains. All the strains were equally able
99

100
to infect persian lime flowers in this study. Type 1 may be
considered as the most virulent form based on observations
made by others (Fagan 1980; Sonoda and Pelosi 1988; Agostini
et al 1992; Gantotti and Davis, personal communication).
The isozyme and genetic diversity (Chapter 3) of
cutinases between type 1 and type 2 strains needs to be
further investigated. Sequencing of cutinase proteins and
genes may provide accurate measurement of diversity of these
phenotypic and genetic characters between type 1 and type 2
strains.
The ribosomal DNA in C. gloeosporioides is polymorphic
(Chapter 3). Within type 1 strains, rDNA appears to be of a
single form. The sequence of type 1 rDNA differs from type
2. The type 1 rDNA specific subclone pCGRIN may be a
suitable probe to detect these strains from PFD epidemics.
The cloning and characterization of rDNA from type 2 strains
was incomplete. Only a 6.8 kb PstI fragment was cloned and
mapped as a form of type 2 rDNA. Observations made in this
study suggest that rDNA may exist in several forms in
morphologically unstable type 2 strains. Other forms of type
2 rDNA need to be cloned and mapped. Perhaps, a sequence
comparison at the NTS region may provide some insight into
the rDNA diversity within type 2 strains and between type 1
and type 2 strains. In addition, the RFLP patterns
associated with four "house keeping genes" also suggest
genetic diversity within type 1 or type 2 strains.

101
The specific RFLP detected by the 6-tubulin gene needs
to be further investigated in light of the differences in
benomyl tolerance of type 1 and type 2 strains (Chapters 2
and 3). Future research may need to be focused on cloning
the 6-tubulin genes from type 1 and type 2 strains. A
sequence comparison may perhaps elucidate the molecular
mechanisms underlying the observed differences in benomyl
tolerance.
The distinct molecular karyotypes (Chapter 3)
representing each subpopulation of C. gloeosporioides
confirm the division of two RFLP types. The two types of
strains differ not only in chromosome number and size but
also in chromosome assignment.
On the whole, all morphological, phenotypic and genetic
markers suggest that type 1 strains are distinct from type
2. The taxonomic position of these type 1 and type 2
strains is not known. They may be considered as subspecies
or races within the species. However, host specificity
corresponding to types was not observed.
This study has conclusively demonstrated that two
genetically distinct pathogen populations of C.
gloeosporioides exist in association with post bloom fruit
drop disease of Tahiti lime and Sweet orange. Strains from
both types are present in Florida. However, of the five
strains obtained from elsewhere outside the United States
all were type 2.

APPENDIX A
STRAINS OF COLLETOTRICHUM GLOEOSPORIOIDES
Strain
Year & place
Host
Type 1
H-l
1989,
Iirunokalee,
FL
Tahiti
lime
H-3
1989,
Immokalee,
FL
Tahiti
lime
H-9
1988,
Homestead,
FL
Tahiti
lime
H-21
1989,
Homestead,
FL
Tahiti
lime
H-22
1989,
Homestead,
FL
Tahiti
lime
H-25B
1989,
Homestead,
FL
Tahiti
lime
H-35a
1989,
Ocala, FL
Sweet
orange
H-36a
1989,
Ft. pierce,
FL
Sweet
orange
LP-1C
1990,
Lake Placid
., FL
sweet
orange
Maranc
1990,
Indiantown,
FL
Sweet
orange
0C0c
1990,
Arcadia, FL
Sweet
orange
TUR-1C
1990,
Lake Alfred
., FL
Sweet
orange
Type 2
H-4
1989,
Immokalee,
FL
Tahiti
lime
H-ll
1988,
Homestead,
FL
Tahiti
lime
H-12
1988,
Homestead,
FL
Tahiti
lime
102

103
Strain
Year & place
Host
H-23
1989,
Homestead,
FL
Tahiti lime
H-24
1989,
Homestead,
FL
Tahiti lime
H-46a
1989,
Vera Cruz,
Mexico
Sweet orange
H-47a
1989,
Vera Cruz,
Mexico
Sweet orange
(0
00
K
1989,
Vera Cruz,
Mexico
Sweet orange
180269b
Belize
Sweet orange
226802b
Belize
Sweet orange
aStrains
were provided by Dr. R.
Sonoda,
University of
Florida, IFAS Agricultural Research and Education Center,
Fort Pierce, FL.
bStrains obtained from Commonwealth Institute of Mycology,
Kew, London, UK; provided by Dr. R. Sonoda
cStrains provided by Dr. L. W. Timmer, University of
Florida, IFAS Citrus Research and Education Center, Lake
Alfred, FL. All the other strains were obtained from the
culture collection at University of Florida, IFAS Tropical
Research and Education Center, Homestead, FL.

APPENDIX B
ANALYSIS OF VARIANCE TABLES
Table B.l ANOVA for growth rates (cm/24h) of type 1 and type
2 strains in potato dextrose agar (experiment 1)
Source
df
SS
MS
Fo
Type-1
vs Type-2
1
0.45
0.45
2250***
Within
Type-1
7a
0.2759
0.0394
197***
Within
type-2
7
0.0225
0.0032
16***
Error
64
0.0172
0.0002
Total
79
0.7656
a9 type 1 isolates were tested. Two isolates 0C0 and H-36
had identical measurements, and were considered as one for
analysis.
*** Significant at a=0.01 level
104

105
Table B.2 ANOVA for growth rates (cm/24h) of type 1 strains
in benomyl treated potato dextrose agar
(experiment 1)
Source
df
SS
MS
F0
Strains
7a
0.2193
0.0313
313 0***
Concentration
2
0.2401
0.12
12000***
Strain X Cone.
14
0.1199
0.0085
_ _ _ *★*
850
Error
96
0.0014
0.00001
Total
119
0.5807
a9 type 1 isolates were tested. Two isolates OCO and H-36
had identical measurements, and were considered as one for
analysis.
*** Significant at a=0.01 level

106
Table B.3 ANOVA for growth rates (mm/h) of type 1 and type
2 strains in potato dextrose agar (experiment 2)
Source
df
SS
MS
Fo
Type-1
vs Type-2
1
0.0367
0.0367
29424***
Within
Type-1
7a
0.0194
0.00277
2216***
Within
type-2
7
0.0034
0.000485
3 88***
Error
16
0.00002
0.00000125
Total
31
0.0596
a9 type 1 isolates were tested. Two isolates OCO and H-36
had identical measurements, and were considered as one for
analysis.
*** Significant at a=0.01 level

107
Table B.4 ANOVA for growth rates (mm/h) of type 1 strains
in benomyl treated potato dextrose agar
(experiment 2)
Source
df
SS
MS
Fo
Strains
7a
0.015
0.002
10***
Concentration
2
0.017
0.008
40***
Strain X Cone.
14
0.004
0.0003
1.5*
Error
24
0.005
0.0002
Total
47
0.041
a9 type 1 isolates were tested. Two isolates OCO and H-36
had identical measurements, and were considered as one for
analysis.
*** Significant at a=0.01 level
* Significant at a=0.25 level

108
Table B.5 ANOVA for growth rates (mm/h) of type 1 and type
2 strains in potato dextrose agar (experiment 1&2
combined)
Source
df
SS
MS
Fo
Type-1
vs Type-2
1
0.10492
0.10492
2098***
Within
Type-1
7a
0.07086
0.01012
202***
Within
type-2
7
0.00235
0.00033
6.6***
Error
96
0.00487
0.00005
Total
111
0.18301
a9 type
1 isolates
were tested. Two isolates OCO
and H-3 6
had identical measurements, and were considered as one for
analysis.
*** Significant at a=0.01 level

109
Table B.6 ANOVA for growth rates (mm/h) of type 1 strains
in benomyl treated potato dextrose agar
(experiment 1 & 2 combined)
Source
df
SS
MS
Fo
Strains
7a
0.05440
0.00777
4450***
Concentration
2
0.06265
0.03132
17938***
Strain X Cone.
14
0.03175
0.00226
1294***
Error
145
0.000253
0.00000174
Total
167
0.014906
a9 type 1 isolates were tested. Two isolates OCO and H-36
had identical measurements, and were considered as one for
analysis.
*** Significant at a=0.01 level

APPENDIX C
PROCEDURES FOR DNA LABELLING AND SOUTHERN HYBRIDIZATION
Probes were labelled with either radioactive a-32P or
digeoxigenin-labelled deoxyuridine-triphosphate (dUTP)
(Boehringer Mannheim Corp. Indianapolis, Indiana) by random
priming according to the manufacturer's directions.
Gels were irradiated with UV light (254 nm) for 1-2 min
before incubating with 0.25 M HCl for 20 min. DNA was
denatured in 0.5 M NaOH-1.5 M NaCl and then neutralized in
0.5 M Tris-HCl-1.5 M NaCl pH 7.0 for 30 m each. DNA was
capillarily transferred (Southern 1975) to a Nytran membrane
(Schleicher & Schuell, Keene, NH) for >24 h in 10X SSC (1 x
SSC=0.5 M NaCl-0.015 M Sodium Citrate pH 7.0) transfer
buffer. DNA on the membrane was immobilized by UV cross
linking (Stratalinker, Stratagene Co., La Jolla, CA).
Labelling of the probes with 32P dCTP was according to
the manufacturer's recommendations (Boehringer Mannheim
Corp. Indianapolis, IN). Labelled nucleotide was
incorporated into probes by random oligonucleotide priming
of the large subunit of DNA polymerase (Klenow fragment).
Non-radioactive DNA labelling and detection were by enzyme-
linked immunoassay of digeoxigenin-labelled deoxyuridine
triphosphate incorporated into probes by random
110

Ill
oligonucleotide priming of the Klenow fragment.
Membranes were pre-hybridized and hybridized according
to the manufacturer's instructions for the nonradioactive
process (Boehringer Mannheim Corp. Indianapolis, IN).
For the radioactive high stringency process, membranes
were pre-hybridized in sealed plastic bags for 3-6 h at 68 C
in 0.2 ml/cm2 of pre-hybridization solution (250 mg dried
milk, 10 jug denatured salmon sperm DNA in 100 ml 6X SSC) .
After removing pre-hybridization solution 0.1 ml/cm2,
hybridization solution (pre-hybridization solution + 10% w/v
dextran sulfate) containing labelled probe was added, and
hybridized at 68 °C for >18 h in sealed plastic bags.
Hybridized blots were washed twice (first, 0.25 g dried
milk, 0.1% SDS in 100 ml 2X SSC; second 0.1X SSC, 0.1% SDS)
at 68 °C for 45 min each. The blots were air dried and
exposed to X-ray films (Kodak X-Omat GR, Kodak co.,
Rochester NY) at -80 °C.
The following are the low stringency conditions for
both radioactive and non radioactive probes: Pre¬
hybridization was the same. Hybridization was at 65 °C and
washes were at 55 °C. Solution for the first wash was same
as high stringency conditions. Only the second wash was at
2x SSC instead of 0.lx SSC.

APPENDIX D
CALCULATED MB SIZES FOR CHROMOSOMAL DNAS IN
COLLETOTRICHUM GLOEOSPORIOIDES
In all figures Sp=Schizosaccharomyces pombe;
Sc=Saccharomyces cerevisiae. All CHEF-gels were run in a
cold room at 4 °C. Gels in figures D.l to D.7 were 0.6%
FastLane agarose (FMC BioProducts, Rockland, ME). The
running buffer was 0.25X TBE, voltage was 40 V, and the
running time was 7-10 d. Gels in figures D.10 and D.ll were
of 1% SeaKem agarose (FMC BioProducts, Rockland, ME) in 0.5X
TBE buffer at 200 V. Running time was 24 h. The beginning
and ending pulse times are indicated under the figure
legends.
The megabase sizes tabulated in this appendix were
calculated from calibration curves developed for each gel
seperately using S. pombe and S. cerevisiae chromosomes as
size standards.
112

113
Figure D.l Chromosome-sized DNAs from C. gloeosporioides
compared to those for yeast (S. cerevisiae) and
fission yeast (S. pombe). DNAs were separated on
agarose gels using CHEF electrophoresis. Gels
were stained with ethidium bromide (0.5 ¿ug/ml)
and fluorescence photographed with UV
transillumination. DNAs from the indicated
strains were run in 0.25X TBE, 0.6% agarose for
168 h at 40 V. Pulse switching times were ramped
from 50-100 min. Numbers at the left indicate
size standards in megabases (Mb). Lanes Sp., H-
12, Maram and H-36 DNA was partially degraded.

114
Mb
O)
CN
VO
O
CO
CQ
CQ
VO
CO
CM
CO
CN
in
in
in
â– 
•
O
VO
rH
n
â– 
(Tv
I
iH
i
(N
I
CN
â– 
i
CN
|
i
x
i
X
1
X
CO
iH
CN
CN
1
X
i
X
1
X
1
X
1
X
1
X
X
X
Figure D.2 Chromosome-sized DNAs from C. gloeosporioides
compared to those for yeast (S. cerevisiae) and
fission yeast (S. pombe). DNAs were separated on
agarose gels using CHEF electrophoresis. Gels
were stained with ethidium bromide (0.5 /xg/ml)
and fluorescence photographed with UV
transillumination. DNAs from the indicated
strains were run in 0.25X TBE, 0.6% agarose for
240 h at 40 V. Pulse switching times were ramped
from 10-180 min. Numbers at the left indicate
size standards in megabases (Mb).

115
[" v£>
*r
t I
K X
CO CM H (M
'T H I Tf H m H
i i ex. i i iii
Mb KKJEE "CEE
cm cn
O VO
co
cn vo
l
a:
CM CO
O M*
CM 00
CM t—i
I
SC
Figure D.3 Chromosome-sized DNAs from C. gloeosporioides
compared to those for yeast (S. cerevisiae) and
fission yeast (S. pombe). DNAs were separated on
agarose gels using CHEF electrophoresis. Gels
were stained with ethidium bromide (0.5 pq/ml)
and fluorescence photographed with UV
transillumination. DNAs from the indicated
strains were run in 0.25X TBE, 0.6% agarose for
240 h at 40 V. Pulse switching times were 40 min
(120 h) and 100 min (120 h). Numbers at the left
indicate size in megabases (Mb).

116
OJ
e o e 03
00 CO (OiriiNVOiH
CTirfUiVO'^C^^fMr-i'T I fOn
uv. OI IfOCMl líOI I ICUl IQ.
nD wissNiESsri^xxw
Figure D.4 Chromosome-sized DNAs from C. gloeosporioides
compared to those for yeast (S. cerevisiae) and
fission yeast (S. pombe). DNAs were separated on
agarose gels using CHEF electrophoresis. Gels
were stained with ethidium bromide (0.5 ¿xg/ml)
and fluorescence photographed with UV
transillumination. DNAs from the indicated
strains were run in 0.25X TBE, 0.6% agarose for
240 h at 40 V. Pulse switching times were ramped
from 30-70 min. Numbers at the left indicate
size standards in megabases (Mb).

117
Mb
4-6
3-5
2-2
cn
VO
O
CO
CQ
«0
p»
co
CO
CM
CO
CM
IT)
in
iH
T
T
O
VO
iH
n
(Ti
rH
CM
CM
I
1
l
l
1
CO
CM
1
1
1
1
1
1
0-
0
X
X
X
rH
CM
SC
SC
SC
X
X
»H
w
Figure D.5 Chromosome-sized DNAs from C. gloeosporioides
compared to those for yeast (S. cerevisiae) and
fission yeast (S. pombe). DNAs were separated on
agarose gels using CHEF electrophoresis. Gels
were stained with ethidium bromide (0.5 /xg/ml)
and fluorescence photographed with UV
transillumination. DNAs from the indicated
strains were run in 0.25X TBE, 0.6% agarose for
240 h at 40 V. Pulse switching times were ramped
from 30-50 min. Numbers at the left indicate
size standards in megabases (Mb).

118
Mb
5-7
4-6
3-5
2'2
o» r\i
n vo o
H VO (N I CM CO VO
r-v m | TTr-tOtTvOOVOn
u a i i a. i i u i s co cm i
W WXSt-JEKOSHr-ifMK
Figure D.6 Chromosome-sized DNAs from C. gloeosporioides
compared to those for yeast (S. cerevisiae) and
fission yeast (S. pombe). DNAs were separated on
agarose gels using CHEF electrophoresis. Gels
were stained with ethidium bromide (0.5 pg/ml)
and fluorescence photographed with UV
transillumination. DNAs from the indicated
strains were run in 0.25X TBE, 0.6% agarose for
168 h at 40 V. Pulse switching times were ramped
from 50-100 min. Numbers at the left indicate
size standards in megabases (Mb). Lanes H-l, H-
H-12, H-9, and 226802 DNA was partially degraded.

119
vo r*
> Uu o a i i
ME) to C/) SC X
av CQ vo O
in cm co
M 0\ (N O VO 'f
| I I 00 X X X «H 04 X
Figure D.7 Chromosome-sized DNAs from C. gloeosporioides
compared to those for yeast (5. cerevisiae) and
fission yeast (S. pombe). DNAs were separated on
agarose gels using CHEF electrophoresis. Gels
were stained with ethidium bromide (0.5 /ng/ml)
and fluorescence photographed with UV
transillumination. DNAs from the indicated
strains were run in 0.25X TBE, 0.6% agarose for
192 h at 40 V. Pulse switching times were ramped
from 40-100 min. Numbers at the left indicate
size standards in megabases (Mb). Lane H-4 DNA
was partially degraded.

120
Mb £
m
s
VO
i
(0
CM
m
m
L
o
rH
l
l
s
u
1
SC
SC
H
a
o
X
Figure D.8 Chromosome-sized DNAs from C. gloeosporioides
compared to those for yeast (S. cerevisiae) and
fission yeast (S. pombe). DNAs were separated on
agarose gels using CHEF electrophoresis. Gels
were stained with ethidium bromide (0.5 /ig/ml)
and fluorescence photographed with UV
transillumination. DNAs from the indicated
strains were run in 0.25X TBE, 0.6% agarose for
192 h at 40 V. Pulse switching times were ramped
from 40-100 rain. Numbers at the left indicate
size standards in megabases (Mb). Lanes H-l, H-3,
H-4, Maram1, H-12, DNA was partially degraded.

121
Mb
5-7
4-6
3-5
2 2
O
CM
VO
O
CQ
X
(N
CO
in
in
CM
00
O'
VO
O
VO
m
Oi
n
r—i
rH
'3-
1
CO
CM
i
l
1
i
i
1
1
l
l
0
a
X
rH
(N
X
33
33
33
52
X
X
X
w
w
Figure D.9 Chromosome-sized DNAs from C. gloeosporioides
compared to those for yeast (S. cerevisiae) and
fission yeast (S. pombe). DNAs were separated on
agarose gels using CHEF electrophoresis. Gels
were stained with ethidium bromide (0.5 /xg/ml)
and fluorescence photographed with UV
transillumination. DNAs from the indicated
strains were run in 0.25X TBE, 0.6% agarose for
264 h at 40 V. Pulse switching times were ramped
from 10-120 min. Numbers at the left indicate
size standards in megabases (Mb). This gel was
used in the Figure 4.6.

122
kb
1125
630
245
O VO g CQ
^ CO (N CO (ClDCNVOrH
Figure D.10 Chromosome-sized DNAs from C. gloeosporioides
compared to those for yeast (S. cerevisiae) and
fission yeast (S. pombe). DNAs were separated on
agarose gels using CHEF electrophoresis. Gels
were stained with ethidium bromide (0.5 /¿g/ml)
and fluorescence photographed with UV
transillumination. DNAs from the indicated
strains were run in 0.5X TBE, 1.0% agarose for
24 h at 200 V. Pulse switching times were ramped
from 50-90 s. Numbers at the left indicate size
standards in kilobases (kb).

123
CTl
CM
CO
O
a
a
VO
r>-
CD
CO
CM
00
(N
in
in
rH
'S'
O
VO
rH
r>
o\
rH
CM
CM
1
1
i
1
1
i
CO
CM
1
i
1
1
1
1
a
X
a
a
a
a
rH
CM
a
a
a
52
a
a
a
850
630
245
Figure D.ll Chromosome-sized DNAs from C. gloeosporioides
compared to those for yeast (S. cerevisiae) and
fission yeast (S. pombe). DNAs were separated on
agarose gels using CHEF electrophoresis. Gels
were stained with ethidium bromide (0.5 /xg/ml)
and fluorescence photographed with UV
transillumination. DNAs from the indicated
strains were run in 0.5X TBE, 1.0% agarose for
18 h at 200 V. Pulse switching times were ramped
from 30-60 s. Numbers at the left indicate size
standards in kilobases (kb).

124
Table D.l Estimated magabase sizes for chromosome size DNA
Chromosome number
Strain I II III IV V VI VII VII Figure
7.6
7.0
4.9
-
-
-
-
-
4.1
>5.7
-
4.5
3.7
3.3
-
-
-
4.2
-
-
-
-
-
1.1
0.63
-
4.3
-
-
-
-
-
-
0.63
-
4.4
-
-
-
-
-
1.1
0.63
-
4.5
7.6
-
4.8
-
-
-
-
-
D.2
7.6
7.1
4.7
-
-
-
-
-
D.3
>5.7
-
4.6
3.7
3.4
-
-
-
D.5
-
-
-
-
-
1.1
0.61
-
D. 10
-
-
-
-
-
-
0.61
-
D. 11
7.6
7.0
4.9
-
-
-
-
-
4.1
>5.7
-
4.5
3.7
3.3
-
-
-
4.2
-
-
-
-
-
1.1
0.63
-
4.3
-
-
-
-
-
-
0.62
-
4.4
-
-
-
-
-
1.1
0.63
-
4.5
7.7
7.1
4.8
3.7
-
-
-
-
D.l
7.6
-
4.8
-
-
-
-
-
D.2
7.6
7.1
4.7
-
-
-
-
-
D.3
>5.7
-
4.6
3.7
3.4
-
-
-
D.5
7.6
7.0
4.7
-
-
-
-
-
D. 6
7.7
7.0
4.7
3.8
_
—
—
—
D. 7

125
Table D.l— Continued
Strain
I
II
III
Chromosome number
IV V VI VII
VII
Figure
H-3
-
-
-
-
-
1.1
0.61
-
D.10
-
-
-
-
-
-
0.61
-
D.ll
H-4
7.9
4.9
-
-
-
-
-
-
4.1
7.8
4.8
-
-
-
-
-
-
D.2
7.8
4.7
-
-
-
-
-
-
D. 3
-
-
-
0.42
0.38
-
-
-
D.ll
H-9
7.6
7.0
4.9
-
-
-
-
-
4.1
>5.7
-
4.5
3.7
3.3
-
-
-
4.2
-
-
-
-
-
1.1
0.63
-
4.3
-
-
-
-
-
-
0.63
-
4.4
-
-
-
-
-
1.1
0.63
-
4.5
7.7
7.1
4.8
3.7
-
-
-
-
D.l
7.6
-
4.8
-
-
-
-
-
D.2
7.6
7.1
4.7
-
-
-
-
-
D. 3
>5.7
>5.7
4.7
3.7
-
-
-
-
D. 4
>5.7
-
4.6
3.7
3.4
-
-
-
D. 5
7.7
7.0
4.7
3.8
-
-
-
-
D. 7
-
-
-
-
-
1.1
0.62
-
D. 10
-
-
-
-
-
-
0.61
-
D.ll
H-12
7.9
4.9
-
-
-
-
-
-
4.1

126
Table D.l— Continued
Chromosome number
Strain I II III IV V VI VII VII Figure
H-12
>5.7
4.5
3.7
—
—
—
—
—
4.2
-
-
-
0.47
0.38
-
-
-
4.4
-
-
-
0.46
0.39
-
-
-
4.5
7.8
4.8
-
-
-
-
-
-
D.2
7.8
4.7
-
-
-
-
-
-
D. 3
>5.7
4.7
3.7
-
-
-
-
-
D.4
>5.7
4.6
3.7
-
-
-
-
-
D. 5
-
-
-
0.46
0.38
-
-
-
D.10
-
-
-
0.46
0.38
-
-
-
D. 11
>5.7
-
4.5
3.7
3.3
-
-
-
4.2
-
-
-
-
-
1.1
0.63
-
4.3
-
-
-
-
-
-
0.63
-
4.4
-
-
-
-
-
1.1
0.63
-
4.5
7.7
7.0
4.8
3.7
-
-
-
-
D.l
7.6
-
4.8
-
-
-
-
-
D.2
>5.7
>5.7
4.7
3.7
-
-
-
-
D.4
>5.7
-
4.6
3.7
3.4
-
-
-
D. 5
7.7
7.0
4.7
3.8
-
-
-
-
D. 7
-
-
-
-
-
1.1
0.62
-
D. 10
_
__
0.61
_
D. 11

127
Table D.l— Continued
Chromosome number
Strain I II III IV V VI VII VII Figure
H-3 6
—
—
—
—
—
0.77
0.63
—
4.3
-
-
-
-
-
0.75
0.63
-
4.4
>5.7
>5.7
4.7
3.7
3.3
-
-
-
D. 4
7.6
7.0
4.7
-
-
-
-
-
D. 8
-
-
-
-
-
0.77
0.63
-
D.10
7.9
4.9
-
-
-
-
-
-
4.1
>5.7
4.5
3.7
-
-
-
-
-
4.2
-
-
-
0.52
0.47
0.43
0.27
-
4.4
-
-
-
0.52
0.47
0.41
0.27
-
4.5
7.8
4.8
3.7
-
-
-
-
-
D.l
7.8
4.8
-
-
-
-
-
-
D.2
7.8
4.7
-
-
-
-
-
-
D.3
>5.7
4.7
3.7
-
-
-
-
-
D. 4
7.8
4.7
-
-
-
-
-
-
D. 6
7.8
4.7
-
-
-
-
-
-
D. 7
-
-
-
0.49
0.46
0.42
0.27
-
D. 10
-
-
-
-
-
-
0.27
-
D. 11
7.9
4.9
-
-
-
-
-
-
4.1
>5.7
4.5
3.7
_
_
—
—
—
4.2

128
Table D.l— Continued
Strain
I
II
Ill
Chromosome number
IV V VI VII
VII
Figure
H-48
-
-
-
0.46
0.43
0.40
-
-
4.4
-
-
-
0.46
0.43
0.40
-
-
4.5
7.8
4.8
-
-
-
-
-
-
D.2
7.8
4.7
-
-
-
-
-
-
D. 3
>5.7
4.7
3.7
-
-
-
-
-
D. 4
>5.7
4.7
3.7
-
-
-
-
-
D. 5
7.8
4.7
-
-
-
-
-
-
D. 7
-
-
-
0.46
0.43
0.40
-
-
D.10
-
-
-
0.45
0.43
0.40
-
-
D. 11
180269
7.9
4.9
-
—
—
—
—
4.1
-
-
-
0.43
0.41
-
-
-
4.4
-
-
-
0.45
0.41
0.39
-
-
4.5
7.8
4.8
-
-
-
-
-
-
D.2
7.8
4.7
-
-
-
-
-
-
D.3
>5.7
4.6
3.7
-
-
-
-
-
D.5
7.8
4.7
3.7
-
-
-
-
-
D. 6
7.8
4.7
3.8
-
-
-
-
-
D. 7
-
-
-
0.44
0.42
0.39
-
-
D. 10
—
-
-
0.43
0.41
0.39
—
—
D. 11

129
Table D.l— Continued
Chromosome number
Strain I II III IV V VI VII VII Figure
226802
Maram
OCO
7.9
4.9
—
—
—
—
—
—
4.1
>5.7
4.5
3.7
-
-
-
-
-
4.2
-
-
-
0.46
0.42
0.39
0.37
-
4.4
-
-
-
0.44
0.42
0.39
-
-
4.5
7.7
4.8
3.7
-
-
-
-
-
D.l
7.8
4.8
-
-
-
-
-
-
D.2
7.8
4.7
-
-
-
-
-
-
D. 3
>5.7
4.7
3.7
-
-
-
-
-
D. 4
>5.7
4.6
3.7
-
-
-
-
-
D. 5
7.8
4.7
-
-
-
-
-
-
D. 7
>5.7
-
4.5
3.7
2.8
-
-
-
4.2
-
-
-
-
-
-
0.63
-
4.3
-
-
-
-
-
-
0.63
-
4.4
>5.7
>5.7
4.7
3.7
2.8
-
-
-
D. 4
7.6
7.0
4.7
-
-
-
-
-
D.8
-
-
-
-
-
-
0.62
-
D. 10
>5.7
-
4.7
3.7
2.8
-
-
-
4.2
-
-
-
-
-
-
0.65
-
4.3
7.6
7.0
4.7
-
-
-
-
-
D. 6
7.6
7.0
4.7
_
—
—
—
D.8

130
Table D.l— Continued
Chromosome number
Strain I II III IV V VI VII VII Figure
IMB-3
>5.7
—
4.5
3.7
3.3
—
—
—
4.2
-
-
-
-
-
1.1
0.63
-
4.3
7.6
7.0
4.7
-
-
-
-
-
D. 6
7.6
7.0
4.7
-
-
-
-
-
D. 8
7.6
7.0
4.9
-
-
1.6
-
-
4.1
>5.7
-
4.5
3.7
3.3
1.6
-
-
4.2
-
-
-
-
-
-
0.63
-
4.3
-
-
-
-
-
-
0.63
-
4.4
-
-
-
-
-
-
0.63
-
4.5
7.7
7.1
4.8
3.7
-
1.6
-
-
D. 1
>5.7
>5.7
4.7
-
-
1.6
-
-
D. 4
>5.7
-
4.6
3.7
3.4
1.6
-
-
D.5
7.6
7.0
4.7
3.7
-
-
-
-
D. 6
7.6
7.0
4.7
-
-
1.6
-
-
D. 7
-
-
-
-
-
-
0.61
-
D. 10
_
_
_
0.60
D. 11

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BIOGRAPHICAL SKETCH
The author was born on November 5, 1958, in Galle, the
capital of southern Sri Lanka. He entered the University of
Peradeniya, Sri Lanka in 1978 and received a B.S. degree in
agriculture with honors in 1982. After four years of work as
an assistant lecturer in the University of Ruhuna, Sri
Lanka, he entered the University of Florida, Gainesville, to
pursue graduate studies. He received an M.S. degree in plant
pathology in 1989 and continued in the Ph.D program in
molecular plant pathology.
His goal is to continue research in molecular plant
pathology.
148

I certify that I have read this study and that in my
opinion it conforms to acceptable standards of scholarly
presentation and is fully adequate, in scope and quality, as
a dissertation for the degree of Doctor of Philosophy.
R. T. McMillan, Jr., Chair \
Associate Professor of Plant Pathology
I certify that I have read this study and that in my
opinion it conforms to acceptable standards of scholarly
presentation and is fully adequate, in scope and quality, as
a dissertation for the degree of Doctor of Philosophy.
/J. iLtj—
H. Corby Kistler, Cochair
Associate Professor of Plant Pathology
I certify that I have read this study and that in my
opinion it conforms to acceptable standards of scholarly
presentation and is fully adequate, in scope and quality, as
a dissertation for the degree of Doctor of Philosophy.
(
D. R. Pring
Professor of Plant Pathology
I certify that I have read this study and that in my
opinion it conforms to acceptable standards of scholarly
presentation and is fully adequate, in scope and quality, as
a dissertation for the degree of Doctor of Philosophy.
J.^W. Kitnbrough
Professor of Plant^Pathology
I certify that I have read this study and that in my
opinion it conforms to acceptable standards of scholarly
presentation and is fully adequate, in scope and quality, as
a dissertation for the degree of Doctor of Philosophy.
&
C.
L. C. Hannah
Professor of Horticultural Science

This dissertation was submitted to the Graduate Faculty
of the College of Agriculture and to the Graduate School and
was accepted as partial fulfillment of the requirements for
the degree of Doctor of Philosophy.
August, 1992
Dean, Graduate School

UNIVERSITY OF FLORIDA




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