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Disease resistance mechanisms in waterhyacinths and their significance in biocontrol programs with phytopathogens /

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Title:
Disease resistance mechanisms in waterhyacinths and their significance in biocontrol programs with phytopathogens /
Creator:
Martyn, Raymond DeWint, 1946-
Publication Date:
Copyright Date:
1977
Language:
English
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xvi, 204 leaves : ill. ; 28 cm.

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Subjects / Keywords:
Cells ( jstor )
Chloroplasts ( jstor )
Diseases ( jstor )
Enzymes ( jstor )
Fungi ( jstor )
Infections ( jstor )
Leaves ( jstor )
Pathogens ( jstor )
Phenols ( jstor )
Plants ( jstor )
Dissertations, Academic -- Plant Pathology -- UF
Plant Pathology thesis Ph. D
Water hyacinth -- Control ( lcsh )
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bibliography ( marcgt )
non-fiction ( marcgt )

Notes

Thesis:
Thesis--University of Florida.
Bibliography:
Bibliography: leaves 187-203.
General Note:
Typescript.
General Note:
Vita.
Statement of Responsibility:
by Raymond DeWint Martyn, Jr.

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University of Florida
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University of Florida
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Copyright [name of dissertation author]. Permission granted to the University of Florida to digitize, archive and distribute this item for non-profit research and educational purposes. Any reuse of this item in excess of fair use or other copyright exemptions requires permission of the copyright holder.
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03386351 ( OCLC )
AAB3883 ( NOTIS )

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DISEASE RESISTANCE MECHANISMS IN WATERHYACINTHS AND THEIR
SIGNIFICANCE IN BIOCONTROL PROGRAMS WITH PHYTOPATHOGENS










By

RAYMOND DEWINT MARTYN, JR.


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





UNIVERSITY OF FLORIDA


1977

































This above all: to thine own self be True


William Shakespeare
Hamlet; Act I, scene iii



























To my parents, who had the wisdom and foresight to
know the difference between "guidance" and "insistence",
and who used as one of the cornerstones of my education,
Robert W. Service's poem "The Quitter" which appears on
the following page .

To my wife, Dickie, whose unyielding faith and many
hours of unselfish help and patience were perhaps the
greatest factors in the completion of this program . .

To my daughter, Susan, whose 6-year-old smile made
it all worthwhile, when I overheard her tell a playmate,
"My Daddy is a plant doctor!"














The Quitter


When you're lost in the wild and you're scared as a child,
And death looks you bang in the eye;
And you're sore as a boil, it's according to Hoyle
To cock your revolver and die.
But the code of a man says fight all you can,
And self-dissolution is barred;
In hunger and woe, oh it's easy to blow --
It's the hell served for breakfast that's hard.

You're sick of the game? Well now, that's a shame!
You're young and you're brave and you're bright.
You've had a raw deal, I know, but don't squeal.
Buck up, do your damnedest and fight!
It's the plugging away that will win you the day,
So don't be a piker, old pard;
Just draw on your grit; it's so easy to quit --
It's the keeping your chin up that's hard.

It's easy to cry that you're beaten and die,
It's easy to crawfish and crawl,
But to fight and to fight when hope's out of sight,
Why, that's the best game of them all.
And though you come out of each grueling bout,
All broken and beaten and scarred --
Just have one more try, it's dead easy to die;
It's the keeping on living that's hard.


Robert W. Service










ACKNOWLEDGEMENTS

I wish to express sincere gratitude to Dr. Thomas E.

Freeman, Chairman of my Supervisory Committee, for his

friendship, advice, guidance, and patience during the course

of this study, and for his criticism and encouragement in

appropriate doses for three years which ultimately made this

dissertation possible.

I also wish to extend thanks to members of my Super-

visory Committee, Dr. T.E. Humphreys, Dr. H.H. Luke, Dr.

D.A. Roberts, and Dr. R.E. Stall for their advice and

friendship, and for their time spent in critical review of

this manuscript.

A special thanks is extended to Mr. D.A. Samuelson for

his many hours of assistance during the ultrastructural and

cytochemical portions of this study, and for the many hours

of help in preparing the electron micrograph plates.

Gratitude is also extended to Dr. H.A. Altrich for his

kindness for allowing use of equipment and facilities of the

Biological Ultrastructure Laboratory, and to Ms. Janet Plaut

for performing the many statistical analyses used throughout

this dissertation.

This research supported in part by the U.S. Army Corps

of Engineers, Florida Department of Natural Resources, U.S.

Department of Interior, Office of Water Resources and

Research Act as amended and by the University of Florida

Cerner for Environmental Programs.










TABLE OF CONTENTS


Page
ACKNOWLEDGEMENTS . . . . . . . . . v

LIST OF TABLES . . . ... . . . . . viii

LIST OF FIGURES . . . . . . . . . ix

ABSTRACT . . ... . . . . . . xiii

GENERAL INTRODUCTION . . . . . . . . 1

Part I The Aquatic Weed Problem . . .. 1
Part II The Potential of Biological Control 5
ParT III Pathogens of Waterhyacinth with
Possible Biocontrol Potential . 8

CHAPTER I RESPONSES OF WATERHYACINTH TO INFECTION
WITH ACREMONIUM ZONATUM AND ITS IMPLI-
CATIONS IN BIOLOGICAL CONTROL . ... 15

Introduction . .. . ..... . 15
Materials and Methods . . . .. 17
Results . . ... . . . . 19
Discussion . . . ... . . . 28

CHAPTER II A CYTOCHEMICAL AND ULTRASTRUCTURAL STUDY
OF THE PHENOL CELLS AND POLYPHENOLOXI-
DASE ACTIVITIES IN HEALTHY AND DISEASED
WATERHYACINTH LEAVES . . . . .. 37

Introduction . . . . . . . 37
Materials and Methods . . . . 43
Results . .. . . .. . 49
Discussion . . . . . . . 84

CHAPTER III A BIOCHEMICAL STUDY OF THE PHENOLIC
ACIDS AND POLYPHENOLOXIDASE RATES IN
HEALTHY AND DISEASED WATERHYACINTH
LEAVES . ... . . . . . . 90

Introduction . . . . . . .. 90
Materials and Methods . . . .. 100
Results . . .. . .. .. . 108
Discussion .... . . . . . . 129










Page


CHAPTER IV AN ULTRASTRUCTURAL STUDY OF PENE-
TRATION AND COLONIZATION OF WATER-
HYACINTH BY ACREMONIUM ZONATUM ... . 139

Introduction .. . . . . . . 139
Materials and Methods . .. . 141
Results . . . . .. . . 144
Discussion . . . . . . 169

SUMMARY AND CONCLUSIONS . . ... .. . . 179

LITERATURE CITED .. . . . ... . . 187

BIOGRAPHICAL SKETCH . . ... . . . . 204










LIST OF TABLES


Table Page

III-1 Free phenolic acids detected in healthy
and A. zonatum-infected waterhyacinths by
thin layer chromatography . . . . .. 114

III-2 Phenolic acids detected in healthy water-
hyacinth leaves by thin layer chromatogra-
phy and various locating reagents after
alkaline hydrolysis . ... . . . . 115

11-3 Phenolic acids detected in A. zonatum-in-
fected waterhyacinth leaves by thin layer
chromatography and various locating rea-
gents after alkaline hydrolysis . . .. 116

II-4 R values and color characteristics of
tne phenolic acids detected in healthy
and A. zonatum-infected waterhyacinth
leaves after alkaline hydrolysis . . . 117

III-5 Growth of A. zonatum on healthy and A.
zonatum-infected waterhyacinth leaf-
extract media . . ... . . . . . 124

TII-6 Growth of A. zonatum on phenolic acid
media . . . . . ... . . .. 125

III-7 Growth of A. zonatum on phenolic acid
media with yeast extract . . . ... 126

S-1 Differences and similarities among
healthy and A. zonatum-infected water-
hyacinth morphotypes . ... ....... 181


viii










LIST OF FIGURES


Page

CHAPTER I

Fig. I-I Symptoms of disease on water-
hyacinths incited by Acremonium
zonatum . . . ..... 23

Fig. 1-2 Quantitation of disease on small,
medium, and large waterhyacinths 25

Fig. I-3 Quantitation of leaf regeneration
rates of small, medium, and large
waterhyacinths .. . ... . . 27

CHAPTER II

Fig. II-1 Biosynthetic pathway for conver-
sion of tyrosine to melanin . 42

Fig. II-2 Flow diagram of procedure for
standard electron microscopy fi-
xation and embedding . . .. 45

Fig. 11-3 Flow diagram of procedure for the
cytochemical localization of po-
lyphenoloxidase . . ... ... 48

Fig. 1-4 Light micrographs of phenol cells
in healthy waterhyacinth leaves 57

Fig. II-5 Number of phenol cells/mm2 leaf
area in small, medium, and large
waterhyacinth leaves . . . 59

Fig. II-6 Electron micrograph of phenol
cell in palisade cell layer of
waterhyacinth leaf tissue . .. 61

Fig. 11-7 Electron micrograph of phenol
cell in vascular tissue area of
waterhyacinth leaf . . . 63

Fig. II-8 Chloroplasts of healthy waterhya-
cinth leaf tissue incubated with-
out DOPA . . . . . . 65










Page


Fig. II-9 Localization of polyphenoloxidase
in healthy waterhyacinth leaf
tissue without lead postaining


Fig. II-10




Fig. II-11




Fig. II-12



Fig. II-13



Fig. II-14



Fig. II-15




Fig. II-16




Fig. II-17.


CHAPTER III


Fig. III-1


Fig. III-2


Localization of polyphenoloxidase
in chloroplasts of xylem paren-
chyma cells in healthy waterhya-
cinth leaves ....

Localization of polyphenoloxidase
in chloroplasts of bundle sheath
cells in healthy waterhyacinth
leaves . . . . . . .

Localization of polyphenoloxidase
in chloroplasts of phenol cells
in healthy waterhyacinth leaves .

Chloroplasts of boiled, healthy
waterhyacinth leaf tissue incu-
bated with DOPA . . . . .

Chloroplasts of healthy waterhya-
cinth leaf tissue incubated in
inhibitor (DDC) and DOPA ..

Localization of polyphenoloxidase
in chloroplasts of palisade cells
from diseased waterhyacinth
leaves . . . . .

Localization of polyphenoloxidase
in chloroplasts of spongy meso-
phyll cells from diseased water-
hyacinth leaves . . . . .

Localization of polyphenoloxidase
in chloroplasts of cells several
centimeters away from infection
center . . .



Principal phenolic acids found in
plants . . . . . .

Shikimic acid pathway for the
biosynthesis of monocyclic phe-
nols and major derivatives










Page


Fig. III-3



Fig. III-4



Fig. III-5



Fig. III-6


Flow diagram of procedure for ex-
traction of ester-linked phenols
in plants . . . . . .

Total phenol concentrations in
healthy and A. zonatum-infected
waterhyacinth morphotypes ..

Polyphenoloxidase activities in
small, medium, and large healthy
waterhyacinth leaves ..

Polyphenoloxidase activities in
small, medium, and large diseased
waterhyacinth leaves ..


Fig. III-7 In vitro synthesis of indoleace-
tic from tryptophan by Acremonium
zonatum . . . . . . .

CHAPTER IV


Flow diagram for testing of car-
bohydrate degrading enzymes pro-
duced by Acremonium zonatum .


Fig. IV-2 Penetration of waterhyacinth leaf
by Acremonium zonatum .. ...


Fig. IV-3a



Fig. IV-3-c



Fig. IV-4a


Fig. IV-4b




Fig. IV-5


Cross-section of Acremonium zona-
tum observed in xylem tissue of
diseased waterhyacinth leaf .

Degradation of wall material in
waterhyacinth by Acremonium
zonatum . . . . . . .

Attachment of Acremonium zonatum
to the cuticle ....

Attachment of Acremonium zonatum
to epidermis and the possible
area of localized enzyme secre-
tion . . .

Penetration of phenol cell by
Acremonium zonatum ...


Fig. IV-1










Page


Phenol cell invaded by Acremonium
zonatum . . . . . . .

Breakdown of starch reserves in
chloroplasts during disease . .


Fig. IV-8 Increase in the number of plasto-
globuli in chloroplasts during
disease . . . . . . .


Increase in the number of micro-
bodies in cytosol as a result of
infection with Acremonium zonatum

Destruction of chloroplast integ-
rity during later stages of
disease . . . . . . .

Diseased palisade cell showing
extent of necrosis and cellular
breakdown . . . . . .


Fig. IV-6


Fig. IV-7


Fig. IV-9a



Fig. IV-9b



Fig. IV-10










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


DISEASE RESISTANCE MECHANISMS IN WATERHYACINTHS AND THEIR
SIGNIFICANCE IN BIOCONTROL PROGRAMS WITH PHYTOPATHOGENS


By

Raymond DeWint Martyn, Jr.

June, 1977

Chairman: Dr. Thomas E. Freeman
Major Department: Plant Pathology

The pathological relationship between the floating

waterhyacinth, Eichhornia crassipes (Mart.) Solms and the

fungus, Acremonium zonatum (Sawada) Gams, was investigated to

determine possible disease resistance mechanisms in the plant

as they relate to potential biocontrol agents. Waterhyacinths

were separated into three morphotypes based upon their leaf

surface area; small plants (leaves < 15 cm ), medium plants
2 2
(leaves 15-40 cm ), and large plants (leaves > 40 cm ) and

used for quantitating symptoms of disease. Inoculated small

plants exhibited fewer lesions/leaf after two weeks than did

either medium or large plants; however, the total percent

diseased leaf area for each morphotype was the same (approxi-

mately 40%). It was observed that large plants regenerated

almost three times as many new leaves after infection deve-

lopment than did either medium or small plants.

Biochemical, histochemical, cytochemical, and ultra-

structural studies were conducted on both healthy and diseased


xiii










morphotypes to determine what role host phenolic com-

pounds had in disease development. Phenolic compounds in

waterhyacinth leaves are localized in specialized idioblasts

(phenol cells) immediately beneath both epidermal surfaces

and also in close association with the vascular tissue. The

concentration of phenol cells increased significantly from

a mean of 33.6/mm2 leaf area in small plants to 48.7/mm2

in large plants.

In healthy plants, polyphenoloxidase (PPO) activity was

greater in small than in large leaves and was restricted to

the thylakoids of chloroplasts in only three cell types:

vascular parenchyma, bundle sheath, and phenol cells. After

infection by A. zonatum, PPO activity decreased in small

leaves but increased over 300% in large leaves. After

infection, PPO activity was observed in all chloroplasts

throughout the leaf.

Chlcrogenic acid was the only free phenolic acid found

in norphotypes of both healthy and diseased plants. Alka-

line hydrolysis of healthy leaf tissue yielded six phenolic

acids from small and medium plants and nine from large

plants. After infection, one additional phenolic acid was

detected from small- and medium-sized leaves. No change in

the types of phenolic acids present in large leaves was

detected after infection. The concentration of total phenols

in healthy plants increased significantly from 92 pg/g fresh

leaf tissue in small to 104 pg/g in large leaves. There was










a significant decrease in total phenols in both small

and medium diseased plants while the concentration remained

constant in large diseased plants.

Acremonium zonatum grew significantly better when cul-

tured on minimal media containing phenolic acids than it did

on media without these compounds. Acremonium zonatum was

inhibited by p-coumaric acid at 1000 ppm, when yeast extract

was added as a growth supplement to the media. In addition,

growth of the fungus on diseased plant-extract media was

stimulated significantly over growth on media containing

extracts from healthy plants.

Penetration of waterhyacinth leaves by A. zonatum

occurred directly through the cuticle or through the sto-

mata. Cellular penetration was aided by the production of

cellulolytic enzymes. Penetration of the phenol cells re-

sulted in death of the invading hyphae. Associated with

disease was the disappearance of starch granules from the

chloroplast, an increase in the number of plastoglobuli

within chloroplasts, and a build-up of microbodies within the

cytosol.

The results presented in this study suggest that phenol

metabolism in waterhyacinth plays a significant role in the

defense against potential pathogens and may account for why

only a few of pathogens have been reported on this plant.

It appears that A. zonatum is capable of causing relatively









severe damage to the waterhyacinth because of its high

tolerance to phenols and warrants continued study as a

potential biocontrol of this noxious aquatic plant.














GENERAL INTRODUCTION


Part I: The Aquatic Weed Problem

All plant and animal species in their native habitats

are subject to natural forces that control their population

levels. Natural enemies along with other environmental

influences maintain a balance among populations of plants

and animals in an ecosystem. There is little question that

the parasites and predators existing in a particular system

are the greatest resource that we have for effective pest

suppression and management (180).

Man steps beyond Nature's boundaries, however, and

thereby sidesteps natural controls by transporting plant

and animal species to new habitats, and in so doing, often

causes disastrous shifts in the ecological balance between

species. Such has been the case with many of the noxious

aquatic plants in Florida. Exotic water plants imported into

this country as aquaria specimens and ornamentals have escaped

into lakes and waterways and, once established, have created

serious control dilemmas. In areas where aquatic plants have

reached high densities, they greatly obstruct the water flow,

dec'.- se the water level through increased rates of evapo-

ration ,nd transpiration, increase the rate of eutrophication,










interfere with navigation, prevent fishing and other water

recreational activities, depress real estate values, and

may, in some instances, present severe health hazards

(52, 75, 201). Infamous examples of these pestiferous

plants include the floating waterhyacinth, Eichhornia cras-

sipes (Mart.) Solms, Florida elodea, Hydrilla verticillata

(Casp.), Eurasian watermilfoil, Myriophyllum spicatum L.,

and alligatorweed, Alternanthera philoxeroides (Mart.)

Griseb.

The rampant growth of exotic water weeds in Florida and

other Gulf states has been attributed to several factors

(78, 118, 139). First, the year-round warm temperature and

extended photoperiod combine to give a growing season

almost the entire year. Secondly, many bodies of water

provide an abundance of inorganic compounds necessary for

luxuriant plant growth. Thirdly, the absence of enemies

normally present in their native habitats does not allow the

natural system of checks and balances to operate. And,

lastly, most aquatic plants are capable of extremely rapid

vegetative reproduction. It is for these reasons that some

160,000 hectares of Florida's fresh water are weed-choked

(5 ).

One of the most Destiferous aquatic plants in tropical

and subtropical climates is the floating waterhyacinth,

E. crassipes, the subject of this dissertation. The










genus Eichhornja is a member of the Pontederiacae family

and includes four other species: E. paniculata, E. paradoxa,

E. azurea, and E. diversifolia (139). Eichhornia crassipes

is the only species which is free floating; all other

members of the genus are rooted either in shallow water

or near shore.

The waterhyacinth reproduces almost entirely by vegeta-

tive means although sexual reproduction does occur. It

reproduces rapidly and will completely fill many lakes and

rivers in a single growing season. Pcnfound and Earle (139)

reported that E. crassipes is capable of doubling its mass

every 11-15 days. Taking an average rate of doubling of

two weeks and a growing season of eight months, then ten

plants given plenty of room and good growing conditions

would produce 655,360 Apants which would cover 0.6 hec-

tares. These figures emphasize the tremendous rate of

colonizaticn of this species and the necessity of good

Cntrcl methods.

It is believed that the waterhyacinth is a native of

Brazil, but has spread from there to nearly all of the

South American and Central American countries and through-

out the world where the climate is favorable for its

development. Few tropical or subtropical countries are free

fror waerrhyacinrhs (97).

The accounts differ somewhat regarding its appearance

in the LUnted States. There is some evidence that it was










cultivated as a greenhouse exotic shortly after the War

Between the States (139); however, the earliest authentic

account details its introduction at the Cotton Centennial

Exposition at New Orleans in 1884 (88). It appeared in

Florida in 1890 (190) and has since become an important

aquatic pest. By the turn of the century it was reported

from all the southeastern coastal states as far north as

Virginia and westward to California (81).

Eichhornia crassipes was officially recognized as a

serious aquatic pest in this country on June 4, 1897, when

Congress passed an act authorizing the Secretary of War to

investigate the extent of obstruction to navigation in the

waters of Florida and Louisiana (139). Since that time, the

U.S. Army Corps of Engineers have been responsible for

clearing it from navigable waterways.

Florida, like many parts of the United States and

world, is in dire need of an efficient and effective means

of controlling noxious aquatic plants. Since their introduc-

tion, millions of dollars, both tax and private, have been

spent on chemical and mechanical control of these weeds. An

estimated 10 to 15 million dollars is being spent per year

for the control of aquatic weeds in Florida alone, and this

figure is increasing every year (64). Despite this huge

financial expenditure, the total infestation continues to

grow and at present there is no end to the increasing costs

unless new control measures are found.









Fart II: The Fotential of Biological Control

In past years, control of waterweeds has been based on

two basic procedures. Both mechanical and chemical controls

are used routinely in maintenance programs. However,

neither method on its own is completely satisfactory and the

weed infestations continue to expand. More recently, the

concept of biological control was proposed for aquatic

weeds. Huffaker and Andres (78) have stated that any or-

ganism which curtails plant growth or reproduction may be

used as a biological control agent. Such could potentially

include animals either higher or lower than insects, and

parasitic higher plants, fungi, bacteria, and viruses. For

this reason the term biological control organism, or agent

is used -o include all suitable phytophagous animals and

plant pathogens on a given weed.

It was generally believed that biological control works

best with agents of foreign origin (75); however, as

Wapshere (188) points out, successful biocontrol with an

organism in one counTry does not necessarily imply that the

organisms) used will be successful elsewhere. For instance,

Chrysclina quadrigemina was relatively ineffective against

Hypericum perforatum in Australia, but bee les of the same

genetic srock were highly successful against the same weed

in California, apparently because of more suitable climatic

conditions there (188).










Many investigations have been undertaken to study

potential uses of macrobiological agents to control noxious

aquatic weeds. In most instances, these studies have

involved insects (13,78,105,165,199) and, to a lesser

extent, other animals (33,39,118,162,164).

Of the insects screened for possible control agents,

one of the most effective appears to be the flea beetle,

Agasicles hygrophila which feeds only on alligatorweed (13,

105, 199). It was successfully introduced into the United

States from Argentina for the control of alligatorweed (13).

In April, 1965, 266 adult beetles were released near Jack-

sonville, Florida, and by June, 1966, there were hundreds of

thousands of them present at the release sites and most of

the floating alligatorweed was dead (105). It has since

spread rapidly throughout the watersheds in northeast

Florida (199). Insects alone, however, are not likely to

control aquatic weed pests because there are relatively few

phytophagous species capable of living beneath the water

(201).

Other biological control agents being investigated

include phytopathogenic fungi, bacteria, and viruses.

Zettler and Freeman (201) list four advantages of using such

control agents: (i) control applications would presumably

require minimal technology and, if successfully established,

the pathogen in theory would be selfmaintaining; (ii) the










overwhelming number of different plant pathogenic species

from which to choose offers an unmatched versatility in

selecting a specific biological control; (iii) virtually

none can attack man or his animals, therefore providing an

important advantage over the use of various animals such as

snails, which may harbor vertebrate pathogens, and (iv)

plant pathogens, although often killing individuals in a

given population, would not be expected to cause the exter-

mination of a species. This last attribute is important

because eradictaion of one aquatic weed species, such as the

waterhyacinth, may create an ecological void that in turn

may allow a population explosion of a different and more

serious species. In addition, Wilson (192) points out three

more advantages of using biological control agents over

chemical control procedures: (i) they can be specific to

the target weed which lessens the chance of damage to

cultivated or desired species, (ii) residue and toxicity

problems created by herbicides would be greatly reduced or

eliminated altogether, and (iii) there would be no accumu-

lation of the herbicide in the soil or underground water.

In essence, then, the use of biocontrol agents has many

advantages over chemical control methods and warrants

continued research.

The use of plant pathogens is not without hazards. Any

study undertaken to introduce or test phytopathogens must










be done with extreme care. Well controlled and monitored

prerelease experiments, however, can greatly reduce any

potential dangers.





Part III. Pathogens of Waterhyacinth with Possible Biocontrol
Potential

The first recorded disease on waterhyacinth caused by a

fungus was reported in 1917 by Tharp (174). He described a

Cercospora sp. as occurring on Piaropus crassipes (= E.

crassipes) in Texas and subsequently identified the causal

agent as C. piaropi Tharp. Thirty-seven years later, in

1954, it was reported on waterhyacinth in India (175) and

was again reported from the United States in 1974 (53).

The disease symptoms are oval leaf spots, 1.5 4.0 mm

in size, on the distal portion of the leaf blade. As with

other leaf spot diseases reported on waterhyacinth (2, 154),

C. piaropi does not appear severe enough to retard the

prodigious growth of the plant significantly; however, its

host specificity enhances its potential as a biocontrol

agert and is being investigated further (53).

The second recorded disease on waterhyacinth was caused

by a rust fungus, Uredo eichhorniae, found in the Dominican

Republic in 1927 (27). A year later, Ciferri (26) reported

the occurence of a smut, Doassansia eichhorniae on E.

crassipes from the same area. Neither of these organisms,










however, had been studied as potential biocontrol agents

until last year (25).

In 1932, a species of Fusarium was reported on water-

hyacinth from India (2). It caused reddish-brown necrotic

spots and streaks on both sides of the petioles and the

infected plant parts gradually shriveled up. The disease

caused only slight injury and the plant rapidly regenerated

new leaves and petioles. This is possibly the first pub-

lished paper concerned with phytopathogens as controls for

waterhyacinth as indicated by the authors' concluding state-

ment:

The infection takes place readily, but
owing to the high resisting power of the
plant, the disease makes very slow pro-
gress. From this it may be inferred
that this fungus cannot be regarded as a
possible remedy against the spread of
waterhyacinth (2).

Ten years later, Banerjee (7) identified the causal

agent as F. equiseti and Snyder and Hansen (169) reduced

this species to synonymy with F. roseum. A recent survey of

Florida for diseases of waterhyacinth resulted in the

isolation of this same species (F. roseum) from diseased

plants in Lake Griffin near Leesburg (154). This report was

the first of a F. roseum isolate affecting waterhyacinth in

the western hemisphere. The disease is characterized by

chlorosis and vascular discoloration in advance of necrosis

which proceeds towards the leaf tip. The leaf spot, however,










did not expand over the entire leaf surface but remained

localized. This is in line with that described by Agharkar

and Banerjee in their original report (2).

In 1946, Padwick (133) reported two species of fungi

pathogenic to waterhyacinth. The first, Rhizoctonia solani

(Corticum solani), was isolated near Dacca, Bengal, from

infected leaves and petioles. It caused extensive blotching

and streaking, often killing individual plants. Some 20 years

later, R. solani was again reported on waterhyacinth from

India by Nag Raj and Ponnappa (124).

During surveys for phytopathogens in the Canal Zone of

Panama, Freeman and Zettler isolated a R. solani from the

anchoring hyacinth (E. azurea) which proved to be extremely

pathogenic on the floating hyacinth (56). In addition,

sclerotia of this fungus were able to maintain their viability

without loss of virulence after being submersed in lake water

for 26 months (56). Disease symptoms on E. crassipes were

severe blighting of the enersed portions of the plant which

frequently resulted in death of the entire plant. Although

R. solani is an aggressive pathogen of waterhyacinth, it

cannot be considered as a biocontrol at this time, because

of its wide pathogenicity to a number of economically

important hosts (133).

The second fungal species reported by Padwick (133) was

Cephalosporium eichhornae Padwick sp. nov. It induced









large, oval, buff-colored spots on the leaves which were

covered with a white mat of mycelium. In 1973, Rintz (153)

reported another Cephalosporium species, C. zonatum, as

causing a zonal leaf spot disease of waterhyacinth in

Louisiana and Florida. His report was the fourth pathogen

described as occurring on waterhyacinth in the United

States. There was some discrepancy as to the synonomy of

these two Cephalosporium species (162) and the Commonwealth

Mycological Institute reduced them to synonomy, with C.

zonatum being the preferred name (123, 153). However,

several years later, C. zonatum was reclassified and is

presently placed in the form genus Acremonium of the class

Hyphomycetes (86). It is this fungus, Acremonium zonatum

(Sawada) Gams, which was studied as a biocontrol agent for

waterhyacinths in the present paper.

A concentrated research program on biological control

of aquatic weeds at the Indian Station of the Commonwealth

of Biological Control in Bangalore has resulted in the

isolation of several species of phytopathogenic fungi. In

1965, Nag Raj (122) reported a thread blight of waterhya-

cinth occurring in Calicut, India. Subsequent isolations

showed the fungus Marasmiellus inoderma (Berk.) Sing. to be

the causal agent (122). The diseased plants in the field

exhibited necrotic areas on the leaves, petioles, and all

aerial Darts. The infection was more evident in dense









stands of the weed and death of individual plants occurred

in irregular patches (122). Infection by M. inoderma under

laboratory conditions spreads very rapidly on host plants

which is a distinct advantage for a potential biocontrol

agent.

In 1970, Ponnappa (142), working at the same Indian

laboratory, isolated the fungus Myrothecium roridum from

waterhyacinth. Although this organism caused extensive

damage to E. crassipes, its usefulness as a biocontrol agent

cannot be considered at this time because of its patho-

genicity on a number of important economic crops (142).

This fungus was also reported on waterhyacinth from India by

Charudattan in 1973 (21).

One fungus which appears to have good potential as a

control agent is Alternaria eichhorniae, isolated and

described by Nag Raj and Ponnappa (125). It was isolated in

India in 1970 and was proved the causal agent of a leaf

blight disease. Leaf spots frequently covered the majority

of the leaf and caused premature death of those leaves. In

culture, A. eichhorniae produces a bright-red diffusable

pigment which deepens with age. In addition, it also

produces a host-specific toxic matabolite that causes

necrotic lesions when placed on leaves or petioles. The

host range of this fungus was tested on 42 genera of plants

in 15 families including aquatics and such important










terrestrial families as Brassicaceae, Fabaceae, and Sola-

naceae. The results showed A. eichhorniae to be non-

pathogenic on all plants tested except the waterhyacinth

(125). Its host specificity along with its specific toxic

metabolite enhances its potential as a biocontrol agent.

A similar species of this fungus was isolated in 1973

by McCorquodale, Martyn, and Sturrock (113) from water

hyacinth in south Florida and tentatively identified as A.

eichhorniae var. floradana (114). It resembled that de-

scribed by Nag Raj and Ponnappa (125) in host specificity,

conidial size, and toxin production, but differed in pigment

production and gemmae formation. This is the first report

of this species in the United States.

Tests indicate that A. eichhorniae has good potential

as a biocontrol agent of waterhyacinth, but because it is

not indigenous to the United States it is under strict

quarantine by the U.S. Department of Agriculture. For this

reason, A. eichhorniae cannot be adequately field tested in

Florida at the present time.

A second Cercospora species, C. rodmanii was isolated

from diseased waterhyacinth in 1973 in Florida (55) and is

currently being evaluated as a biocontrol agent

(30). Symptoms of the disease on waterhyacinth include

general chlorosis of the plant, failure to produce off-

shoots, spindly petioles and a root rot. Field trials









indicated that the fungus greatly reduced the waterhyacinth

population in test plots, but did not eradicate it since new

growth appeared which continued to spread (30).

In summary, among the phytopathogens reported on

waterhyacinth, some are capable of inducing severe damage

and even death of the plant. The fact remains, however,

that there are relatively few capable of causing such severe

diseases. Most of those that do, however, are also patho-

genic to important cash crops and therefore unacceDtable as

biocontrol agents at the present time. Consequently, it

would be a great advantage if one or more of the pathogens

with a narrow or restricted host range could be utilized.

With this in mind, the intent of this study was to examine

the pathological relationship of E. crassipes and A. zonatum

in an effort to more fully understand the basis of disease

resistance and pathogenesis in this host-parasite couplet.














CHAPTER I
RESPONSES OF WATERHYACINTHS TO INFECTION WITH ACREMONIUM
ZONATUM AND ITS IMPLICATIONS IN BIOLOGICAL CONTROL



Introduction

Research into biological control of noxious aquatic

plants was initiated at the University of Florida, Depart-

ment of Plant Pathology, in 1970. Major emphasis was placed

on finding diseases of waterhyacinth, alligatorweed, hydril-

la, and Eurasian watermilfoil. Surveys for diseases of

these plants were made throughout Florida and portions of

Alabama, Maryland, Louisiana, Georgia, South Carolina, the

Chesapeake Bay, and the Tennessee Valley areas (55).

Surveys were also made in ten other countries including most

of the Caribbean and eight states in India (55). During

these surveys, several diseases were found and the causal

agents isolated for further study (22,23,24,83,95).

In 1971, a zonal leafspot of waterhyacinth was first

noted in Fuerto Rico where it caused considerable damage to

the plant (55). The causal organism was not isolated.

However, a similar disorder was subsequently found in the

Spring Bayou region of Louisiana. A species of the fungus,

Cephalosporium, was isolated from those plants, and upon

inoculation onto healthy plants, induced symptoms typical










of those observed under natural conditions. The causal

agent was ultimately identified as Cephalosporium zonatum

Sawada (153) and was originally described as the causal

agent of zonal leafspot disease of figs in Louisiana (177).

This disease was found since to occur on waterhyacinths in

El Salvador, India, Panama, and at two locations in Florida

(55). The causal agent, Cephalosporium zonatum, (Sawada)

recently was reclassified to Acremonium zonatum (Sawada)

Gams (86).

The disease is first evident as small sunken lesions on

both leaf surfaces and the petiole (153). Under conditions

of high humidity, A. zonatum causes severe spotting and

death of leaves (107). The lesions are characteristically

zonate, oval to irregular in shape, and often coalesce

covering the entire surface. Alternating light- and dark-

brown bands are typical of the lesions. Under conditions of

prolonged high humidity the fungus produces abundant white

mycelia on the leaf surfaces and sporulates intermittently.

Rintz (153) reported that A. zonatum can attack a wide

range of plants under artificial conditions. Despite this

apparent wide host range, reports of its occurrence on hosts

other than fig in North America are unknown. Consequently,

this fungus need not necessarily be excluded from consi-

deration as a possible biocontrol agent of waterhyacinth

(55).










During field trials with this fungus in Gainesville, it

was observed that small, young plants displayed fewer

lesions after infection than did larger plants in the same

plot. (T.E. Freeman, personal communication, 1974). In

addition, it was observed that some of the infected plants

appeared to produce more new leaf growth than did either

other diseased plants or control plants. The present study

was initiated to determine if small plants were more resis-

tant to A. zonatum than large plants and also if there was

an accelerated leaf regeneration in response to infection.



Materials and Methods

Quantitation of disease

Waterhyacinths were collected from natural infestations

in south Florida and maintained under greenhouse conditions

in Gainesville. Plants were separated into three size cate-

gories based upon leaf surface area: (i) small plants, with

leaves less than 15 cm (ii) medium plants, with leaves 15-
2
40 cm and (iii) large plants, with leaves greater than 40
2
cm The plants were inoculated by swabbing the leaves with

a 10% (wt./vol.) slurry of A. zonatum (grown on potato

dextrose agar) and 0.75% water agar. Plants were maintained

in ten-gallon glass aquaria half-filled with tap water with

plastic covers to maintain the humidity at 99-100%. Control

plants were inoculated with sterile 0.75% water agar and


I










maintained under identical conditions. Two weeks post-

inoculation, leaves were excised and used for subsequent

tests.

Twenty-five to seventy-five leaves from each plant size

group were removed and the number of lesions/leaf counted.

Mean percentage figures were determined for (i) number of

leaves with one or more lesions/leaf, (ii) number of leaves

with ten or more lesions/leaf, and (iii) mean number of

lesions/ leaf. The total diseased area on each leaf was

calculated by the dot counting method ("Stippentelplaatje",

J.C. Zadoks, unpublished) and the mean percent diseased area

determined for each plant size.

Quantitation of leaf regeneration

Plants from each size category were selected at random,

trimmed of any necrotic or senescent leaves, and inoculated

as before. The total number of leaves on each plant was

noted prior to inoculation. The plants were maintained in

ten-gallon glass aquaria half-filled with tap water and

fitted with plastic covers as before. Control plants from

each size category were painted with 0.75% sterile water

agar and maintained under identical conditions. After two

weeks, the total number of leaves on each plant was counted

and percent new leaf growth figure was calculated for each

plant size group.

In addition, ren plants from each size category were










selected and the number of leaves/plant noted. Each leaf

was then excised and the plants placed in aquaria. After

two weeks, a percent new leaf growth figure was determined

for each plant size.



Results

Quantitation of disease

Inoculated waterhyacinths kept under conditions of high

humidity were severely damaged by A. zonatum (Fig. I-1).

Necrotic lesions varying in size and number occurred on both

leaf surfaces and the petioles. On the average, 70.1% of

the leaves on small plants had at least one lesion while

98.2% of medium leaves had at least one lesion (Fig. 1-2).

Large plants had 133% of their leaves infected indicating a

secondary spread to the new growth during the course of

disease development. These percentages decreased when the

number of leaves exhibiting ten or more lesions was calcu-

lated but the trend was the same. Large plants had signifi-

cantly more leaves with ten lesions (83.3%) than did either

medium (53.6%) or small (29.8%) plants. The average number

of lesions/leaf also followed the same pattern. Small

plants averaged 3.7 lesions/leaf while medium and large

averaged 12.8 and 18.3 respectively. However, when the

total diseased leaf area was measured after two weeks, there

was no significant difference among small, medium, and large










plants, each exhibiting approximately 40% diseased leaf area

(Fig. I-2).

Quantitation of leaf regeneration

On the average, after two weeks of growth, small

healthy waterhyacinths regenerated 27.3% new leaves or one

new leaf/plant (Fig 1-3). There was no significant diffe-

rence at the 0.01 confidence level in the percentage of new

leaves produced by small diseased plants. Small plants,

after infection, regenerated 21.6% new leaves or 0.95

leaves/plant. Likewise, there was no significant difference

between the new leaves produced by healthy and diseased

medium-sized plants. Medium-sized control plants produced

28.5% new leaves during the two weeks while diseased plants

of the same size regenerated 33.9% new leaves. A trend was

noted, however, that as the plant increased in size, its

rate of new leaf production also increased.

When the number of new leaves produced by large healthy

plants was compared to that from large diseased plants,

there was a significant difference. Large healthy plants

normally regenerated 46.1% of their leaves over the two week

period; however, diseased large plants produced 93.3% new

leaves, an increase of almost 50% (Fig. 1-3). In addition,

the average number of leaves/plant for large plants in-

creased from 2.0 in healthy to 4.6 in diseased plants.

Small and large plants which had their leaves excised










prior to the test displayed little variation in the number

of new leaves when compared to the controls (Fig. 1-3).

Small plants regenerated 25.2% of their leaves compared to

27.3% new growth in normal small plants. Likewise, large

plants displayed little difference in new leaf production

between control plants and those in which the leaves were

excised (46.1% and 50.0% respectively). A notable exception

was observed with medium-sized plants. Controls produced

28.5% new leaves in two weeks while plants of the same size

whose leaves were removed first regenerated only 19.3% new

leaves.


































Figure I-1 (a d). Symptoms of disease on water-
hyacinths incited by Acremonium zonatum.

a. Waterhyacinth with zonate lesions on leaves and
petioles.

b. Close-up of waterhyacinth plants two weeks post-
inoculation.

c. Waterhyacinth leaf showing coalescence of leaf
spots.

d. Large waterhyacinth leaf with abundant white my-
celia of A. zonatum.

































Figure 1-2. Quantitation of disease on small, medium
and large waterhyacinths. Small = plants with leaves 2 15
cm surface area; medium = plants wilh leaves 15-40 cm
surface area; large = plants > 40 cm surface area. a= %
leaves with 1 or more lesions/leaf; b= % leaves with 10 or
more lesions/leaf; c= mean number of lesions/leaf; d= %
total diseased leaf area.
































































a b c d a b c d 'a b c d

SMALL MEDIUM LARGE


125






100




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Lu 75






50






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20







LL
15
Id




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d
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-I
0
z






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Figure I-3. Quantitation of leaf regeneration rates
of small, medium, and large waterhyacinths. C= control
plants; I= inoculated plants; E= plants with leaves excised
prior to test.









I I I I I I


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Discussion

The concept of biological control, the use of one

organism to control another, although not new in practice is

relatively new in its wide-scale applications. Debach (40)

cites that the introduction of the mynah bird from India to

Mauritues in 1762 to control the red locust was the first

successful attempt at biological control. Perhaps the first

successful deliberate control of one organism with another

in the United States was the introduction of the vedalia

beetle into California in 1888 to control cottony-cushion

scale of citrus (6). Control of one organism by another has

been referred to as "parasitic control" and "the biological

method" but it wasn't until 1919 that H.S. Smith referred to

it as "biological control" (40).

It is a difficult task to impart a precise definition

to the term "biological control" since there is little

unanimity on this point among plant pathologist. Perhaps

the best definition is that given by Baker and Cook (6)

Biological control is the reduction of
inoculum density or disease-producing
activities of a pathogen or parasite in
its active or dormant state, by one or
more organisms, accomplished naturally
or through manipulation of the environ-
ment, host, or antagonist, or by mass
introduction of one or more antagonists.

The above definition encompasses several points not

dealt with in this dissertation. For convenience and ease

of understanding in the present discussion, the term "bio-

logical control" will be used to imply the use of native or










introduced organisms to control or reduce the population of

another organism through an antagonistic or parasitic

relationship. Thus, the use of the fungus A. zonatum to

parasitize the waterhyacinth and thereby reduce its popula-

tion size is well within the scope of the definition by

Baker and Cook (6).

Plant pathologists are generally concerned with con-

trolling epiphytotics--not starting them. However, this is

not the case when control of a noxious weed such as the

waterhyacinth is desired through biological methods.

Therefore, it takes some adjustment in one's own thinking

when the initial idea is presented.

When an alien plant establishes itself in a particular

habitat it may mean several things: (i) it is better suited

to a particular niche than are the residents, (ii) that it

was introduced in such numbers as to temporarily or perma-

nently "swamp" the residents, or, (iii) that it may modify

the environment in some way favorable to itself. It usually

means, however, that man has upset the natural balance in

some way, making the environment more favorable to the alien

than to the resident. Such has been the case with the

waterhyacinth. Over-nutrification of our waters by man's

increasing agricultural and urban demands has been the

single most contributing factor to the aquatic weed problem.

Thus a weed is "a generally unwanted organism that thrives

in habitats disturbed by man" (6).









The first step in a biological control program is, in

most cases, the evaluation of the potential biocontrol

agent. This usually involves the introduction of the

control agent onto its target host and/or additional poten-

tial hosts under greenhouse conditions. Thus the effec-

tiveness of the control agent on its target host can be

determined as well as its potential to parasitize other

crops for which it was not intended. If a potential bio-

control agent passes the initial greenhouse tests, field

trials are usually initiated. In these studies, evaluations

as to how the control agent manifests itself and its ability

to compete with the other biotic agents present can be

made. In some instances, it may be necessary to bring the

control agent back into the laboratory and greenhouse to

further evaluate situations observed in the field.

The above description depicts the studies conducted on

A. zonatum over the past five years. Isolation and patho-

genicity tests of the fungus under greenhouse conditions

were initially done by Rintz (152). Field trials with A.

zonatum were initiated in 1973 by Freeman, et al. (54) on

well established stands of waterhyacinths in Lake Alice on

the campus of the University of Florida. It was during

these studies that apparent differences in symptoms and

growth rates were noticed on the plants.

In 1974, greenhouse tests were initiated once again to









see how host plant size influenced A. zonatum as to infec-

tion and subsequent disease.

Disease measurement is often regarded as a synonym for

"estimation of losses," but this is misleading (92). There

is a great need for some reasonably simple but critical

parameters that can be used consistently and systematically

to measure the prevalence and severity of plant diseases in

the field. On the other hand, there are no portmanteau

methods that will serve for all plant diseases. Some of the

currently accepted disease assessment techniques are dis-

cussed by Large (92) and include such things as standard

diagrams, the Horsfall and Barratt grading system, and

disease progress curves.

Perhaps one of the easiest techniques to use is the

standard diagran method. This, of course, assumes that

standard diagrams for the particular host-parasite couplet

in qesrion have been constructed. If not, then this method

requires the researcher to work out such diagrams. In The

case of E. crassipes A. zonatum, standard disease diagrams

have not been constructed. For this reason, disease assess-

ment was based on two criteria: (i) number of lesions/leaf,

and (ii) total percent of diseased area/leaf. Both of these

methods have been used routinely with other host-parasite

combinations and are the basis of standard diagram keys.

Lesion counts on different size waterhyacinths indi-

cated initially that small plants were more resistant than









large plants since they exhibited fewer lesions/leaf.

However, when the mean percent diseased area of each leaf

was measured, there were no significant differences among

any of the three sizes, all showing approximately 40%

disease severity. This allows for two possibilities.

First, small plants are more resistant to initial attack,

but over the two week infection period gradually lose this

resistance and obtain a level of susceptibility shown by the

larger plants, or secondly, large plants are more suscep-

tible initially but gradually build up a resistance. Based

upon data presented elsewhere in this dissertation, i.e.

polyphenoloxidase rates and phenolic acid concentrations

(see Chapter III), it is believed that a combination of both

mechanisms is involved. That is, small plants gradually

lose some of their initial resistance while larger plants

gain various degrees of resistance.

That plants may increase or decrease in susceptibility

to a particular pathogen with age is well documented (197).

It has been suggested (196) that susceptibility to faculta-

tive saprophytes increases with age of host tissues, whereas

isceptibility to obligate parasites decreases with age

although this does not always hold true.

Based upon the results of the present study with water-

hyacinths, susceptibility to attack by A. zonatum increases

with plant size. Generally, plant size can be correlated









with ontogenetic development, that is, the older the plant,

the larger is its size. However, this may not always be a

correct assumption with waterhyacinths since growth rate

depends upon environmental conditions of its habitat (light

intensity, nutrients, and temperature). For this reason,

then, predisposition to A. zonatum in nature due to host age

may be only part of the answer. Differences in symptom

expression during field trials with this fungus may then be

the result of several predisposing factors operating in

conjunction with one another.

Water quality was not monitored during field trials

with this fungus so the effect of environmental predisposing

factors cannot be discussed. However, waterhyacinths used

in the greenhouse studies were all maintained under the same

environmental parameters. Since the only variable in these

tests was the age of the host, it can be stated that suscep-

tibility of waterhyacinths to A. zonatum increases with the

ontogenetic development of the plant. This is an important

criterion when considering the use of any agent as a control

measure. Time of application is extremely important in

order to obtain the most effective control.

Another very important observation made during these

studies was that of the leaf regeneration rates of different

plant sizes after infection. As healthy plants increase in

size (small to medium to large) their leaf regeneration rate









increases. That is, small plants regenerate approximately

27% of their leaves in two weeks or about one leaf/plant.

Medium-sized plants produced a slightly higher, but insig-

nificant, percentage rate of 28.5 or 1.5 leaves/plant.

Large plants, however, are able to reproduce almost half of

their total leaves within a two week period (46.1%).

When plants are inoculated with A. zonatum, their leaf

regeneration rates are altered. There is a slight reduction

in new leaf production exhibited by infected small plants

(5.7%) and a slight increase shown by infected medium-

sized plants (5.4%). But the significant difference is

demonstrated by infected large plants. With these there is

a two-fold increase in new leaves after two weeks. The rise

from 46.1% to 93.5% in large plants represents an increase

on the average from 2.0 new leaves/plant to 4.6 new leaves/

Dlant.

Because A. zonatum is a leafspotting pathogen, it was

postulated that accelerated leaf production was a response

to photosynthetic stress placed upon it by infection which

resulted in the destruction of most of its photosyntheti-

cally active tissue. In order to test this idea leaves were

excised from a set of each of the three plant sizes and

monitored for new leaf growth. There was little variation

in the percentages of new leaf growth when compared to their

respective controls. In one case (medium-sized plants)









there appeared to be a deleterious effect on the plant's

normal leaf production rate. It would appear, then, that

the accelerated new leaf production observed in waterhya-

cinths after infection by A. zonatum is not a response to

the destruction of photosynthetically active tissue, but

one of interaction between the host and the pathogen.

Accelerated growth rates in diseased plants has often

been correlated with increased activity of growth regulators

(61). Normal growth in plants is under hormonal control by

such biologically active endogenous compounds as 8-indole-

3-acetic acid (IAA, auxin), gibberellins, cytokinins, and

others (61). A departure from the normal levels of these

compounds in the plant, such as might be caused by

pathogenic attack, could alter the growth habit of the host.

Data presented in Chapter III show that A. zonatum is

capable of synthesizing high amounts of auxin in vitro when

given the amino acid tryptophan as a precursor. Even though

this does not represent conclusive evidence for the produc-

tion of auxin in vivo by this organism, it does suggest its

possibility. In addition, it has been suggested (87,132)

that increased levels of auxin in diseased tissue may be

correlated with the inhibition of IAA oxidase in the plants

by phenolic inhibitors. Further implications on the pos-

sible roles of auxin, IAA oxidase, and phenolic compounds

during pathogenesis are discussed in Chapter III.










In contemplating A. zonatum as a biocontrol agent of

waterhyacinths, several criteria must be considered. Fore-

most is the proper time at which to apply the inoculum.

Results in this study have indicated that small, young

plants are more resistant to fungal attack than are larger,

older plants. Eased on this, the fungus should perhaps be

applied late in the spring or summer when the plants have

reached maturity. On the other hand, data indicated that

the plants respond to infection by accelerating their rate

of leaf regeneration and that large plants do this more

quickly than do smaller ones. In essence, then, application

of the fungus to large plants would appear to negate or

minimize any control afforded by the pathogen. When, then

would be the best time to apply the control agent? Since

disease severity proceeds to approximately 40% within two

weeks, regardless of the plant size, application early in

the spring, as the new season's growth is beginning, would

appear To be the best time. In this manner one could avoid

the accelerated leaf growth response displayed by larger

planTs while at the same time expect substantial damage to

the plant














CHAPTER II
A CYTOCHEMICAL AND ULTRASTRUCTURAL STUDY OF THE PHENOL
CELLS AND POLYPHENOLOXIDASE ACTIVITIES IN HEALTHY AND
DISEASED WATERHYACINTH LEAVES



Introduction

Phenolic compounds are among the most widespread and

varied compounds in plants. Perhaps the best known role for

plant phenolics is their assimilation into the anthocyanins

and flavone pigments (150). However, as many authors have

indicated, phenolic compounds have nearly unlimited potential

in accounting for the many differences that occur in disease

resistance (12,34,35,50,90,157,179).

Phenols are particularly abundant in the leaves of many

plants. They are also found in the xylem, phloem, and

periderm of stems and roots; in unripe fruits; in the testa

of seeds; and in pathological growths such as galls (49).

Phenolic compounds in plants may be present in individual

cells or in specialized idioblasts termed tannin sacs (49) or

phenol-storing cells (119). Recent studies have shown that

specialized phenol-storing cells occur randomly in several

plant species (10,11,100,107,119,120). Phenols may be a

common ingredient of the vacuoles or they may occur in the

cytoplasm proper in the form of small droplets which even-

tually fuse (49).


__










In many plant tissues, phenols become oxidized to poly-

meric dark red or brown compounds (phlobaphenes), which are

sometimes microscopically visible in the cell contents of

fresh sections. Oxidation of phenolic compound accounts for

the pathological darkening in plant tissues (38).

Histochemical detection of naturally occurring phenols

is difficult because few reagents that react with them to

form characteristic color compounds are adaptable to his-

tological methods (148). In addition, the natural enzymatic

browning may not be sufficiently intense for easy detection

microscopically. In 1951, Reeve (148) described a histo-

chemical test for phenols in fresh plant tissue. It is

based upon a colorimetric method for phenols using a nitrous

acid reaction. The method has become widely accepted and

used and is often referred to as the "nitroso reaction."

One of the enzymes associated with the oxidation of

phenolic compounds is polyphenoloxidase (PPO). The term

polyphenoloxidase has been used extensively in the litera-

ture, although the names phenolase, phenoloxidase, catecho-

loxidase, and tyrosinase have been used as synonyms.

Classification of this enzyme is difficult because several

different activities have been described for it. The enzyme

'as originally termed tyrosinase since the aromatic amino

id, tyrosine, was the first experimental substrate (38).

ever, p-cresol and catechol have been most frequently










employed as experimental substrates. Consequently, two

activities have been ascribed and have come to be known as

the "cresolase" activity when referring to monohydric phenol

oxidation and "catecholase" activity when referring to o-

dihydric phenol oxidation (38).

Many different phenolic compounds can serve as sub-

strates for polyphenoloxidases. For sake of convenience

these enzymes have been divided into three main groups (155)

based upon their affinity for certain substrates, response

to inhibitors, and type of reaction catalyzed: (i) Tyro-

sinases enzymes of this group catalyze both o-hydroxyla-

tion of monophenols and the oxidation of o-diphenols. (ii)

Ortho-diphenoloxidases these enzymes, unlike the tyro-

sinases, are devoid of hydroxylation properties and act only

on o-diphenols. (iii) Para-diphenoloxidases members of

this group act primarily on p-diphenols but may also have

some affinity for the oxidation of certain o-diphenols. The

laccases can be classified in this category.

In the present discussion, the term polyphenoloxidase

has been retained whenever the oxidation activity is being

described regardless of whether it is acting upon an o- or

p-diphenol. For a detailed review of the polyphenoloxi-

dases, the reader is referred to Dawson and Magie (38),

Nelson and Dawson (128), and Patil and Zucker (138).

Some cells are capable of converting tyrosine into a










brown or black pigment called melanin (48). The pathway for

this conversion is depicted in Figure II-1. The first step

involves an o-hydroxylation of tyrosine thereby forming

dihydroxyphenylalanine dopaA). The enzyme that catalyzes

this conversion is in the tyrosinase group and consequently

can also oxidize DOPA in the second step to dopaquinone.

Polyphenoloxidases are devoid of any hydroxylation proper-

ties and therefore cannot convert tyrosine to DOPA but are

capable of oxidizing it to dopaquinone. It is this property

which has been investigated as a marker for this enzyme in

vivo.

Polyphenoloxidase activity has long been thought to

reside within the chloroplasts of plant cells (5), but until

recently cytochemical localization had not been demonstrated.

Based on techniques developed by Novikoff et al. (129) and

Okun et al. (132) for the localization of tyrosinase in

animal tissues, Czaninski and Catesson (36,37) have recently

demonstrated the cytochemical localization of PPO in plant

cells. Since 1972, several investigators (72,74,107,134,135)

have shown that PPO activity is localized within the thyla-

koids of chloroplasts in several plant species.

This chapter presents the results of a histochemical

and ultrastructural study of the phenol cells in water-

hyacinth leaves and the cytochemical localization of PPO in

healthy and diseased plants.































Figure II-i. Biosynthetic pathway for conversion of
tyrosine to melanin [after Eppig (48)].











I
0









\ a
\ 0




















D


c-
U U
































-- C
0---- ------nu


























0 a











om o
0 Z







S I I o




















O
o
0 -
U) U ,




I-



















UU U
/ 4 0











Zr
x )










Materials and Methods

Histochemical localization of phenols

Cross sections of fresh waterhyacinth leaf tissue (12-

24p) from small, medium, and large plants were made with a

Hooker plant microtome, tested for phenols by the nitroso

reaction (148), and observed with the light microscope.

With this method a nitroso derivative of the phenolic

compound is formed and after addition of the base, a bright-

red salt is formed.

Spatial distribution of phenol cells

The spatial distribution of the subepidermal phenol

cells from each size category was determined from tangential

sections made along the vascular bundles. Sections of the

leaves (10 x 15 mm) were taken from areas selected at random

and the epidermal surfaces separated from each other with a

razor blade. Each half was then stained for phenols as

previously described and observed with the light microscope.

The mean number of phenol cells/mm2 leaf tissue was calcu-

lated for the top and bottom surfaces of each plant size

group.

Electron microscopy

Standard fixation and embedding procedures were used

throughout with slight modifications as presented below. A

flow diagram for the basic technique is presented in Figure

11-2. Fresh waterhyacinth leaf tissue was placed in a

































Figure 11-2. Flow diagram of procedure for standard
electron microscopy fixation and embedding.

2% glutaraldehyde paraformaldehyde

0.2 M sodium cacodylate, pH 7.2

Spurr, 1969 (172)




fresh tissue


fix in Karnovsky's fixative'
(2hr- 22 C)


I
wash in buffer (4x)

post-fix in 1% OsO4
(Ihr-22 C)
I
wash in buffer (4x)
I
dehydrate in 25% EtOH series
1
transfer to 100% acetone

embed in epoxy resin

section
1
post-stain w/ UrAc (10min.)

post- stain w/ PbCi (5min.)


Flow Diagram for Electron


Microscopy


Fixation and Embedding









buffered (0.2 M sodium cacodylate, pH 7.2) solution of 2.0%

glutaraldehyde and 2.0% paraformaldehyde (85). Each leaf

was cut into 3-5 mm pieces and fixed for two hours at room

temperature. The material was washed in 50% buffer 50%

distilled water solution for a minimum of 30 minutes before

being postfixed in 1.0% osmium tetraoxide for one hour at 22

C. Sections were then rinsed several times with the aqueous-

buffer mixture and passed through an ethanol graded dehydra-

tion series at 25% increments and finally into 100% acetone.

After dehydration the sections were infiltrated with a

graded acetone-plastic series and embedded in a 100% low

viscosity epoxy resin (170). The embedded sections were

then placed under vacuum for five minutes to remove bubbles

and the resin was polymerized for 18 hours in a 60 C oven.

Thin sections were cut on a Sorvall MT-2 ultramicrotome with

a diamond knife and placed on single-hole, Formvar coated

grids. Sections were then poststained in 0.5% uranyl

acetate for ten minutes and in 1.0% lead citrate for five

minutes. The sections were examined with a Hitachi HU 11E

electron microscope.

Syt>chemical localization of polyphenoloxidase

The procedure for the localization of PPO activity in

hyacinth leaves follows closely that described by

iC.-. nski and Catesson (37). A flow diagram of this pro-

ceduce is presented in Figure 11-3. Fresh leaf tissue, both
































Figure II-3. Flow diagram of procedure for the
cytochemical localization of polyphenoloxidase.


redistilled glutaraldehyde

0.2 M sodium cacodylate, pH 7.2

0.02 M sodium diethyldithiocarbamate

4L-dihydroxyphenylalanine (50 mg/10 ml 0.067 M
phosphate buffer, pH 7.0)




fresh leaf sections


fix in 5% glut.'

wash in b er(x)
wash in buffer(5x)
I


treat w/ DDC3

wash in buffer(5x)
I -


boil sections
(10 mir)
t


pre-incube w/DOPA4
pre-incubate w/DOPA

incubate w/DOPA
(Ihr.-37 C)
wash in d.w.-sucrose (5x)

post-fix w/ 2% Os04
(2hr. -22C)

dehydrate in EtOH

embed in epoxy resin


section
t t


post-stain w/
PbCi

Flow Diagram for C


no post-stain


ytochemical Localization of


PPO Activity













healthy and diseased, was placed in buffer as before and cut

into 2-4 mm pieces. The sections were fixed in 5.0% gluta-

raldehyde for 1 1/2 hours at room temperature and washed in

buffer 5 times for 15 minutes each. The sections were then

separated into three groups and treated by one of the

following methods: (i) boiled for ten minutes, (ii) incu-

bated in 0.02 M DDC (sodium diethyldithiocarbarate) for 20

minutes at 22 C and then washed 5 times in buffer, and (iii)

no treatment. After their respective treatments, each group

was preincubated in a DOPA substrate solution (50 mg DOPA in

10 ml of 0.067 M phosphate buffer, pH 7.0, made up fresh) at

4 C overnight. After the preincubation period, the sections

were incubated in fresh DOPA for one hour (fresh solution

added after 30 minutes) at 37 C, followed by five washings

in distilled water made to 0.5 M with sucrose. After

postfixing in 1.0% osmium tetraoxide they were dehydrated,

embedded in epoxy resin, sectioned, and examined with the

electron microscope as previously described.



Results

Histochemical localization of phenols

When waterhyacinth leaves were stained for phenols by

the nitroso reaction, these compounds were found in large,

specialized idioblasts or phenol cells immediately beneath

both epidermal surfaces (Figs. II-4a & b) and in cells










closely associated with the vascular bundles (Fig. II-4c).

The size of these cells in the palisade layer varied consi-

derably, often exceeding several hundred microns in length

and extending down to the vascular elements. Those phenol

cells near the vascular tissue were much more isodiametric

and varied much less in size. There was no significant

difference in morphology of the cells among the three plant

sizes examined.

Spatial distribution of phenol cells

Phenol cells occurred randomly beneath both leaf

surfaces in all plant sizes and were found throughout the

entire leaf (Fig. II-4d). There were significantly more

phenol cells beneath the adaxial leaf surface (40.6/mm2)

than on the abaxial surface (26.6/mm2) in small plants but

the reverse was true for medium and large plants (Fig. II-

5). Medium and large plants exhibited a more equal dis-

tribution of phenol cells between the two surfaces but there

was a significantly greater number on the top (51.8/41.8 in

medium vs 54.2/48.7 in large). The total number of phenol
2
cells/mm2, both adaxial and abaxial surfaces, significantly

increased as the leaf increased in area with a mean of

33.6/mm2 for small, 41.8/mm2 for medium, and 48.7/mm2 for

*i .ge .

Ulirastructure of phenol cells

Electron micrographs indicate that in most cases the










subepidermal phenol cells were two to three times longer

than the adjacent palisade cells (Fig. 11-6). The phenolic

compounds appeared in close association with the tonoplast

and as discrete bodies within the cells. These were ac-

tively metabolizing cells containing nuclei, mitochondria,

and plastids. In contrast, the phenol cells near the level

of the vascular tissue were much more circular, had a

thicker wall, and the phenolic compounds were in amorphous

masses as opposed to discrete globules (Fig. 11-7). There

were no morphological differences observed between phenol

cells of the same type in any of the plant sizes examined.

Cytochemical localization of polyphenoloxidase

The principle of the reaction for the cytochemical

localization of PPO activity involves obtaining an insol-

uble, electron dense reaction product (dopaquinone) from the

synthetic substrate at the point where enzyme activity is

proceeding (37). Although the reaction can be observed

without additional staining, the intensity of the reaction

and the clarity of the surrounding material is enhanced by

poststaining with lead citrate. When examined by this

.nique, a positive PPO reaction product was absent in all

cle oplasts of small and large healthy waterhyacinth leaves

in-abated without DOPA. Chloroplasts in palisade cells

(Fit;. II-8a), have distinctly clear thylakoid spaces and

fret channels. Similar observations were made for










chloroplasts of bundle sheath cells (Fig. Ii-Sb), vascular

parenchyma (Fig. II-8c), and phenol cells (Fig. II-Sd). The

thylakoids within the chloroplasts of phenol cells were not

readily detected until poststained with lead citrate.

Sections from both small and large healthy leaves incu-

bated with DOPA reacted in an identical manner for the

localization of PPO. Chloroplasts of the palisade cells

(Fig. II-9a) and spongy mesophyl cells (Fig. II-9b) did not

stain for PPO activity. On the other hand, PPO activity was

localized in the thylakoids of chloroplasts in three other

cell types, two of which were associated with the vascular

tissue. In each instance, the thylakoid spaces and fret

channels were the only areas stained for P0O activity.

In contrast to other cells, chloroplasts of the vascu-

lar parenchyma, both phloem parenchyma (Fig. II-9d) and xylem

parenchyma (Fig. II-10) were PPO positive. The chloroplasts

in these cells appeared black or electron-dense. These

elecTron-dense areas were restricted to the Thylakoids

within the chloroplasts (Fig. 11-10b). Chloroplasts which

were not poststained (Figs. II-9d S II-lOc) also showed a

positive reaction but the inLensity and clarity was not as

good.

Another type of cell having PPO positive chloroplasts

were the bundle sheath cells (Figs. II-Sc and II-lla E b).

Waterhvacinths are typical monocots and have a large bundle










sheath surrounding the vascular elements. Chloroplasts in

these bundle sheath cells were PPO positive, although

perhaps not as intense as those in the vascular parenchyma.

The phenol cell itself also showed PPO activity (Figs.

II-9c and 11-12). The reaction in these cells was the most

intense of the three. In this cell type, the chloroplasts

are extremely electron-dense (Fig. II-12a), and examination

under higher magnification revealed that not only were the

thylakoids positive, but the entire organelle was electron-

dense (Fig. II-12b).

Leaf material that was boiled prior to incubation in

DOPA did not give a positive PPC reaction, in any chloro-

plasts, indicating heat inactivation of the enzyme after

boiling (Fig. 11-13). The thylakoids became distorted after

boiling and starch granules swelled forming large lacunae

(Fig. II-13a & c).

When the inhibitor, DDC, was added to the sections

prior to incubation in DOPA, no reaction product could be

detected in the thylakoids of any chloroplasts (Fig. II-14).

When sections were poststained with lead citrate (Fig. II-

14a), the thylakoid spaces and fret channels contrasted

sharply with the stroma. Only the partitions were notably

electron-dense. Thus, the electron density of lead citrate

cannot be confused with the electron-dense product of a

positive PPO reaction. Consequently, use of the poststain









acts to heighten the observed reactions and surrounding

material. In addition, PPO activity was not observed in any

cell organelle other than chloroplasts. These observations

were consistent for each of the plant sizes examined.

When diseased leaves were examined for enzyme localiza-

tion, PPO activity was found to be no longer restricted to

vascular parenchyma, bundle sheath, and phenol cells rather

every chloroplast in every cell was positive. Palisade

cells were now positive (Fig. II-15) and there was an

increase in the number of plastoglobuli in those chloro-

plasts. Likewise, spongy mesophyl cells, which in healthy

cells were negative, became positive after infection (Fig.

11-16). These chloroplasts also showed an increase in the

number and size of the plastoglobuli.

The changes in PPO localization were apparent in chloro-

plasts in cells immediately surrounding the lesions.

Sections taken several centimeters away from the lesion were

examined to determine if periphery cells also showed a

"turn-on" in enzyme activity. Electron micrographs indicate

that even those cells which are two to five centimeters

removed from the center of infection were also positive for

PPO activity. Thus, palisade cells became positive (Fig.

II-17a E b), spongy mesophyl cells became positive (Fig. II-

17 c 6 d), and chloroplasts in cells normally positive such

as bundle sheath cells became very intense (Fig. II-17d).





55



In essence, PPO activity was found in the chloroplasts in

only three cell types in healthy leaves: (i) vascular

parenchyma, (ii) bundle sheath, and (iii) phenol cells

proper. However, during disease, there was a turn-on of PPO

activity in all cells which contain chloroplasts. Whether

this turn-on in enzyme activity is host-induced or pathogen-

induced is not known at this time.
































Figure 11-4 (a d). Light micrographs of phenol
cells in healthy waterhyacinth leaves.

a. Cross section of waterhyacinth leaf showing
arrangement of phenol cells in upper and
lower palisade cell layers. (375 X).

b. Cross section of waterhyacinth leaf showing phc
and vascular bundle (vb). (1,500 X).

c. Cross section of waterhyacinth leaf showing phc
in relation to vb and bundle sheath cells (bsc).
(1,500 X).

d. Tangential section of waterhyacinth leaf showing
spatial arrangement of phc. (375 X).
































Figure II-5. Number of phenol cells/mm2 leaf area
in small, medium, and large waterhyacinth leaves. ST= small
plants, top surface of leaf; SB= small plants, bottom surface
of leaf; MT= medium plants, top surface of leaf; MB= medium
plants, bottom surface of leaf; LT= large plants, top sur-
face of leaf; LB=large plants, bo tom surface of leaf;
p= mean number of phenol cells/mm leaf (both surfaces).
























50




U)
W














10
cU 40


LL
LJ

N
E 30
E

C)
_J
_J
w

20
2
z





10






0


PLANT SIZE































Figure II-6. Electron micrograph of phenol cell in
palisade cell layer of waterhyacinth leaf tissue. Phenol
bodies (pb) appear in close association with the plasmalemma
and as discrete globules within the tonoplast (t). Post-
stained with PbCi. (2,140 X).
































Figure 11-7. Electron micrograph of phenol cell
in vascular tissue area of waterhyacinth leaf. Phenol
bodies (pb) appear as an amorphous mass within the cell.
x = xylem. Poststained with PbCi. (9,400 X).

































Figure 11-8 (a d). Chloroplasts of healthy water
hyacinth leaf tissue incubated without DOPA.


a. Palisade cell chloroplast with clear thylakoids
(th). s= starch (29,400 X).

b. Bundle sheath cell chloroplast. cw= cell wall
th= thylakoids (37,500 X).

c. Vascular parenchyma cell chloroplast. th= thy-
lakoids (30,000 X).

d. Phenol cell chloroplast. c= chloroplast, th=
thylakoids, pb= phenol body. Foststained with
PbCi (24,000 X).































Figure 11-9 (a e). Localization of polypheno-
loxidase in healthy waterhyacinth leaf tissue without
lead poststaining.

a. Palisade cell chloroplast. Negative PPO activity
in thylakoids (th). s= starch (45,500 X).

b. Spongy mesophyll cell chloroplast. Negative PPO
activity in thylakoids (th). (45,000 X).

c. Phenol cell chloroplast (phc). Positive PPO
activity in thylakoids (th). (33,000 X).

d. Vascular parenchyma cell chloroplast. Positive
PPO activity in thylakoids (th). pl= plasto-
globuli (57,500 X).

e. Bundle sheath cell chloroplast. Positive PPO
activity in thylakoids (th). (75,000 X).































Figure II-10 (a c). Localization of polyphenol-
oxidase in chloroplasts of xylem parenchyma cells in
healthy waterhyacinth leaves.

a. Cross section of leaf showing a xylem element (x)
and surrounding xylem parenchyma cells (xp).
Chloroplasts (c) in the xp cells are positive for
PPO activity. Poststained with PbCi. (4,800 X).

b. Close-up of chloroplasts in xp showing positive
PPO activity between the thylakoids (th) and
several plastoglobuli (pl). cw= cell wall. Post-
stained with PbCi. (26,000 X).

c. Chloroplast in xp cell showing positive PPO acti-
vity without PbCi poststaining. (16,500 X).
































Figure II-11 (a c). Localization of polyphenol-
oxidase in chloroplasts of bundle sheath cells in healthy
waterhyacinth leaves.

a. Chloroplasts (c) in bundle sheath cells (bsc)
showing positive PPO activity. Poststained with
PbCi. (6,200 X).

b. Close-up of chloroplast in bsc showing positive
PPO activity in thylakoids. Poststained with
PbCi. cw= cell wall. (32,000 X).

c. Chloroplasts in bsc incubated in diethyldithio-
carbamate (DDC) prior to incubation in DOPA.
Thylakoids (th) are negative for PPO activity.
Poststained with PbCi. pm= plasmalemma.
(40,000 X).
































Figure II-12 (a b). Localization of polyphenol-
oxidase in chloroplasts of phenol cells in healthy water-
hyacinth leaves.

a. Ultrastructure of phenol cell in palisade cell
layer showing nucleus (n), mitochondrion (m),
chloroplasts (c), and phenol bodies (pb).
Chloroplasts are positive for PPO activity. Post-
stained with PbCi. (2,200 X).

b. Enlargement of chloroplast in phenol cell showing PPO
positive thylakoids (th) and large phenol body (pb)
in association with the chloroplast. s= starch.
Poststained with PbCi. (57,500 X).
































Figure II-13 (a d). Chloroplasts of boiled, healthy
waterhyacinth leaf tissue incubated with DOPA.

a. Spongy mesophyll cell chloroplast showing distended
thylakoids (th). cw= cell wall, sl= starch lacuna
(37,500 X).

b. Vascular parenchyma cell chloroplast showing thy-
lakoids (th) negative for PPO activity. pl= plas-
toglobuli, m= mitochondrion (69,000 X).

c. Bundle sheath cell (bsc) chloroplast (c) with nega-
tive PPO activity. mc= mesophyll cell. (7,000 X).

d. Enlargement of bsc chloroplast with negative PPO
activity. th= thylakoids, pl= plastoglobuli,
sl= starch lacuna, cw= cell wall (56,000 X).































Figure II-14 (a d). Chloroplasts of healthy water-
hyacinth leaf tissue incubated in inhibitor (DDC) and DOPA.

a. Palisade cell chloroplast with distinct thylakoid
spaces (th) and fret channels. m= mitochondrion;
Poststained with PbCi (40,000 X).

b. Vascular parenchyma cell chloroplast with negative
PPO activity. th= thylakoids (28,000 X).

c. Bundle sheath cell chloroplast with negative PPO
activity. th= thylakoids, s= starch (46,000 X).

d. Phenol cell chloroplast with negative PPO activity.
th= thylakoids, pl= plastoglobuli (55,000 X).

































Figure II-15 (a b). Localization of polyphenol-
oxidase in chloroplasts of palisade cells from diseased
waterhyacinth leaves.

a. Necrotic palisade cells (pc) showing positive PPO
activity in their chloroplasts and an increase in
the size and number of plastoglobuli. Chloro-
plasts in palisade cells in healthy leaf tissue
are negative for PPO activity. Poststained with
PbCi. (7,820 X).

b. Enlargement of -chloroplasts in palisade cells showing
PPO activity in the thylakoids (th). Poststained
with PbCi. (24,300 X).


__
































Figure II-16 (a c). Localization of polyphenol-
oxidase in chloroplasts of spongy mesophyll cells from
diseased waterhyacinth leaves.

a. Mesophyll cells (mc) showing PPO positive chloro-
plasts. Hyphae (h) shown in upper right corner.
Poststained with PbCi. Chloroplasts in mesophyll
cells in healthy leaf tissue are negative for PPO
activity. (8,280 X).

b. Enlargement of mesophyll chloroplast showing positive
PPO reaction in thylakoids. m= mitochondrion. Post-
stained with PbCi. (35,400 X).

c. Enlargement of positive PFO chloroplast in mesophyll
cell without PbCi poststain. (40,000 X).
































Figure II-17 (a d). Localization of polyphenoloxi-
dase in chloroplasts of cells several centimeters away from
infection center.

a. Palisade cells (pc) with positive PPO activity in
their chloroplasts. e= epidermis. Poststained
with PbCi. (6,200 X).

b. Enlargement of palisade chloroplast showing PPO acti-
vity in the thylakoids (th). Poststained with
PbCi. (17,400 X).

c. Mesophyll cell (mc) showing positive PPO activity
in the chloroplast. n= nucleus, m= mitochondrion,
th= thylakoids. Poststained with PbCi. (27,500 X).

d. Electron micrograph showing PPO positive chloroplasts
in mesophyll cell (mc) and very intense reaction in
the bundle sheath cell (bsc) chloroplast. cw= cell
wall. Poststained with PbCi. (18,900 X).









Discussion

A wide variety of simple and complex compounds pos-

sessing phenolic hydroxyl groups occur in plant tissues and

the importance of these compounds during the life cycle of

the plant has become increasingly evident (143). Plant

pathologists and physiologists have a keen interest in

phenolics as the "antiseptics" of the Plant Kingdom (143)

and many investigations have been made on disease resistance

and interaction of microorganisms with phenols.

As indicated previously, specialized cells containing

phenolic compounds have been reported in tissues from many

plant species. These cells are often called "tannin cells"

when the nature of the phenolic substances is not known, or

the substances have become decompartmented, oxidized, and

polymerized to varying degrees (120).

Common, nonspecific tests for tannins usually consist

of treatment with ferric chloride solutions followed by

treatment with dilute bases (148). A blue-green precipitate

is usually formed but not all phenolics give such a reaction

and the results may be influenced by other materials pre-

sent. The Gibbs indophenol reaction (59) is a dependable

test for the detection of phenols (51), but appears to be of

little or no value in determining the number of hydroxyl

groups on the benzene ring (51,100). On the other hand, the

nitroso reaction (148) forms a cherry-red nitroso derivative




Full Text

PAGE 1

DISEASE RESISTANCE MECHANISMS IN WATERHYACINTHS AND THEIR SIGNIFICANCE IN BIOCONTROL PROGRAMS WITH PHYTOPATHOGENS By RAYMOND DEWINT MARTYN, JR, A DISSERTATION PRESENTED TO THE GRADUATE COUNCIL OF THE UNIVERSITY OF FLORIDA IN PARTIAL FULFILLMENT OF THE REQUIREMENTS FOR THE DEGREE OF DOCTOR OF PHILOSOPHY UNIVERSITY OF FLORIDA 1977

PAGE 2

This above all: to thine own self be true William Shakespeare Hamlet ; Act I, scene iii

PAGE 3

To my parents, who had the wisdom and foresight to know the difference between "guidance" and "insistence", and who used as one of the cornerstones of my education, Robert W. Service's poem "The Quitter" which appears on the following page . . . To my wife, Dickie, whose unyielding faith and many hours of unselfish help and patience were perhaps the greatest factors in the completion of this program . . . To my daughter, Susan, whose 6-year-old smile made it all worthwhile, when I overheard her tell a playmate, "My Daddy is a plant doctor!"

PAGE 4

The Quitter When you're lost in the wild and you're scared as a child, And death looks you bang in the eye; And you're sore as a boil, it's according to Hoyle To cock your revolver and die. But the code of a man says fight all you can, And self-dissolution is barred; In hunger and woe, oh it's easy to blow -It's the hell served for breakfast that's hard. You're sick of the game? Well now, that's a shame! You're young and you're brave and you're bright. You've had a raw deal, I know, but don't squeal. Buck up, do your damnedest and fight! It's the plugging away that will win you the day, So don't be a piker, old pard ; Just draw on your grit; it's so easy to quit -It's the keeping your chin up that's hard. It's easy to cry that you're beaten and die, It's easy to crawfish and crawl, But to fight and to fight when hope's out of sight, Why, that's the best game of them all. And though you come out of each grueling bout, All broken and beaten and scarred -Just have one more try, it's dead easy to die; It's the keeping on living that's hare. Robert W . Service

PAGE 5

ACKNOWLEDGEMENTS I wish to express sincere gratitude to Dr. Thomas E. Freeman, Chairman of my Supervisory Committee, for his friendship, advice, guidance, and patience during the course of this study, and for his criticism and encouragement in appropriate doses for three years which ultimately made this dissertation possible. I also wish to extend thanks to members of my Supervisory Committee, Dr. T.E. Humphreys, Dr. H.H. Luke, Dr. D.A. Roberts, and Dr. R.E. Stall for their advice and friendship, and for their time spent in critical review of this manuscript. A special thanks is extended to Mr. D.A. Samuelson for his many hours of assistance during the ultrastructural and cytochemical portions of this study, and for the many hours of help in preparing the electron micrograph plates. Gratitude is also extended to Dr. H.A. Aldrich for his kindness for allowing use of equipment and facilities of the Biological Ultrastructure Laboratory, and to Ms. Janet Plaut for performing the many statistical analyses used throughout this dissertation. This research supported in part by the U.S. Army Corps of Engineers, Florida Department of Natural Resources, U.S. Department of Interior, Office of Water Resources and Research Act as amended and by the University of Florida Cer_~er for Environmental Programs.

PAGE 6

TABLE OF CONTENTS Page ACKNOWLEDGEMENTS LIST OF TABLES . . . LIST OF FIGURES . . ABSTRACT GENERAL INTRODUCTION Part I The Aquatic Weed Problem Part II The Potential of Biological C Part III Pathogens of Waterhyacinth wi ontrol th Possible Biocontrol Potential CHAPTER I RESPONSES OF WATERHYACINTH TO INFECTION WITH ACREMONIUM ZONATUM AND ITS IMPLICATIONS IN BIOLOGICAL CONTROL Introduction .... Materials and Methods Results Discussion CHAPTER II A CYTOCHEMICAL AND ULTRASTRUCTURAL STUDY OF THE PHENOL CELLS AND POLYPHENOLOXIDASE ACTIVITIES IN HEALTHY AND DISEASED WATERHYACINTH LEAVES . . Introduction .... Materials and Methods Results Discussion ..... CHAPTER III A BIOCHEMICAL STUDY OF THE PHENOLIC ACIDS AND POLYPHENOLOXIDASE RATES IN HEALTHY AND DISEASED WATERHYACINTH LEAVES Introduction Materials and Results . . Discussion . , iethods xiii 1 1 5 15 15 17 19 23 37 37 43 49 84 90 90 100 108 129

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Page CHAPTER IV AN ULTRASTRUCTURAL STUDY OF PENETRATION AND CODONIZATION OF WATERHYACINTH BY ACREMONIUM ZONATUM 13 9 Introduction 139 Materials and Methods 141 Results 144 Discussion .169 SUMMARY AND CONCLUSIONS 17 9 LITERATURE CITED 18 7 BIOGRAPHICAL SKETCH 204

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LIST OF TABLES Table Page III-l Free phenolic acids detected in healthy and A. zona turn in fee ted waterhyacinths by thin layer chromatography 114 III-2 Phenolic acids detected in healthy waterhyacinth leaves by thin layer chromatography and various locating reagents after alkaline hydrolysis 115 III-3 Phenolic acids detected in A. zonatum -infected waterhyacinth leaves by thin layer chromatography and various locating reagents after alkaline hydrolysis 116 III-4 R^ values and color characteristics of the phenolic acids detected in healthy and A. zonatum -inf ected waterhyacinth leaves after alkaline hydrolysis 117 III-5 Growth of A. zonatum on healthy and A. zonatum -inf ected waterhyacinth leafextract media 124 III-6 Growth of A. zonatum on phenolic acid media 125 III-7 Growth of A. zonatum on phenolic acid media with yeast extract 125 S-l Differences and similarities among healthy and A. zonatum -inf ected waterhyacinth morphotypes 181

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LIST OF FIGURES CHAPTER I CHAPTER II Page Fig. 1-1 Symptoms of disease on waterhyacinths incited by Acremonium zonatum ~ ~ '. . '. . . 23 Fig. 1-2 Quantitation of disease on small, medium, and large waterhyacinths . 25 Fig. 1-3 Quantitation of leaf regeneration rates of small, medium, and large waterhyacinths 27 Fig. II-l Biosynthetic pathway for conversion of tyrosine to melanin ... 42 Fig. II-2 Flow diagram of procedure for standard electron microscopy fixation and embedding 4 5 Fig. II-3 Flow diagram of procedure for the cytochemlcal localization of polyphenoloxidase 48 Fig. II-4 Light micrographs of phenol cells in healthy waterhyacinth leaves . 57 Fig. II-5 Number of phenol cells/mm leaf area in small, medium, and large waterhyacinth leaves 5 9 Fig. II-6 Electron micrograph of phenol cell in palisade cell layer of waterhyacinth leaf tissue .... 61 Fig. II-7 Electron micrograph of phenol cell in vascular tissue area of waterhyacinth leaf 63 Fig. II-8 Chloroplasts of healthy waterhyacinth leaf tissue incubated without DOPA 6 5

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Page Fig. II-9 Localization of polyphenoloxidase in healthy waterhyacinth leaf tissue without lead postaining . r 67 Fig. 11-10 Localization of polyphenoloxidase in chloroplasts of xylem parenchyma cells in healthy waterhyacinth leaves 69 Fig. 11-11 Localization of polyphenoloxidase in chloroplasts of bundle sheath cells in healthy waterhyacinth leaves 71 Fig. 11-12 Localization of polyphenoloxidase in chloroplasts of phenol cells in healthy waterhyacinth leaves . 7 3 Fig. 11-13 Chloroplasts of boiled, healthy waterhyacinth leaf tissue incubated with DOPA 75 Fig. 11-14 Chloroplasts of healthy waterhyacinth leaf tissue incubated in inhibitor (DDC) and DOPA .... 77 Fig. 11-15 Localization of polyphenoloxidase in chloroplasts of palisade cells from diseased waterhyacinth leaves 7 9 Fig. 11-16 Localization of polyphenoloxidase in chloroplasts of spongy mesophyll cells from diseased waterhyacinth leaves 81 Fig. 11-17. Localization of polyphenoloxidase in chloroplasts of cells several centimeters away from infection center 83 CHAPTER III Fig. III-l Principal phenolic acids found in plants 9 4 Fig. III-2 Shikimic acid pathway for the biosynthesis of monocyclic phenols and maior derivatives ... 98

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Page Fig. III-3 Flow diagram of procedure for extraction of ester-linked phenols in plants 1^3 Fig. III-4 Total phenol concentrations in healthy and A. zonatum infected waterhyacinth morphotypes .... 11° Fig. III-5 Polyphenoloxidase activities in small, medium, and large healthy waterhyacinth leaves 121 Fig. III-6 Polyphenoloxidase activities in small, medium, and large diseased waterhyacinth leaves I 23 Fig. III-7 In vitro synthesis of indoleacetic from tryptophan by Acremonium zonatum CHAPTER IV t ion 128 Fig. IV-1 Flow diagram for testing of carbohydrate degrading enzymes produced by Acremonium zonatum . . . 150 vie IV-2 Penetration of waterhyacinth leaf by Acremonium zona i_um J3 * Fig. IV-3a Cross-section of Acremonium zonatum observed in xylem tissue of diseased waterhyacinth leaf . . 154 Fig. IV-3-c Degradation of wall material in waterhyacinth by Ac remonium zonatum xo • Fig. IV-4a Attachment of Acremonium zonatum to the cuticle 156 Fig. IV-Ub Attachment of Acremonium zonatum to epidermis and the possible area of localized enzyme secre156 \g. IV-5 Penetration of phenol cell by Acremonium zonatum 15 8

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Page Fig. IV-6 Phenol cell invaded by Acremonium zonatum 160 Fig. IV-7 Breakdown of starch reserves in chloroplasts during disease . . . 162 Fig. IV-8 Increase in the number of plastoglobuli in chloroplasts during disease 164 Fig. IV-9a Increase in the number of microbodies in cytosol as a result of infection with Acremonium zonatum 166 Fig. IV-9b Destruction of chloroplast integrity during later stages of disease 166 Fig. IV-10 Diseased palisade cell showing extent of necrosis and cellular breakdown 168

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Abstract of Dissertation Presented to the Graduate Council of the University of Florida in Partial Fulfillment of the Requirements for the Degree of Doctor of Philosophy DISEASE RESISTANCE MECHANISMS IN WATERHYACINTHS AND THEIR SIGNIFICANCE IN BIOCONTROL PROGRAMS WITH PHYTOPATHOGENS By Raymond DeWint Martyn , Jr. June, 19 7 7 Chairman: Dr. Thomas E. Freeman Major Department: Plant Pathology The pathological relationship between the floating waterhyacinth , Eichhornia crassipes (Mart.) Solms and the fungus, Acremonium zonatum (Sawada) Gams, was investigated to determine possible disease resistance mechanisms in the plant as they relate to potential biocontrol agents. Waterhyacinths were separated into three morphotypes based upon their leaf 2 surface area; small plants (leaves < 15 cm ), medium plants 2 2 (leaves 15-40 cm ), and large plants (leaves > 40 cm ) and used for quantitating symptoms of disease. Inoculated small plants exhibited fewer lesions/leaf after two weeks than did either medium or large plants; however, the total percent diseased leaf area for each morphotype was the same (approximately 40%). It was observed that large plants regenerated almost three times as many new leaves after infection development than did either medium or small plants. Biochemical, histochemlcal , cytochemlcal , and ultrastructural studies were conducted on both healthy and diseased

PAGE 14

morphotypes to determine what role host phenolic compounds had in disease development. Phenolic compounds in waterhyacinth leaves are localized in specialized idioblasts (phenol cells) immediately beneath both epidermal surfaces and also in close association with the vascular tissue. The concentration of phenol cells increased significantly from 2 . 2 a mean of 33.6/mm leaf area in small plants to 48.7/mm in large plants. In healthy plants, polyphenoloxidase (PPO) activity was greater in small than in large leaves and was restricted to the thylakoids of chloroplasts in only three cell types: vascular parenchyma, bundle sheath, and phenol cells. After infection by A. zonatum , PPO activity decreased in small leaves but increased over 300% in large leaves. After infection, PPO activity was observed in all chloroplasts throughout the leaf. Chlorogenic acid was the only free phenolic acid found in morphotypes of both healthy and diseased plants. Alkaline hydrolysis of healthy leaf tissue yielded six phenolic acids from small and medium plants and nine from large plants. After infection, one additional phenolic acid was detected from smalland medium-sized leaves. No change in the types of phenolic acids present in large leaves was detected after infection. The concentration of total phenols in healthy plants increased significantly from 92 ug/g fresh leaf tissue in small to 104 ug/g in large leaves. There was

PAGE 15

a significant decrease in total phenols in both small and medium diseased plants while the concentration remained constant in large diseased plants. Acremonium zonatum grew significantly better when cultured on minimal media containing phenolic acids than it did on media without these compounds. Acremonium zonatum was inhibited by p-coumaric acid at 1000 ppm, when yeast extract was added as a growth supplement to the media. In addition, growth of the fungus on diseased plant-extract media was stimulated significantly over growth on media containing extracts from healthy plants. Penetration of waterhyacinth leaves by A. zonatum occurred directly through the cuticle or through the stomata. Cellular penetration was aided by the production of cellulolytic enzymes. Penetration of the phenol cells resulted in death of the invading hyphae. Associated with disease was the disappearance of starch granules from the chloroplast, an increase in the number of plastoglobuli within chloroplasts , and a build-up of microbodies within the cytosol . The results presented in this study suggest that phenol metabolism in waterhyacinth plays a significant role in the defense against potential pathogens and may account for why only a few of pathogens have been reported on this plant. It appears that A. zonatum is capable of causing relatively

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severe damage to the waterhyacinth because of Its high tolerance to phenols and warrants continued study as a potential biocontrol of this noxious aquatic plant.

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GENERAL INTRODUCTION Part I: The Aquatic Weed Problem All plant and animal species in their native habitats are subject to natural forces that control their population levels. Natural enemies along with other environmental influences maintain a balance among populations of plants and animals in an ecosystem. There is little question that the parasites and predators existing in a particular system are the greatest resource that we have for effective pest suppression and management (180). Man steps beyond Nature's boundaries, however, and thereby sidesteps natural controls by transporting plant and animal species to new habitats, and in so doing, often causes disastrous shifts in the ecological balance between species. Such has been the case with many of the noxious aquatic plants in Florida. Exotic water plants imported into this country as aquaria specimens and ornamentals have escaped into lakes and waterways and, once established, have created serious control dilemmas. In areas where aquatic plants have reached high densities, they greatly obstruct the water flow, decrease the water level through increased rates of evaporation and transpiration, increase the rate of eutrophication,

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interfere with navigation, prevent fishing and other water recreational activities, depress real estate values, and may, in some instances, present severe health hazards (52, 75, 201). Infamous examples of these pestiferous plants include the floating waterhyacinth , Eichhornia cras sipes (Mart.) Solms , Florida elodea, Hydrilla verticillata (Casp.), Eurasian watermilf oil , Myriophyllum spicatum L. , and alligatorweed , Alternanthera philoxeroides (Mart.) Griseb. The rampant growth of exotic water weeds in Florida and other Gulf states has been attributed to several factors (78, 118, 139). First, the year-round warm temperature and extended photoperiod combine to give a growing season almost the entire year. Secondly, many bodies of water provide an abundance of inorganic compounds necessary for luxuriant plant growth. Thirdly, the absence of enemies normally present in their native habitats does not allow the natural system of checks and balances to operate. And, lastly, most aquatic plants are capable of extremely rapid vegetative reproduction. It is for these reasons that some 160,000 hectares of Florida's fresh water are weed-choked (54). One of the most pestiferous aquatic plants in tropical and subtropical climates is the floating waterhyacinth, E. crassipes, the subject of this dissertation. The

PAGE 19

genus Eichhornia is a member of the Pontederiacae family and includes four other species: E. paniculata , E. paradoxa , E. azurea , and E. diversifolia (139). Eichhornia crassipes is the only species which is free floating; all other members of the genus are rooted either in shallow water or near shore. The wat erhyacinth reproduces almost entirely by vegetative means although sexual reproduction does occur. It reproduces rapidly and will completely fill many lakes and rivers in a single growing season. Penfound and Earle (139) reported that E. crassipes is capable of doubling its mass every 11-15 days. Taking an average rate of doubling of two weeks and a growing season of eight months, then ten plants given plenty of room and good growing conditions would produce 65 5,36 plants which would cover 0.6 hectares. These figures emphasize the tremendous rate of colonization of this species and the necessity of good control methods. It is believed that the wat erhyacinth is a native of Brazil , but has spread from there tc nearly all of the South American and Central American countries and throughout the world where the climate is favorable for its development. few tropical or subtropical countries are free from water-hyacinths (97). The accounts differ somewhat regarding its appearance in the United States. There is some evidence that it was

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cultivated as a greenhouse exotic shortly after the War Between the States (139); however, the earliest authentic account details its introduction at the Cotton Centennial Exposition at New Orleans in 1884 (88). It appeared in Florida in 1890 (190) and has since become an important aquatic pest. By the turn of the century it was reported from all the southeastern coastal states as far north as Virginia and westward to California (81). Eichhornia crassipes was officially recognized as a serious aquatic pest in this country on June 4, 1897, when Congress passed an act authorizing the Secretary of War to investigate the extent of obstruction to navigation in the waters of Florida and Louisiana (139). Since that time, the U.S. Army Corps of Engineers have been responsible for clearing it from navigable waterways. Florida, like many parts of the United States and world, is in dire need of an efficient and effective means of controlling noxious aquatic plants. Since their introduction, millions of dollars, both tax and private, have been spent on chemical and mechanical control of these weeds. An estimated 10 to 15 million dollars is being spent per year for the control of aquatic weeds In Florida alone, and this figure is increasing every year (64). Despite this huge financial expenditure, the total infestation continues to grow and at present there is no end to the increasing costs unless new control measures are found.

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Fart II: The Fotential of Biological Control In past years, control of waterweeds has been based on two basic procedures. Both mechanical and chemical controls are used routinely in maintenance programs. However, neither method on its own is completely satisfactory and the weed infestations continue to expand. More recently, the concept of biological control was proposed for aquatic weeds. Huf faker and Andres (78) have stated that any organism which curtails plant growth or reproduction may be used as a biological control agent. Such could potentially Include animals either higher or lower than insects, and parasitic higher plants, fungi, bacteria, and viruses. For this reason the term biological control organism, or agent is used to include all suitable phytophagous animals and plant pathogens on a given weed. It was generally believed that biological control works best with agents of foreign origin (75); however, as Wapshere (18 3) points out, successful biocontrol with an organism in one country does not necessarily imply that the organism(s) used will be successful elsewhere. For instance, Chrysolina quadrigemina was relatively Ineffective against Hypericum perforatum in Australia, but beetles of the same genetic stock were highly successful against the same weed in California, apparently because of more suitable climatic conditions there (168).

PAGE 22

Many investigations have been undertaken to study potential uses of macrobiological agents to control noxious aquatic weeds. In most instances, these studies have involved insects (13,78,105,165,199) and, to a lesser extent, other animals (33,39,118,162,164). Of the insects screened for possible control agents, one of the most effective appears to be the flea beetle, Agasicles hygrophila which feeds only on alligatorweed (13, 105, 199). It was successfully introduced into the United States from Argentina for the control of alligatorweed (13). In April, 1965, 266 adult beetles were released near Jacksonville, Florida, and by June, 1966, there were hundreds of thousands of them present at the release sites and most of the floating alligatorweed was dead (105). It has since spread rapidly throughout the watersheds in northeast Florida (199). Insects alone, however, are not likely to control aquatic weed pests because there are relatively few phytophagous species capable of living beneath the water (201). Other biological control agents being investigated include phytopathogenic fungi, bacteria, and viruses. Zettler and Freeman (201) list four advantages of using such control agents: (i) control applications would presumably require minimal technology and, if successfully established, the pathogen in theory would be self maintaining ; (ii) the

PAGE 23

overwhelming number of different plant pathogenic species from which to choose offers an unmatched versatility in selecting a specific biological control; (iii) virtually none can attack man or his animals, therefore providing an important advantage over the use of various animals such as snails, which may harbor vertebrate pathogens, and (iv) plant pathogens, although often killing individuals in a given population, would not be expected to cause the extermination of a species. This last attribute is important because eradictaion of one aquatic weed species, such as the waterhyacinth , may create an ecological void that in turn may allow a population explosion of a different and more serious species. In addition, Wilson (192) points out three more advantages of using biological control agents over chemical control procedures: (i) they can be specific to the target weed which lessens the chance of damage to cultivated or desired species, ( ii ) residue and toxicity problems created by herbicides would be greatly reduced or eliminated altogether, and (iii) there would be no accumulation of the herbicide in the soil or underground water. In essence, then, the use of biocontrol agents has many advantages over chemical control methods and warrants continued research. The use of plant pathogens Is not without hazards. Any study undertaken to introduce or test phytopathogens must

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be done with extreme care. Well controlled and monitored prerelease experiments, however, can greatly reduce any potential dangers. Part III. Pathogens of Waterhyacinth with Possible Biocontrol Potential " ' The first recorded disease on waterhyacinth caused by a fungus was reported in 1917 by Tharp (174). He described a Cercospora sp . as occurring on Piaropus crassipes (= E. crassipes ) in Texas and subsequently identified the causal agent as C. piaropi Tharp. Thirty-seven years later, in 1954, it was reported on waterhyacinth in India (175) and was again reported from the United States in 1974 (53). The disease symptoms are oval leaf spots, 1.5 4.0 mm in size, on the distal portion of the leaf blade. As with other leaf spot diseases reported on waterhyacinth (2, 154), C. piaropi does not appear severe enough to retard the prodigious growth of the plant significantly; however, its host specificity enhances its potential as a biocontrol agent and is being investigated further (53). The second recorded disease on waterhyacinth was caused by a rust fungus, Uredo eichhorniae , found in the Dominican Republic in 1927 (27). A year later, Ciferri (26) reported the occurence of a smut, Doassansia eichhorniae on E. crassipes from the same area. Neither of these organisms,

PAGE 25

however, had been studied as potential biocontrol agents until last year (25). In 1932, a species of Fusarium was reported on waterhyacinth from India (2). It caused reddish-brown necrotic spots and streaks on both sides of the petioles and the infected plant parts gradually shriveled up. The disease caused only slight injury and the plant rapidly regenerated new leaves and petioles. This is possibly the first published paper concerned with phytopathogens as controls for waterhyacinth as indicated by the authors' concluding statement : The infection takes place readily, but owing to the high resisting power of the plant, the disease makes very slow progress. From this it may be inferred that this fungus cannot be regarded as a possible remedy against the spread of waterhyacinth (2). Ten years later, Banerjee (7) identified the causal agent as F. equiseti and Snyder and Hansen (169) reduced this species to synonymy with F. roseum. A recent survey of Florida for diseases of waterhyacinth resulted in the isolation of this same species (F. roseum ) from diseased plants in Lake Griffin near Leesburg (154). This report was the first of a F. roseum isolate affecting waterhyacinth in the western hemisphere. The disease is characterized by chlorosis and vascular discoloration in advance of necrosis which proceeds towards the leaf tip. The leaf spot, however

PAGE 26

10 did not expand over the entire leaf surface but remained localized. This is in line with that described by Agharkar and Eanerjee in their original report (2). In 1946, Padwlck (133) reported two species of fungi pathogenic to waterhyacinth . The first, Rhizoctonia solani ( Corticum solani ) , was isolated near Dacca, Bengal, from infected leaves and petioles. It caused extensive blotching and streaking, often killing individual plants. Some 20 years later, R. solani was again reported on waterhyacinth from India by Nag Raj and Ponnappa (124). During surveys for phytopathogens in the Canal Zone of Panama, Freeman and Zettler isolated a R. solani from the anchoring hyacinth (E. azurea ) which proved to be extremely pathogenic on the floating hyacinth (56). In addition, sclerotia of this fungus were able to maintain their viability without loss of virulence after being submersed in lake water for 26 months (56). Disease symptoms on E. crassipes were severe blighting of the emersed portions of the plant which frequently resulted in death of the entire plant. Although R. solani is an agressive pathogen of waterhyacinth, it cannot be considered as a biocontrol at this time, because of its wide pathogenicity to a number of economically Important hosts (133). The second fungal species reported by Padwick (133) was Ceohalosporlum eichhornae Padwick sp . nov. It induced

PAGE 27

11 large, oval, buff-colored spots on the leaves which were covered with a white mat of mycelium. In 1973, Rintz (153) reported another Cephalosporium species, C. zonatum , as causing a zonal leaf spot disease of waterhyacinth in Louisiana and Florida. His report was the fourth pathogen described as occurring on waterhyacinth in the United States. There was some discrepancy as to the synonomy of these two Cephalosporium species (162) and the Commonwealth Mycological Institute reduced them to synonomy, with C. zonatum being the preferred name (123, 153). However, several years later, C_. zonatum was reclassified and is presently placed in the form genus Acremonium of the class Hyphomycetes (86). It is this fungus, Acremonium zonatum (Sawada) Gams, which was studied as a biocontrol agent for waterhyacinths in the present paper. A concentrated research program on biological control of aquatic weeds at the Indian Station of the Commonwealth of Biological Control in Bangalore has resulted in the isolation of several species of phytopathogenic fungi. In 1965, Nag Raj (122) reported a thread blight of waterhyacinth occurring in Calicut, India. Subsequent isolations showed the fungus Marasmiellus inoderma (Berk.) Sing, to be the causal agent (122). The diseased plants in the field exhibited necrotic areas on the leaves, petioles, and all aerial Darts. The infection was more evident in dense

PAGE 28

12 stands of the weed and death of individual plants occurred in irregular patches (122). Infection by M. inoderma under laboratory conditions spreads very rapidly on host plants which is a distinct advantage for a potential biocontrol agent . In 1970, Ponnappa (142), working at the same Indian laboratory, isolated the fungus Myrothecium roridum from waterhyacinth . Although this organism caused extensive damage to E. crassipes , its usefulness as a biocontrol agent cannot be considered at this time because of its pathogenicity on a number of important economic crops (142). This fungus was also reported on waterhyacinth from India by Charudattan in 1973 (21). One fungus which appears to have good potential as a control agent is Alternaria eichhorniae , isolated and described by Nag Raj and Ponnappa (125). It was isolated in India in 1970 and was proved the causal agent of a leaf blight disease. Leaf spots frequently covered the majority of the leaf and caused premature death of those leaves. In culture, A. eichhorniae produces a bright-red diffusable pigment which deepens with age. In addition, it also produces a host-specific toxic matabolite that causes necrotic lesions when placed on leaves or petioles. The host range of this fungus was tested on 42 genera of plants in 15 families Including aquatics and such important

PAGE 29

13 terrestrial familes as Brassicaceae , Fabaceae , and Solanaceae. The results showed A. eichhorniae to be nonpathogenic on all plants tested except the waterhyacinth (125). Its host specificity along with its specific toxic metabolite enhances its potential as a biocontrol agent. A similar species of this fungus was isolated in 1973 by McCorquodale , Martyn, and Sturrock (113) from water hyacinth in south Florida and tentatively identified as A. eichhorniae var . floradana (114). It resembled that described by Nag Raj and Ponnappa (125) in host specificity, conidial size, and toxin production, but differed in pigment production and gemmae formation. This is the first report of this species in the United States. Tests indicate that A. eichhorniae has good potential as a biocontrol agent of waterhyacinth, but because it is not indigenous to the United States it is under strict quarantine by the U.S. Department of Agriculture. For this reason, A. eichhorniae cannot be adequately field tested in Florida at the present time. A second Cercospora species, C. rodmanii was isolated from diseased waterhyacinth in 1973 in Florida (55) and is currently being evaluated as a biocontrol agent (30). Symptoms of the disease on waterhyacinth include general chlorosis of the plant, failure to produce offshoots, spindly petioles and a root rot. Field trials

PAGE 30

14 indicated that the fungus greatly reduced the waterhyacinth population in test plots, but did not eradicate it since new growth appeared which continued to spread (30). In summary, among the phytopathogens reported on waterhyacinth, some are capable of inducing severe damage and even death of the plant. The fact remains, however, that there are relatively few capable of causing such severe diseases. Most of those that do, however, are also pathogenic to important cash crops and therefore unacceptable as biocontrol agents at the present time. Consequently, it would be a great advantage if one or more of the pathogens with a narrow or restricted host range could be utilized. With this in mind, the intent of this study was to examine the pathological relationship of E. crassipes and A. zonatum in an effort to more fully understand the basis of disease resistance and pathogenesis in this host-parasite couplet.

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CHAPTER I RESPONSES OP WATERHYACINTHS TO INFECTION WITH ACREMONIUM ZONATUM AND ITS IMPLICATIONS IN BIOLOGICAL CONTROL Introduction Research into biological control of noxious aquatic plants was initiated at the University of Florida, Department of Plant Pathology, in 1970. Major emphasis was placed on finding diseases of waterhyacinth , alligatorweed , hydrilla, and Eurasian watermilfoil . Surveys for diseases of these plants were made throughout Florida and portions of Alabama, Maryland, Louisiana, Georgia, South Carolina, the Chesapeake Bay, and the Tennessee Valley areas (55). Surveys were also made In ten other countries including most of the Caribbean and eight states In India (55). During these surveys, several diseases were found and the causal agents Isolated for further study (22,23,24,83,95). In 1971, a zonal leaf spot of waterhyacinth was first noted in Puerto Rico where it caused considerable damage to the plant (55). The causal organism was not isolated. However, a similar disorder was subsequently found in the Spring Bayou region of Louisiana. A species of the fungus, Cephalospcrium, was isolated from those plants, and upon inoculation, onto healthy plants, induced symptoms typical 15

PAGE 32

of those observed under natural conditions. The causal agent was ultimately identified as Cephalosporium zonatum Sawada (153) and was originally described as the causal agent of zonal leafspot disease of figs in Louisiana (177). This disease was found since to occur on waterhyacinths in El Salvador, India, Panama, and at two locations in Florida (55). The causal agent, Cephalosporium zonatum, (Sawada) recently was reclassified to Acremonium zonatum (Sawada) Gams (86). The disease is first evident as small sunken lesions on both leaf surfaces and the petiole (153). Under conditions of high humidity, A. zonatum causes severe spotting and death of leaves (107). The lesions are characteristically zonate , oval to irregular in shape, and often coalesce covering the entire surface. Alternating lightand darkbrown bands are typical of the lesions. Under conditions of prolonged high humidity the fungus produces abundant white mycelia on the leaf surfaces and sporulates intermittently. Rintz (153) reported that A. zonatum can attack a wide range of plants under artificial conditions. Despite this apparent wide host range, reports of its occurrence on hosts other than fig in North America are unknown. Consequently, this fungus need not necessarily be excluded from consideration as a possible biocontrol agent of waterhyacinth (55).

PAGE 33

17 During field trials with this fungus in Gainesville, it was observed that small, young plants displayed fewer lesions after infection than did larger plants in the same plot. (T.E. Freeman, personal communication, 1974). In addition, it was observed that some of the infected plants appeared to produce more new leaf growth than did either other diseased plants or control plants. The present study was initiated to determine if small plants were more resistant to A. zonatum than large plants and also if there was an accelerated leaf regeneration in response to infection. Materials and Methods Quantitation of disease Waterhyacinths were collected from natural infestations in south Florida and maintained under greenhouse conditions in Gainesville. Plants were separated into three size categories based upon leaf surface area: (i) small plants, with 2 leaves less than 15 cm , (ii) medium plants, with leaves 152 40 cm , and (iii) large plants, with leaves greater than 40 2 cm . The plants were inoculated by swabbing the leaves with a 10% (wt./vol.) slurry of A. zonatum (grown on potato dextrose agar) and 0.75% water agar. Flants were maintained in ten-gallon glass aquaria half-filled with tap water with plastic covers to maintain the humidity at 99-100%. Control plants were inoculated with sterile 0.75% water agar and

PAGE 34

maintained under identical conditions. Two weeks postinoculation, leaves were excised and used for subsequent tests . Twenty-five to seventy-five leaves from each plant size group were removed and the number of lesions/leaf counted. Mean percentage figures were determined for (i) number of leaves with one or more lesions/leaf, (ii) number of leaves with ten or more lesions/leaf, and (iii) mean number of lesions/ leaf. The total diseased area on each leaf was calculated by the dot counting method ( "Stippentelplaat j e" , J.C. Zadoks , unpublished) and the mean percent diseased area determined for each plant size. Quantitation of leaf regeneration Plants from each size category were selected at random, trimmed of any necrotic or senescent leaves, and inoculated as before. The total number of leaves on each plant was noted prior to inoculation. The plants were maintained in ten-gallon glass aquaria half-filled with tap water and fitted with plastic covers as before. Control plants from each size category were painted with 0.75% sterile water agar and maintained under identical conditions. After two weeks, the total number of leaves on each plant was counted and percent new leaf growth figure was calculated for each plant size group. In addition, ten plants from each size category were

PAGE 35

19 selected and the number of leaves/plant noted. Each leaf was then excised and the plants placed in aquaria. After two weeks, a percent new leaf growth figure was determined for each plant size. Results Quantitation of disease Inoculated waterhyacinths kept under conditions of high humidity were severely damaged by A. zonatum (Fig. 1-1). Necrotic lesions varying in size and number occurred on both leaf surfaces and the petioles. On the average, 70.1% of the leaves on small plants had at least one lesion while 98.2% of medium leaves had at least one lesion (Fig. 1-2). Large plants had 133% of their leaves infected indicating a secondary spread to the new growth during the course of disease development. These percentages decreased when the number of leaves exhibiting ten or more lesions was calculated but the trend was the same. Large plants had significantly more leaves with ten lesions (83.3%) than did either medium (53.6%) or small (29.8%) plants. The average number of lesions/leaf also followed the same pattern. Small plants averaged 3.7 lesions/leaf while medium and large averaged 12.8 and 18.3 respectively. However, when the total diseased leaf area was measured after two weeks, there was no significant difference among small, medium, and large

PAGE 36

20 plants, each exhibiting approximately 40% diseased leaf area (Fig. 1-2). Quantitation of leaf regeneration On the average, after two weeks of growth, small healthy waterhyacinths regenerated 27.3% new leaves or one new leaf/plant (Fig 1-3). There was no significant difference at the 0.01 confidence level in the percentage of new leaves produced by small diseased plants. Small plants, after infection, regenerated 21.6% new leaves or 0.95 leaves/plant. Likewise, there was no significant difference between the new leaves produced by healthy and diseased medium-sized plants. Medium-sized control plants produced 28.5% new leaves during the two weeks while diseased plants of the same size regenerated 33.9% new leaves. A trend was noted, however, that as the plant increased in size, its rate of new leaf production also increased. When the number of new leaves produced by large healthy plants was compared to that from large diseased plants, there was a significant difference. Large healthy plants normally regenerated 46.1% of their leaves over the two week period; however, diseased large plants produced 93.3% new leaves, an increase of almost 50% (Fig. 1-3). In addition, the average number of leaves/plant for large plants increased from 2.0 in healthy to 4 . 6 in diseased plants. Small and large plants which had their leaves excised

PAGE 37

21 prior to the test displayed little variation in the number of new leaves when compared to the controls (Fig. 1-3). Small plants regenerated 25.2% of their leaves compared to 27.3% new growth in normal small plants. Likewise, large plants displayed little difference in new leaf production between control plants and those in which the leaves were excised (46.1% and 50.0% respectively). A notable exception was observed with medium-sized plants. Controls produced 28.5% new leaves in two weeks while plants of the same size whose leaves were removed first regenerated only 19.3% new leaves .

PAGE 38

Figure 1-1 (a d). Symptoms of disease on waterhyacinths incited by Acremonium zonatum . a. Waterhyacinth with zonate lesions on leaves and petioles . b. Close-up of waterhyacinth plants two weeks postinoculation . c. Waterhyacinth leaf showing coalescence of leaf spots . d. Large waterhyacinth leaf with abundant white mycelia of A. zonatum.

PAGE 39

n

PAGE 40

Figure 1-2. Quantitation of disease on small, medium and large waterhyacinths . Small = plants with leaves < 15 cm surface area; medium = plants wi£;h leaves 15-40 cm surface area; large = plants > 40 cm surface area. a= % leaves with 1 or more lesions/leaf; b= % leaves with 10 or more lesions/leaf: c = mean number of lesions/leaf; d= % total diseased leaf area.

PAGE 41

25 125 100 UJ < uj 75 oo O 50 25 / 1 5 ±: 1 abed SMALL bed MEDIUM *^v 5 abed LARGE 20 U. 15 < O Ul > <

PAGE 42

Figure 1-3. Quantitation of leaf regeneration rates of small, medium, and large waterhyacinths C= control _ plants; 1= inoculated plants; E= plants with leaves excised prior to test.

PAGE 43

27 O O t 1 1 1 1 r o en o CD i r KWAWWWW Ld LU ! O F^^ LU ID Q LU KWWW^ LJ o ] < CO J I I I L J I « o o o to o o ro o o CM — HIMOcdO JV31 M3N %

PAGE 44

28 Discussion The concept of biological control, the use of one organism to control another, although not new in practice is relatively new in its wide-scale applications. Debach (40) cites that the introduction of the mynah bird from India to Mauritues in 1762 to control the red locust was the first successful attempt at biological control. Perhaps the first successful deliberate control of one organism with another in the United States was the introduction of the vedalia beetle into California in 1888 to control cottony-cushion scale of citrus (6). Control of one organism by another has been referred to as "parasitic control" and "the biological method" but it wasn't until 1919 that H.S. Smith referred to it as "biological control" (40). It is a difficult task to impart a precise definition to the term "biological control" since there is little unanimity on this point among plant pathologist. Perhaps the best definition is that given by Baker and Cook (6) . . Biological control is the reduction of inoculum density or disease-producing activities of a pathogen or parasite in its active or dormant state, by one or more organisms, accomplished naturally or through manipulation of the environment, host, or antagonist, or by mass introduction of one or more antagonists. The above definition encompasses several points not dealt with in this dissertation. For convenience and ease of understanding in the present discussion, the term "biological control" will be used to imply the use of native or

PAGE 45

29 introduced organisms to control or reduce the population of another organism through an antagonistic or parasitic relationship. Thus, the use of the fungus A. zonatum to parasitize the waterhyacinth and thereby reduce its population size is well within the scope of the definition by Baker and Cook ( 6 ) . Plant pathologists are generally concerned with controlling epiphytotics--not starting them. However, this is not the case when control of a noxious weed such as the waterhyacinth is desired through biological methods. Therefore, it takes some adjustment in one's own thinking when the initial idea is presented. When an alien plant establishes itself in a particular habitat it may mean several things: (i) it is better suited to a particular niche than are the residents, (ii) that it was introduced in such numbers as to temporarily or permanently "swamp" the residents, or, (iii) that it may modify the environment in some way favorable to itself. It usually means, however, that man has upset the natural balance in some way, making the environment more favorable to the alien than to the resident. Such has been the case with the waterhyacinth. Over-nutrif ication of our waters by man's increasing agricultural and urban demands has been the single most contributing factor to the aquatic weed problem. Thus a weed is "a generally unwanted organism that thrives in habitats disturbed by man" (6).

PAGE 46

30 The first step in a biological control program is, in most cases, the evaluation of the potential biocontrol agent. This usually involves the introduction of the control agent onto its target host and/or additional potential hosts under greenhouse conditions. Thus the effectiveness of the control agent on its target host can be determined as well as its potential to parasitize other crops for which it was not intended. If a potential biocontrol agent passes the initial greenhouse tests, field trials are usually initiated. In these studies, evaluations as to how the control agent manifests itself and its ability to compete with the other biotic agents present can be made. In some instances, it may be necessary to bring the control agent back into the laboratory and greenhouse to further evaluate situations observed in the field. The above description depicts the studies conducted on A. zonatum over the past five years. Isolation and pathogenicity tests of the fungus under greenhouse conditions were initially done by Rintz (152). Field trials with A. zonatum were initiated in 1973 by Freeman, et al . (54) on well established stands of waterhyacinths In Lake Alice on the campus of the University of Florida. It was during these studies that apparent differences in symptoms and growth rates were noticed on the plants. In 1974, greenhouse tests were initiated once again to

PAGE 47

31 see how host plant size influenced A. zonatum as to infection and subsequent disease. Disease measurement is often regarded as a synonym for "estimation of losses," but this is misleading (92). There is a great need for some reasonably simple but critical parameters that can be used consistently and systematically to measure the prevalence and severity of plant diseases in the field. On the other hand, there are no portmanteau methods that will serve for all plant diseases. Some of the currently accepted disease assessment techniques are discussed by Large (92) and include such things as standard diagrams, the Korsfall and Barratt gracing system, and disease progress curves. Perhaps one of the easiest techniques to use is the standard diagram method. This, of course, assumes that standard diagrams for the particular host-parasite couplet in qestion have been constructed. If not, then this method requires the researcher to work cut such diagrams. In the case cf E. crassipes A. zonatum , standard disease diagrams have not been constructed. For this reason, disease assessment was based on two criteria: (i) number of lesions/leaf, and (ii) total percent of diseased area/leaf. Both of these methods have been used routinely with other hcst-parasite combinations and are the basis of standard diagram keys. Lesion counts on different size waterhyacinths indicated initially that small plants were more resistant than

PAGE 48

32 large plants since they exhibited fewer lesions/leaf. However, when the mean percent diseased area of each leaf was measured, there were no significant differences among any of the three sizes, all showing approximately 40% disease severity. This allows for two possibilities. First, small plants are more resistant to initial attack, but over the two week infection period gradually lose this resistance and obtain a level of susceptibility shown by the larger plants, or secondly, large plants are more susceptible initially but gradually build up a resistance. Based upon data presented elsewhere in this dissertation, i.e. polyphenoloxidase rates and phenolic acid concentrations (see Chapter III), it is believed that a combination of both mechanisms is involved. That is, small plants gradually lose some of their initial resistance while larger plants gain various degrees of resistance. That plants may increase or decrease in susceptibility to a particular pathogen with age is well documented (197). It has been suggested (196) that susceptibility to facultative saprophytes increases with age of host tissues, whereas isceptibility to obligate parasites decreases with age although this does not always hold true. Based upon the results of the present study with waterhyacinths, susceptibility to attack by A. zonatum increases with plant size. Generally, plant size can be correlated

PAGE 49

33 with ontogenetic development, that is, the older the plant, the larger is its size. However, this may not always be a correct assumption with waterhyacinths since growth rate depends upon environmental conditions of its habitat (light intensity, nutrients, and temperature). For this reason, then, predisposition to A. zonatum in nature due to host age may be only part of the answer. Differences in symptom expression during field trials with this fungus may then be the result of several predisposing factors operating in conjunction with one another. Water quality was not monitored during field trials with this fungus so the effect of environmental predisposing factors cannot be discussed. However, waterhyacinths used in the greenhouse studies were all maintained under the same environmental parameters. Since the only variable in these tests was the age of the host, it can be stated that susceptibility of waterhyacinths to A. zonatum increases with the ontogenetic development of the plant. This is an important criterion when considering the use of any agent as a control measure. Time of application is extremely important in order to obtain the most effective control. Another very important observation made during these studies was that of the leaf regeneration rates of different plant sizes after infection. As healthy plants increase in size (small to medium to large) their leaf regeneration rate

PAGE 50

34 increases. That is, small plants regenerate approximately 27% of their leaves in two weeks or about one leaf /plant. Medium-sized plants produced a slightly higher, but insignificant, percentage rate of 28.5 or 1.5 leaves/plant. Large plants, however, are able to reproduce almost half of their total leaves within a two week period (46.1%). When plants are inoculated with A. zonatum , their leaf regeneration rates are altered. There is a slight reduction in new leaf production exhibited by infected small plants (5.7%) and a slight increase shown by infected mediumsized plants (5.4%). But the significant difference is demonstrated by infected large plants. With these there is a two-fold increase in new leaves after two weeks. The rise from 46.1% to 93.5% in large plants represents an increase on the average from 2.0 new leaves/plant to 4 . 6 new leaves/ plant . Because A. zonatum is a leaf spotting pathogen, it was postulated that accelerated leaf production was a response to photosynthetic stress placed upon it by infection which resulted in the destruction of most of its photosynthetically active tissue. In order to test this idea leaves were excised from a set of each of the three plant sizes and monitored for new leaf growth. There was little variation in the percentages of new leaf growth when compared to their respective controls. In one case (medium-sized plants)

PAGE 51

35 there appeared to be a deleterious effect on the plant's normal leaf production rate. It would appear, then, that the accelerated new leaf production observed in waterhyacinths after infection by A. zonatum is not a response to the destruction of photosynthetically active tissue, but one of interaction between the host and the pathogen. Accelerated growth rates in diseased plants has often been correlated with increased activity of growth regulators (61). Normal growth in plants is under hormonal control by such biologically active endogenous compounds as 6-indole3-acetic acid (IAA, auxin), gibberellins , cytokinins , and others (61). A departure from the normal levels of these compounds in the plant, such as might be caused by pathogenic attack, could alter the growth habit of the host. Data presented in Chapter III show that A. zonatum is capable of synthesizing high amounts of auxin in vitro when given the amino acid tryptophan as a precursor. Even though this does not represent conclusive evidence for the production of auxin in vivo by this organism, it does suggest its possibility. In addition, it has been suggested (87,132) that Increased levels of auxin in diseased tissue may be correlated with the inhibition of IAA oxidase in the plants by phenolic inhibitors. Further implications on the possible roles of auxin, IAA oxidase, and phenolic compounds during pathogenesis are discussed in Chapter III.

PAGE 52

36 In contemplating A. zonatum as a biocontrol agent of waterhyacinths , several criteria must be considered. Foremost is the proper time at which to apply the inoculum. Results in this study have indicated that small, young plants are more resistant to fungal attack than are larger, older plants. Eased on this, the fungus should perhaps be applied late in the spring or summer when the plants have reached maturity. On the other hand, data indicated that the plants respond to infection by accelerating their rate of leaf regeneration and that large plants do this more quickly than do smaller ones. In essence, then, application of the fungus to large plants would appear to negate or minimize any control afforded by the pathogen. When, then would be the best time to apply the control agent? Since disease severity proceeds to approximately 40% within two weeks, regardless of the plant size, application early In the spring, as the new season's growth is beginning, would appear to be the best time. In this manner one could avoid the accelerated leaf growth response displayed by larger plants while ai the same time expect substantial damage to the plant.

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CHAPTER II A CYTOCHEMICAL AND ULTRASTRUCTURAL STUDY OF THE PHENOL CELLS AND POLYPHENOLOXIDASE ACTIVITIES IN HEALTHY AND DISEASED WATERHYACINTH LEAVES Introduction Phenolic compounds are among the most widespread and varied compounds in plants. Perhaps the best known role for plant phenolics is their assimilation Into the anthocyanins and flavone pigments (150). However, as many authors have indicated, phenolic compounds have nearly unlimited potential in accounting for the many differences that occur in disease resistance (12,34,35,50,90,157,179). Phenols are particularly abundant in the leaves of many plants. They are also found in the xylem, phloem, and periderm of stems and roots; In unripe fruits; in the testa of seeds; and in pathological growths such as galls (49). Phenolic compounds in plants may be present in individual cells or in specialized idioblasts termed tannin sacs (49) or phenol-storing cells (119). Recent studies have shown that specialized phenol-storing cells occur randomly in several plant species (10,11,100,107,119,120). Phenols may be a common Ingredient of the vacuoles or they may occur In the cytoplasm proper in the form of small droplets which eventually fuse (49). 37

PAGE 54

38 In many plant tissues, phenols become oxidized to polymeric dark red or brown compounds (phlobaphenes ) , which are sometimes microscopically visible in the cell contents of fresh sections. Oxidation of phenolic compound accounts for the pathological darkening in plant tissues (38). Histochemical detection of naturally occurring phenols is difficult because few reagents that react with them to form characteristic color compounds are adaptable to histological methods (148). In addition, the natural enzymatic browning may not be sufficiently intense for easy detection microscopically. In 1951, Reeve (148) described a histochemical test for phenols in fresh plant tissue. It is based upon a colorimetric method for phenols using a nitrous acid reaction. The method has become widely accepted and used and is often referred to as the "nitroso reaction." One of the enzymes associated with the oxidation of phenolic compounds is polyphenoloxidase (PPO). The term polyphenoloxidase has been used extensively in the literature, although the names phenolase, phenoloxidase , catecholoxidase, and tyrosinase have been used as synonyms. Classification of this enzyme is difficult because several different activities have been described for it. The enzyme as originally termed tyrosinase since the aromatic amino Id, tyrosine, was the first experimental substrate (38). : .'-ever, p-cresol and catechol have been most frequently

PAGE 55

39 employed as experimental substrates. Consequently, two activities have been ascribed and have come to be known as the "cresolase" activity when referring to monohydric phenol oxidation and "catecholase" activity when referring to odihydric phenol oxidation (38). Many different phenolic compounds can serve as substrates for polyphenoloxidases . For sake of convenience these enzymes have been divided into three main groups (155) based upon their affinity for certain substrates, response to inhibitors, and type of reaction catalyzed: (i) Tyrosinases enzymes of this group catalyze both o-hydroxylation of monophenols and the oxidation of o-diphenols. (ii) Ortho-diphenoloxidases these enzymes, unlike the tyrosinases, are devoid of hydroxylation properties and act only on o-diphenols. (iii) Para-diphenoloxidases members of this group act primarily on p-diphenols but may also have some affinity for the oxidation of certain o-diphenols. The laccases can be classified in this category. In the present discussion, the term polyphenoloxidase has been retained whenever the oxidation activity is being described regardless of whether it is acting upon an oor p-diphenol. For a detailed review of the polyphenoloxidases, the reader is referred to Dawson and Magie (38), Nelson and Dawson (128), and Patil and Zucker (138). Some cells are capable of converting tyrosine into a

PAGE 56

40 brown or black pigment called melanin (48). The pathway for this conversion is depicted in Figure II-l. The first step involves an o-hydroxylation of tyrosine thereby forming dihydroxyphenylalanine (DOPA). The enzyme that catalyzes this conversion is in the tyrosinase group and consequently can also oxidize DOPA in the second step to dopaquinone. Polyphenoloxidases are devoid of any hydroxylat ion properties and therefore cannot convert tyrosine to DOPA but are capable of oxidizing it to dopaquinone. It is this property which has been investigated as a marker for this enzyme in vivo . Polyphenoloxidase activity has long been thought to reside within the chloroplasts of plant cells (5), but until recently cytochemical localization had not been demonstrated. Based on techniques developed by Novikoff et al . (129) and Okun et al . (132) for the localization of tyrosinase in animal tissues, Czaninski and Catesson (36,37) have recently demonstrated the cytochemical localization of PPO in plant cells. Since 1972, several investigators (72,74,107,134,135) have shown that PPO activity is localized within the thylakoids of chloroplasts in several plant species. This chapter presents the results of a histochemical and ultrastructural study of the phenol cells in waterhyacinth leaves and the cytochemical localization of PPO in healthy and diseased plants.

PAGE 57

Figure II-l. Biosynthetic pathway for conversion of tyrosine to melanin [after Eppig (48)].

PAGE 58

M2

PAGE 59

43 Materials and Methods Histochemical localization of phen ols Cross sections of fresh waterhyacinth leaf tissue (1224y) from small, medium, and large plants were made with a Hooker plant microtome, tested for phenols by the nitroso reaction (148), and observed with the light microscope. With this method a nitroso derivative of the phenolic compound is formed and after addition of the base, a brightred salt is formed. Spatial distribution of phenol cells The spatial distribution of the subepidermal phenol cells from each size category was determined from tangential sections made along the vascular bundles. Sections of the leaves (10 x 15 mm) were taken from areas selected at random and the epidermal surfaces separated from each other with a razor blade. Each half was then stained for phenols as previously described and observed with the light microscope. 2 The mean number of phenol cells/mm leaf tissue was calculated for the top and bottom surfaces of each plant size group. Electron microscopy Standard fixation and embedding procedures were used throughout with slight modifications as presented below. A flow diagram for the basic technique is presented in Figure II-2 . Fresh waterhyacinth leaf tissue was placed in a

PAGE 60

Figure II-2 . Flow diagram of procedure for standard electron microscopy fixation and embedding. 1 2% glutaraldehyde paraformaldehyde 2 0.2 M sodium cacodylate , pH 7.2 3 Spurr, 1969 (172)

PAGE 61

fresh tissue fix in Karnovsky's fixative (2hr-22C) I 2 wash in buffer (4x) post-fix in 1% 0s0 4 (lhr.-22C) 45 wash in buffer (4x) i dehydrate in 25%EtOH series transfer to 100% acetone embed in epoxy resin section post-stain w/ UrAc (lOmin.) poststain w/ PbCi (5min.) Flow Diagram for Electron Microscopy Fixation and Embedding

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46 buffered (0.2 M sodium cacodylate, pH 7.2) solution of 2.0% glutaraldehyde and 2.0% paraformaldehyde (85). Each leaf was cut into 3-5 mm pieces and fixed for two hours at room temperature. The material was washed in 50% buffer 50% distilled water solution for a minimum of 30 minutes before being postfixed in 1.0% osmium tetraoxide for one hour at 22 C. Sections were then rinsed several times with the aqueousbuffer mixture and passed through an ethanol graded dehydration series at 25% increments and finally into 100% acetone. After dehydration the sections were infiltrated with a graded acetone-plastic series and embedded in a 100% low viscosity epoxy resin (170). The embedded sections were then placed under vacuum for five minutes to remove bubbles and the resin was polymerized for 18 hours in a 60 C oven. Thin sections were cut on a Sorvall KT-2 ultramicrotome with a diamond knife and placed on single-hole, Formvar coated grids. Sections were then poststained in 0.5% uranyl acetate for ten minutes and in 1.0% lead citrate for five minutes. The sections were examined with a Hitachi HU HE electron microscope. Cyto chemical localization of polyphenoloxidase The procedure for the localization of PPO activity in wati ohyacinth leaves follows closely that described by C,i.;:ninski and Catesson (37). A flow diagram of this procedure is presented in Figure II-3. Fresh leaf tissue, both

PAGE 63

Figure II-3. Flow diagram of procedure for the cytochemical localization of polyphenoloxidase . redistilled glutaraldehyde n 0.2 M sodium cacodylate , pH 7 . 2 0.0 2 M sodium diethyldithiocarbamate 4 L-dihydroxyphenylalanine (50 mg/10 ml 0.067 M phosphate buffer, pH 7.0)

PAGE 64

fresh leaf sections fix in 5% glut. 1 i wash in buffer(5x) treat w/ DDC wash in bu fer(5x) boil sections OOmin.) pre-incubate w/DOPA incubate w/DOPA (lhr.-37C) wash in d.w.sucrose (5x) U8 r post-stain w/ PbCi post-fix w/ 2% 0s0 4 (2hr. -22C) I dehydrate in EtOH I embed in epoxy resin I section } no poststain Flow Diagram for Cytochemical Localization of PPO Activity

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49 healthy and diseased, was placed in buffer as before and cut into 2-4 mm pieces. The sections were fixed in 5.0% glutaraldehyde for 11/2 hours at room temparature and washed in buffer 5 times for 15 minutes each. The sections were then separated into three groups and treated by one of the following methods: (i) boiled for ten minutes, (ii) incubated in 0.02 M DDC (sodium diethyldithiocarbamate ) for 20 minutes at 22 C and then washed 5 times in buffer, and (iii) no treatment. After their respective treatments, each group was preincubated in a DOPA substrate solution (50 mg DOPA in 10 ml of 0.067 M phosphate buffer, pH 7.0, made up fresh) at 4 C overnight. After the preincubation period, the sections were incubated in fresh DOPA for one hour (fresh solution added after 30 minutes) at 37 C, followed by five washings in distilled water made to 0.5 M with sucrose. After postfixing in 1.0% osmium tetraoxide they were dehydrated, embedded in epoxy resin, sectioned, and examined with the electron microscope as previously described. Results Histcchemical localization of phenols When waterhyacinth leaves were stained for phenols by the nitroso reaction, these compounds were found in large, specialized idioblasts or phenol cells immediately beneath both epidermal surfaces (Figs. II-4a £ b) and in cells

PAGE 66

50 closely associated with the vascular bundles (Fig. II-4c). The size of these cells in the palisade layer varied considerably, often exceeding several hundred microns in length and extending down to the vascular elements. Those phenol cells near the vascular tissue were much more isodiametric and varied much less in size. There was no significant difference in morphology of the cells among the three plant sizes examined. Spatial distribution of phenol cells Phenol cells occurred randomly beneath both leaf surfaces in all plant sizes and were found throughout the entire leaf (Fig. II-4d). There were significantly more 2 phenol cells beneath the adaxial leaf surface (40.6/mm ) 2 than on the abaxial surface (26.6/mm ) in small plants but the reverse was true for medium and large plants (Fig. II5). Medium and large plants exhibited a more equal distribution of phenol cells between the two surfaces but there was a significantly greater number on the top (51.8/41.8 in medium vs 54.2/4 8.7 in large). The total number of phenol 2 . . cells/mm , both adaxial and abaxial surfaces, significantly increased as the leaf increased in area with a mean of 2 2 2 3?.6/mm for small, 41.8/mm for medium, and 48.7/mm for J n-ge. !J I trastructure of phenol cells Electron micrographs indicate that in most cases the

PAGE 67

51 subepidermal phenol cells were two to three times longer than the adjacent palisade cells (Fig. II-6). The phenolic compounds appeared in close association with the tonoplast and as discrete bodies within the cells. These were actively metabolizing cells containing nuclei, mitochondria, and plastids. In contrast, the phenol cells near the level of the vascular tissue were much more circular, had a thicker wall, and the phenolic compounds were in amorphous masses as opposed to discrete globules (Fig. II-7). There were no morphological differences observed between phenol cells of the same type in any of the plant sizes examined. Cytochemical localization of polyphenoloxidase The principle of the reaction for the cytochemical localization of PPO activity involves obtaining an insoluble, electron dense reaction product ( dopaquinone ) from the synthetic substrate at the point where enzyme activity is proceeding (37). Although the reaction can be observed without additional staining, the intensity of the reaction and the clarity of the surrounding material is enhanced by poststaining with lead citrate. When examined by this hnique , a positive PPO reaction product was absent in all cl'" oplasts of small and large healthy waterhyacinth leaves incubated without DOPA. Chloroplasts in palisade cells (Pi;-;. II-8a) , have distinctly clear thylakoid spaces and fret channels. Similar observations were made for

PAGE 68

52 chloroplasts of bundle sheath cells (Fig. II-8b), vascular parenchyma (Fig. II-8c), and phenol cells (Fig. II-8d). The thylakoids within the chloroplasts of phenol cells were not readily detected until poststained with lead citrate. Sections from both small and large healthy leaves incubated with DOPA reacted in an identical manner for the localization of PPO . Chloroplasts of the palisade cells (Fig. II-9a) and spongy mesophyl cells (Fig. II-9b) did not stain for PPO activity. On the other hand, PPO activity was localized in the thylakoids of chloroplasts in three other cell types, two of which were associated with the vascular tissue. In each instance, the thylakoid spaces and fret channels were the only areas stained for PPO activity. In contrast to other cells, chloroplasts of the vascular parenchyma, both phloem parenchyma (Fig. IISd) and xylem parenchyma (Fig. 11-10) were PPO positive. The chloroplasts in these cells appeared black or electron-dense. These electron-dense areas were restricted to the thylakoids within rhe chloroplasts (Fig. II-10b). Chloroplasts which were not poststained (Figs. II-9d 5 II-10c) also showed a positive reaction but the intensity and clarity was not as good . Another type of cell having PPO positive chloroplasts were the bundle sheath cells (Figs. II-9c and II-lla £ b). Waterhyscinths are typical monocots and have a large bundle

PAGE 69

53 sheath surrounding the vascular elements. Chloroplasts in these bundle sheath cells were PPO positive, although perhaps not as intense as those in the vascular parenchyma. The phenol cell itself also showed PPO activity (Figs. II-9c and 11-12). The reaction in these cells was the most intense of the three. In this cell type, the chloroplasts are extremely electron-dense (Fig. II-12a), and examination under higher magnification revealed that not only were the thylakoids positive, but the entire organelle was electrondense (Fig. II-12b). Leaf material that was boiled prior to incubation in DOPA did not give a positive PPO reaction, in any chloroplasts, indicating heat inactivation of the enzyme after boiling (Fig. 11-13). The thylakoids became distorted after boiling and starch granules swelled forming large lacunae (Fig. II-13a £ c). When the inhibitor, DDC , was added to the sections prior to incubation in DOPA, no reaction product could be detected in the thylakoids of any chloroplasts (Fig. 11-14). When sections were poststained with lead citrate (Fig. II14a), the thylakoid spaces and fret channels contrasted sharply with the stroma. Only the partitions were notably electron-dense. Thus, the electron density of lead citrate cannot be confused with the electron-dense product of a positive PPO reaction. Consequently, use of the poststain

PAGE 70

54 acts to heighten the observed reactions and surrounding material. In addition, PPO activity was not observed in any cell organelle other than chloroplasts . These observations were consistent for each of the plant sizes examined. When diseased leaves were examined for enzyme localization, PPO activity was found to be no longer restricted to vascular parenchyma, bundle sheath, and phenol cells rather every chloroplast in every cell was positive. Palisade cells were now positive (Fig. 11-15) and there was an increase in the number of plastoglobuli in those chloroplasts. Likewise, spongy mesophyl cells, which in healthy cells were negative, became positive after infection (Fig. 11-16). These chloroplasts also showed an increase in the number and size of the plastoglobuli. The changes in PPO localization were apparent in chloroplasts in cells immediately surrounding the lesions. Sections taken several centimeters away from the lesion were examined to determine if periphery cells also showed a "turn-on" in enzyme activity. Electron micrographs indicate that even those cells which are two to five centimeters removed from the center of infection were also positive for PPO activity. Thus, palisade cells became positive (Fig. II-17a £ b), spongy mesophyl cells became positive (Fig. II17 c £ c), and chloroplasts In cells normally positive such as bundle sheath cells became very intense (Fig. II-17d).

PAGE 71

55 In essence, PPO activity was found in the chloroplasts in only three cell types in healthy leaves: (i) vascular parenchyma, (ii) bundle sheath, and (iii) phenol cells proper. However, during disease, there was a turn-on of PPO activity in all cells which contain chloroplasts. Whether this turn-on in enzyme activity is host-induced or pathogeninduced is not known at this time.

PAGE 72

Figure II-U (a d). Light micrographs of phenol cells in healthy waterhyacinth leaves. a. Cross section of waterhyacinth leaf showing arrangement of phenol cells in upper and lower palisade cell layers. (375 X). b. Cross section of waterhyacinth leaf showing phc and vascular bundle (vb). (1,500 X). c. Cross section of waterhyacinth leaf showing phc in relation to vb and bundle sheath cells (bsc). (1,500 X ) . d. Tangential section of waterhyacinth leaf showing spatial arrangement of phc. (37 5 X).

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2 Figure II-5. Number of phenol cells/mm leaf area in small, medium, and large waterhyacinth leaves. ST= small plants, top surface of leaf; SB= small plants, bottom surface of leaf; MT= medium plants, top surface of leaf; ME= medium plants, bottom surface of leaf; LT= large plants, top surface of leaf; LB=large plants, bottom surface of leaf; y= mean number of phenol cells/mm leaf (both surxaces).

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60 50 59 UJ z> CO CO H < LLi £ E v. CO UJ o 40 30 20 10 I ST SB I : ^V S MT MB ]X LW : i LT .-; LB Ji PLANT SIZE

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Figure II-6. Electron micrograph of phenol cell in palisade cell layer of waterhyacinth leaf tissue. Phenol bodies (pb) appear in close association with the plasmalemma and as discrete globules within the tonoplast (t). Poststained with PbCi. (2,140 X).

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61 r / 4 f v \ -,*

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Figure II-7. Electron micrograph of phenol cell in vascular tissue area of waterhyacinth leaf. Phenol bodies (pb) appear as an amorphous mass within the cell x = xylem. Fcststained with PbCi. (9,400 X).

PAGE 79

63

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Figure II-8 (a d). Chloroplasts of healthy waterhyacinth leaf tissue incubated without DOPA. a. Palisade cell chloroplast with clear thylakoids (th). s= starch (29,400 X) . b. Bundle sheath cell chloroplast. cw= cell wall th= thylakoids (37,500 X). c. Vascular parenchyma cell chloroplast. th = thylakoids (30,000 X). d. Phenol cell chloroplast. c= chloroplast, th= thylakoids, pb= phenol body. Poststained with PbCi (24,000 X) .

PAGE 81

65 \ * / cw t

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Figure II-9 (a e). Localization of polyphenoloxidase in healthy waterhyacinth leaf tissue without lead poststaining . a. Palisade ceil chloroplast. Negative PPO activity in thylakoids (th). s= starch (45,500 X). b. Spongy mesophyll cell chloroplast. Negative PPO activity in thylakoids (th). (45,000 X). c. Phenol cell chloroplast (phc). Fositive PPO activity in thylakoids (th). (33,000 X). d. Vascular parenchyma cell chloroplast. Positive PPO activity in thylakoids (th). pl= plastoglcbuli (57,500 X) . e. Bundle sheath cell chloroplast. Positive PPO activity in thylakoids (th). (75,000 X).

PAGE 83

6? th © © J © th. ©

PAGE 84

Figure 11-10 (a c). Localization of polyphenoloxidase in chloroplasts of xylem parenchyma cells in healthy waterhyacinth leaves. a. Cross section of leaf showing a xylem element (x) and surrounding xylem parenchyma cells (xp). Chloroplasts (c) in the xp cells are positive for PPO activity. Poststained with PbCi. (4,800 X) . b. Close-up of chloroplasts in xp showing positive PPO activity between the thylakoids (th) and several plastoglobuli (pi). cw= cell wall. Poststained with PbCi. (26^,000 X). c. Chloroplast in xp cell showing positive PPO activity without PbCi post staining . (16,5 00 X).

PAGE 85

69 % fc v. r?^ — '_ ..-.

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Figure 11-11 (a c). Localization of polyphenoloxidase in chloroplasts of bundle sheath cells in healthy waterhyacinth leaves. a. Chloroplasts (c) in bundle sheath cells (bsc) showing positive PPO activity. Poststained with PbCi. (6,200 X) . b. Close-up of chloroplast in bsc showing positive PPO activity in thylakoids. Poststained with PbCi. cw= cell wall. (32,000 X). c. Chloroplasts in bsc incubated in diethyldithiocarbamate (DDC) prior to incubation in DOPA. Thylakoids (th) are negative for PPO activity. Poststained with PbCi. pm= plasmalemma . ( 4 , X ) .

PAGE 87

71

PAGE 88

Figure 11-12 (a b). Localization of polyphenoloxidase in chloroplasts of phenol cells in healthy waterhyacinth leaves. a. infrastructure of phenol cell in palisade cell layer showing nucleus (n), mitochondrion (in), chloroplasts (c), and phenol bodies (pb). Chloroplasts are positive for PPO activity. Poststained with PbCi. (2,200 X). b. Enlargement of chloroplast in phenol cell showing PPO positive thylakoids (th) and large phenol body (pb) in association with the chloroplast. s= starch. Poststained with PbCi. (57,500 X).

PAGE 89

73 ©

PAGE 90

Figure 11-13 (a d). Chloroplasts of boiled, healthy waterhyacinth leaf tissue incubated with DOPA. a. Spongy mesophyll cell chloroplast showing distended thylakoids (th). cw= cell wall, sl= starch lacuna (37 ,500 X) . b. Vascular parenchyma cell chloroplast showing thylakoids (th) negative for PPO activity. pl= plastoglobuli, m= mitochondrion (69,000 X). c. Bundle sheath cell (bsc) chloroplast (c) with negative PPO activity. mc= mesophyll cell. (7,000 X) . d. Enlargement of bsc chloroplast with negative _ PPO activity. th= thylakoids, pl= plastoglobuli , sl= starch lacuna, cw= cell wall (56,000 X).

PAGE 91

75 th^ -th cw si • m ^ © bsc . • • mc cw © ©

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Figure 11-14 (a d). Chloroplasts of healthy waterhyacinth leaf tissue incubated in inhibitor (DDC) and DOPA. a. Palisade cell chloroplast with distinct thylakoid spaces (th) and fret channels. m= mitochondrion; Poststained with PbCi (40,000 X). b. Vascular parenchyma cell chloroplast with negative PPO activity. th= thylakoids (28,000 X). c. Bundle sheath cell chloroplast with negative PPO activity. th= thylakoids, s= starch (46,000 X). d. Phenol cell chloroplast with negative PPO activity, th= thylakoids, pl= plastoglobuli (55,000 X).

PAGE 93

77 I
PAGE 94

Figure 11-15 (a b). Localization of polyphenoloxidase in chloroplasts of palisade cells from diseased waterhyacinth leaves. a. Necrotic palisade cells (pc) showing positive PPO activity in their chloroplasts and an increase in the size and number of plastoglobuli . Chloroplasts in palisade cells in healthy leaf tissue are negative for PPO activity. Poststained with PbCi. (7,820 X). b. Enlargement of -chloroplasts in palisade cells showing PPO activity in the thylakoids (th). Poststained with PbCi. (24 ,300 X) .

PAGE 95

79

PAGE 96

Figure 11-16 (a c). Localization of polyphenoloxidase in chloroplasts of spongy mesophyll cells from diseased waterhyacinth leaves. a. Mesophyll cells (mc) showing PPO positive chloroplasts. Hyphae (h) shown in upper right corner. Poststained with PbCi. Chloroplasts in mesophyll cells in healthy leaf tissue are negative for PPO activity. (8,280 X). b. Enlargement of mesophyll chloroplast showing positive PPO reaction in thylakoids . m= mitochondrion. Poststained with PbCi. (35,400 X). c. Enlargement of positive PPO chloroplast in mesophyll cell without PbCi poststain. (40,000 X).

PAGE 97

81 _«. * h ^ -<+>

PAGE 98

Figure 11-17 (a d). Localization of polyphenoloxidase in chloroplasts of cells several centimeters away from infection center. a. Palisade cells (pc) with positive PPO activity in their chloroplasts. e = epidermis. Post stained with PbCi. (6,200 X) . b. Enlargement of palisade chloroplast showing PPO activity in the thylakoids (th). Poststained with PbCi. (17,400 X). Mesophyll cell (mc) showing positive PPO activity in the chloroplast. n= nucleus, m= mitochondrion, xh= thylakoids. Poststained with PbCi. (27,500 X). Electron micrograph showing PPO positive chloroplasts in mesophyll cell (mc) and very intense reaction in the bundle sheath cell (bsc) chloroplast. cwcell wall. Poststained with PbCi. (18,900 X).

PAGE 99

®

PAGE 100

84 Discussion A wide variety of simple and complex compounds possessing phenolic hydroxyl groups occur in plant tissues and the importance of these compounds during the life cycle of the plant has become increasingly evident (143). Plant pathologists and physiologists have a keen interest in phenolics as the "antiseptics" of the Plant Kingdom (143) and many investigations have been made on disease resistance and interaction of microorganisms with phenols. As indicated previously, specialized cells containing phenolic compounds have been reported in tissues from many plant species. These cells are often called "tannin cells" when the nature of the phenolic substances is not known, or the substances have become decompartmented , oxidized, and polymerized to varying degrees (120). Common, nonspecific tests for tannins usually consist of treatment with ferric chloride solutions followed by treatment with dilute bases (148). A blue-green precipitate is usually formed but not all phenolics give such a reaction and the results may be influenced by other materials present. The Gibbs indophenol reaction (59) is a dependable test for the detection of phenols (51), but appears to be of little or no value in determining the number of hydroxyl groups on the benzene ring (51,100). On the other hand, the nitroso reaction (148) forms a cherry-red nitroso derivative

PAGE 101

85 of o-dihydric phenols and appears to be a generally reliable means of differentiating these compounds from other phenols. Work with a large number of phenols has shown that the color of the nitroso derivatives other than those of o-dihydric phenols are yellow, brown, red-brown, and green (100). Results with waterhyacinths show that phenolic compounds are localized in phenol-storing cells in both the palisade cell layer and in close association with the vascular bundles in leaf tissue. After staining by the nitroso reaction these cells appear bright red and, therefore, probably contain o-dihydric phenols and their derivatives. Using this histochemical method in conjunction with chromotographic and ultraviolet absorption data, Mace (100) was able to identify 3-hydroxytyramine (dopamine) as the major o-dihydric phenol in cells of banana roots. Results obtained by similar methods are presented in Chapter III of this dissertation and show the presence of several odihydric phenols in waterhyacinth phenol cells. Several articles have appeared recently on the ultrastructure of phenol cells in plants (119,120). In each case, the cells described were In root tissue. The data presented here are the first reported from an ultrastructural investigation of phenol cells as they occur in leaf tissue. Two morphologically distinct phenol cells occur in waterhyacinth leaves. Both contain nuclei, mitochondria,

PAGE 102

86 and plastids, and presumably are actively metabolizing cells. Phenol cells in the palisade cell layer of waterhyacinth leaves are elongated and vary greatly in size, often exceeding several hundred microns in length. In contrast, those in the vascular region of the leaf are much more isodiametric , vary much less in size, and the phenolic compounds appear in an amorphous mass. The vascular phenol cells closely resemble those in root tissue of cotton (120) and banana (119). Despite the widespread occurrence and the substantial evidence for the importance of phenolic substances in plant metabolism, the mechanics by which the phenolics are synthesized, stored, and released have not been established (120). Mueller and Beckman (120) concluded that the origin and formation of phenolic material could not be determined by microscopic examination alone, although they suggested a role for plastids In this process. Wardrop and Cronshaw (189) suggested that phenolic material is synthesized in modified amyloplasts. The plastids disintegrate shortly afterwards and the phenolic compounds aggregate in vacuoles. Studies with waterhyacinth support Wardrop £ Cronshaw' s hypothesis because (i) phenolic compounds are consistently observed in close association with plastids in the phenol cells; (ii) plastids in those cells tend to be smaller and more compact than those in adjacent cell types suggesting a

PAGE 103

87 possible degradation process; and (iii) the turn-on of PPO activity in plastids in additional cells after infection with A. zonatum indicate a likely role for phenol oxidation in response to pathogenic attack. Results of this study also indicate that the phenolic compounds are released from these cells after disruption of the membranes as a result of damage inflicted by the pathogen (see Chapter IV). Subsequent oxidation of these compounds by PPO to fungitoxic quinones would likely occur in chloroplasts of adjacent cells. Thus, the phenol cells serve as a mechanisms of defense by acting as miniature "time bombs" which are activated during pathogenesis. The spatial distribution of foliar phenol cells in waterhyacinths is random with the highest concentrations occurring in the leaves of larger plants. They occur in greater numbers on the top surface of small leaves but the opposite is true for medium and large leaves. The reason for this distribution is unknown and any explanation at this point would only be speculation. Since waterhyacinths have an almost equal distribution of stomata on both the adaxial and abaxial leaf surfaces (139), correlation of phenol cell occurence with the presence or absence of stomata can be ruled out . Previous investigations (36,37,72,73,74,135) on the cytochemical localization of foliar PPO have shown this

PAGE 104

enzyme to be present in the thylakoids of all chloroplasts examined. Henry (74) reiterated the possibility that PPO is ubiquitously associated with certain phenolic compounds which participate in electron transport. The present study, however, describes for the first time a restricted localization of this enzyme in healthy leaf tissue and a subsequent turn-on in other cells after infection. The presence of this enzyme in particular cell types suggests a more specific action than that implied by Henry. The PPO found in chloroplasts of phenol cells is possibly responsible for the oxidation of certain stored phenols. However, PPO observed in vascular bundle sheath chloroplasts may be related to electron transport, while that in vascular parenchyma cells is possibly involved with lignin synthesis during secondary wall thickening of the xylem elements. The turn-on in PPO activity after infection with A. zonatum in cells normally devoid of any such activity is highly suggestive of an active role for this enzyme in the disease reaction. Cook and Wilson (31) were the first to suggest that the f ungitoxicity of tannin was due to the action of an oxidase which formed a germicidal fluid. Since their work in 1915, numerous investigations have correlated increased phenol-oxidizing enzyme activities with disease resistance (9,68,101,102,121,144,146,149,181,182). Whether the turn-on in PPO activity in waterhyacinths is pathogeninduced or host-induced is not known at the present time

PAGE 105

89 but it is probably a host-mediated response to the pathogen' s attack .

PAGE 106

CHAPTER III A BIOCHEMICAL STUDY OF THE PHENOLIC ACIDS AND POLYPHENOLOXIDASE RATES IN HEALTHY AND DISEASED WATERHYACINTH LEAVES Introduction A potential biocontrol agent must meet several criteria if it is to be successful. Perhaps the most important of these is its specificity of the pathogen to a given host. One of the factors which determines the host-parasite specificity is the biochemical relationship between the two organisms. Lewis (94) attempted to explain this biochemical relationship on the basis of nutrition in his "Balanced Nutrition Hypothesis of Parasitism." His assumption was that in order for one organism to parasitize another, a correct balance of nutrients must be present. Snell (168) classified these "nutriolytes" on the basis of whether they were essential to growth of the parasite or stimulatory to its growth. In essence, both Lev/is and Snell postulated that the outcome of a host-parsite relation was determined by the interaction of water diffusable substances from both organisms . One of the major objections to this hypothesis is that it does not take into account the effect of any inhibitory substances which might be produced by the host either prior 90

PAGE 107

91 to or in response to infection. In 1954, Garber (58) expanded the idea to include inhibitory substances as another factor in pathogenesis. He states that there are two environments which affect the outcome of a disease relationship, the balanced nutrition environment and the inhibitory substance environment. According to Garber, there are four possible combinations of these two environments, only one of which results in disease, i.e. a relationship where the correct nutrients are supplied and no inhibitory compounds are present. In many instances, phenolic compounds are inhibitory to the growth of microorganisms and it has been suggested that they may be involved in disease resistance mechanisms. It was therefore, the intent of this study, to examine the phenol chemistry of waterhyacinths to determine what role, if any, host phenolic compounds play in disease resistance to Acremonium zonatum . Our knowledge of the phenolic compounds of plant origin had its beginnings in industry. The earliest recognized class of these compounds, the tannins, have been employed since ancient times in the tanning of skins, the manufacture of inks, and in the fining of wines (150). Fhenolics are a vast group of compounds, comprising the anthocyanins (red and blue pigments), the flavones (yellow pigments), the coumarins, tannins, lignins, and phenolic

PAGE 108

92 acids and their esters. The term "tannin" has generally been used to describe this wide array of organic compounds; however, it is now restricted to those compounds which have the specific property of tanning leathers. Recently, the term "polyphenol" which implies the presence of more than one hydroxyl group on a benzene ring has been used to include the plant phenolic compounds (61). The principal polyphenols are not present in a free state in nature, but exist primarily in the form of esters or glycosides (150). Hydrolysis of these compounds liberate the aglycones or phenolic moiety from their respective glycosidic or ester linkage. Extraction at low pH ' s reduces them to their acidic state. As a group, phenolic acids comprise the benzoic acids (C fi -C. ) and the cinnamic acids (Cg-Cg). The structures of the principal benzoic and cinnamic acids are given in Fig. III-l. The benzoic acids are widely distributed both in angiosperms and gymnosperms (69). For instance, Tomaszewski [in Ribereau-Gayon (150)] has identified two benzoic acids, p-hydroxybenzoic (p-HBA) and gentisic acids in the leaves of 97% of plants sampled from 86 families. Likewise, vanillic and syringic acids are widespread as they are constituents of lignin along with p-HBA. In general, plants which do not contain lignin do not contain these acids either (69). The phenylpropane skeleton (Cg-Cg) is unquestionably

PAGE 109

Figure III-l. Principal phenolic acids found in plants. [After Ribereau Gayon (150)].

PAGE 110

94 o Ld o o cr o -J X o Q_ Q O < o < z; o CO -g 'o < o "o e cr> o Cl Q_ CO Q O < o o N z: LjJ CD

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95 the commonest and most important of the polyphenols (150). This group includes the cinnamic acids and their derivatives; the aromatic amino acids, tyrosine, phenylalanine, and DOPA; the constituents of essential oils; and the lignins. Like the benzoic acids, the cinnamic acids are widely distributed in plants. p-Coumaric acid is the most common of all phenolic constitutents and is found in practically all higher plants (150). Because cinnamic acids possess a double bond, they are capable of existing in two isomeric forms, i.e. cis and trans -cinnamic acids. Different biological properties have been assigned to the cis and trans forms, however, the naturally occurring cinnamic acids are the trans isomers, which are more stable (150). The cinnamic acids, especially caffeic acid, have been known for a long time. One of their properties is their affinity to form esters with other phenolic acids. They were first studied in coffee, which is particularly rich in these compounds (140) and from which caffeic acid derives its name. Perhaps the most well known and studied of these esters is chlorogenic acid ( 3-caf f eoylquinic acid) which was discovered in coffee by Payen [in Ribereau-Gayon (150)] in 1846. It is an ester of caffeic acid with a cyclic acidalcohol, quinic acid. Many ester-derivatives of the cinnamic acids have been described (66,112,156,200). The major pathway for the biosynthesis of monocyclic

PAGE 112

96 phenols is the shikimic acid pathway (Fig. III-2) (126,127, 145,156). The elucidation of this pathway was was done largely with mutant strains of bacteria using radioactive tracers (145). The basic sequence of reactions is also believed to occur in higher plants. The initial step involves the condensation of the three-carbon compound, phosphoenolpyruvate, derived from glycolysis, with the four-carbon compound, erythrose-4-phosphate , derived via the pentosephosphate pathway. The initial branch in phenol synthesis yields the benzoic acids. The second branch provides for the synthesis of the amino acid tryptophan which can undergo conversion to indoleacetic acid ( IAA , auxin). The third branch provides for synthesis of the aromatic amino acids phenylalanine, tyrosine, and DOPA, the later two involving PPO. Deamination of phenylalanine by phenylalanineammonialyase (PAL) yields the C R -C_ skeleton, trans -cinnamic acid (32). The action of several enzymes on trans -cinnamic acids yields a variety of hydroxylated and methoxylated cinnamic acids (29) i.e. p-coumaric , caffeic, and ferulic. All of these acids can undergo ^-oxidation to yield their corresponding benzoic acids (66), be incorporated into the B-ring of flavonoids (61,66,172), undergo isomerization and intramolecular condensation to form coumarins and phytoalexins (45,172), or undergo reduction to cinnamyl alcohols and subsequent oxidative polymerization to lignin (61,126,200).

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Figure III-2 . Shikimic acid pathway for the biosynthesis of monocyclic phenols and major derivatives. Adapted from Neish (126,127) and Robinson (156).

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Carbohydrate Metabolism phosphoenolerythrose-4pyruvate (pep) phosphate 2-keto-3-deoxy-7-phosphoglucoheptonic acid 5-dehydroquinic acid © 5-dehydroshikimic acid — protocatechuic acid shikimic acid pep -J ) chorismic acid — -anthranilic — -tryptophanacid — vanillic acid IAA prephenic acid •> tyrosine DOPA phenyla anine (a) cinnamic acid — » — «• — ~ coumarins a phytoalexins p-coumaric acid quinic acid caffeic acid " — chlorogenic acid (6) ferulic acid — isocoumanns lignin Shikimic Acid Pathway and Major Derivatives

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99 The biosynthesis of polyphenols may also be accomplished by the polyketide (150) or acetate pathway (61). Compounds derived via this pathway are formed from the headto-tail condensation of acetate units. There is good reason to believe that this pathway also operates in higher plants, although to a lesser extent than the shikimic acid pathway, and has a role in the biosynthesis of the A-ring of flavonoids ( 61 ) . The importance of PPO to phenol synthesis lies in its ability to oxidize monophenols to polyphenols (108). Through these reactions, the flavonoids, tannins, lignins , and melanins are formed, all of which have been implicated in disease resistance (76,131,151). Numerous studies have shown that host polyphenols and their oxidizing enzymes typically increase in diseased plants (35,50,110). Their role in pathogenesis has been attributed to several mechanisms including inhibition of spore germination (91), antibiosis (96,104,161), initiation of the hypersensitive response (98, 179), inhibition of pectolytic enzymes (14,28,42, 79), and the inhibition of indoleacetic acid oxidase (50,89). This chapter presents the results of a study of the benzoic and cinnamic acids present in healthy and diseased waterhyacinths and the subsequent changes in PPO activities. The effect of these compounds on the growth of A. zonatum , as well as the fungus' ability to synthesize auxin in_ vitro is also presented.

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100 Materials and Methods Assay for "free" phenolic acids Extraction . Fifty g (fresh weight) of leaves from each of small, medium, and large healthy, greenhouse-maintained waterhyacinths were excised, washed in distilled water, and macerated in 500 ml 95% ethanol in a Waring blender for five minutes. The slurry was filtered through cheese cloth and then through Whatman no. 1 filter paper in a Blichner funnel. Total volume after filtration was 465 ml. Two hundred ml of the filtrate were concentrated to approximately 5 ml under vacuum at 40 C in a rotary evaporator. The concentrate was made to 10 ml with 95% ethanol and centrifuged for three minutes at 2500 rpm . The clear supernatnant was stored at -20 C unless used immediately . Identical extraction procedures were used with leaves from small, medium, and large diseased plants. Thin layer chromatography . Twenty yl (5 yl at a time) of each supernatnant were spotted onto silica-gel, thinlayer, chromatography sheets (TLC) with fluorescent indicator added (20 x 20 cm, Kodak) and developed ascending in nbutanol: acetic acid: water (EuAW) (40:10:20 v/v, organic phase). TLC sheets spotted with 10 yl of nine different phenolic acid standards (5 mg/ml in 95% ethanol) were developed simutaneously and used as markers for identification of unknowns. The chromatograms were dried at 100 C

PAGE 117

101 and phenolic acids located under ultraviolet light (253 nm) . Fluorescent spots were noted as to color and marked for later identification. Phenolic acids on replica chromatograms were located by uniformly spraying with one of the following reagents (167): (i) sulphanilic acid, (ii) pnitroaniline , and (iii) p-nitroaniline oversprayed with 2N NaOH. The R f values and colors obtained with the different locating reagents were compared to standards. Assay for "ester-linked" phenolic acids Extraction . The extraction procedure used for waterhyacinth is an adaptation of the methods described by Isamil and Brown (80) and Woodward (195). A flow diagram of the basic procedure is shown in Fig. III-3. Fifty g (fresh weight) of leaves from each plant morphotype were washed and macerated as before. The ethanolic extract was then boiled for 30 minutes and filtered. The emerald-green filtrate was reduced to dryness in a rotary evaporator under vacuum at 4 C, and then was redissolved in 50 ml hot, distilled water. After cooling to room temperature, the extract was extracted 3 times in 50 ml petroleum ether to remove chlorophyll. The golden-yellow aqueous phase was hydrolyzed by adjusting to 2N with NaOH pellets (approximately 4 g) and boiling for three minutes. The resulting dark-brown hydrolyzate was placed immediately into an ice-bath and adjusted to pH 1.0 with concentrated HC1. The acidified extracts were then

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Figure III-3. Flow diagram of procedure for extraction of°ester-linked phenols in plants. Adapted from Isamil and Brown (81) and Woodward (199). Step 1. Disruption from entact tissue Step 2. Extraction of chlorophyll Step 3. Hydrolysis of ester linkage Step H . Purification and concentration of phenolic acids .

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© 50 q. fresh tissue I macerate in hot EtOH i . boil for 30mins. filter 103 residue discard filtrate reduce to dryness resuspend in hot d. w. wash w/ pet. ether aqueous phase I adjust to 2N w/ NaOH I boil for 3mins acidify to pH 1.0 w/ HC extract w/ diethyl ether (3x) 'organic phase 1 discard aqueous phase discard "organic phase I reduce to dryness resuspend in 5ml. EtOH use for TLC © Flow Diagram for Extraction of Phenols

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104 extracted 3 times with 30 ml diethyl ether (v;ashings combined) and 0.5 ml distilled water added. They were reduced to dryness as before, resuspended in 5.0 ml 95% ethanol and stored at -20 C unless used immediately. Identical procedures were used for the extraction of phenolic acids from diseased plants. Thin layer chromatography . Twenty yl (5 yl at a time) of each extract were spotted onto TLC sheets and developed two-dimensionally ; ascending in benzene : acetic acid:water (BzAW) (10:7:3 v/v, organic phase) and after drying and rotating the sheets 90°, ascending in 2% formic acid (FA). Chromatograms of the nine phenolic acid standards were run simultaneously. Phenolic acids were located with ultraviolet light and by spraying as previously described. R f values were calculated and compared to standards. Assay for total phenols The concentration of total phenols in healthy and diseased waterhyacinth morphotypes was determined by the Folin-Denis colorimetric method (150). A portion (0.1 ml) of the hydrolyzed extract was diluted to 5.0 ml in photometrically-matched cuvettes by the addition of 4.0 ml disi lied water, 0.1 ml Folin-Denis reagent, and 0.8 ml satu•ited Na„C0_. The contents of the cuvettes were mixed for five seconds on a Vortex mixer and the optical density of each was recored at 74 nm on a Bausch and Lomb Spectronic

PAGE 121

105 20. The concentration of each was determined as phenol equivalents (phe) by extrapolation from a standard curve prepared from nine different phenolic acids ( trans -cinnamic , o-coumaric, p-coumaric , caffeic, f erulic , vanillic, protocatechuic, p-hydroxybenzoic , and chlorogenic ) . Polyphenoloxidase assay Fifteen g (fresh weight) of healthy or diseased waterhyacinth leaves from each size category were excised and placed immediately in the cold (4 C). Each group was macerated in a previously chilled Waring blender with 100 ml of cold 0.01 M phosphate buffer, pH 7.0 for five minutes. The leaf slurry was centrifuged in a precooled enclosed Sorvall superspeed centrifuge at 10,000 rpm for five minutes to remove excess leaf debris. The supernatant fraction was decanted and separated into two equal portions (approximately 30 ml each). One portion was kept in an ice bath while the other was boiled for one minute to serve as the inactivated enzyme control. Two drops of each portion was added to separate micro-cuvettes containing 1.0 ml of 0.001 M L-dihydroxyphenylalanine (L-DOPA) made up fresh in 0.01 M phosphate buffer, pH 7.0. After diluting 1:1 and mixing, the cuvettes were immediately placed into a Beckman Model 25 recording spectrophotometer. The optical densities at 246 nm were recored at 15 second intervals for the first minute, 30 second intervals for the next four minutes, and one

PAGE 122

106 minute intervals thereafter for ten minutes. The boiled fraction containing DOPA was used to zero the instrument and for the reference sample. Polyphenoloxidase activity rates of each sample were calculated as the change in O.D./time. Growth of A. zonatum on waterhyacinth-extract media Waterhyacinth-extract media were prepared from healthy and diseased plants from each size category in the following manner: Ten g (fresh weight) of leaves were macerated in 100 ml hot, 95% ethanol and filtered. The filtrate was reduced to dryness under vacuum and the residue redissolved in 15 ml boiling distilled water. The redissolved residue was extracted three times with petroleum ether and the aqueous phase was filter-sterilized through 0.45 y Millipore filters. One ml of each filtrate was added to separate tubes containing 9.0 ml Czapek-dox agar (45 C) (Difco) with and without yeast-extract added (0.5%). The media were poured into sterile petri dishes, allowed to solidify, and seeded with a 7 mm plug of A. zonatum . Control plates contained 1.0 ml distilled water in place of the plant extract. Five replicas of each were prepared and incubated at 22 C for two weeks. After the incubation period, the mean colony diameter of each was determined. Growth of A. zonatum on phenolic acid media Czapek-dox agar, with and without yeast extract, was made to final concentrations of 10, 100, and 1000 ppm of

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107 seven different phenolic acids ( chlorogenic , p-coumaric , vanillic, ferulic, caffeic, protocatechuic , and p-hydroxybenzoic). Ten ml of each medium was poured into sterile petri dishes, allowed to solidify, and seeded with a 7 mm plug of A. zonatum . Five replicas of each were prepared and incubated at 22 C. The mean colony diameter was determined after two weeks. Indoleacetic acid assay Colorimetric detection. The ability of A. zonatum to synthesize indole-3-acetic acid ( IAA , auxin) in_ vitro was determined using the colorimetric technique described by Gordon and Weber (62). Cultures were grown in 30 ml of modified Czapek's liquid medium at 22 C (193) with either NaNCor tryptophan as the sole nitrogen source. At two-day intervals, two flasks of each culture medium were removed, filter-sterilized through 0.45 y Millipore filters and assayed for auxin content in a Bausch and Lomb Spectronic 20 at 530 nm. Auxin concentration (ppm) was determined by extrapolation from a standard curve using purified IAA. Chromatographic detection . Synthesis of IAA from tryptophan medium was confirmed by thin layer chromatography. Cellulose TLC sheets with fluorescent indicator (20 x 20 cm, Kodak) were spotted with 10 yl of filtrate from ten-day-old cultures grown with and without tryptophan. Ten yl of a 100 ppm IAA standard was also spotted onto the TLC

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108 sheets as a comparison. Ascending chromato grams were developed in 70% ethanol for four hours, air-dried, and examined under a Wood's mineral light (253 nm) . Fluorescent spots were noted as to color, marked for identification, and the chromatograms then uniformly sprayed with ferric chloride-perchloric acid reagent (163). Spots which were ash colored under UV light and which subsequently yielded a pink color after spraying were assumed to be IAA. R^ values were calculated and compared to the standards. Results "Free" phenolic acids There were no differences in the "free" phenolic acids found among any of the plant sizes, either healthy or diseased (Table III-l). In each case, a spot which was indistinguishable from chlorogenic acid, was the only phenolic acid detected on the chromatograms. Mean R f values, fluorescence under UV light, and color after spraying were all similar to authentic samples of chlorogenic acid. Ester-linked phenolic acids Healthy waterhyacinth leaves . Three benzoic acids, (protocatechuic , p-hydroxybenzoic , and vanillic), and three cinnamic acids (ferulic, caffeic, and p-coumaric) were detected in all three sizes of healthy waterhyacinth leaves (Table III-2 ) . Three additional acids (spots #7,8, S 9)

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109 were observed on chromatograms from large healthy leaves which were not seen in small or medium plants. The identity of these additional phenolic acids was not determined. Mean R f values and color characteristics for each of these are listed in Table III-4. p-Coumaric acid (spot #6) was present in very small amounts in all plant sizes tested. It could not be detected under UV light in any of the chromatograms but was detected using either p-nitroaniline or sulphanilic acid reagent. Spot size and color intensity suggested that caffeic acid (spot #1) was in the greatest concentration in all samples. Infected waterhyacinth leaves . The same six phenolic acids that occurred in healthy leaves were also detected in leaves infected with A. zonatum (Table III-3). p-Coumaric acid was again in the smallest amounts while caffeic acid was the most concentrated. The only qualitative change in phenolic acids noted was the appearance of an unknown (spot #7) in smalland medium-sized leaves after infection which was not observed in healthy plants of the same size. This unknown phenolic acid, along with the other two (spots #8 £ 9), were also present in infected large leaves. Mean R^ values and color characteristics suggest that these unknown acids are the same ones which are present in large healthy leaves. The mean R f values in each solvent and the color characteristics of each of the phenolic acids detected in healthy

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110 and diseased waterhyacinth leaves are listed in Table III-4 . In each case, spots #1-6 were indistinguishable from the authentic reference samples and consequently considered to be the same. Total Phenols In healthy waterhyacinths , medium and large plants had a significantly greater concentration of total phenols in their leaves than did small plants (Fig. III-4). Data show that leaves from small plants contained 92 ug phe/g fresh weight while medium and large leaves contained 106 ug and 104 ug respectively. There was no significant difference between the total phenols in medium and large healthy leaves . After infection with A. zonatum , the total phenol concentration dropped significantly in both smalland medium-sized plants, but remained constant in large plants. Medium-sized diseased plants retained a significantly greater concentration over small diseased plants (96 ug/g vs 80 Ug/g) but was not significantly different from the concentration in large diseased plants (96 ug/g vs 105 ug/g). Pholyphenoloxidase assay In healthy plants, small leaves had a much greater PPO activity than large leaves. Data show that the activity of the enzyme in large leaves was 325% greater in small leaves than in large leaves (Fig. III-5). Medium-sized leaves

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Ill exhibited a PPO activity between that of small and large leaves. After infection with A. zonatum , the PPO rate in small leaves decreased almost 40% (Fig. III-6). In large leaves, the rate increased 300% to near the level found in healthy small plants. Medium-sized infected leaves showed little change in PPO activity. Growth of A. zonatum on waterhyacinth-extract media When A. zonatum was cultured on minimal media containing extracts from either small, medium, or large, healthy waterhyacinth leaves it grew significantly better than it did on minimal media without plant extracts (Table III-5). When yeast extract was added to the medium as a growth supplement, the fungus again grew significantly better than it did on medium without healthy plant extracts. Fungal growth on media containing extracts from diseased leaves were similar (Table III-5). Growth was increased on all media containing yeast and diseased-leaf extracts, whether from small, medium, or large plants. However, growth on the media which did not contain the yeast supplement increased significantly only on plates containing extracts from large diseased plants. Comparisons of fungal growth rates between extracts from healthy and diseased plants of each morphotype revealed no significant differences among any of the media which did not contain yeast extract (Table III-5). However, each of

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112 the cultures on diseased leaf -extract s grew significantly better on media with yeast than their respective healthy plant-extract counterparts. Growth of A. zonatum on phenolic acid media When A. zonatum was tested for its ability to grow on media containing phenolic acids, it grew either as well or better on each phenolic acid and concentration level tested than it did on the controls (Table III-6). p-Coumaric acid was the only phenolic compound tested which did not significantly increase growth of the fungus. In all other instances, the higher concentrations of phenolic acids (100 or 1000 ppm) significantly stimulated the growth of A. zonatum. In only one case did the lower concentrations increase growth and that was with p-hydroxybenzoic acid at 10 ppm. When yeast extract was added to the phenolic acid media fungal growth was similar to that on phenolic acid media without yeast extract with only a few noteable exceptions (Table III-7). Most cultures grew better on the phenolic acid media than they did on control plates, however, neither ferulic or chlorogenic acids stimulated the growth of A. zonatum . In addition, p-coumaric acid at 1000 ppm significantly decreased its growth compared to the controls. This was the only acid tested which was inhibitory to fungal growth in vitro .

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113 Indoleacetic acid assay Colorimetric detection . Cultures of A. zonatum grown in Czapek broth with NaNO„ as the sole nitrogen source did not synthesize auxin. When NaNO„ was replaced with tryptophan as the sole nitrogen source, auxin was detected in the filtrate after two days of growth (Fig. III-7). The auxin concentration increased in the filtrates reaching a maximum of 25 ppm after ten days and then decreased slightly after 14 days. Chromatographic detection . Specificity of the GordonWeber test for auxin, was verified by TLC , using filtrate samples from ten-day-old cultures. Results showed two distinct spots from the filtrates containing tryptophan. Spot #1, which was not identified, had an R f value of 0.71 and turned brown after spraying with ferric chloride-perchloric acid. A similar spot was observed from the filtrates containing NaN0„ (R f = 0.73). Spot #2 was ash colored under UV light and turned pink when sprayed with the locating reagent. It had an R f value of 0.87. The IAA standard displayed identical colors under UV and after spraying as spot #2, had a similar R^ value (0.91), and was otherwise indistinguishable from the authentic sample. It was concluded from this that spot #2 was most probably IAA. A corresponding spot was not observed from filtrates containing NaN0~.

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114 x < O cc 2: cd < o >< x s: H O _i or < x LU CD X CC 2: LU -" > < o —> LU X IhUJ a >CQ < 2: u u — < _l >O X 2: cr UJ LU X H 0< 2 LU LU Q CC LU Ll_ H O LU PQ <
PAGE 131

115 X CO

PAGE 132

116 o

PAGE 133

o < cc lu u h—i u_ _i < o 2: in LU LU X > Q< LU LU _J X IX hU_ 2T O -" O 00 < CJ >— X I— CC 00 LU — Icc < cj

PAGE 134

Figure III-4. Total phenol concentrations in healthy and A. zonatum-inf ected waterhyacinth morphotypes .

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119 en 120 110 100 I90 80 70 -J 60 O z LiJ X 50 a. _j < 40 \— O J30 20 10 i V -.'.: E' > • Hd 'V g^ 'V i; H I Ha •-;: 1 p 1 SH SD MH MD LH LD PLANT SIZE

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Figure III-5 . Polyphenoloxidase activities in small, medium, and large healthy waterhyacinth leaves. PPO rate calculated as the change in optical density at 246 nm/time.

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-i — i — i — r 121 o "t H "-' d E • * (ujugt-2) A1ISN3Q IVOIldO V

PAGE 138

Figure III-6. Polyphenoloxidase activities in small, medium, and large diseased waterhyacinth leaves. PPO rate calculated as the change in optical density at 246 nm/time.

PAGE 139

"i — r 123 — o 01 s & en o k> b"" — — -— o LU (wugt^) A1ISN3Q IVOIldO V

PAGE 140

124 Q Z. < z <

PAGE 141

< O N LL. O X CD Q cm lu C!D Q oo CD — s: n^ CD LU I Q_ CD
PAGE 142

Q X o

PAGE 143

Figure III-7. In vitro synthesis of indoleacetic acid from tryptophan by Acremonium zonatum .

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128 E Q. Q. ^ 10 4 6 8 10 Incubation time (days)

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129 Discussion Varietal resistance of plants to pathogens has been rs i i.ted to polyphenol content on numerous occasions. The olct^sic works of Walker (18 3,184) and Walker and Link (185) were perhaps the first positive evidence for a role of phenols in disease resistance. Working with the onion pathogen, Colletotrichum circinans , they demonstrated that the presence of certain phenolic compounds, mainly protocatechuic acid and catechol, in the scales of red onions imparted resistance to the smudge pathogen. Since then numerous investigators have correlated host phenolic compounds to disease resistance (12,35,50,145,159,179,186). Condensed and hydrolyzable tannins have been implicated in resistance of cotton, strawberry, and apricot to Verticil lium wilt and the resistance of chestnut to Endothia parasitica (12). Other compounds such as 3 , 4-dihydroxyphenylalanine (DOPA) and benzoic acid have been implicated in the resistance of banana to Fusarium wilt (102) and apples to Nectria rot (17) respectively. Chlorogenic acid content has been correlated with resistance to such disease as coffee canker (44), potato scab (82), and Verticillium wilt of potatoes (93 ) . The idea that phenolic compounds may be involved in disease resistance stems from the many articles which appear in the literature demonstrating a biostatic or biocidal

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130 effect on microorganisms. Microorganisms, however, very considerably in their sensitivity to different phenolic acids. Bell (12) states that the order of activity of fungal pathogens to polyphenols is obligate parasites > facultative saprophytes > facultative parasites > saprophytes. Thus certain pathogens such as the mildew and rust fungi are frequently inhibited by polyphenols while many saprophytes are stimulated by moderate concentrations of polyphenols as i carbon source. The toxicity of phenols is generally believed to be due to their oxidation products, the quinones, brought about either by enzymatic action or autooxidation (12). Thus, Schaal and Johnson (161) demonstrated that chlorogenic acid, caffeic acid, catechol, and tetrahydroxybenzoin were toxic to Streptomyces scabies in vitro. p-Hydroxybenzoic acid was not as an effective inhibitor as were the other four. They concluded that the inhibition effect of these phenolic compounds was due to autooxidation to their respective quinones, and although p-hydroxybenzoic acid also autooxidizes, it does so much more slowly since it is a monophenol whereas the others are polyphenols. The effectiveness of polyphenols in disease resistance depends on many factors (12): (i) the quantity and type of polyphenols present in healthy tissue, (ii) the speed and quantity of polyphenol synthesis induced by infection, (iii)

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131 the quantity and type of oxidases present in healthy tissue, (iv) the speed and quantity of oxidase synthesis induced by infection, (v) location of the polyphenols and oxidases in the host, (vi) the sensitivity of the pathogen to these compounds, and (vii) the cellular environment in which these occur. The studies presented in this chapter were designed to evaluate most of these criteria as they relate to waterhyacinth . The only phenolic acid detected in the free state was chlorogenic, which is an ester of caffeic and quinic acids. Alkaline hydrolysis liberates the phenols which are bound in plants as esters while acid hydrolysis, on the other hand, liberates those phenols which are bound as glycosides (152). The six phenolic acids identified in waterhyacinth after alkaline hydrolysis and the three unidentified ones, are therefore most probably present in the plant as esters, either with quinic acid (152) or with sugars (70,71). Liberation of the aglycones in vivo from their respective linkages, either by host-mediated or pathogen-mediated reactions could then make them available to the host during pathogenic attack and possibly serve in defensive reactions. The mechanism by which this is accomplished, has not been fully investigated. It has been demonstrated that polyphenols increase, either qualitatively or quantitatively, or both, in many plants following infection (159). The decrease in

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132 concentration of total phenols in smalland medium-sized plants after infection was therefore, unexpected. However, this decrease in total phenols may be indicative of a decrease in resistance of small plants. Results similar to those obtained with waterhyacinths were obtained by McLean et al . (115) working with potatoes and Verticillium wilt. They observed that wilt developed more rapidly and more severly in susceptible varieties, coincident with or following the decrease in phenol compounds in the vascular system. Patil et al . (136,137) also showed that young potato roots which are partically resistant to infection by Verticillium have a relatively high level of phenols until five weeks after sprouting. From the time of sprouting, the chlorogenic acid content decreased continuously and was correlated with an increase in susceptibility to infection. The decrease in total phenols in small diseased waterhyacinths also correlates with the decrease in PPO activity after infection. It would be logical to assume that reduced levels of substrate would lead to reduced levels of enzyme activity. In large waterhyacinths, however, the total phenol content did not decrease after infection, but remained at the level found in healthy plants. In this case, increased resistance can best be correlated with the greatly accelerated PPO rate. The concentrations of polyphenols in

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133 large healthy plants is significantly higher than that in small healthy plants, but the PPO rate is much lower. This would account for a reduced rate of enzymatic oxidation. After infection, however, PPO rates increase, thereby increasing the oxidation of phenols to the quinones. In contrast to small plants, large plants are initally more susceptible to fungal attack by virtue of their lower PPO rate but gradually build up resistance as the PPO activity increases over 300%. Thus, disease severity in each plant morphotype balances at about 40% diseased leaf area, small plants by an increasing susceptibility and large plants by an increasing resistance. On first inspection, the qualitative data on the phenolic acids present in small waterhyacinth might not appear to coincide with the total phenol content. An additional phenolic acid was detected on chromatograms from diseased small plants which was not detected on chromatograms from the same morphotype s of healthy plants. This could result from two possibilities. First, the compound is in healthy tissues but in concentrations too low to detect and after infection, synthesis is increased, or second, it is not in healthy tissues, but is synthesized de_ novo during disease. In either case, its concentration should increase, resulting in an increase in total phenols. It is believed, however, that even though this compound is increasing, the remaining

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134 phenolic acids are decreasing, thereby reducing the total phenolic content. Preliminary results with caffeic acid support this hypothesis (R. Martyn, unpublished). Caffeic acid content, which is in the highest concentrations of all the phenolic acids in waterhyacinth , decreased greatly in small diseased plants from that present in healthy plants. The higher content of total phenols in large healthy plants, as opposed to small healthy plants, is most probably due to increased concentrations of each acid plus the presence of three additional ones not found in smaller plants. One of the most striking results of this study was the high tolerance of A. zonatum to phenolic acids, in_ vitro . Acremonium zonatum is a facultative parasite, and according to Bell's sensitivity ranking of pathogens, it should be relatively tolerant of polyphenols. Results presented in this chapter support this concept. Its tolerance was evidenced by its ability to grow significantly better on minimal media with various concentrations of phenolic acids incorporated into them than it did on minimal media alone. This fact could explain why A. zonatum is able to cause relatively severe damage on waterhyacinth. The above finding is in agreement with the results of fungal growth on plant-extract media. Healthy waterhyacinths of each morphotype have a relatively high phenol content in their leaves and fungal growth was stimulated on media

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135 containing such plant extracts. The significance of this can be noted in the growth of A. zonatum on media containing extracts from diseased plant morphotypes. An increase in growth was only observed on media containing extracts from large diseased leaves. Extracts from both small and medium diseased leaves failed to stimulate fungal growth which may be indicative of their reduced phenol content after infection. Oddly, p-coumaric was the only phenolic acid which did not stimulate fungal growth in vitro on minimal media. When additional growth supplements were added to the media, pcoumaric was the only phenol found to be inhibitory. This suggests that in the absence of additional nutrients, A. zonatum is capable of metabolizing several different phenolic acids as a carbon source. In the presence of an enriched medium, however, these same phenolic acids lose some of their simulatory effect, noticeably ferulic and chlorogenic acids, while one (p-coumaric) becomes inhibitory. The reason for the inhibitory effects of p-coumaric acid is not known but a recent study by Elstner and Heupel (46) may add some information as to its mode of action. Working with isolated cell walls from horseradish, they demonstrated that hydrogen peroxide production was inhibited by dihycroxyphenols but stimulated by monohydroxyphenols such as p-coumaric acid. If p-coumaric acid stimulated hydrogen peroxide production in A. zonatum as it does in horseradish,

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136 it may well account for the inhibitory effect. Studies on the peroxidase system of this fungus should be done to test this hypothesis. Data from electron microscopy studies during pathogensis (see Chapter IV) revealed that the penetration of a phenol cell by A. zonatum resulted in death of the invading hyphae . It is presumed that either the concentrations of phenols in those cells are such that they are no longer stimulatory but toxic to the fungus or that some other factor of metabolism, such as PPO activity, is increased to the point where the fungus can no longer tolerate it. Unfortunately, data from this study do not permit a conclusion on either possibility . In the present discussion, it has been suggested that host phenols play a major role in the defense against potential pathogens of waterhyacinths by being biocidal or biostatic. This concept seems plausible when dealing with an organism that is susceptible to the toxic properties of phenols. However, this is not the case with A. zonatum , and it appears that some waterhyacinth plants have an additional defensive mechanism which Is indirectly linked to phenol metabolism. It was observed that after infection, large plants displayed almost a three-fold increase in new leaf production over that of either small or medium plants. Increased meristematic activity in plants is not uncommon

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137 during disease (61). It has been demonstrated many times that auxin content markedly increases in diseased tissue (16,173). The increased levels of auxin in diseased tissue may result from either (i) increased synthesis of IAA by either host or pathogen or (ii) a decreased rate of auxin degradation. There is no direct evidence that reveals whether the host plant or the pathogen is the actual source of increased IAA levels in infected tissues (61), but a number of studies indicate that pathogens are capable of synthesizing auxin in vitro (103,106,171,193, 194). Acremonium zonatum was also able to synthesize high concentrations of IAA in vitro which is a possible source of extra growth hormone necessary to promote accelerated rates of leaf regeneration. If the fungal-IAA was responsible for the accelerated leaf production, then higher growth rates would be expected to occur in all infected plant morphotypes but it did not; only large plants increased leaf production . A second possible means of increased levels of auxin in the hosi suggested was a decrease in the rate of auxin degradation. This may be correlated with an inhibition of IAA oxidase activity in the plant. IAA oxidase is a peroxidase mediated system responsible for keeping a balanced level of auxin in the plant and phenolic compounds are known inhibitors of this system (61). In large diseased

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138 waterhyacinths , phenolic compounds do not decrease as they do in small and medium, but remain at their preinfection level. It is highly possible that these phenols are inhibiting the normal "checks and balances" system of auxin regulation in large plants consequently allowing it to build up to abnormal levels resulting in a faster growth. The waterhyacinth ' s primary disease defense system appears to lie in its unique phenol cell-PPO complex and operates effectively against numerous potential pathogens. However, this system breaks down somewhat when the plant is attacked by an organism such as A. zonatum which can utilize the plant's phenolics for its own growth. In such event, the role of the phenols is to block auxin degradation allowing the plant to "outgrow" the infection.

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CHAPTER IV AN ULTRASTRUCTURAL STUDY OF PENETRATION AND COLONIZATION OF WATERHYACINTH BY ACREMONIUM ZONATUM AND CYTOLOGICAL CHANGES ASSOCIATED WITH INFECTION Introduction The process of pathogenesis can be viewed as a battle between a plant and a pathogen refereed by the environment (191). A small change in a single environmental factor, such as temperature or moisture, often can mean the difference between crop success and crop failure. When studying plant diseases, one should always be conscious of the climatic conditions under which the disease is evident. In nature, the environmental variables are numerous, and as Matta (109) has suggested, plant-pathogen interactions can sometimes be best studied in the laboratory where some of the variables can be controlled. It is axiomatic that before disease can ensue, a virulent pathogen must come into contact with a susceptible host. Although some pathogens are brought to their hosts through a vector relationship, most fungal pathogens make contact with their hosts fortuitously in the form of windblown or water-borne spores. Merely establishing contact is not enough in most cases to ensure a parasitic relationship. 139

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140 Entry into the host by the parasite must usually be accomplished. The manner in which this is achieved has been debated for many years. In general, there are two mechanisms through which a pathogen can enter its host: (i) direct penetration through natural openings, wounds, or through unbroken surfaces by mechanical pressure and (ii) penetration facilitated by enzymatic degradation. Although there is little argument against direct penetration through openings in the plant surfaces, it is generally believed that penetration through unbroken surfaces involve a combination of both mechanisms (61). Pressure for direct penetration is presumably supplied by the appresorium of the fungus which serves as an anchoring device and from which the infection peg emerges. Cuticle and cell wall degrading enzymes are secreted from the hyphal tip, facilitating entrance of the fungus into the host. After penetration fungal pathogens may spread from the site of infection throughout the host. Only if the fungus enters into a parasitic relationship with its host is colonization successful, and hence pathogenesis is initiated. Unlike bacteria, most fungi invade their hosts intracellular and obtain nutrients from those cells. This may be accomplished by distinctive fungal structures termed haustoria which penetrate the cell wall and absorb nutrients through the host cells' plasmalemma, or by secretion of

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141 toxins which may act on the cell membrane causing its disruption and electrolyte leakage. Those fungi that do not form haustoria or produce toxins may penetrate within the host cell and gain nutrients directly from the hosts' cytoplasm. This appears to be the case with A. zonatum . In general, after penetration has occurred, the protoplasm of the host cell becomes granular and the nucleus migrates towards the infecting hyphae . Later the nucleus increases in size considerably (61). In advanced stages of disease the nucleus, as well as other cell organelles, begins to collapse and degenerate. This chapter presents the results of an ultrastructural study of the penetration and colonization of waterhyacinth leaves by A. zonatum . Special attention was given to the possible method(s) by which the fungus was able to gain entrance into its host cells and the cytological features which changed as a result of the infection. Materials and Methods Electron microscopy Leaves, displaying characteristic symptoms of disease, from both small and large waterhyacinth plants were excised and used for the ultrastructural study. The leaves were placed in petri dishes fitted with filter paper and moisten with approximarely 5 ml of Karnovsky's fixative.

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142 Sections (2-4 mm) were cut from three areas on each leaf: (i) center of the lesion, (ii) periphary of the lesion and, (iii) asymptomatic tissue several centimeters (2-5) from the lesion. Each section was then fixed, dehydrated, and embedded as previously described (see Chapter II). Thin sections were cut using a diamond knife, placed on single hole grids, poststained in uranyl acetate and lead citrate, and observed on an Hitachi HU-11E electron microscope. Production of pectinases . The procedure used for the detection of extracellular pectinases produced by A. zonatum follows closely that described by English (47). Five hundred grams of peeled sweet potatoes were boiled in one liter of distilled water until soft (approximately 20 minutes). The solution was filtered through successive layers of cheese cloth and 100 ml of the filtrate placed into each of ten 250 ml sterile Erlenmeyer flasks. Each flask was seeded with a 7 mm plug of A. zonatum and incubated for ten days at room temperature. After incubation the media were filter-sterilized through 0.45 u Millipore filters and each filtrate divided into two fractions. One fraction was then boiled for ten minutes while the other fraction was left unboiled. Fresh sweet potato discs (0.5 x 7 mm) were placed into sterile petri dishes fitted with moisten filter paper and 1.0 ml of either the boiled or unboiled filtrate pipetted onto each disc.

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143 Five replicates of each plate were made. In addition, small pieces of the mycelial growth were also placed onto additional sweet potato discs. All plates were incubated at room temperature and examined after 4, 8, 12, 16, and 24 hours for signs of tissue maceration. Cultures of Rhizopus stolonif er , supplied by J.W. Kimbrough , were grown and treated in an identical manner and were used as positive controls . Production of cellulase Carboxymethylcellulose (CMC) medium (63, and R.E. Stall, personal communication) was prepared in the following manner. The CMC gum (Hercules Chemical Co.) was autoclaved for ten minutes at 15 psi in 80% ethanol and washed twice in fresh 80% ethanol for 15 minutes each. The resin was dried at room temperature overnight and the dried powder added to hot distilled water to give a final concentration of 2.0%. Bacto-agar (Difco) was added to a concentration of 0.5% and yeast extract added to 0.3%. The entire mixture was then autoclaved and poured into standard size (100 x 14 mm) sterile petri dishes. The CMC agar was allowed to solidify and seeded with a 7 mm plug of A. zonatum . The plates were incubated at room temperature and monitored daily for fungal growth and liquefaction of the gel around the colonies (pit formation ) .

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144 Results Production of pectinases and cellulases Acremonium zonatum did not produce detectable amounts of extracellular pectinases when tested with the potato assay method (Fig. IV-1). The fungus grew well in the sweet potato broth and on the sweet potato discs, but it failed to cause any noticeable signs of tissue maceration. Similarly, the unboiled filtrates failed to cause any detectable breakdown of the discs. After 24 hours the potato discs became dehydrated and were discarded. Potato pieces treated with R. stolonif er filtrate showed tissue breakdown after eight hours which increased over the next 16 hours. Electron micrographs of infected waterhyacinth leaves revealed that A. zonatum penetrated the middle lamellae of cells (Fig. IV-2c £ d) but did so without destroying the integrity of the surrounding portions. If extracellular pectinases were produced by A. zonatum as an aid in hostcell penetration, they were localized at the tip of the invading hypha and hence were produced in concentrations too low to be detected in_ vitro by the sweet potato disc maceration test . On the contrary, A. zonatum produced detectable amounts of an extracellular cellulase in_ vitro as evidenced by the pit formation or liquefaction when grown on CMC medium (Fig. IV-1). This is supported by electron micrographs which

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145 revealed the hyphae within the cell walls and by large areas of lysis and wall breakdown well away from the advancing mycelia (Fig. IV-3b and c). infrastructure of penetration and colonization Penetration of waterhyacinth leaves by A. zonatum occurred either through the stomata (Fig. IV-2a) or directly through the cuticle (Fig. IV-2b) . In most cases the hyphae firmly cemented themselves to the leaf surface by secretion of a mucilaginous substance (Fig. IV-2a). In other cases, however, there did not appear to be a cementing matrix (Fig. IV-2b) . Acremonium zonatum did not appear to produce appresoria. However, direct penetration of the cuticle by mechanical force did apparently occur, at least in part, as evidenced by the inward displacement of the cuticle at the .site of penetration (Fig. IV-2b). Similar results were . ;ained with Collet otrichum graminicola on maize (141). Additional evidence for direct penetration by A. zonatum was the formation of papillae (Fig. IV-2b). Papillae were seen to form immediately opposite the infection peg in the epidermal cell wall. There is good evidence (3) that papillae are formed in response to mechanical pressure during penetration. Penetration through either the epidernal walls (Fig. IV-2b and c) or xylem walls (Fig. IV-2d) was apparently

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146 accomplished with equal ease. There was no sign of wall tearing at the penetration site or disturbance of adjacent wall material which indicated that penetration was enhanced by localized secretions of cellulases and other wall degrading enzymes. The resolution of the micrographs did not permit a clear interpretation as to whether a haustorium was formed, or not since it was not possible to distinguish between fungal cell wall and host-cell plasmalemma. Figure IV-M-a is a cross-section of a collection of hyphal strands which pulled away from the epidermis but illustrates the extent to which this fungus secretes a mucilaginous matrix to cement itself to the host cell surface. Micrograph IV-Ub also illustrates this matrix and perhaps the first stage of penetration through the epidermis. In the area of immediate contact between the host cell wall and the fungus there was an area of wall material which appeared to be undergoing degradation. This may be the point of localized enzyme secretion. Penetration of the phenol cell by A. zonatum is shown in Figs. IV-5 and IV-6 . The hyphae were cemented to the cell surface and penetration through the wall occurred as it did in other cells. Once inside the phenol cell, however, the hyphae appeared extremely vesiculated, granulated, and distorted. It appeared that the hyphae were killed, by the high concentrations of polyphenolic compounds within the cell.

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147 An additional observation made from electron micrographs of diseased waterhyacinths was the presence of hyphae within the xylem (Fig. IV-3a). Disease symptoms incited by \. zonatum do not include those indicative of vascular i Issue infection such as wilting and it was therefore unexpected when hyphae were consistently observed within the ^acheary elements. vtological alterations induced by infection One of the most striking changes in waterhyacinth cells infected with A. zonatum was the disappearance of starch granules in the chloroplasts . The chloroplasts in palisade cells of large healthy waterhyacinth leaves had an abundance of starch granules in them (Fig. IV-7a). After infection, there was a noticeable absence of starch in the chloroplasts (Fig. IV-7b). A second noticeable change after infection was the build up of plastoglobuli within the chloroplasts. Chloroplasts of healthy waterhyacinth leaves consistently had several plastoglobuli (Fig. IV-8a). After infecton, however, the number and size of these plastoglobuli increased greatly (Fig IV-8b). A third cytological change after infection was a build up of microbodies within the cells. Healthy waterhyacinth leaf cells had only a few microbodies but after infection they increased In number (Fig. IV-9a). These

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148 generally appeared during the later stages of disease after starch disappeared and the plastoglobuli started to increase . Associated with the appearance of microbodies in the cell, the chloroplasts began to distort and the thylakoids started to distend (Fig. IV-9a). During the final stages of disease, the chlorplasts, as well as other organelles, lost their integrity completely, plastoglobuli fused forming large, irregular complexes, and the cytoplasm took on a very granular appearance (Fig. IV-9b). Eventually, the entire cell became convoluted and filled with dark, electron-dense material which was indicative of necrosis and cell death (Fig. IV-10).

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Figure IV-1. Flow diagram for testing of carbohydrate degrading enzymes produced by Acremonium zonatum .

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150 T3 CD E CO LU CO <
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Figure IV-2 (a d). Penetration of waterhyacinth leaf by Acremonium zonatum . a. Hyphae (h) penetrating through an open stoma. sc= substomatal cavity, gc= guard cell. Arrow points to mucilaginous matrix secreted by the fungus (6,200 X). Hyphae (h) penetrating the cuticle (cu) and epidermal cell wall (cw). ip= infection peg, p= papillae (14,400 X). c Hyphae (h) penetrating the epidermis of a waterhyacinth leaf. cw= cell wall (22,400 X). d. tf< ,) :e penetrating the cell wall (cw) of a xylary ele." i-'30,000 X).

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152

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Figure IV-3a. Cross-section of Acremonium zonatum observed in xylem tissue of diseased waterhyacmth leaf. h= hyphae, x= xylem. (5,200 X). Figure IV-3 (b c). Degradation of wall material in i.terhyacinth by Acremonium zonatum . b. Growth of A. zonatum within the cell wall. h= hyphae, cw= cell wall, ml= middle lamellae (10,000 X). Growth of A. zonatum within the cell wall showing large areas of lysis (arrows). Middle lamellae (ml) remains intact (27,500 X).

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15^

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Figure IV-M-a. Attachment of Acremonium zonatum to the cuticle. h= hyphae , E= epidermis. Arrow points to mucilaginous cementing substance (9,200 X). Figure IV-4b. Attachment of Acremonium zonatum to eoidermis and the possible area of localized enzyme secretion h = hyphae, cw= cell wall. Part of the cell wall immediately beneath the hyphae is eroded and suggestive of enzymatic degradation (32,0 00 X).

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156

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Figure IV-5 (a c). Penetration of phenol cell by Acremonium zonatum . Kyphae on the external cell surface appear viable while those inside the phenol cell are highly vesiculated and appear nonviable. h=hyphae, phc= phenol cell, pbphenol body, sp= septum. a= 6,200 X; b= 27,000 X; c= uo ,noo x.

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158

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Figure IV-6. Phenol cell invaded by Acremonium zonatum . Hyphae (h) appear granulated and nonviable, phc= phenol cell, pb= phenol body. (9,200 X).

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160

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Figure IV-7 (a b) . Breakdown of starch reserves in chloroplasts during disease. a. Chloroplasts in healthy palisade cell (pc) showing an abundance of starch (s) (6,800 X). b. Choroplasts in diseased palisade cell (pc) showing the absence of starch. c= chloroplast , n= nucleus, pl= plastoglobuli (6,200 X).

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162 s . •* •\ «• ©

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Figure IV-8 (a b). Increase in the number of plastoglobuli in chloroplasts during disease. a. Chloroplast of a healthy cell depicting only a few plastoglobuli (pi). (48,000 X). b. Chloroplasts in an infected cell showing the increase in plastoglobuli (pi). m= mitochondrion. Arrow points to area of thylakoid disruption (2 500 X) .

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164

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Figure IV-9a. Increase in the number of microbodies in cytosol as a result of infection with Acremonium zonatum. mb= microbody, m= mitochondria, th= thylakoids (16,000 X) . Figure IV-9b. Destruction of chloroplast integrity during later stages of disease. c= chloroplast, cw= cell wall (18,000 X).

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166

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Figure IV-10. Diseased palisade cell showing extent of necrosis and cellular breakdown. E= epidermis, cw= cell wall, h= hyphae. (9,200 X).

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168

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169 Discussion When a fungal cell contacts a susceptible host prior to penetration, an amorphous, electron-dense layer is often found between the walls of the host and the pathogen (15). This material is probably a fungal secretion and is assumed to aid adhesion of the pathogen to the host cell. Electron micrographs of infected waterhyacinth leaf tissue indicate that A. zonatum produces abundant secretions which serve as a cementing matrix. It is quite common at almost any interface between the pathogen and host cell but is apparently in greatest amounts on the surface of epidermal cells and stomata. Since the outer surface of a leaf would be expected to be subjected to greater physical stresses than the internal surface, it is quite logical that a fungus would require greater adhension forces externally than it would internally. It is also probable that the mucilaginous matrix serves as a buffer or insulator from the external environment during the course of penetration, thereby protecting the pathogen from dessication or other deleterious conditions. Entry of A. zonatum into waterhyacinth leaves was through rhe stomata or by direct penetration of the unbroken cuticle. Under natural conditions it is presumed that entry is by way of the stomata since conditions of high humidity favor disease development and the substomatal cavities would

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170 conceivably provide this needed moisture. Similar results were obtained by Rintz (152). However, when the relative humidity was maintained at 99-100% after inoculation, A. zonatum was capable of penetrating the cuticle directly. The method by which fungi penetrate their hosts has been a topic of considerable interest. For many years, direct penetration of plant cuticles and cell walls by pathogens was thought to be primarily a mechanical process (61). More recently, this idea has been modified to include enzymatic activity in addition or in combination with mechanical force (191). The idea that enzymes may be involved in plant tissue breakdown during pathogenesis was initiated by DeBary in 18 86 [see Bateman and Millar, (8)]. Working with the fungus Sclerotinia libertiana , DeBary demonstrated that a thermolabile substance from the fungus brought about disorganization of the host tissue. Brown (18) subsequently described a similar substance from Botrytis cinerea . Tissue macerating unzymes have since been described from numerous microi'^anisms. For a detailed discussion of these enzymes see . reviews by Brown (19), Albersheim et al . (4), and J, Ojman and Millar (8). Pectic enzymes are implicated almost routinely as a feature of host-pathogen interactions. Their involvement in the degradation of pectic constitutents of cell walls and of the middle lamella in plant tissues has been reported for

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171 such diverse types of diseases as soft rots, dry rots, wilts, blights, and leaf spots and for many types of pathogenic agents such as fungi, bacteria, and nematodes (8). It is perhaps significant that many of the pathogenic fungi and bacteria examined have been found capable of producing pectic enzymes. On the other hand, that a pathogen has this property does not explain why the organism is pathogenic (8). Rather, pectic enzyme production is likely to be but one of several properties of the pathogen, all of which are acting in concert to determine the pathogenecity of the organism. In regard to A. zonatum , pectic enzymes were not detected in_ vitro ; however, micrographs of this fungus penetrating cell walls indicated that they were perhaps produced, at least in small concentrations, and most probably were localized near the hyphal tip. Although some fungi are capable of excerting enough force to cause localized indentations in films of cured resins (117) there is little evidence that such pressure alone is responsible for penetration of plant cell walls. Mechanical pressure alone ...tjuld be expected to show evidence of tearing and distortion of the cell wall around the penetration site and these signs were not observed. Instead, penetration was accomplished i : i-oi.'sJh smooth-bordered holes in the cell wall which indicaii-M a presoftening of the wall constitutents prior to

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172 penetration. Mechanical pressure was involved to some extent, however, during penetration of the cuticle. Electron micrographs showed an inward bending of the cuticle which can best be explained by the application of force. In addition, there was no excessive damage to the middle lamella, which is composed chiefly of pectin substances, while extensive damage occurred to the more cellulosic walls. In this regard, A. zonatum did produce extensive amounts of a cellulolytic enzyme in vitro as evidenced by its ability to cause pit formation on CMC medium. Likewise, in vivo cellulolytic activity of the fungus was demonstrated by the extensive degradation and lysis of the cell walls . It is not possible from this study to determine which cellulolytic enzyme(s) is involved but Reese et al . (147) have reported that the ability of microorganisms to develop an enzyme capable of hydrolyzing the B, 1-4 glucosidic linkage found in cellulose and its derivatives is widespread anv'iig microorganisms. The degradation of native cellulose, hoVv-^/er, is less common. Thus, microorganisms which are c ^d as non-cellulolytic may develop the C enzyme, the activity of which is measured by its capacity to degrade carboxymethylcellulose , but are unable to breakdown native cellulose . The earliest and most consistent morphological response

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173 to pathogens which penetrate directly is the formation of structures called papillae (191). Several lines of evidence suggest that mechanical forces applied by the pathogen during penetration may provide the stimulus for papilla formation (3). First, papillae are restricted to areas beneath or immediately adjacent to the point of penetration. Second, papilla-like structures can be induced by gentle pin-pricks. Third, when no evidence of localized mechanical force is found during penetration, palillae are not formed. Finally, papilla-like structures are not formed in response to treatment with the pathotoxin victorin which causes other morphological changes typical of disease. The proposed function of papillae is to impede or block penetration by some pathogens but evidence for such a role is far from conclusive (191). Papillae are formed in epidermal cells of waterhyacinths during penetration by A. zonatum but are not formed in other cell types. This is additional evidence to support mechanical force as the most probable means of initial entry i u.o its host. . In summary it can be stated that entry of A. zonatum thnmifrh the cuticle was mechanical; penetration through cell walls involved production of cellulases and to a lesser extent pectinases; and penetration from cell to cell was also enhanced by localized pectinase production.

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174 One of the proposed functions of the phenol cell, is defense against invading pathogens. Electron micrographs of invaded phenol cells support this theory. The hyphae penetrate the phenol cells, perhaps as they would any other cell, but once inside are confronted with a collection of phenolic compounds in concentrations that are toxic. The hyphae appear less electron-dense than those in other cell types, highly vesiculated, and cytoplasmic organelles such as mitochondria are not evident. In other words, the hyphae are nonviable. It would appear that the concentration of phenols in those cells is such that they are no longer stimulatory to the fungus' growth but are toxic to it. In this case the phenol cell is able to stop the advancing mycelia at various points within the leaf, thereby limiting the infection. This could also help explain why disease severity in this plant is limited to only 40% of the leaf. The presence of hyphae in the tracheary elements was unexpected. However, A. zonatum is closely related to members of the genus Cephalosporium , which has many species that are vascular pathogens of other plants. It is postulated from the results of this study, that A. zonatum is •'. anslocated through its host in the xylem without causing tissue destruction or blockage. On the other hand, it does cause extensive wall degradation of the surrounding mesophyll cells. This suggests that this fungus lacks the

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175 necessary enzymes to degrade the lignified materials present in the secondary wall thickenings of xylary cells. Several cytological changes due to infection were observed in the cells of waterhyacinth leaves. The most noticeable was the disappearance of starch granules from the chloroplasts of palisade cells. Changes in starch content following infection have been observed in many foliar diseases (191). The general pattern is an initial decrease followed by a marked increase with heavy accumulations around the margins of the lesions. This presumably is brought about by the increased respiration which occurs soon after infection and serves to increase anabolic pathways (61). The accumulation of starch shortly after infection has been attributed to an increase on C0„ fixation in the dark by plants (198). Similar results were obtained by Luke and Freeman (99) in victorin-treated oat leaves and by Wang (137) in Uromyces phaseoli infected bean leaves. Later in infection, however, starch content declines drastically until most or all of it is gone. A second noticeable cytological change during pathogenesis was the build up of plastoglobuli in chloroplasts. A similar increase in plastoglobuli was reported in spinach plants infected with Albugo occidentales (60). The function of plastoglobuli is unknown but their occurrence in the stroma is a characteristic feature of chloroplasts fixed

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176 with osmium tetraoxide (176). Several lines of evidence indicate that they are not artifacts (65). Studies also suggest that they may be a product of senesence since they increase in size and number during aging (43). The exact chemical nature of plastoglobuli is not clear although it is generally believed that they represent a reservoir of excess lipid (65). An interesting observation has been made by Adams and Strain (1) in a study of the drought-deciduous desert plant Cercidium . They found that chloroplasts in the rather ephemeral leaves which appear after a heavy rainfall contained starch and are ultrastructurally similar to those of higher plants. The chloroplasts in the green bark tissue, which evidently provides a major source of photosynthetically fixed carbon to the plant, resemble those in the ephemeral leaves of other plants, except they lacked starch and have numerous large plastoglobuli. They suggested that the plastoglobuli may represent a form in which photosynthetically fixed carbon is stored in these chloroplasts. Thus, plastoglobuli may have the same general role as starch in most other plants. Thomson (176) considers this an attractive hypothesis, particularly in regard to p] ? nts where limited water may be available for metabolic pri :esses and from the point of view of effecient energy conservation and utilization. It is unlikely that a freefloating aquatic plant such as the waterhyacinth would be

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177 subjected to limited water supplies under natural growing conditions. Therefore, plastoglobuli in this plant must have a different function. The last major cytological change observed in infected waterhyacinths was the increase in microbodies within the cells. The term "microbody" was introduced into the literature by Rhodin in 1954 [see DeDuve (41)] to designate a special type of cytoplasmic body present in the convoluted tubule cells of the mouse kidney, characterized by a single membrane and finely granular matrix. Similar structures have since been found in yeast cells (57), other fungi (111,116) green algae (168) and higher plants (20,67,84). These structures have been called by various names (peroxisomes, glyoxysomes and crystalloids) depending on the type of function proposed for them (178). The biological significance of microbodies has been attributed to several things, including serving as mitochondrial precursors (158). The main functions appear to be involved with gluconeogensis (178). In germinating fatty seeds, glyoxysomes are involved in the conversion of fats to sugars via B-oxidation and the c.lyoxylate cycle (178). Leaf peroxisomes, however, are /olved in the conversion of nonphosphorylated compounds d: : 'ved from photosynthesis to glycine, serine, and C-, cc pounds via the glycolate pathway (178). From glycolate, glycine, or serine, the pathway is gluconeogenic in the

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178 light since it leads to phosphoglycerate and then to sugars (178). For a detailed discussion of microbodies the reader is referred to the excellent review by Tolbert (178). The appearance of microbodies in infected waterhyacinth leaves at about the time starch is lost in the chloroplasts represents, perhaps, an alternate method of producing needed carbohydrates for energy production. During the final stages of pathogenesis, chloroplasts were observed to break down along with other cell organelles. The grana distend, plastoglobuli may coalesce, and organelle structure is lost completely. Plasmolysis occurs, oxidation of cellular components takes place, and the end result is the formation of dark brown melanin pigments typical of necrosis and death.

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SUMMARY AND CONCLUSIONS The waterhyacinth [ Eichhornia crassipes (Mart.) Solms.] is a free-floating vascular hydrophyte that has colonized much of Florida's inland waters. In 1970, a program was initiated at the University of Florida to study biological control of this noxious plant with phytopathogens . One of the pathogens currently being studied is the fungus Acremonium zonatum (Sawada) Gams. It causes severe spotting on both leaves and petioles of this plant under conditions of high humidity. During field trials with this fungus, it was observed than small, young, plants displayed fewer symptoms after infection than did larger plants in the same plots. It also appeared that large plants infected with A. zonatum exhibited a faster rate of leaf regeneration than did smaller plants. The present study was initiated to determine if small plants were in fact more resistant to A. zonatum than large plants; if meristematic activity in the plants was altered after infection; and, if so, to what extent host phenolic compounds and their oxidizing enzymes, namely polyphenoloxidase (PPO), were responsible. Waterhyacinths displayed various degrees of resistance to A. zonatum depending on their morphotypic state of development. Results of this study indicate that these differences in resistance are due to the variations in phenol 179

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180 chemistry among plants of different sizes and to subsequent changes induced by infection (Table S-l). Small plants are more resistant to fungal attack than are medium or large plants, based upon the number of lesions/ leaf after infection. It appears that the presence of high concentrations of phenolic compounds does not itself impart resistance to the pathogen. Rather it is the oxidation of these compounds by enzymes, such as polyphenoloxidase (PPO), which is responsible for the resistance. This view is supported by qualitative and quantitative data on the phenols in plant morphotypes and is coincident with the observed differences in resistance. o Small plants, by virtue of having fewer phenol cells/mm leaf area, have less total phenol content/leaf than larger plants. If phenol content alone, was responsible for disease resistance, then small plants would be more susceptible than large plants but they were not. In this case PPO activity is apparently the mediating factor. The rate of enzyme activity in small plants is three-fold that in large plants; presumably therefore, oxidation of polyphenols to quinones is much greater in small plants. Thus, small plants are initially more resistant to pathogenic attack than are larger plants. After the disease has progressed for several weeks the differences in resistance among the morphotypes is no

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181 Q

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182 longer evident. Each plant size exhibits a percenttotal-diseased leaf area which is statistically the same (approximately 40%). It is believed that this equalization of disease severity results from a gradual loss in resistance by small plants while at the same time there is a gradual increase in resistance by large plants. Again, quantitative data of the phenol metabolism can be correlated with this change. The total phenol content decreased significantly after infection in smalland medium-sized plants. This is coincident with a reduction in PPO activity. The coupling of these two phenomena may account for the decrease in resistance of small plants. Large plants, on the other hand, retain their total phenol content and at the same time exhibit a three-fold increase in FPO activity. Therefore, an increase in polyphenol oxidation would be expected to occur and could account for the increase in resistance in large plants. In essence, then, the point being made is: if infected -small plants retained the phenol content and PPO activity of ; -filthy plants, then disease severity would probably be ll'"ii:ed to much less than 40%. Similarly, if infected large plrini--: retained the PPO activity of healthy plants, disease would progress to a much higher percentage, perhaps 60-70%. Howev-i , because each morphotype responds to infection

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183 differently (in most cases in contrast to each other) disease severity balances among the plant sizes at approximately 40% of the leaf-surface area. If disease severity is viewed, not from a percentage of leaf-area infected, but as a reduction in plant growth, then data on leaf regeneration rates among the morphotypes becomes of prime importance. It has been observed that infected large plants regenerate two to three times as many new leaves as do infected small plants. This too, is correlated with the plant's phenol chemistry. It has been shown that A. zonatum is capable of synthesizing indoleacetic acid in vitro and that this is one explanation for the increased growth observed in large plants. More important, however, is the fact that phenols are known inhibitors of IAA oxidase , the enzyme responsible for controlling the IAA level in the plant. It has already been pointed out that the different waterhyacinth morphotypes vary in phenol content, both prior to and after infection. The higher phenol content in large plants could account for the increased growth observed in large plants by inhibition of the IAA oxidase system. Perhaps the most significant data supporting a positive .? e for phenols in disease resistance comes from the location studies of PPO in healthy and diseased plants. !.' i/,-..ne activity is localized in the thylakoids of

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184 chloroplasts in only three cell types in healthy plants. After infection there is a "turn on" in PPO activity in all cells which contain chloroplasts. This turn on in FPO activity is highly suggestive of a vital role for enzymatic oxidation of polyphenols during disease. Before disease can ensue, the pathogen must come into contact with and penetrate its host. In this regard, A. zonatum can enter the waterhyacinth by either of two ways: through open stomata or by directly penetrating the unbroken cuticle of the leaf. Intracellular colonization is enhanced by the diffuse secretion of cellulolytic enzymes and perhaps by the localized secretion of pectolytic enzymes. Growth of A. zonatum was either unaffected or stimulated by seven different phenolic acids in concentrations up to 1000 ppm in minimal media. When yeast extract was added to the media as a growth supplement, one phenolic acid, pcoumaric, was found to be inhibitory. In addition, fungal growth was enhanced on media containing yeast extract and extracts from diseased leaves over that on media containing healthy leaf extracts. Several cytological changes were observed in the cells om infected waterhyacinth leaves. First, chloroplasts in cLis of healthy leaves have an abundance of starch granules which disappear after infection. Second, there are only a few plastoglobuli in chloroplasts in healthy cells, but

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185 after infection, they increase both in size and in number. Third, there is a noticeable increase in microbodies in the cytosol of infected cells. It is believed that each of these cytological changes is the result of a shift in host metabolism induced by infection. It is concluded that waterhyacinths have at least two distinct biochemical defense mechanisms that are related to phenol metabolism and plant size. The first is the presence of high concentrations of polyphenols in specialized phenolcells which, under the proper conditions, can serve as toxicants to potential pathogens. The second proposed defense mechanism of waterhyacinths is an acceleration of its growth rate brought about by the inhibition of IAA oxidase by the phenolic compounds. Which of the above mechanisms is operational is depen-nt upon the plant's morphotypic stage of development. It i:j believed that initially small plants defend against p.< isogenic attack by virture of their high PPO activity whereas large plants respond by increased leaf production. Mi Hum-sized plants appear to have a combination of both me i. nisms. In consideration of A. zonatum as a potential biocontrol agent for waterhyacinths, it is concluded that best control would be achieved with small, young, plants rather than with larger, more mature plants. In this regard,

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186 control procedures should be initiated early in the spring when new plants start to grow and colonize the body of water.

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10 LITERATURE CITED Adams, M.S. and B.S. Strain. 1969. Seasonal photosynthetic rates in stems of Ceridum floridum Benth. Photosynthetica 3:55-62. Agharkar, S.P. and S.N. Banerjee. 1932. Fusarium sp . causing disease of Eichhornia crassipes Solms . Proc. Indian Sci. Cong. 19:298. Aist, J.R. 1976. Papillae and related wound plugs of plant cells. Ann. Rev. Phytopathol . 14:145-163. Albersheim, P., T.M. Jones, and P.D. English. 1969. Biochemistry of the cell wall in relation to infective processes. Ann. Rev. Phytopathol. 7:171-194. Arnon, D. 1949. Copper enzymes in isolated chloroplasts. Polyphenoloxidase in Beta vulgaris . Plant Physiol. 24:1-15. Baker, K.F. and R.J. Cook. 1974. Biological Control of Plant Pathogens. W.H. Freeman Co., San Francisco. 433 p. Banerjee, S.N. 1942. Fusarium equiseti Sacc . Causing a leaf spot on Eichhornia crassipes . J. Dept . Sci. Calcultta Univ. 1:29-37. Bateman, D.F. and R.L. Millar. 1966. Pectic enzymes in tissue degradation. Ann. Rev. Phytopathol. 4:119146. Batra, G.K. and C.W. Kuhn . 1975. Polyphenoloxidase and peroxidase activities associated with acquired resistance and its inhibition by 2-thiouracil in virus-infected soybean. Physiol. Plant Pathol. 5:239-248. Beckman, C.K. and W.C. Mueller. 1970. Distribution of phenols in specialized cells of banana roots. Phytopathology 60:79-82. 11. Beckman, C.H., W.C. Mueller and W.E. McHardy. _1972. The localization of stored phenols in plant hairs. Physiol. Plant Pathol. 2:69-74. 12. Bell, A. A. 1974. Biochemical basis of resistance of plants to pathogens. In F.G. Maxwell and F.A. Harris 187

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(Eds.) Proc . of the Summer Institute of Biological Control of Plant Insects and Diseases, pp. 403-462. Univ. Press, Jackson, Miss. 13. Bennett, P.D. 1967. Notes on the possibility of biological control of the water hyacinth Eichhornia crassipes . PANS (C) 13:304-309. 14. Black, H.S. 1968. Pectolytic enzyme product by Phymatotrichum omnivorum . (Abstr.) Phytopathology 58 :1044. 15. Bracker, C.E. and L.J. Littlefield. 1973. Structural concepts of host-pathogen interfaces. In R.J.W. Byrde and C.V. Cutting (Eds.) Eungal Pathogenicity and the Plant's Response. pp. 159-318. Academic Press, N.Y. 16. Braun, A.C. 1959. Growth is affected. In J.G. Horsfall and A.E. Dimond (Eds.) Plant Pathology. An Advanced Treatise. Vol. 1. pp. 189-248. Academic Press, N.Y. 17. Brown, A.E. and T.R. Swinburne. 1973. Factors affecting the accumulation of benzoic acid in Bramley's seedling apples infected with Nectria galligena . Physiol. Plant Pathol. 3:91-99. 18. Brown, W. 1915. Studies in the physiology of parasitism. I. The action of Botrytis cinerea . Ann. Bot . 29: 313-348. 19. Brown, W. 1965. Toxins and cell-wall dissolving enzymes in relation to plant disease. Ann. Rev. Phytopathol. 3:1-18. 20. Chabot, J.F. and B.E. Chabot . 1974. Microbodies in conifer needle mesophyl. Protoplasma 79:349-358. 21. Charudattan, R. 1973. Pathogenicity of fungi and bacteria from India to hydrilla and waterhyacinth. Hyacinth Contr. J. 11:44-4 8. 22. Charudattan, R. and K.E. Conway. 197 6. Mycolepto discus terrestris leaf spot on waterhyacinth. Plant Dis. Rep. 60:77-80. 23. Charudattan, R., K.E. Conway and T.E. Ereeman. 1975. A blight of waterhyacinth, Eichhornia crassipes caused by Bipolaris stenospila ( Helminthosporium stenospilum ) (Abstr. ) . Proc. Amer . Phytopathol. Soc . 2:65.

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203 189. Wardrop , A.B. and J. Cronshaw. 1962. Formation of phenolic substances in the ray parenchyma of angiosperms. Nature. 193:90-92. 190. Webber, H.J. 1897. The waterhyacinth and its relation to navigation in Florida. Bull. 18. U.S. Dept . Agr. Div. Bot . 20 p. 191. Wheeler, H. 1975. Plant Pathogenesis. SpringerVerlag, N.Y. 106 p. 192. Wilson, C.L. 1969. Use of plant pathogens in weed control. Ann. Rev. Phytopathol. 7:411-434. 193. Wolf, F.T. 1952. The production of indoleacetic acid by Ust ilago zea and its possible significance in tumor formation. Proc . Nat. Acad. Sci. U.S. 38:106-111. 194. Wolf, F.T. 1955. The production of indoleacetic acid by the cedar apple rust fungus and its identification by paper chromatography. Phytopathol. Z. 26:219-22 3. 195. Woodward, R.E. 1975. Phenolic compounds of Hydrilla verticillata (L.f.) Royle and some herbicide related fluctuations in their content. M.S. Thesis, Univ. South Florida, Tampa. 61 p. 196. Yarwood , C.E. 1934. The comparative behavior of four clover-leaf parasites on excised leaves. Phytopathology. 24:797-806. 197. Yarwood, C.E. 1959. Predisposition. In J.G. Horsfall and A.E. Dimond (Eds.) Plant Pathology. An Advanced Treatise. Vol. I. pp. 521-562. Academic Press, N.Y. 198. Zaki, A.I. and C.J. Mirocha. 1965. Carbon dioxide fixation by rust-infected bean plants in the dark. Phytopathology. 55:130 3-1308. 199. Zeiger, C.F. 1976. Biological control of alligatorweed with Agasicles n. sp . in Florida. Proc. Southern Weed Conf. 20:299-303. 200. Zenk, M.H. and C.C. Gross. 1972. The enzymatic reduction of cinnamic acids. Rec . Adv. Phytochem. 4:87-106. 201. Zettler, F.W. and T.E. Freeman. 1972. Plant pathogen as biocontrols of aquatic weeds. Ann. Rev. Phytopathol. 10:455-470.

PAGE 220

BIOGRAPHICAL SKETCH Raymond D. Martyn , Jr. was born in Washington, B.C. on December 15, 1946. He was graduated from Pompano Beach Senior High School in June, 1964, and obtained an Associate of Arts degree in Engineering from Palm Beach Junior College in April, 1966. He was awarded the degrees of Bachelor of Science in Biology in June, 1969 and Master of Science in Microbiology in June, 1971, from Florida Atlantic University . From September, 1971 to August, 1973, he served as laboratory supervisor at the Biological Control Laboratory at Florida Atlantic University. From September, 1973 to May, 1974, he was a biology instructor at Palm Cove Beach High School . In June, 1974, he entered the Plant Pathology Department at the University of Florida to pursue graduate studies towards the degree Doctor of Philosophy and is presently a candidate for that degree in June, 1977. He is married to Jane D. Brooks and has a six-year-old daughter, Susan, by a former marriage. 204

PAGE 221

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. I L-Cc,. Thomas E. Freeman, Chairman 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. 6a.™&G. ^cS^tU Daniel A. Roberts 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. Rob/e'rt E. Stall Professor of Plant Pathology I certify that I have read this study and that in my 'pinion 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. ' >*tA Herbert H. Luke Plant Pathologist, U.S.D.A,

PAGE 222

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. ..... f il„. l CO Thomas E. Humphreys Professor of Botany This dissertation was submitted to the Dean of the College of Agriculture and to the Graduate Council, and was accepted as partial fulfillment of the requirements for the degree of Doctor of Philosophy. June, 1977 f ,' i College of Agrj-culti Dean ,' < College of Ag£; Dean, Graduate School

PAGE 223

iilii" 3 1262 08553 1»3T


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DISEASE RESISTANCE MECHANISMS IN WATERHYACINTHS AND THEIR
SIGNIFICANCE IN BIOCONTROL PROGRAMS WITH PHYTOPATKOGENS
By
RAYMOND DEWINT MARTYN, JR.
A DISSERTATION PRESENTED TO THE GRADUATE COUNCIL OF
THE UNIVERSITY OF FLORIDA
IN PARTIAL FULFILLMENT OF THE REQUIREMENTS FOR THE DEGREE
OF DOCTOR OF PHILOSOPHY
UNIVERSITY OF FLORIDA
1977

This above all: to thine own self be true . .
William Shakespeare
Hamlet; Act I, scene iii

To ir.y parents, who had the wisdom and foresight to
know the difference between "guidance" and "insistence",
and who used as one of the cornerstones of my education,
Robert W. Service's poem "The Quitter" which appears on
the following page . . .
To my wife, Dickie, whose unyielding faith and many
hours of unselfish help and patience were perhaps the
greatest factors in the completion of this program . . .
To my daughter, Susan, whose 6-year-old smile made
it all worthwhile, when I overheard her tell a playmate,
"My Daddy is a plant doctor!"

The Quitter
When you're lost in the wild and you're scared as a child,
And death looks you bang in the eye;
And you're sore as a boil, it's according to Hoyle
To cock your revolver and die.
But the code of a man says fight all you can,
And self-dissolution is barred;
In hunger and woe, oh it's easy to blow --
It's the hell served for breakfast that's hard.
You're sick of the game? Well now, that's a shame!
You're young and you're brave and you're bright.
You've had a raw deal, I know, but don't squeal.
Buck up, do your damnedest and fight!
It's the plugging away that will win you the day,
So don't be a piker, old pard;
Just draw on your grit; it's so easy to quit --
It's the keeping your chin up that's hard.
It's easy to cry that you're beaten and die,
It's easy to crawfish and crawl,
But to fight and to fight when hope's out of sight,
Why, that's the best game of them all.
And though you come out of each grueling bout,
All broken and beaten and scarred --
Just have one more try, it's dead easy to are;
It's the keeping on living that's hard.
Robert W. Service
IV

ACKNOWLEDGEMENTS
I wish to express sincere gratitude to Dr. Thomas E.
Freeman, Chairman of my Supervisory Committee, for his
friendship, advice, guidance, and patience during the course
of this study, and for his criticism and encouragement in
appropriate doses for three years which ultimately made this
dissertation possible.
I also wish to extend thanks to members of my Super¬
visory Committee, Dr. T.E. Humphreys, Dr. H.H. Luke, Dr.
D.A. Roberts, and Dr. R.E. Stall for their advice and
friendship, and for their time spent in critical review of
this manuscript.
A special thanks is extended to Mr. D.A. Samuelson for
his many hours of assistance during the ultrastructural and
cytochemical portions of this study, and for the many hours
of help in preparing the electron micrograph plates.
Gratitude is also extended to Dr. H.A. Aldrich for his
kindness for allowing use of equipment and facilities of the
Biological Ultrastructure Laboratory, and to Ms. Janet Plaut
for performing the many statistical analyses used throughout
this dissertation.
This research supported in part by the U.S. Army Corps
of Engineers, Florida Department of Natural Resources, U.S.
Department of Interior, Office of Water Resources and
Research Act as amended and by the University of Florida
Cer.xer for Environmental Pro gleams.
v

TABLE OF CONTENTS
Page
ACKNOWLEDGEMENTS v
LIST OF TABLES viii
LIST OF FIGURES ±x
ABSTRACT xiii
GENERAL INTRODUCTION 1
Part I The Aquatic Weed Problem I
Fart II The Potential of Biological Control . 5
Parr in Pathogens of Waterhyacinth with
Possible Biocontrol Potential ... 8
CHAPTER I RESPONSES OF WATERHYACINTH TO INFECTION
WITH ACREMONIUM ZONATUM AND ITS IMPLI¬
CATIONS IN BIOLOGICAL CONTROL 15
Introduction 15
Materials and Methods 17
Results T9
Discussion 28
CHAPTER II A CYTOCHEMICAL AND ULTRASTRUCTURAL STUDY
OF THE PHENOL CELLS AND POLYPHEKOLOXI-
DASE ACTIVITIES IN HEALTHY AND DISEASED
WATERHYACINTH LEAVES 37
Introduction 37
Materials and Methods 43
Results 4g
Discussion 84
CHAPTER III A BIOCHEMICAL STUDY OF THE PHENOLIC
ACIDS AND POLYPHENOLOXIDASE RATES IN
HEALTHY AND DISEASED WATERHYACINTH
LEAVES 9 0
Introduction 90
Materials and Methods 100
Results 108
Discussion 129
vi

Page
CHAPTER IV AN ULTRASTRUCTURAL STUDY OF PENE¬
TRATION AND COLONIZATION OF WATER-
HYACINTK BY ACREMONIUM ZONATUM 139
Introduction 139
Materials and Methods 141
Results 144
Discussion 159
SUMMARY AND CONCLUSIONS 179
LITERATURE CITED 187
BIOGRAPHICAL SKETCH 204
vii

LIST OF TABLES
Table Page
III-l Free phenolic acids detected in healthy
and A. zonatum-infected waterhyacinths by
thin layer chromatography 114
III-2 Phenolic acids detected in healthy water-
hyacinth leaves by thin layer chromatogra¬
phy and various locating reagents after
alkaline hydrolysis 115
II1-3 Fhenolic acids detected in A. zonatum-in¬
fected waterhyacinth leaves-by thin layer
chromatography and various locating rea¬
gents after alkaline hydrolysis 116
1II-4 R, values and color characteristics of
the phenolic acids detected in healthy
and A. zonatum-infected waterhyacinth
leaves after alkaline hydrolysis 117
III-5 Growth of A. zonatum or. healthy and A.
zonacum-Infected waterhyacinth leaf-
extract media 124
III-6 Growth of A. zonatum on phenolic acid
media 125
III-7 Growth of A. zonatum on phenolic acid
media with yeast extract 126
S-l Differences and similarities among
healthy and A. zonatum-infected water¬
hyacinth morphotypes 181
vi i i

LIST OF FIGURES
Page
CHAPTER I
Pig-
1-1
Symptoms of disease on water-
hyacinths incited by Acremonium
zonatum
23
Pig-
1-2
Quantitation of disease on small,
medium, and large waterhyacinths .
25
Fig.
1-3
Quantitation of leaf regeneration
rates of small, medium, and large
waterhyacinths
27
CHAPTER II
Pig-
II-l
Biosynthetic pathway for conver¬
sion cf ryrcsine to melanin . . .
42
Pig-
II-2
Flow diagram of procedure for
standard electron microscopy fi¬
xation and embedding
45
Fig-
II-3
Flow diagram of procedure for the
cytochemicai localization of po-
'yphenoloxidase
48
Pig-
II-4
Light micrographs of phenol cells
in healthy water-hyacinth leaves .
57
Pig-
II- 5
2
Number of phenol cells/mm leaf
area in small, medium, and large
waterhyacinth leaves
59
Pig-
II-6
Electron micrograph of phenol
cell in palisade cell layer of
waterhyacinth leaf tissue ....
61
Pig-
II- 7
Electron micrograph of phenol
cell in vascular tissue area of
waterhyacinth leaf
63
Fig.
II-S
Chloroplasts of healthy waterhya¬
cinth leaf tissue incubated with¬
out DOPA
65
IX

Page
Fig. II-9
Fig. 11-10
Fig. 11-11
Fig. 11-12
Fig. 11-13
Fig. 11-14
Fig. 114.5
Fig. 11-15
Fig. Itel7.
CHAPTER III
Fig. III-l
Fig. Ill-2
Localization of polyphenoloxidase
in healthy waterhyacinth leaf
tissue without lead postaining . 67
Localization of polyphenoloxidase
in chloroplasts of xylem paren¬
chyma cells in healthy waterhya¬
cinth leaves 69
Localization of polyphenoloxidase
in chloroplasts of bundle sheath
cells in healthy waterhyacinth
leaves 71
Localization of polyphenoloxidase
in chloroplasts of phenol cells
in healthy waterhyacinth leaves . 73
Chloroplasts of boiled, healthy
waterhyacinth leaf tissue incu¬
bated with DOPA 7 5
Chloroplasts of healthy waterhya¬
cinth leaf tissue incubated in
inhibitor (DDC) and DOPA .... 77
Localization of polyphenoloxidase
in chloroplasts of palisade cells
from diseased waterhyacinth
leaves 79
Localization of polyphenoloxidase
in chloroplasts of spongy meso-
phyll cells from diseased water¬
hyacinth leaves 81
Localization of polyphenoloxidase
in chloroplasts of cells several
centimeters away from infection
center 8 3
Principal phenolic acids found in
plants 94
Shikimic acid pathway for the
biosynthesis of monocyclic phe¬
nols and major derivatives ... 98
x

Page
Pig-
III-3
Flow diagram of procedure for ex¬
traction of ester-linked phenols
in plants
103
Pig-
111-4
Total phenol concentrations in
healthy and A. zonatum-infected
waterhyacinth morphotypes ....
119
Pig-
III- 5
Polyphenoloxidase activities in
small, medium, and large healthy
waterhyacinth leaves
121
Pig-
III-6
Polyphenoloxidase activities in
small, medium, and large diseased
waterhyacinth leaves
123
Pig-
III-7
In vitro synthesis of indoleace-
tTc from tryptophan by Acremonium
zonatum
128
CRAPTER IV
Pig-
IV-1
Flow diagram for testing of car¬
bohydrate degrading enzymes pro¬
duced by Acremonium zonatum . . .
150
Pig-
IV-2
Penetration of waterhyacinth leaf
by Acremonium zonatum
152
Pig-
IV-3a
Cross-section of Acremonium zona-
turn observed in xylem tissue of
diseased waterhyacinth leaf . . .
154
Pig-
IV-3-c
Degradation of wall material in
waterhyacinth by Acremonium
zonatum
154
Pig-
IV-4a
Attachment of Acremonium zonatum
to the cuticle
156
Pig-
IV-4b
Attachment of Acremonium zonatum
to epidermis and the possible
area of localized enzyme secre-
tion
156
Pig-
IV-5
Penetration of phenol cell by
Acremonium zonatum
158

Page
Pig-
IV-6
Phenol cell invaded by Acremonium
zonatum
160
Fig-
IV-7
Breakdown of starch reserves in
chloroplasts during disease . . .
162
Pig-
IV-8
Increase in the number of plasto-
globuli in chloroplasts during
disease
164
Fig-
IV-9a
Increase In the number of micro¬
bodies in cytosol as a result of
infection with Acremonium zonatum
166
Fig-
IV-9b
Destruction of chloroplast integ¬
rity during later stages of
disease
166
Fig-
IV-10
Diseased palisade cell showing
extent of necrosis and cellular
breakdown
168
XI1

Abstract of Dissertation Presented to the Graduate Council
of the University of Florida in Partial Fulfillment of the
Requirements for the Degree of Doctor of Philosophy
DISEASE RESISTANCE MECHANISMS IN WATERHYACINTHS AND THEIR
SIGNIFICANCE IN BIOCONTROL PROGRAMS WITH PHYTOPATHOGENS
By
Raymond DeWint Martyn, Jr.
June, 1977
Chairman: Dr. Thomas E. Freeman
Major Department: Plant Pathology
The pathological relationship between the floating
waterhyacinth, Eichhornia erasslpes (Mart.) Solms and the
fungus, Acremonium zonatum (Sawada) Gams, was investigated to
determine possible disease resistance mechanisms in the plant
as they relate to potential biocontrol agents. Waterhyacinths
were separated into three morphotypes based upon their leaf
2
surface area; small plants (leaves < 15 cm ), medium plants
2 2
(leaves 15-40 cm ), and large plants (leaves > 40 cm ) and
used for quantitailing symptoms of disease. Inoculated small
plants exhibited fewei lesions/leaf after two weeks than did
either medium or large plants; however, tile total percent
diseased leaf area for each morphotype was the same (approxi¬
mately 40%). It was observed that large plants regenerated
almost three times as many new leaves after infection deve¬
lopment than did either medium or small plants.
Biochemical, histochemical, cytochemical, and ultra-
structural studies were conducted on both healthy and diseased
xiii

morphotypes to determine what role host phenolic com¬
pounds had in disease development. Phenolic compounds in
waterhyacinth leaves are localized in specialized idioblasts
(phenol cells) immediately beneath both epidermal surfaces
and also in close association with the vascular tissue. The
concentration of phenol cells increased significantly from
2 . 2
a mean of 33.6/mm leaf area in small plants to 48.7/mm
in large plants.
In healthy plants, polyphenoloxidase (PPO) activity was
greater in small than in large leaves and was restricted to
the thylakoids of chloroplasts in only three cell types:
vascular parenchyma, bundle sheath, and phenol cells. After
infection by A. zonatum, PPO activity decreased in small
leaves but increased over 300% in large leaves. After
infection, PPO activity was observed in all chloroplasts
throughout the leaf.
Chlorogenic acid was the only free phenolic acid found
morphotypes of both healthy and diseased plants. Alka¬
line hydrolysis of healthy leaf tissue yielded six phenolic
acids from small and medium plants and nine from large
plants. After infection, one additional phenolic acid was
detected from small- and medium-sized leaves. No change in
the types of phenolic acids present in large leaves was
detected after infection. The concentration of total phenols
in healthy plants increased significantly from 92 yg/g fresh
leaf tissue in small to 104 yg/g in large leaves. There was
xiv

a significant decrease in total phenols |_n both small
and medium diseased plants while the concentration remained
constant in large diseased plants.
Acremonium zonatum grew significantly better when cul¬
tured on minimal media containing phenolic acids than it did
on media without these compounds. Acremonium zonatum was
inhibited by p-coumaric acid at 1000 ppm, when yeast extract
was added as a growth supplement to the media. In addition,
growth of the fungus on diseased plant-extract media was
stimulated significantly over growth on media containing
extracts from healthy plants.
Penetration of waterhyacinth leaves by A. zonatum
occurred directly through the cuticle or through the sto¬
mata. Cellular penetration was aided by the production of
cellulolytic enzymes. Penetration of the phenol cells re¬
sulted in death of the invading hyphae. Associated with
disease was the disappearance of starch granules from the
chloroplast, an increase in the number of plasfoglobuli
within chloroplasts, and a build-up of microbodies within the
cytosol.
The results presented in this study suggest that phenol
metabolism in waterhyacinth plays a significant role in the
defense against potential pathogens and may account for why
only a few of pathogens have been reported on this plant.
It appears that A. zonatum is capable of causing relatively
xv

severe damage to the waterhyacinth because of its high
tolerance to phenols and warrants continued study as a
potential biocontrol of this noxious aquatic plant.
xvt

GENERAL INTRODUCTION
Part I: The Aquatic Weed Problem
All plant and animal species in their native habitats
are subject to natural forces that control their population
levels. Natural enemies along with other environmental
Influences maintain a balance among populations of plants
and animals in an ecosystem. There is little question that
the parasites and predators existing in a particular system
are the greatest resource that we have for effective pest
suppression and management (180).
Man steps beyond Nature's boundaries, however, and
thereby sidesteps natural controls by transporting plant
and animal species to new habitats, and in so doing, often
causes disastrous shifts in the ecological balance between
species. Such has been the case with many of the noxious
aquatic plants in Florida. Exotic water plants imported into
this country as aquaria specimens and ornamentals have escaped
Into lakes and waterways and, once established, have created
sfeffiious control dilemmas. In areas where aquatic plants have
reached high densities, they greatly obstruct the water flow,
decrease the water level through increased rates of evapo¬
ration and transpiration, increase the rate of eutrophication,
1

2
interfere with. inavigation, prevent fishing and other water
recreational activities, depress real estate values, and
may, intone instances, present severe health hazards
(52, 75, 201). Infamous examples of these pestiferous
plants include the floating waterhyacinth, Eichhornia cras-
sipes (Mart.) Solms, Florida elodea, Hydrilla verticillata
(Casp.), Eurasian watermilfoil, Myriophyllum spicatum L.,
and alligatorweed, Alternanthera philoxeroides (Mart.)
Griseb.
The rampant growth of exotic water weeds in Florida and
other Gulf states has been attributed to several factors
(78, 118, 139). First, the year-round warm temperature and
extended photoperiod combine to give a growing season
almost the entire year. Secondly, many bodies of water
provide an abundance of inorganic compounds necessary for
luxuriant: plant growth. Thirdly, the absence of enemies
normally present in their native habitats does not allow the
natural system of checks and balances to operate. And,
lastly, most aquatic plants are capable of extremely rapid
vegetative reproduction. It is for these reasons that some
160,000 hectares of Florida's fresh water are weed-choked
(54 ) .
One of the most pestiferous aquatic plants in tropical
and subtropical climates is the floating waterhyacinth,
E. crassipes, the subject of this dissertation. The

3
genus Eichhornia is a member of the Fontederiacae family
and incHudespfour other species: E. paniculate, E. paradoxa,
E. azurea, and E. diversi fol|^| (13 9). Eichhornia crassipes
is the only speciefpwhich is free floating; all other
members of the genus are rooted either in shallow water
or near shore.
fjie waterHfifflcinth reproduces almost entirely by vegeta¬
tive meHns although sexual reproduction does occur. It
reproduces rapidly and will completely fill many lakes and
rivers in a single growing season, tenfound and Earle (139)
reported that E. crassifes is capable of doubling its mass
every 13-15. Taking an average rate of doubling of
two weeks and a gy-owing Season of •■SftK'r.t months, then ten
plants given plenty of room and good growing conditions
would produce 655 , Bo plants which would cover 0.6 hec¬
tares. These figure® Mphasize the tremendous rate of
colonisation of this species and the necessity of good
centrel^Htnods .
It is belJfiSged that the waterhyaciMáik is a native of
Brazil, Jyut has spread from there to nearly all of the
South Attrican and Central America» countries and through¬
out the worJÜ where the climate is favorable for its
development. Few7 tropical or subtrogBcJR countries are free
.from, waterhyacinths (97).
The accounts differ somewhat regarding its appej^Bnce
in the United States. There is seme evidence that it was

4
cultivated as a greenhouse exotic shortly after the War
Between the States (139); however, the earliest authentic
account details its introduction at the Cotton Centennial
Exposition at New Orleans in 1884 (88). It appeared in
Florida in 1890 (190) and has since become an important
aquatic pest. By the turn of the century it was reported
from all the southeastern coastal states as far north as
Virginia and westward to California (81).
Eichhornia crassipes was officially recognized as a
serious aquatic pest in this country on June 4, 1897, when
Congress passed an act authorizing the Secretary of War to
investigate the extent of obstruction to navigation in the
waters of Florida and Louisiana (139). Since that time, the
U.S. Army Corps of Engineers have been responsible for
clearing it from navigable waterways.
Florida, like many parts of the United States and
world, is in dire need of an efficient and effective means
of controlling noxious aquatic plants. Since their introduc¬
tion, millions of dollars, both tax and private, have been
spent on chemical and mechanical control of these weeds. An
estimated 1G to 15 million dollars is being spent per year
for the control of aquatic weeds in Florida alone, and this
figure is increasing every year (64). Despite this huge
financial expenditure, the total infestation continues to
grow and at present there is no end to the increasing costs
unless new control measures are found.

5
Pari XI: The Potential of Biological Control
In past years, control of waterweeds has ba&m on
two basic procesares. Both mechani^H and chemical controls
are used routinely in maintenance pregrams. However,
neither W’.cc on its own is completely satisfactory and the
w® 1 infestations continue to expand. McHe recently, the
concept of biological control was proposesd for aquatic
weeds. Kuffaker and Andres (78) have sta'ifed that any or¬
ganism which curtails plant growth or reproduction may be
used as a biological corHrol agent. Such could potentially
include animals either higher or lower than insects, and
parasitic higher plants, fungi, bacteria, and viruses. Fpf*:
this ajasen the term, biological ccntrojL ©rganism, or agent
is used to include all suitable phytophagous animals and
plant pafftoMftg Bn a given weed.
It was generally believed that biological control works
test with agents of foreign origin (75); however, as
Wapshere (188) noints out, successful bioccntrol with an
organism in one country does not necessarily imply tftStr the
organis^H) used will be successful elsewhere. For instance,
Chrysclina quadrigemina was relatively ineffective against
Hypericum, perfot genetic stock were highly successful against the same weed
ia California, apparently because of mere suitable climatic
conditions there (168).

6
Many investigations have been undertaken to study
potential uses of macrobiological agents to control noxious
aquatic weeds. In most instances, these studies have
involved insects (13,78,105,165,199) and, to a lesser
extent, other animals (33,39,118,162,164).
Of the insects screened for possible control agents,
one of the most effective appears to be the flea beetle,
Agasicles hygrophila which feeds only on alligatorweed (13,
105, 199). It was successfully introduced into the United
States from Argentina for the control of alligatorweed (13).
In April, 1965, 266 adult beetles were released near Jack¬
sonville, Florida, and by June, 1966, there were hundreds of
thousands of them present at the release sites and most of
the floating alligatorweed was dead (105). It has since
spread rapidly throughout the watersheds In northeast
Florida (199). Insects alone, however, are not likely to
control aquatic weed pests because there are relatively few
phytophagous species capable of living beneath the water
(201).
Other biological control agents being investigated
include phytopathogenic fungi, bacteria, and viruses.
Zettler and Freeman (201) list four advantages of using such
control agents: (i) control applications would presumably
require minimal technology and, if successfully established,
the pathogen in theory would be selfmaintaining; (ii) the

7
overwhelming number of different plant pathogenic species
from which to choose offers an unmatched versatility in
selecting a specific biological control; (iii) virtually
none can attack man or his animals, therefore providing an
important advantage over the use of various animals such as
snails, which may harbor vertebrate pathogens, and (iv)
plant pathogens, although often killing individuals in a
given population, would not be expected to cause the exter¬
mination of a species. This last attribute is important
because eracictaion of one aquatic weed species, such as the
waterhyacinth, may create an ecological void that in turn
may allow a population explosion of a different and more
serious species. In addition, Wilson (192) points out three
more advantages of using biological control agents over
chemical control procedures: (i) they can be specific to
the target weed which lessens the chance of damage to
cultivated or desired species, (ii) residue and toxicity
problems created by herbicides would be greatly reduced or
eliminated altogether, and (iii) there would be no accumu¬
lation of the herbicide in the soil or underground water.
In essence, then, the use of biocontrol agents has many
advantages over chemical control methods and warrants
continued research.
The use of plant pathogens is not without hazards. Any
study undertaken to introduce or test phytopathogens must

8
be done with extreme care. Well controlled and monitored
prerelease experiments, however, can greatly reduce any
potential dangers.
Part III. Pathogens of Waterhyacinth with Possible Biocontrol
Potential
The first recorded disease on waterhyacinth caused by a
fungus was reported in 1917 by Tharp (174). He described a
Cercospora sp. as occurring on Piaropus crassipes (= E.
crassipes) in Texas and subsequently identified the causal
agent as C. piaropi Tharp. Thirty-seven years later, in
1954, it was reported on waterhyacinth in India (175) and
was again reported from the United States in 1974 (53).
The disease symptoms are oval leaf spots, 1.5 - 4.0 mm
in size, on the distal portion of the leaf blade. As with
other leaf spot diseases reported on waterhyacinth (2, 154),
C. piaropi does not appear severe enough to retard the
prodigious growth of the plant significantly; however, its
host specificity enhances its potential as a biocontrol
agent and is being investigated further (53).
The second recorded disease on waterhyacinth was caused
by a rust fungus, Uredo eichhorniae, found in the Dominican
Republic in 1927 (27). A year later, Ciferri (26) reported
the occurence of a smut, Doassansia eichhorniae on E.
crassipes from the same area.
Neither of these organisms,

however, had been studied as potential biocontrol agents
until last year (25).
9
In 1932, a species of fusarium was reported on water-
hyacinth from India (2). It caused reddish-brown necrotic
spots and streaks on both sides of the petioles and the
infected plant parts gradually shriveled up. The disease
caused only slight injury and the plant rapidly regenerated
new leaves and petioles. This is possibly the first pub¬
lished paper concerned with phytopathogens as controls for
waterhyacinth as indicated by the authors' concluding state¬
ment :
The infection takes place readily, but
owing to the high resisting power of the
plant, the disease makes very slow pro¬
gress. From this it may be inferred
that this fungus cannot be regarded as a
possible remedy against the spread of
waterhyacinth (2).
Ten years later, Banerjee (7) identified the causal
agent as F. equiseti and Snyder and Hansen (169) reduced
this species to synonymy with F. róseme. A recent survey of
Florida for diseases of waterhyacinth resulted in the
isolation of this same species (F. roseum) from diseased
plants in Lake Griffin near Leesburg (154). This report was
the first of a F. roseum isolate affecting waterhyacinth in
the western hemisphere. The disease Is characterized by
chlorosis and vascular discoloration in advance of necrosis
which proceeds towards the leaf tip. The leaf spot, however,

10
did not expand over the entire leaf surface but remained
localized. This is in line with that described by Agharkar
and Bar.erjee in their original report (2).
In 1946, Padwick (133) reported two species of fungi
pathogenic to waterhyacinth. The first, Rhizoctonia solani
(Corticum solani), was isolated near Dacca, Bengal, from
infected leaves and petioles. It caused extensive blotching
and streaking, often killing individual plants. Some 20 years
later, R. solani was again reported on waterhyacinth from
India by Nag Raj and Ponnappa (124).
During surveys for phytopathogens in the Canal Zone of
Panama, Freeman and Zettler isolated a R. solani from the
anchoring hyacinth (E. azurea) which proved to be extremely
pathogenic on the floating hyacinth (56). In addition,
sclerotia of this fungus were able to maintain their viability
without loss of virulence after being submersed in lake water
for 26 months (56). Disease symptoms on E. crassipes were
severe blighting of the enersed portions of the plant which
frequently resulted in death of the entire plant. Although
R. solani is an agressive pathogen of waterhyacinth, it
cannot be considered as a biocontrol at this time, because
of Its wide pathogenicity to a number of economically
important hosts (133).
The second fungal species reported by Padwick (133) was
Cenhalosporium eichhornae Padwick sp. nov. It induced

11
large, oval, buff-colored spots on the leaves which were
covered with a white mat of mycelium. In 1973, Rintz (153)
reported another Cephalosporium species, C. zonatum, as
causing a zonal leaf spot disease of waterhyacinth in
Louisiana and Florida. His report was the fourth pathogen
described as occurring on waterhyacinth in the United
States. There was some discrepancy as to the synonomy of
these two Cephalosporium species (162) and the Commonwealth
Mycological Institute reduced them to synonomy, with C.
zonatum being the preferred name (123, 153). However,
several years later, C. zonatum was reclassJHfed and is
presently placed in the form genus Acremoniun of the class
Hyphomycetes (86). It is this fungus, Acremonium zonatum
(Sawada) Gams, which was studied as a biocontrol agent for
waterhyacinths in the present paper.
A concentrated research program on biological control
of aquatic weeds at the Indian. Station of the Commonwealth
of Biological Control in Bangalore has resulted in the
isolation of several species of phytopathogenic fungi. In
1965, Nag Raj (122) reported a thread blight of waterhya¬
cinth occurring in Calicut, India. Subsequent isolations
showed the fungus Marasmiellus inoderma. (Berk.) Sing, to be
the causal agent (122). The diseased plants in the field
exhibited necrotic areas on the leaves, petioles, and all
aerial carts. The infection was more evident In dense

12
stands of the weed and death of individual plants occurred
in irregular patches (122). Infection by M. inoderma under
laboratory conditions spreads very rapidly on host plants
which is a distinct advantage for a potential biocontrol
agent.
In 1970, Ponnappa (142), working at the same Indian
laboratory, isolated the fungus Myrothecium roridum from
waterhyacinth. Although this organism caused extensive
damage to E. crassipes, its usefulness as a biocontrol agent
cannot be considered at this time because of its patho¬
genicity on a number of important economic crops (142).
This fungus was also reported on waterhyacinth from India by
Charudattan in 1973 (21).
One fungus which appears to have good potential as a
control agent Is Aiternaria eichhorniae, isolated and
described by Nag Raj and Ponnappa (125). It was isolated in
India in 1970 and was proved the causal agent of a leaf
blight disease. Leaf spots frequently covered the majority
of the, leaf and caused premature death of those leaves. In
culture, A. eichhorniae produces a bright-red diffusable
pigment which deepens with age. In addition, it also
produces a host-specific toxic metabolite that causes
necrotic lesions when placed on leaves or petioles. The
host range of this fungus was tested on 42 genera of plants
in 15 families including aquatics and such important

13
terrestrial familes as Brassicaceae, Fabaceae, and Sola-
naceae. The results showed A. eichhorniae, to be non-
pathogenic on all plants tested except the waterhyacinth
(125). Its host specificity along with its specific toxic
metabolite enhances its potential as a biocontrol agent.
A similar species of this fungus was isolated in 1973
by McCorquodale , Martyn, and Sturrock (113) from water
hyacinth in south Florida and tentatively identified as A.
eichhorniae var. floradana (114). It resembled that de¬
scribed by Nag Raj and Ponnappa (125) in host specificity,
conidial size, and toxin production, but differed in pigment
production and gemmae formation. This is the first report
of this species in the United States.
Tests indicate that A. eichhorniae has good potential
as a biocontrol agent of waterhyacinth, but because it is
not indigenous to the United States it is under strict
quarantine by the U.S. Department of Agriculture. For this
reason, A. eichhorniae cannot be adequately field tested in
Florida at the present time.
A second Cercospora species, C. rodmanii was isolated
from diseased waterhyacinth in 1973 in Florida (55) and is
currently being evaluated as a biocontrol agent
(30). Symptoms of the disease on waterhyacinth include
general chlorosis of the plant, failure to produce off¬
shoots, spindly petioles ana a root rot. Field trials

14
indicated that the fungus greatly reduced the waterhyacinth
population in test plots, but did not eradicate it since new
growth appeared which continued to spread (30).
In summary, among the phytopathogens reported on
waterhyacinth, some are capable of inducing severe damage
and even death of the plant. The fact remains, however,
that there are relatively few capable of causing such severe
diseases. Most of those that do, however, are also patho¬
genic to important cash crops and therefore unacceptable as
biocontrol agents at the present time. Consequently, it
would be a great advantage if one or more of the pathogens
with a narrow or restricted host range could be utilized.
With this in mind, the intent of this study was to examine
the pathological relationship of E. crassipes and A. zonatum
in an effort to more fully understand the basis of disease
resistance and pathogenesis in this host-parasite couplet.

CHAPTER I
RESPONSES fF WATERKYACINTHS TO INFECTION WITH ACREMONIUM
ZONATUM AND ITS IMPLICATIONS IN BIOLOGICAL CONTROL
Introduction
Research into biological control of noxious aquatic
plants was Initiated at the University of Florida, Depart¬
ment of Plant Pathology, in 1970. Major emphasis was placed
on finding diseases of waterhyacinth, alligatorweed, hydril-
la, and Eurasian watermilfoil. Surveys for diseases of
these plants were made throughout Florida and portions of
Alabama, Maryland, Louisiana, Georgia, South Carolina, the
Chesapeake Bay, and the Tennessee Valley areas (55).
Surveys were also made in ten other countries including most
of the Caribbean and eight states in India (55). During
these surveys, several diseases were found and the causal
agents isolated for further study (22,23,24,83,95).
In 1971, a zonal leafspot of waterhyacinth was first
tinted in Puerto Rico where it caused considerable damage to
thA plant (55). The causal organism was not isolated.
However, a similar disorder was subsequently found in the
Spring Bayou region of Louisiana. A species of the fungus,
Cenhalosporium, was isolated from those plants, and upon
inoculation onto healthy plants, induced symptoms typical
15

16
of those observed under natural conditions. The causal
agent was ultimately identified as Cephalosporium zonatum
Sawada (153) and was originally described as the causal
agent of zonal leafspot disease of figs in Louisiana (177).
This disease was found since to occur on waterhyacinths in
El Salvador, India, Panama, and at two locations in Florida
(55). The causal agent, Cephalosporium zonatum, (Sawada)
recently was reclassified to Acremonium zonatum (Sawada)
Gams (86).
The disease is first evident as small sunken lesions on
both leaf surfaces and the petiole (153). Under conditions
of high hijfflidity, A. zonatum causes severe spotting and
death of leaves (107). The lesions are characteristically
zonate, oval to irregular in shape, and often coalesce
covering the entire surface. Alternating light- and dark-
brown bands are typical of the lesions. Under conditions of
prolonged high humidity the fungus produces abundant white
mycelia on the leaf surfaces and sporulates intermittently.
Rinflt (153) reported that A. zonatum can attack a wide
range of plants under artificial conditions. Despite this
apparent wide host range, reports of its occurrence on hosts
other than fig in North America are unknown. Consequently,
this fungus need not necessarily be excluded from consi¬
deration as a possible biocontrol agent of waterhyacinth
(55) .

17
During field trials with this fungus in Gainesville, it
was observed that small, young plants displayed fewer
lesions after infection than did larger plants in the same
plot. (T.E. Freeman, personal communication, 1974). In
addition, it was observed that some of the infected plants
appeared to produce more new leaf growth than did either
other diseased plants or control plants. The present study
was initiated to determine if small plants were more resis¬
tant to A. zonatum than large plants and also if there was
an accelerated leaf regeneration in response to infection.
Materials and Methods
Quantitation of disease
Waterhyacinths were collected from natural infestations
in south Florida and maintained under greenhouse conditions
in Gainesville. Plants were separated into three size cate¬
gories based upon leaf surface area: (i) small plants, with
2
leaves less than 15 cm , (ii) medium plants, with leaves 15-
2
40 cm , and (iii) large plants, with leaves greater than 40
2
cm . The plants were inoculated by swabbing the leaves with
a 10% (wt./vol.) slurry of A. zonatum (grown on potato
dextrose agar) and 0.75% water agar. Flants were maintained
in ten-gallon glass aquaria half-filled with tap water with
plastic covers to maintain the humidity at 99-100%. Control
plants were inoculated with sterile 0.75% water agar and

18
maintained under identical conditions. Two weeks post¬
inoculation, leaves were excised and used for subsequent
tests.
Tweaty-five to seventy-five leaves from each plant size
group were removed and the number of lesions/leaf counted.
Mean percentage figures were determined for Ci) number of
leaves with one or more lesions/leaf, (ii) number of leaves
with ten or more lesions/leaf, and (iii) mean number of
lesions/ leaf. The total diseased area on each leaf was
calculated by the dot counting method ("Stippentelplaatje" ,
<9-C. Zadoks, unpublished) and the mean percent diseased area
determined for each plant size.
Quantitation of leaf regeneration
Plants from each size category were selected at random,
trimmed of any necrotic or senescent leaves, and inoculated
as before. The total number of leaves on each plant was
noted prior to inoculation. The plants were maintained in
ten-gallon glass aquaria half-filled with tap water and
fitted with plastic covers as before. Control plants from
each size category -were painted with 0.75% sterile water
■«jpr and maintained under identical conditions. After two
weeks, the Btal number of leaves on each plant was counted
and percent new» leaf growth figure was calculated for each
plant size group.
In addition, ten plants from each size category were

19
selected and the number of leaves/plant noted. Each leaf
was then excised and the plants placed in aquaria. After
two weeks, a percent new leaf growth figure was determined
for each plant size.
Results
Quantitation of disease
Inoculated waterhyacinths kept under conditions of high
humidity were severely damaged by A. zonatum (Fig. 1-1).
Necrotic lesions varying in size and number occurred on both
leaf surfaces and the petioles. On the average, 70.1% of
the leaves on small plants had at least one lesion while
98.2% of medium leaves had at least one lesion (Fig. 1-2).
Large plants had 133% of their leaves infected indicating a
secondary spread to the new growth during the course of
disease development. These percentages decreased when the
number of leaves exhibiting ten or more lesions was calcu¬
lated but the trend was the same. Large plants had signifi¬
cantly more leaves with ten lesions (83.3%) than did either
medium (53.6%) or small (29.8%) plants. The average number
of lesions/leaf also followed the same pattern. Small
giants averaged 3.7 lesions/leaf while medium and large
averaged 12.8 and 18.3 respectively. However, when the
total diseased leaf area was measured after two weeks, there
was no significant difference among small, medium, and large

20
plants, each exhibiting approximately 40% diseased leaf area
(Fig. 1-2).
Quantitation of leaf regeneration
On the average, after two weeks of growth, small
healthy waterhyacinths regenerated 27.3% new leaves or one
new leaf/plant (Fig 1-3). There was no significant diffe¬
rence at the 0.01 confidence level in the percentage of new
leaves produced by small diseased plants. Small plants,
after infection, regenerated 21.6% new leaves or 0.95
leaves/plant. Likewise, there was no significant difference
between the new leaves produced by healthy and diseased
medium-sized plants. Medium-sized control plants produced
28.5% new leaves during the two weeks while diseased plants
of the same size regenerated 33.9% new leaves. A trend was
noted, however, that as the plant increased in size, its
rate of new leaf production also increased.
When the number of new leaves produced by large healthy
plants was compared to that from large diseased plants,
there was a significant difference. Large healthy plants
normally regenerated 46.1% of their leaves over the two week
period; however, diseased large plants produced 93.3% new
leaves, an increase of almost 50% (Fig. 1-3). In addition,
the average number of leaves/plant for large plants in¬
creased from 2.0 in healthy to 4.6 in diseased plants.
Small and large plants which had their leaves excised

21
prior to the test displayed little variation in the number
of new leaves when compared to the controls (Fig. 1-3).
Small plants regenerated 25.2% of their leaves compared to
27.3% new growth in normal small plants. Likewise, large
plants displayed little difference in new leaf production
between control plants and those in which the leaves were
excised (46.1% and 50.0% respectively). A notable exception
was observed with medium-sized plants. Controls produced
28.5% new leaves in two weeks while plants of the same size
whose leaves were removed first regenerated only 19.3% new
leaves.

Figure 1-1 (a - d). Symptoms of disease on water-
hyacinths incited by Acremonium zonatum.
a. Waterhyacinth with zonate lesions on leaves and
petioles.
b. Close-up of waterhyacinth plants two weeks post
inoculation.
c. Waterhyacinth leaf showing coalescence of leaf
spots.
d. Large waterhyacinth leaf with abundant white my
celia of A. zonatum.

23

Figure 1-2. Quantitation of disease on small, medium
and large waterhyacinths. Small = plants with leaves ^ 15
crn surface area; medium = plants wi^h leaves 15-40 cm
surface area; large = plants > 40 cm surface area. a= %
leaves with 1 or more lesions/leaf; b= % leaves with 10 or
more lesions/leaf; c= mean number of lesions/leaf; d= %
total diseased leaf area.

DISEASE
25
abed
SMALL
abed
MEDIUM
abed
LARGE
AVG. NO. LESIONS / LEAF

Figure 1-3. Quantitation of leaf regeneration rates
of small, medium, and large waterhyacinths. C- control
plants; 1= inoculated plants; E- plants with leaves excised
prior to test.

X
H
$
O
OH
C9
<
UJ
100
90
80
70
60
50
üj 40
° 30
20
10
0
C I E
SMALL
n\\\\\\N

28
Discussion
The concept of biological control, the use of one
organism to control another, although not new in practice is
relatively new in its wide-scale applications. Debach (40)
cites that the introduction of the mynah bird from India to
Mauritues in 1762 to control the red locust was the first
successful attempt at biological control. Perhaps the first
successful deliberate control of one organism with another
in the United States was the introduction of the vedalia
beetle into California in 1888 to control cottony-cushion
scale of citrus (6). Control of one organism by another has
been referred to as "parasitic control" and "the biological
method" but it wasn't until 1919 that H.S. Smith referred to
it as "biological control" (40).
It is a difficult task to impart a precise definition
to the term "biological control" since there is little
unanimity on this point among plant pathologist. Perhaps
the best definition is that given by Baker and Cook (6) . .
Biological control is the reduction of
inoculum density or disease-producing
activities of a pathogen or parasite in
its active or dormant state, by one or
more organisms, accomplished naturally
or through, manipulation of the environ¬
ment, host, or antagonist, or by mass
introduction of one or more antagonists.
The above definition encompasses several points not
dealt with in this dissertation. For convenience and ease
of understanding In the present discussion, the term "bio¬
logical control" will be used to imply the use of native or

29
introduced organisms to control or reduce the population of
another organism through an antagonistic or parasitic
relationship. Thus, the use of the fungus A. zonatum to
parasitize the waterhyacinth and thereby reduce its popula¬
tion size is well within the scope of the definition by
Baker and Cook (6).
Plant' pathologists are generally concerned with con¬
trolling epiphytotics--not starting them. However, this is
not the case when control of a noxious weed such as the
waterhyacinth is desired through biological methods.
Therefore, it takes some adjustment in one’s own thinking
when the initial idea is presented.
When an alien plant establishes itself in a particular
habitat it may mean several things: (i) it is better suited
to a particular niche than are the residents, (ii) that it
was introduced in such numbers as to temporarily or perma¬
nently "swamp" the residents, or, (iii) that it may modify
the environment in some way favorable to itself. It usually
means, however, that man has upset the natural balance in
some way, making the environment more favorable to the alien
than to the resident. Such has been the case with the
waterhyacinth. Over-nutrification of our waters by man’s
increasing agricultural and urban demands has been the
single most contributing factor to the aquatic weed problem.
Thus a weed is "a generally unwanted organism that thrives
in habitats disturbed by man" (6).

30
The first step in a biological control program is, in
most cases, the evaluation of the potential biocontrol
agent. This usually involves the introduction of the
control agent onto its target host and/or additional poten¬
tial hosts under greenhouse conditions. Thus the effec¬
tiveness of the control agent on its target host can be
determined as well as its potential to parasitize other
crops for which it was not intended. If a potential bio¬
control agent passes the initial greenhouse tests, field
trials are usually initiated. In these studies, evaluations
as to how the control agent manifests itself and its ability
to compete with the other biotic agents present can be
made. In some instances, it may be necessary to bring the
control agent back Into the laboratory and greenhouse to
further evaluate situations observed in the field.
The above description depicts the studies conducted on
A. zonatum over the past five years. Isolation and patho¬
genicity tests of the fungus under greenhouse conditions
were initially done by Rintz (152). Field trials with A.
zonatum were initiated in 1973 by Freeman, et al. (54) on
well established stands of waterhyacinths in Lake Alice on
the campus of the University of Florida. It was during
these studies that apparent differences in symptoms and
growth rates were noticed on the plants.
In 1974, greenhouse tests were Initiated once again to

31
See how host plant size influenced A. zcnatum as to infec¬
tion and subsequent disease.
Disease measurement is often regarded a^ a synonym for
"estimation of losses," but this is misleading (92). There
is a great need for some reasonably simple but critical
parameters that can be used consistently aftid systematically
to mesure the prevalence and severity of plant diseases in
the field. On the other there are no portmanteau
methods that will serve for all plant diseases. Some of the
currently accepted disease assessment techniques are dis¬
cussed bf Large (SR and include such things as standard
diagjpfcms, thJ Horsfall and Barratt gracing system, and
dífeease progress curves.
Perhaps one of the es/§isest (techniques to use is the
staisJSferd diRran method. This, of course, assumes that
standard diagrams for the particular host-para «fee couplet
in qesti.on have been constructed. If not, then this method
requires the researcher to Work out gjSch diagrams. In. the
case of E. crassipes - A. zonatum, stanBrc c';J.A#ase diagrams
have not been constructed. For this reasofe disease assess¬
ment was cased on two criteria: (i) number of lesijjr.s/leaf,
and Ui) total percent of diseased area/leaf. Beth of these
metliodsBiave Keen used routinely with other hcst-psrasbffe
combinations and are-’the basis of standard diagram keys.
Lesion counts on different size waterhyaciifths indi¬
cated initially that small plants were more resistant than

32
large plants since they exhibited fewer lesions/leaf.
However, when the mean percent diseased area of each leaf
was measured, there were no significant differences among
any of the three sizes, all showing approximately 40%
disease severity. This allows for two possibilities.
First, small plants are more resistant to initial attack,
but over the two week infection period gradually lose this
resistance and obtain a level of susceptibility shown by the
larger plants, or secondly, large plants are more suscep¬
tible initially but gradually build up a resistance. Based
upon data presented elsewhere in this dissertation, i.e.
polyphenoloxidase rates and phenolic acid concentrations
(see Chapter III), it is believed that a combination of both
mechanisms is involved. That is, small plants gradually
lose some of their initial resistance while larger plants
gain various degrees of resistance.
That plants may increase or decrease in susceptibility
to a particular pathogen with age is well documented (197).
It has been suggested (196) that susceptibility to faculta¬
tive saprophytes increases with age of host tissues, whereas
isceptibility to obligate parasites decreases with age
although this does not always hold true.
Based upon the results of the present study with water-
hyacinths, susceptibility to attack by A. zonatum increases
with plant size. Generally, plant size can be correlated

33
with ontogenetic development, that is, the older the plant,
the larger is its size. However, this may not always be a
correct assumption with waterhyacinths since growth rate
depends upon environmental conditions of its habitat (light
intensity, nutrients, and temperature). For this reason,
then, predisposition to A. zonatum in nature due to host age
may be only part of the answer. Differences in symptom
expression during field trials with this fungus may then be
the result of several predisposing factors operating in
conjunction with one another.
Water quality was not monitored during field trials
with this fungus so the effect of environmental predisposing
factors cannot be discussed. However, waterhyacinths used
in the greenhouse studies were all maintained under the same
environmental parameters. Since the only variable in these
tests was the age of the host , it can be stated that suscep¬
tibility of waterhyacinths to A. zonatum increases with the
ontogenetic development of the plant. This is an important
criterion when considering the use of any agent as a control
measure. Time of application is extremely important in
creer to obtain the most effective control.
Another very important observation made during these
studies was that of the leaf regeneration rates of different
plant sizes after infection. As healthy plants increase in
size (small to medium to large) their leaf regeneration rate

34
increases. That is, small plants regenerate approximately
21% of their leaves in two weeks or about one leaf/plant.
Medium-sized plants produced a slightly higher, but insig¬
nificant, percentage rate of 28.5 or 1.5 leaves/plant.
Large plants, however, are able to reproduce almost half of
their total leaves within a two week period (46.1%).
When plants are inoculated with A. zonatum, their leaf
regeneration rates are altered. There is a slight reduction
in new leaf production exhibited by infected small plants
(5.7%) and a slight increase shown by infected medium¬
sized plants (5.4%). But the significant difference is
demonstrated by infected large plants. With these there is
a two-fold increase in new leaves after two weeks. The rise
from 46.1% to 93.5% in large plants represents an increase
on the average from 2.0 new leaves/plant to 4.6 new leaves/
plant.
Because A. zonatum is a leafspotting pathogen, it was
postulated that accelerated leaf production was a response
to photosynthetic stress placed upon it by infection which
resulted in the destruction of most of its photosyntheti-
cally active tissue. In order to test this idea leaves were
excised from a set of each cf the three plant sizes and
monitored for new leaf growth. There was little variation
in the percentages of new leaf growth when compared to their
respective controls. In one case (medium-sized plants)

35
there appeared to be a deleterious effect on the plant's
normal leaf production rate. It would appear, then, that
the accelerated new leaf production observed in waterhya-
oinths after infection by A. zonatum is not a response to
the destruction of photosynthetically active tissue, but
one of interaction between the host and the pathogen.
Accelerated growth rates in diseased plants has often
been correlated with increased activity of growth regulators
(61). Normal growth in plants is under hormonal control by
such biologically active endogenous compounds as B-indole-
3-acetic acid (IAA, auxin), gibberellins, cytokinins, and
others (61). A departure from the normal levels of these
compounds in the plant, such as might be caused by
pathogenic attack, could alter the growth habit of the host.
Data presented in Chapter III show that A. zonatum is
capable of synthesizing high amounts of auxin in vitro when
given the amino acid tryptophan as a precursor. Even though
this does not represent conclusive evidence for the produc¬
tion of auxin in_ vivo by this organism, it does suggest its
possibility. In addition, it has been suggested (87,132)
that increased levels of auxin in diseased tissue may be
correlated with the inhibition of IAA oxidase in the plants
by phenolic inhibitors. Further implications on the pos¬
sible roles of auxin, IAA oxidase, and phenolic compounds
during pathogenesis are discussed in Chapter III.

36
In contemplating A. zonatum as a biocontrol agent of
waterhyacinths, several criteria must be considered. Fore¬
most is the proper time at which to apply the inoculum.
Results in this study have indicated that small, young
plants are more resistant to fungal attack than are larger,
older plants. Eased on this, the fungus should perhaps be
applied .late in the spring or summer when the plants have
reached maturity. On the other hand, data indicated that
the plants respond to infection by accelerating their rate
of leaf regeneration and that large plants do this more
quickly than do smaller ones. In essence, then, application
of the fungus to large plants would appear to negate or
minimize any control afforded by the pathogen. When, then
would be the best time to apply the control agent? Since
disease severity proceeds to approximately 40% within two
weeks, regardless of the plant size, application early in
the spring, as the new season's growth is beginning, would
appear to be the best time. In this manner one could avoid
the accelerated leaf growth response displayed by larger
plants while at the same time expect substantial damage to

CHAPTER II
A CYTOCHEMICAL AND ULTRASTRUCTURAL STUDY OF THE PHENOL
CELLS AND POLYPÍÍNOLOXIDASE ACTIVITIES IN HEALTHY AND
DISEASED WATERHYACINTH LEAVES
Introduction
Phenolic compounds are among the most widespread and
varied compounds in plants. Perhaps the best known role for
plant phenolics is their assimilation into the anthocyanins
and flavone pigments (150). However, as many authors have
indicated, phenolic compounds have nearly unlimited potential
in accounting for the many differences that occur in disease
resista^gis. (12,34,35,50,90,157,179).
Phenols are particularly abundant in the leaves of many
plants. They are also found in the xylem, phloem, and
periderm of stems and roots; in unripe fruits; in the testa
of seeds; and in pathological growths such as galls (49).
Phenolic compounds in plants may be present in individual
cells or in specialized idioblasts termed tannin sacs (49) or
phenol-storing cells (119). Recent studies have shown that
specialized phenol-storing cells occur randomly in several
plant species (10,11,100,107,119,120). Phenols may be a
common ingredient of the vacuoles or they may occur in the
cytoplasm proper in the form of small droplets which even¬
tually fuse (49).
37

38
In many plant tissues, phenols become oxidized to poly¬
meric dark red or brown compounds (phlobaphenes), which are
sometimes microscopically visible in the cell contents of
fresh sections. Oxidation of phenolic compound accounts for
the pathological darkening in plant tissues (38).
Histochemical detection of naturally occurring phenols
is difficult because few reagents that react with them to
form characteristic color compounds are adaptable to his¬
tological methods (148). In addition, the natural enzymatic
browning may not be sufficiently intense for easy detection
microscopically. In 1951, Reeve (148) described a histo¬
chemical test for phenols in fresh plant tissue. It is
based upon a colorimetric method for phenols using a nitrous
acid reaction. The method has become widely accepted and
used and is often referred to as the "nitroso reaction."
One of the enzymes associated with the oxidation of
phenolic compounds is polyphenoloxidase (PPO). The term
polyphenoloxidase has been used extensively In the litera¬
ture, although the names phenolase, phenoloxidase , catecho-
loxidase, and tyrosinase have been used as synonyms.
Classification of this enzyme is difficult because several
different activities have been described for it. The enzyme
' 'as originally termed tyrosinase since the aromatic amino
id, tyrosine, was the first experimental substrate (38).
â– ever, p-cresol and catechol have been most frequently

39
employed, as experimental substrates. Consequently, two
activities have been ascribed and have come to be known as
the "cresolase" activity when referring to monohydric phenol
oxidation and "catecholase" activity when referring to o-
dihydric phenol oxidation (38).
Many different phenolic compounds can serve as sub¬
strates for polyphenoloxidases. For sake of convenience
these enzymes have been divided into three main groups (155)
based upon their affinity for certain substrates, response
to inhibitors, and type of reaction catalyzed: (i) Tyro-
sinases - enzymes of this group catalyze both o-hydroxyla-
tion of monophenols and the oxidation of o-diphenols. (ii)
Grtho-diphenoloxidases - these enzymes, unlike the tyro-
sinases, are devoid of hydroxylation properties and act only
on o-diphenols. (iii) Para-diphenoloxidases - members of
this group act primarily on p-diphenols but may also ¡have
some affinity for the oxidation of certain o-diphenols. The
laceases can be classified in this category.
In the present discussion, the term polyphenoloxidase
has been retained whenever the oxidation activity is being
described regardless of whether it is acting upon an o- or
p-diphenol. For a detailed review of the polyphenoloxi¬
dases, the reader Is referred to Dawson and Kagie (38),
Nelson and Dawson (128), and Pat^l and Zucker (138).
Some cells are capable of converting tyrosine into a

40
brown or black pigment called melanin (48). The pathway for
this conversion is depicted in Figure II-l. The first step
involves an o-hydroxylation of tyrosine thereby forming
dihydroxyphenylalanine (DOPA). The enzyme that catalyzes
this conversion is in the tyrosinase group and consequently
can also oxidize DOPA in the second step to dopaquinone.
Polyphenoloxidases are devoid of any hydroxylation proper¬
ties and therefore cannot convert tyrosine to DOPA but are
capable of oxidizing it to dopaquinone. It is this property
which has been investigated as a marker for this enzyme in
vivo.
Polyphenoloxidase activity has long been thought to
reside within the chloroplasts of plant cells (5), but until
recently cytochemical localization had not been demonstrated.
Based on techniques developed by Novikoff et al. (129) and
Okun et al. (132) for the localization of tyrosinase in
animal tissues, Czaninski and Catesson (36,37) have recently
demonstrated the cytochemical localization of PPO in plant
cells. Since 1972, several investigators (72,74,107,134,135)
have shown that PPO activity is localized within the thyla-
koids of chloroplasts in several plant species.
This chapter presents the results of a histochemical
and ultrastrucTural study of the phenol cells in water-
hyacinth leaves and the cytochemical localization of PPO in
healthy and diseased plants.

pathway for conversion of
g (48)].
Figure II-l. Biosynthetic
tyrosine to melanin [after Eppi

HO
TYROSINE DIHYDROXYPHENYLALANINE DOPAQUINONE
(DOPA) |
*
INTERMEDIATES
Pathway for Melanin Synthesis
jr

43
Materials and Methods
Histochemical localization of phenols
Cross sections of fresh waterhyacipth leaf tissue (12—
24y) from small, medium, and large plants were made with a
Hooker plant microtome, tested for phenols by the nitroso
reaction (148), and observed with the light microscope.
With this method a nitroso derivative of the phenolic
compound is formed and after addition of the base, a bright-
red salt is formed.
Spatial distribution of phenol cells
The spatial distribution of the subepidermal phenol
cells from each size category was determined from tangential
sections made along the vascular bundles. Sections of the
leaves (10 x 15 mm) were taken from areas selected at random
and the epidermal surfaces separated from each other with a
razor blade. Each half was then stained for phenols as
previously described and observed with the light microscope.
• 2
The mean number of phenol cells/mm leaf tissue was calcu¬
lated for the top and bottom surfaces of each plant size
group.
Electron microscopy
Standard fixation and embedding procedures were used
throughout with slight modifications as presented below. A
flow diagram for the basic technique is presented in Figure
II-2. Fresh waterhyacinth leaf tissue was placed in a

Figure II-2. Flow diagram of procedure for standard
electron microscopy fixation and embedding.
12% glutaraldehyde - paraformaldehyde
n
"0.2 M sodium cacodylate, pH 7.2

fresh tissue
fix in Karnovsky's fixative1
(2hr- 22 C)
I ,
wash in buffer (4x)
post-fix in 1% 0s04
(! hr—22. C)
wash in buffer (4x)
I
dehydrate in 25%EtOH series
transfer to I007o acetone
embed in epoxy resin3
section
post-stain w/UrAc (lOmin.)
post-stain w/ PbCi (5min.)
Flow Diagram for Electron Microscopy
Fixation and Embedding

46
buffered (0.2 M sodium cacodylate, pH 7.2) solution of 2.0%
glutaraldehyde and 2.0% paraformaldehyde (85). Each leaf
was cut into 3-5 mm pieces and fixed for two hours at room
temperature. The material was washed in 50% buffer - 50%
distilled water solution for a minimum of 30 minutes before
being postfixed in 1.0% osmium tetraoxide for one hour at 22
C. Sections were then rinsed several times with the aqueous-
buffer mixture and passed through an ethanol graded dehydra¬
tion series at 25% increments and finally into 100% acetone.
After dehydration the sections were infiltrated with a
graded acetone-plastic series and embedded in a 100% low
viscosity epoxy resin (170). The embedded sections were
then placed under vacuum for five minutes to remove bubbles
and the resin was polymerized for 18 hours in a 60 C oven.
Thin sections were cut on a Sorvall MT-2 ultramicrotome with
a diamond knife and placed on single-hole, Formvar coated
grids. Sections were then poststained in 0.5% uranyl
acetate for ten minutes and in 1.0% lead citrate for five
minutes. The sections were examined with a Hitachi HU - HE
electron microscope.
Cy •t - .'chemical localization of polyphenoloxidase
The procedure for the localizar ion of PPO activity in
wuh rhyacinth leaves follows closely that described by
■- tLnski and Catesson (37). A flow diagram of this pro¬
cedure is presented in Figure II-3. Fresh leaf tissue, both

Figure II-3. Flow diagram of procedure for the
cytochemical localization of polyphenoloxidase.
"''redistilled glutaraldehyde
o
"0.2 M sodium cacodylate , pH 7.2
Q
0.02 M sodium diethyldithiocarbamate
4L-dihydroxyphenylalanine (50 ng/10 ml 0.067 M
phosphate buffer, pH 7.0)

fresh leaf sections
fix in 5% glut.1
I
wash in buffer(5x)
treat w/ DDC
wash in buffer (5x)
pre-incubate w/DOPA
incubate w/DOPA
(I hr. -37 C)
wash in d.w.-sucrose (5x)
post-fix w/ 2% 0s04
(2hr. _22C)
post-stain w/
PbCi
dehydrate in EtOH
I
embed in epoxy resin
I
j section 1
no post-stain
Flow Diagram for Cytochemical Localization of
PPO Activity

M 9
healthy and diseased, was placed in buffer as before and cut
into 2-4 mm pieces. The sections were fixed in 5.0% gluta-
raldehyde for 1 1/2 hours at room temparature and washed in
buffer 5 times for 15 minutes each. The sections were then
separated into three groups and treated by one of the
following methods: (i) boiled for ten minutes, Cii) incu¬
bated in 0.02 M DDC (sodium diethyldithiocarbanate) for 20
minutes at 22 C and then washed 5 times in buffer, and (iii)
no treatment. After their respective treatments, each group
was preincubated in a DOPA substrate solution (50 mg DOPA in
10 ml of 0.067 M phosphate buffer, pH 7.0, made up fresh) at
4 C overnight. After the preincubation period, the sections
were incubated in fresh DOPA for one hour (fresh solution
added after 30 minutes) at 37 C, followed by five washings
in distilled water made to 0.5 M with sucrose. After
postfixing in 1.0% osmium tetraoxide they were dehydrated,
embedded in epoxy resin, sectioned, and examined with the
electron microscope as previously described.
Results
Histochemical localization of phenols
When waterhyacinth leaves were stained for phenols by
the nitroso reaction, these compounds were found in large,
specialized idioblasts or phenol cells immediately beneath
both epidermal surfaces (Figs. II-4a £ b) and in cells

50
closely associated with the vascular bundles (Fig. II-4c).
The size of these cells in the palisade layer varied consi¬
derably, often exceeding several hundred microns in length
and extending down to the vascular elements. Those phenol
cells near the vascular tissue were much more isodiametric
and varied much less in size. There was no significant
difference in morphology of the cells among the three plant
sizes examined.
Spatial distribution of phenol cells
Phenol cells occurred randomly beneath both leaf
surfaces in all plant sizes and were found throughout the
entire leaf (Fig. II-4d). There were significantly more
2
phenol cells beneath the adaxial leaf surface (40.6/mm )
2
than on the abaxial surface (26.6/mm ) in small plants but
the reverse was true for medium and large plants (Fig. II-
5). Medium and large plants exhibited a more equal dis¬
tribution of phenol cells between the two surfaces but there
was a significantly greater number on the top (51.8/41.8 In
medium vs 54.2/48.7 in large). The total number of phenol
2
cells/mm , both adaxial and abaxial surfaces, significantly
increased as the leaf increased in area with a mean of
2 2 2
33.6/mm for small, 41.8/mm for medium, and 48.7/mm for
¡(ge,
U I trastructure of phenol cells
Electron micrographs indicate that in most cases the

51
subepidermal phenol cells were two to three times longer
than the adjacent palisade cells (Fig. II-6). The phenolic
compounds appeared in close association with the tonoplast
and as discrete bodies within the cells. These were ac¬
tively metabolizing cells containing nuclei, mitochondria,
and plastids. In contrast, the phenol cells near the level
of the vascular tissue were much more circular, had a
thicker wall, and the phenolic compounds were in amorphous
masses as opposed to discrete globules (Fig. II-7). There
were no morphological differences observed between phenol
cells of the same type in any of the plant sizes examined.
Cytochemical localization of polyphenolcxidase
The principle of the reaction for the cytochemical
localization of PPO activity involves obtaining an insol¬
uble, electron dense reaction product (dcpaquinone) from the
synthetic substrate at the point where enzyme activity is
proceeding (37). Although the reaction can be observed
without additional staining, the Intensity of the reaction
and the clarity of the surrounding material is enhanced by
poststaining with lead citrate. When examined by this
unique, a positive PPO reaction product was absent in all
eP| oplasts of small and large healthy waterhyacinth leaves
incubated without DOPA. Chloroplasts in palisade cells
(Fig . II-8a), have distinctly clear thylakoid spaces and
fret channels. Similar observations were made for

chloroplasts of bundle sheath cells (Fig. II-8b), vascular
parenchyma (Fig. II-8c), and phenol cells (Fig. ll-8d). The
thylakoids within the chloroplasts of phenol cells were not
readily detected until pcststained with lead citrate.
Sections from both small and large healthy leaves incu¬
bated with DOPA reacted in an identical manner for the
localization of PPO. Chloroplasts of the palisade cells
(Fig. II-9a) and spongy mesophyl cells (Fig. II-9b) did not
stain for PPO activity. On the other hand, PPO activity was
localized in the thylakoids of chloroplasts in three other
cell types, two of which were associated with the vascular
tissue. In each instance, the thylakoid spaces and fret
channels were the only areas stained for PPO activity.
In contrast to other cells, chloroplasts of the vascu¬
lar parenchyma, both phloem parenchyma (Fig. II-9d) and xylem
parenchyma (Fig. 11-10) were PPO positive. The chloroplasts
in these cells appeared black or electron-dense. These
electron-dense areas were restricted to the Thylakoids
within the chloroplasts (Fig. ll-10b). Chloroplasts which
were not poststained (Figs. II-9d £ II-10c) also showed a
positive reaction but the intensity and clarity was not as
good.
Another type cf cell having PPO positive chloroplasts
were the bundle sheath cells (Figs. II-Sc and II-lla £ b).
Waterhyacinths are typical monocots and have a large bundle

53
sheath surrounding the vascular elements. Chloroplasts in
these bundle sheath cells were PPO positive, although
perhaps not as intense as those in the vascular parenchyma.
The phenol cell itself also showed PPO activity (Figs.
II-9c and 11-12). The reaction in these cells was the most
intense of the three. In this cell type, the chloroplasts
are extremely electron-dense (Fig. II-12a), and examination
under higher magnification revealed that not only were the
thylakoids positive, but the entire organelle was electron-
dense (Fig. II-12b).
Leaf material that was boiled prior to incubation in
DOPA did not give a positive PPG reaction, in any chloro¬
plasts, indicating heat inactivation of the enzyme after
boiling (Fig. 11-13). The thylakoids became distorted after
boiling and starch granules swelled forming large lacunae
(Fig. I1-13a S c).
When the inhibitor, DDC, was added to the sections
prior to incubation in DOPA, no reaction product could be
detected in the thylakoids of any chloroplasts (Fig. 11-14).
When sections were poststained with lead citrate (Fig. II-
14a), the thylakoid spaces and fret channels contrasted
sharply with the stroma. Only the partitions were notably
electron-dense. Thus, the electron density of lead citrate
cannot be confused with the electron-dense product of a
positive PPO reaction. Consequently, use of the poststain

54
acts to heighten the observed reactions and surrounding
material. In addition, PPO activity was not observed in any
cell organelle other than chloroplasts. These observations
were consistent for each of the plant sizes examined.
VJhen diseased leaves were examined for enzyme localiza¬
tion, PPO activity was found to be no longer restricted to
vascular parenchyma, bundle sheath, and phenol cells rather
every chloroplast in every cell was positive. Palisade
cells were now positive (Fig. 11-15) and there was an
increase in the number of plastoglobuli in those chloro¬
plasts. Likewise, spongy mesophyl cells, which in healthy
cells were negative, became positive after infection (Fig.
11-16). These chloroplasts also showed an increase in the
number and size of the plastoglobuli.
The changes in PPO localization were apparent in chloro¬
plasts in cells immediately surrounding the lesions.
Sections taken several centimeters away from the lesion were
examined to determine if periphery cells also showed a
"turn-on" in enzyme activity. Electron micrographs indicate
that even those cells which are two to five centimeters
removed from the center of infection were also positive for
PPO activity. Thus, palisade cells became positive (Fig.
II-17a £ b), spongy mesophyl cells became positive (Fig. IT-
17 c £ d), and chloroplasts in cells normally positive such
as bundle sheath cells became very Intense (Fig. II-17d).

55
In essence, PPO activity was found in the chloroplasts in
only three cell types in healthy leaves: (i) vascular
parenchyma, (ii) bundle sheath, and (iii) phenol cells
proper. However, during disease, there was a turn-on of PPO
activity in all cells which contain chloroplasts. Whether
this turn-on in enzyme activity is host-induced or pathogen-
induced is not known at this time.

Figure II-4 (a - d), Light micrographs of phenol
cells in healthy waterhyacinth leaves.
a. Cross section of waterhyacinth leaf showing
arrangement of phenol cells in upper and
lower palisade cell layers. (375 X).
b. Cross section of waterhyacinth leaf showing phc
and vascular bundle (vb). (1,500 X).
c. Cross section of waterhyacinth leaf showing phc
in relation to vb $nd bundle sheath cells (bsc).
(1,500 X).
d. Tangential section of waterhyacinth leaf showing
spatial arrangement of phc. (375 X).

5?

2
Figure II-5. Number of phenol cells/mm leaf area
in small, medium, and large waterhyacinth leaves. ST= small
plants, top surface of leaf; SB- small plants, bottom surface
of leaf; MT= medium plants, top surface of leaf; MB- medium
plants, bottom surface of leaf; LT = large plants, top sur¬
face of leaf; LB=large plants, bottom surface of leaf;
mean number of phenol cells/mm leaf (both surfaces).

PLANT SIZE
NO. CELLS/mm2 LEAF TISSUE
o
oi -fc Ui (T>
O O O O
Ln
LO

Figure 11-6. Electron micrograph of phenol cell in
palisade cell layer of waterhyacinth leaf tissue. Phenol
bodies (pb) appear in close association with the plasmalemma
and as discrete globules within the tonoplast (t). Post-
stained with PbCi. (2,140 X).

6l

Figure
in vascular
bodies (pb)
x = xylem.
II-7. Electron micrograph of phenol cell
tissue area of waterhyacinth leaf. Phenol
appear as an amorphous mass within the cell.
Poststained with PbCi. (9,400 X).

X

Figure II-8 (a - d). Chloroplasts of healthy water-
hyacinth leaf tissue incubated without DOPA.
a. Palisade cell chloroplast with clear thylakoids
(th) . s= starch (29,400 X).
b. Bundle sheath cell chloroplast. cw= cell wall
th= thylakoids (37,500 X).
c. Vascular parenchyma cell chloroplast. th- thy¬
lakoids (30,000 X).
d. Phenol cell chloroplast. c= chloroplast, th=
thylakoids, pb = phenol body. Foststained with
PbCi (24,000 X).

65

Figure II-9 (a - e). Localization of polypheno-
loxidase in healthy waterhyacinth leaf tissue without
lead post staining.
a. Palisade cell chloroplast. Negative PPO activity
in thylakoids (th). s- starch (45,500 X).
b. Spongy mesophyll cell chlorcplast. Negative PPO
activity in thylakoids (th). (45,000 X).
c. Phenol cell chloroplast (phc). Positive PPO
activity in thylakoids (th). (33,000 X).
d. Vascular parenchyma cell chloroplast. Positive
PPO activity in thylakoids (th). pl= plasto-
globuli (57,500 X).
e. Bundle sheath cell chloroplast. Positive PPO
activity in thylakoids (th). (75,000 X).

('1

Figure 11-10 (a - c). Localization of polyphenol-
oxidase in chloroplasts of xylem parenchyma cells in
healthy waterhyacinth leaves.
a. Cross section of leaf showing a xylem element (x)
and surrounding xylem parenchyma cells (xp).
Chloroplasts (c) in the xp cells are positive for
PPO activity. Poststained with PbCi. (4,800 X).
b. Close-up of chloroplasts in xp showing positive
PPO activity between the thylakoids (th) and
several plastoglobuli (pi). cw= cell wall. Post-
stained with PbCi. (26,000 X).
c. Chloroplast in xp cell showing positive FPO acti¬
vity without PbCi poststaining. (16,500 X).

69
/>>'A

Figure 11-11 (a - c). Localization of polyphenol-
oxidase in chloroplasts of bundle sheath cells in healthy
waterhyacinth leaves.
a. Chloroplasts (c) in bundle sheath cells (bsc)
showing positive PPO activity. Poststained with
PbCi. (6,200 X).
b. Close-up of chloroplast in bsc showing positive
PPO activity in thylakoids. Poststained with
PbCi. cw= cell wall. (32,000 X).
c. Chloroplasts in bsc incubated in diethyldithio-
carbamate (DDC) prior to incubation in DOPA.
Thylakoids (th) are negative for PPO activity.
Poststained with PbCi. pm= olasmalemma.
(40,000 X).

71
cw

Figure 11-12 (a - b). Localization of polyphenol-
oxidase in chloroplasts of phenol cells in healthy water-
hyacinth leaves.
a. Ultrastructure of phenol cell in palisade cell
layer showing nucleus (n), mitochondrion (m),
chloroplasts Cc), and phenol bodies (pb).
Chloroplasts are positive for PPO activity. Post-
stained with PbCi. (2,200 X).
b. Enlargement of chloroplast in phenol cell showing PPO
positive thylakoids (th) and large phenol body (pb)
in association with the chloroplast. s= starch.
Poststained with PbCi. (57,500 X).

73

Figure 11-13 (a - d). Chloroplasts cf boiled, healthy
waterhyacinth leaf tissue incubated with DOPA.
a. Spongy mesophyll cell chlorcplast showing distended
thylakoids (th). cw- cell wall, sl= starch lacuna
(37,500 X).
b. Vascular parenchyma cell chloroplast showing thy¬
lakoids (th) negative for PPO activity. pl= plas-
toglobuli, m= mitochondrion (69,000 X).
c. Bundle sheath cell (bsc) chloroplast (c) with nega¬
tive PPO activity. mc= mesophyll cell. (7,000 X).
d. Enlargement of bsc chloroplast with negative PPO
activity. th= thylakoids, pl= plastoglobuli,
sl= starch lacuna, cw- cell wall (56,000 X).


Figure 11-14 (a - d). Chloroplasts of healthy water-
hyacinth leaf tissue incubated in inhibitor (DDC) and DOPA.
a. Palisade cell chloroplast with distinct thylakoid
spaces (th) and fret channels. m= mitochondrion;
Poststained with PbCi (40,000 X).
b. Vascular parenchyma cell chloroplast with negative
?P0 activity. th= thylakoids (28,000 X).
c. Bundle sheath cell chloroplast with negative PPO
activity. th= thylakoids, s= starch (46,000 X).
c. Phenol cell chloroplast with negative PPO activity,
th- thylakoids, pl= plastoglobuíi (55,000 X).

7?

Figure 11-15 (a - b). Localization of polyphenol-
oxidase in chloroplasts of palisade cells from diseased
waterhyacinth leaves.
a. Necrotic palisade cells (pc) showing positive PPO
activity in their chloroplasts and an increase in
the size and number of plastoglobuli. Chloro¬
plasts in palisade cells in healthy leaf tissue
are negative for PPC activity. Poststained with
PbCi. '(7,820 X).
b. Enlargement of -chloroplasts in palisade cells showing
PPO activity in the tnylakoids (th). Poststained
with PbCi. (24,300 X).

79

Figure 11-16 (a - c). Localization of polyphenol-
oxidase in chloroplasts of spongy mesophyll cells from
diseased waterhyacinth leaves.
a. Mesophyll cells (me) showing PPO positive chloro-
plasts. Hyphae (h) shown in upper right corner.
Foststained with PbCi. Chloroplasts in mesophyll
cells in healthy leaf tissue are negative for PPO
activity. (8,280 X).
b. Enlargement of mesophyll chloroplast showing positive
PPO reaction in thylakoids. m= mitochondrion. Post-
stained with PbCi. (35,400 X).
c. Enlargement of positive PPO chloroplast in mesophyll
cell without PbCi poststain. (40,000 X).

81

Figure 11-17 (a - d). Localization of polyphenoloxi-
dase in chioroplasts of cells several centimeters away from
infection center.
a. Palisade cells (pc) with positive PPO activity in
their chloroplasts. e- epidermis. Poststained
with PbCi. (6,200 X).
b. Enlargement of palisade chloroplast showing PPO acti¬
vity in the thylahoids (th). Poststained with
PbCi. (17,400 X).
c. Mesophyl1 cell (me) showing positive PPO activity
in the chloroplast. n= nucleus, m= mitochondrion,
th- thylahoids. Poststained with PbCi. (27,500 X).
d. Electron micrograph showing PPO positive chloroplasts
in mesophyll cell (me) and very intense reaction in
the bundle sheath cell (bsc) chloroplast. cw= cell
wall. Poststained with PbCi. (18,900 X).

83

84
Discussion
A wide variety of simple and complex compounds pos¬
sessing phenolic hydroxyl groups occur in plant tissues and
the importance of these compounds during the life cycle of
the plant has become increasingly evident (143). Plant
pathologists and physiologists have a keen interest in
phenolics as the "antiseptics" of the Plant Kingdom (143)
and many investigations have been made on disease resistance
and interaction of microorganisms with phenols.
As indicated previously, specialized cells containing
phenolic compounds have been reported in tissues from many
plant species. These cells are often called "tannin cells"
when the nature of the phenolic substances is not known, or
the substances have become decompartmented, oxidized, and
polymerized to varying degrees (120).
Common, nonspecific tests for tannins usually consist
of treatment with ferric chloride solutions followed by
treatment with dilute bases (148). A blue-green precipitate
is usually formed but not all phenolics give such a reaction
and the results may be influenced by other materials pre¬
sent. The Gibbs indophenol reaction (59) is a dependable
test for the detection of phenols (51), but appears to be of
little or no value in determining the number of hydroxyl
groups on the benzene ring (51,100). On the other hand, the
nitroso reaction (148) ¡¡forms a cherry-red nitroso derivative

85
of o-dihydric phenols and appears to be a generally reliable
means of differentiating these compounds from other phenols.
Work with a large number of phenols has shown that the color
of the nitroso derivatives other than those of o-dihydric
phenols are yellow, brown, red-brown, and green (100).
Results with waterhyacinths show that phenolic com¬
pounds are localized in phenol-storing cells in both the
palisade cell layer and in close association with the
vascular bundles in leaf tissue. After staining by the
nitroso reaction these cells appear bright red and, there¬
fore, probably contain o-dihydric phenols and their deri¬
vatives. Using this histochemical method in conjunction
with chromotographic and ultraviolet absorption data, Mace
(100) was able to identify 3-hydroxytyramine (dopamine) as
the major o-dihydric phenol in cells of banana roots.
Results obtained by similar methods are presented in Chapter-
Ill of this dissertation and show the presence of several o-
dihydric phenols in waterhyacinth phenol cells.
Several articles have appeared recently on the ultra¬
structure of phenol cells in plants (119,120). In each
case, the cells described, were in root tissue. The data
presented here are the first reported from an ultrastruc-
tural investigation of phenol cells as they occur in leaf
tissue. Two morphologically distinct phenol cells occur in
waterhyacinth leaves. Both contain nuclei, mitochondria,

86
and plastids, and presumably are actively metabolizing
cells. Phenol cells in the palisade cell, layer of waier-
hyacinth leaves are elongated and vary greatly in size,
often exceeding several hundred microns in length. In
contrast, those in the vascular region of the leaf are much
more isodiametric, vary much less in size, and the phenolic
compounds appear in an amorphous mass. The vascular phenol
cells closely resemble those in root tissue of cotton (120)
and banana (119).
Despite the widespread occurrence and the substantial
evidence for the importance of phenolic substances in plant
metabolism, the mechanics by which the phenolics are synthe
sized, stored, and released have not been established (120)
Mueller and Beckman (120) concluded that the origin and
formation of phenolic material could not be determined by
microscopic examination alone, although they suggested a
role for plastids in this process. Wardrop and.Cronshaw
(189) suggested that phenolic material is synthesized in
modified amyloplasts. The plastids disintegrate shortly
afterwards and the phenolic compounds aggregate in vacuoles
Studies with waterhyacinth support Wardrop £ Cronshaw's
hypothesis because (i) phenolic compounds are consistently
observed in close association with plastids in the phenol
cells; (ii) plastids in those cells tend to be smaller and
more compact than those in adjacent cell types Suggesting a

87
possible degradation process; and (iii) the turn-on of PPO
activity in plastids in additional cells after infection
with A. zonatum indicate a likely role for phenol oxidation
in response to pathogenic attack.
Results of this study also indicate that the phenolic
compounds are released from these cells after disruption of
the membranes as a result of damage inflicted by the patho¬
gen (see Chapter IV). Subsequent oxidation of these com¬
pounds by PPO to fungitoxic quiñones would likely occur in
chloroplasts of adjacent cells. Thus, she phenol cells
serve as a mechanisms of defense by acting as miniature
"time bombs" which are activated during pathogenesis.
The spatial distribution of foliar phenol cells in
waterhyacinths is random with the highest concentrations
occurring in the leaves of larger plants. They occur in
greater numbers on the top surface of small leaves but the
opposite is true for medium and large leaves. The reason
for this distribution is unknown and any explanation at this
point would only be speculation. Since waterhyacinths have
an almost equal distribution of stomata on both the adaxial
and abaxzal leaf surfaces (139), correlation of phenol cell
occurence with the presence or absence of stomata can be
ruled out.
Previous investigations (36,37,72,73,74,135) on the
cytochemical localization of foliar PPO have shown this

enzyme to be present in the thylakoids of all chloroplasts
examined. Henry (74) reiterated the possibility that PPO is
ubiquitously associated with certain phenolic compounds
which participate in electron transport. The present study,
however, describes for the first time a restricted localiza¬
tion of this enzyme in healthy leaf tissue and a subsequent
turn-on in other cells after infection. The presence of
this enzyme in particular cell types suggests a more speci¬
fic action than that implied by Henry. The PPO found in
chloroplasts of phenol cells is possibly responsible for the
oxidation of certain stored phenols. However, PPO observed
in vascular bundle sheath chloroplasts may be related to
electron transport, while that in vascular parenchyma cells
is possibly involved with lignin synthesis during secondary
wall thickening of the xylem elements.
The turn-on in PPO activity after infection with A.
zonatum in cells normally devoid of any such activity is
highly suggestive of an active role for this enzyme in the
disease reaction. Cook and Wilson (31) were the first to
suggest that the fungitoxicity of tannin was due to the
action of an oxidase which formed a germicidal fluid. Since
their work it} 1915, numerousqj-hvestigations have correlated
increased phenol-oxidizing enzyme activities with disease
resistance (9,68,101,102,121,144,146,149,181,182). Whether
the turn-on in PPO activity in waterhyacinths is pathogen-
induced or host-induced is not known at the present time

89
but it is probably a host-mediated response to the patho¬
gen 1 s at tack.

CHAPTER III
A BIOCHEMICAL STUDY OF THE PHENOLIC ACIDS AND
POLYPHENOLOXIDASE RATES IN HEALTHY AND DISEASED
WATEKHYACINTH LEAVES
Introduction
A potential biocontrol agent must meet several criteria
if it is to be successful. Perhaps the most important of
these is its specificity of the pathogen to a given host.
One of the factors which determines the host-parasite
specificity is the biochemical relationship between the two
organisms. Lewis (94) attempted to explain this biochemical
relationship on the basis of nutrition in his "Balanced
Nutrition Hypothesis of Parasitism." His assumption was
that in order for one organism to parasitize another, a
correct balance of nutrients must be present. Snell (168)
classified these •"nutriolytes" on the basis of whether they
were essential to growth of the parasite or stimulatory to
its growth. In essence, both Lewis and Snell postulated
that the outcome of a host-parsite relation was determined
by the interaction of water diffusable substances from both
organisms.
One of the major objections to this hypothesis is that
it does not take into account the effect of any inhibitory
substances which might be produced by the host either prior
90

91
to or in response to infection. In 1954, Garber (58) ex¬
panded the idea to include inhibitory substances as another
factor in pathogenesis. He states that there are two environ¬
ments which affect the outcome of a disease relationship,
the balanced nutrition environment and the inhibitory sub¬
stance environment. According to Garber, there are four
possible combinations of these two environments, only one of
which results in disease, i.e. a relationship where the
correct nutrients are supplied and no inhibitory compounds
are present.
In many instances, phenolic compounds are inhibitory to
the growth of microorganisms and it has been suggested that
they may be involved in disease resistance mechanisms. It
was therefore, the intent of this study, to examine the
phenol chemistry of waterhyacinths to determine what role,
if any, host phenolic compounds play in disease resistance
to Acremonium zonatumâ– 
Our knowledge of the phenolic compounds of plant origin
had its beginnings in industry. The earliest recognized
class of these compounds, the tannins, have been employed
since ancient times in the tanning of skins, the manufacture
of inks, and in the fining of wines (150).
Phenolics are a vast group of compounds, comprising the
anthocyanins (red and blue pigments), the flavones (yellow
pigments), the coumarins, tannins, lignins, and phenolic

92
acids and their esters. The term "tannin" has generally
been used to describe this wide array of organic compounds;
however, it is now restricted to those compounds which have
the specific property of tanning leathers. Recently, the
term "polyphenol" which implies the presence of more than
one hydroxyl group on a benzene ring has been used to in¬
clude the plant phenolic compounds (61).
The principal polyphenols are not present in a free
state in nature, but exist primarily in the form of esters
or glycosides (150). Hydrolysis of these compounds liberate
the aglycones or phenolic moiety from their respective gly-
cosidic or ester linkage. Extraction at low pH's reduces
them to their acidic state.
As a group, phenolic acids comprise the benzoic acids
(Cg-C-j) and the cinnamic acids (Cg-C,,). The structures of
the principal benzoic and cinnamic acids are given in Fig.
III-l. The benzoic acids are widely distributed both in
angiosperms and gymnosperms (69). For instance, Tomaszewski
[in Ribereau-Gayon (ISO)] has identified two benzoic acids,
p-hydroxybenzoic (p-HBA) and gentisic acids in the leaves of
97% of plants sampled from 86 families. Likewise, vanillic
and syringic acids are widespread as they are constituents
of lignin along with p-HBA. In general, plants which do not
contain lignin do not contain these acids either (69).
The pher.ylpropane skeleton (Cg-Cg) is unquestionably

Figure III-l. Principal phenolic acids found in plants
[After Ribereau - Gayón (150)].

BENZOIC ACIDS
CINNAMIC ACIDS
CHLOROGENIC ACID
HO-
COOH
R-R1- H; p-HYDROXYBENZOlC ACID
R-OH; R1-H: PROTOCATECHUIC ACID
R-OCHj -, R1" Hâ–  VANILLIC ACID
HO-
-CH-CH"COOH
R-R'-H: p-COUMARIC ACID
R-OH; R'- H: CAFFEIC ACID
R-OCH3 •, R'- H: FERULIC ACID
Principal Phenolic Acids in Plants
CD
-tr

95
the commonest and most important of the polyphenols (150).
This group includes the cinnamic acids and their deriva¬
tives; the aromatic amino acids, tyrosine, phenylalanine,
and DOPA; the constituents of essential oils; and the
lignins. Like the benzoic acids, the cinnamic acids are
widely distributed in plants. p-Coumaric acid is the most
common of all phenolic constiturents and is found in practi¬
cally all higher plants (150). Because cinnamic acids
possess a double bond, they are capable of existing in two
isomeric forms, i.e. cis- and trans-cinnamic acids. Diffe¬
rent biological properties have been assigned to the cis and
trans forms, however, the naturally occurring cinnamic acids
are the trans isomers, which are more stable (150).
The cinnamic acids, especially caffeic acid, have been
known for a long time. One of their properties is their
affinity to form esters with other phenolic acids. They
were first studied in coffee, which is particularly rich in
these compounds (140) and from which caffeic acid derives
its name. Perhaps the most well known and studied of these
esters is chlorogenic acid (3-caffeoylquinic acid) which was
discovered in coffee by Payen [in Ribereau-Gayon (150)] in
1846. It is an ester of caffeic acid with a cyclic acid-
alcohol, quinic acid. Kany ester-derivatives of the cinnamic
acids have been described (66,112,156,200).
The major pathway for the biosynthesis of monocyclic

96
phenols is the shikimic acid pathway (Fig. III-2) (126,127,
145,156). The elucidation of this pathway was was done
largely with mutant strains of bacteria using radioactive
tracers (145). The basic sequence of reactions is also
believed to occur in higher plants. The initial step invol¬
ves the condensation of the three-carbon compound, phosphcenol-
pyruvate, derived from glycolysis, with the four-carbon
compound, erythrose-4-phosphate, derived via the pentose-
phosphate pathway. The initial branch in phenol synthesis
yields the benzoic acids. The second branch provides for
the synthesis of the amino acid tryptophan which can undergo
conversion to indoleacetic acid (IAA, auxin) The third
branch provides for synthesis of the aromatic amino acids
phenylalanine, tyrosine, and DOPA, the later two involving
PPO. Deamination of phenylalanine by phenylalanineammonia-
lyase (PAL) yields the Cg-Cg skeleton, -trans-einnamic-acid
(32). The action of several enzymes on trans-cinnamic acids
yields a variety of hydroxylated and methoxylated cinnamic
acids (29) i.e. p-coumaric, caffeic, and ferulic. All of
these acids can undergo B-oxidation to yield their corresponding
benzoic acids (66), be incorporated into the B-ring of
flavonoids (61,66,172), undergo isomerization and intramo¬
lecular condensation to form coumarins and phytoalexins
(45,172), or undergo reduction to cinnamyl alcohols and
subsequent oxidative polymerization to lignin (61,126,200).

Figure III-2. Shikimic acid pathway for the b
thesis of monocyclic phenols and major derivatives,
from Neish (126,127) and Robinson (156).
Losyn-
Adapt

98
Carbohydrate Metabolism
phosphoenol- erythrose-4-
pyruvate (pep) phosphate
2-keto-3-deoxy-7-phosphogluco-
heptonic acid
5-dehydroquinic acid
©
5-dehydroshikimic
acid
vanillic acid
shikimic acid
pep -J
(2) chorismic acid—-anthranilic—- tryptophan—- IAA
acid
© prephenic acid tyrosine dopa
phenyla!
anine
@ cinnamic acid —- —- —► coumarins a phytoalexins
p-coumaric acid
(5) caffeic acid
quinic acid
1
chlorogenic acid
(§) ferulic acid —
—- isocoumarins
—» lignin
Shikimic Acid Pathway and Major Derivatives

99
The biosynthesis of polyphenols may also be accom¬
plished by the polyketide (150) or acetate pathway (61).
Compounds derived via this pathway are formed from the head-
to-tail condensation of acetate units. There is good reason
to believe that this pathway also operates in higher plants,
although to a lesser extent than the shikimic acid pathway,
and has a role in the biosynthesis of the A-ring of flavo-
noids (61) .
The importance of PPO to phenol synthesis lies in its
ability to oxidize monophenols to polyphenols (108).
Through these reactions, the flavonoids, tannins, lignins,
and melanins are formed, all of which have been implicated
In disease resistance (76,131,151). Numerous studies have
shown that host polyphenols and their oxidizing enzymes
typically increase in diseased plants (35,50,110). Their
role in pathogenesis, has been attributed to several mecha¬
nisms including inhibition of spore germination (91), anti¬
biosis (96,104,161), initiation of the hypersensitive re¬
sponse (98, 179), inhibition of pectolytic enzymes (14,28,42,
79), and the inhibition of indoleacetic acid oxidase (50,89).
JThis chapter presents the results of a study of the
benzoic and cinnamic acids present in healthy and diseased
waterhyacinths and the subsequent changes in PPO activities.
The effect of these compounds on the growth of A. zonatum,
as well as the fungus' ability to synthesize auxin in_ vitro
is also presented.

100
Materials and Methods
Assay for "free" phenolic acids
Extraction. Fifty g (fresh weight) of leaves from each
of small, medium, and large healthy, greenhouse-maintained
waterhyacinths were excised, washed in distilled water, and
macerated in 500 ml 95% ethanol in a Waring blender for five
minutes. The slurry was filtered through cheese cloth and
then through Whatman no. 1 filter paper in a Buchner funnel.
Total volume after filtration was 465 ml. Two hundred ml of
the filtrate were concentrated to approximately 5 ml under
vacuum at 40 C in a rotary evaporator. The concentrate was
made to 10 ml with 95% ethanol and centrifuged for three
minutes at 2500 rpm. The clear supernatnant was stored at
-20 C unless used immediately. Identical extraction proce¬
dures were used with leaves from small, medium, and large
diseased plants.
Thin layer chromatography. Twenty yl (5 pi at a time)
of each supernatnant were spotted onto silioa-gel, thin-
layer, chromatography sheets (TLC) with fluorescent indica¬
tor added (20 x 20 cm, Kodak) and developed ascending in n-
butanol: acetic acid: water (BuAW) (40:10:20 v/v, organic
phase). TLC sheets spotted with 10 yil of nine different
phenolic acid standards (5 mg/ml in 95% ethanol) were
developed simutaneously and used as markers for identifi¬
cation of unknowns. The chromatograms were dried at 100 C

101
and phenolic acids located under ultraviolet light (253 run).
Fluorescent spots were noted as to color and marked for
later identification. Phenolic acids on replica chroma¬
tograms were located by uniformly spraying with one of the
following reagents (167): (i) sulphanilic acid, (ii) p-
nitroaniline, and (iii) p-nitroaniline oversprayed with 2N
NaOH. The values and colors obtained with the different
locating reagents were compared to standards.
Assay for "ester-linked" phenolic acids
Extraction. The extraction procedure used for water-
hyacinth is an adaptation of the methods described by Isamil
and BrowS (80) and Woodward (195). A flow diagram of the
basic procedure is shown in Fig. III-3. Fifty g (fresh
weight) of leaves from each plant morphotype were washed and
macerated as before. The ethanolic extract was then boiled
for 30 minutes and filtered. The emerald-green filtrate was
reduced to dryness in a rotary evaporator under vacuum at 40
C, and then was redissolved in 50 ml hot, distilled water.
After cooling to room temperature, the extract was extracted
3 times in 50 ml petroleum ether to remove chlorophyll.
The golden-yellow aqueous phase was hydrolyzed by adjusting
to 2N with NaOH pellets (approximately 4 g) and boiling for
three minutes. The resulting dark-brown hydrolyzate was
placed immediately into an ice-bath and adjusted to pH 1.0
with concentrated HC1. The acidified extracts were then

Figure III-3. Flow diagram of procedure for extrac¬
tion of ester-linked phenols in plants. Adapted from Isamil
and Brown (81) and Woodward (199).
Step 1.
Step 2.
Step 3.
Step 4 .
Disruption from entact tissue
Extraction of chlorophyll
Hydrolysis of ester linkage
Purification and concentration of phenolic
acids.

103
©
50 q fresh tissue
t
macerate in hot EtOH
i .
boil for.30 mins.
filter
residue
discard
filtrate
reduce to dryness
resuspend in hot d. w.
wash w/ pet. ether
aqueous phase
I
adjust to 2N w/ NaOH
I
boil for 3 mins
acidify to pH 1.0 w/ HCI
extract w/ diethyl ether (3x)
"organic phase
discard
aqueous phase
discard
^organic phase
I
reduce to dryness
resuspend in 5ml. EtOH
use for TLC
@
Flow Diagram for Extraction of Phenols

104
extracted 3 times with 30 ml diethyl ether (washings com¬
bined) and 0.5 ml distilled water added. They were reduced
to dryness as before, resuspended in 5.0 ml 95% ethanol and
stored at -20 C unless used immediately. Identical pro¬
cedures were used for the extraction of phenolic acids from
diseased plants.
Thin layer chromatographyâ–  Twenty pi (5 pi at a time)
of each extract were spotted onto TLC sheets and developed
two-dimensionally; ascending in benzene:acetic acid:water
(BzAW) (10:7:3 v/v, organic phase) and after drying and
rotating the sheets 90°, ascending in 2% formic acid (FA).
Chromatograms of the nine phenolic acid standards were run
simultaneously. Phenolic acids were located with ultravio¬
let light and by spraying as previously described.
±
values were calculated and compared to standards.
Assay for total phenols
The concentration of total phenols in healthy and
diseased waterhyacinth morphotypes was determined by the
Folin-Denis colorimetric method (150). A portion (0.1 ml)
of the hydrolyzed extract was diluted to 5.0 ml in photome¬
trically-matched cuvettes by the addition of 4.0 ml dis-
ilied water, 0.1 ml Folin-Denis reagent, and 0.8 ml satu¬
rated NajCOg. The contents of the cuvettes were mixed for
five seconds on a Vortex mixer and the optical density of
each was recored at 740 nm on a Bausch and Lomb Spectronic

105
20. The concentration of each was determined as phenol
equivalents (phe) by extrapolation from a standard curve
prepared from nine different phenolic acids (trans-cinnamic,
o-coumaric, p-coumaric, caffeic, ferulic , vanillic, proto-
catechuic, p-hydroxybenzoic, and chlorogenic).
Pclyphenoloxidase assay
Fifteen g (fresh weight) of healthy or diseased water-
hyacinth leaves from each size category were excised and
placed immediately in the cold (4 C). Each group was mace¬
rated in a previously chilled Waring blender with 100 ml of
cold 0.01 M phosphate buffer, pK 7.0 for five minutes. The
leaf slurry was centrifuged in a precooled enclosed Sorvall
superspeed centrifuge at 10,000 rpm for five minutes to
remove excess leaf debris. The supernatant fraction was
decanted and separated into two equal portions (approxi¬
mately 30 ml each). One portion was kept in an ice bath
while the other was boiled for one minute to serve as the
inactivated enzyme control. Two drops of each portion was
added to separate micro-cuvettes containing 1.0 ml of 0.001
M L-dihydroxyphenylalanine (L-DOPA) made up fresh in 0.01 M
phosphate buffer, pH 7.0. After diluting 1:1 and mixing,
the cuvettes were immediately placed into a Beckman Model 25
recording spectrophotometer. The optical densities at 246
nm were recored at 15 second intervals for the first minute,
30 second intervals for the next four minutes, and one

106
minute intervals thereafter for ten minutes. The boiled
fraction containing DOPA was used to zero the instrument and
for the reference sample. Polyphenoloxidase activity rates
of each sample were calculated as the change in O.D./time.
Growth of A. zonatum on waterhyacinth-extract media
Waterhyacinth-extract media were prepared from healthy
and diseased plants from each size category in the following
manner: Ten g (fresh weight) of leaves were macerated in
100 ml hot, 95% ethanol and filtered. The filtrate was
reduced to dryness under vacuum and the residue redissolved
in 15 ml boiling distilled water. The redissolved residue was
extracted three times with petroleum ether and the aqueous
phase was filter-sterilized through 0.45 y Millipore fil¬
ters. One ml of each filtrate Was added to separate tubes
containing 9.0 ml Czapek-dox agar (45 C) (Difco) with and
without yeast-extract added (0.5%). The media were poured
into sterile petri dishes, allowed to solidify, and seeded
with a 7 mar. plug of A. zonatum. Control plates contained
1.0 ml distilled water in place of the plant extract. Five
replicas of each were prepared and incubated at 22 C for two
weeks. After the incubation period, the mean colony dia¬
meter of each was determined.
Growth of A., zonatum oh phenolic acid media
Czapek-dox agar, with and without yeast extract, was
made to final concentrations of 10, 100, and 1000 ppm of

107
seven different phenolic acids (chlorogenic, p-coumaric,
var.illie, ferulic, caffeic, protocatechuic, and p-hydroxy-
benzoic). Ten ml of each medium was poured into sterile
petri dishes, allowed to solidify, and seeded with a 7 mm
plug of A. zonatum. Five replicas of"each were prepared and
incubated at 22 C. The mean colony diameter was determined
after two weeks.
Indoleacetic acid assay
Colorimetric detection. The ability of A. zonatum to
synthesize indole-3-acetic acid (IAA, auxin) in_ vitro was
determined using the colorimetric technique described by
Gordon and Weber (62). Cultures were grown in 30 ml of
modified Czapek's liquid medium at 22 C (193) with either
NaNOg or tryptophan as the sole nitrogen source. At two-day
intervals, two flasks of each culture medium were removed,
filter-sterilized through 0.45 y Millipore filters and
assayed for auxin content in a Bausch and Lomb Spectronic 20
at 530 nm. Auxin concentration (ppm) was determined by
extrapolation from a standard curve using purified IAA.
Chromatographic detection. Synthesis of IAA from
tryptophan medium was confirmed by thin layer chromato¬
graphy. Cellulose TLC sheets with fluorescent indicator (20
x 20 cm, Kodak) were spotted with 10 yl of filtrate from
ten-cay-old cultures grown with and without tryptophan. Ten
yl of a 100 ppm IAA standard was also spotted onto the TLC

108
sheets as a comparison. Ascending chromatograms were
developed in 70% ethanol for four hours, air-dried, and
examined under a Wood's mineral light (253 nm). Fluorescent
spots were noted as to color, marked for identification, and
the chromatograms then uniformly sprayed with ferric chlo¬
ride-perchloric acid reagent (163). Spots which were ash
colored under UV light and which subsequently yielded a pink
color after spraying were assumed to be IAA. Rf values were
calculated and compared to the standards.
Results
"Free" phenolic acids
There were no differences in the "free" phenolic acids
found among any of the plant sizes, either healthy or dis¬
eased (Table III-l). In each case, a spot which was indis¬
tinguishable from chlorogenic acid, was the only phenolic
acid detected on the chromatograms. Mean values, fluores¬
cence under UV light, and color after spraying were all
similar to authentic samples of chlorogenic acid.
Ester-linked phenolic acids
Healthy waterhyacinth leaves. Three benzoic acids,
(protocatechuic, p-hydroxybenzoic, and vanillic), and three
cinnamic acids (ferulic, caffeic, and p-coumaric) were
detected in all three sizes of healthy waterhyacinth leaves
(Table III-2). Three additional acids (spots #7,8, £ 9)

109
were observed on chromatograms from large healthy leaves
which were not seen in small or medium plants. The identity
of these additional phenolic acids was not determined. Mean
Rj values and color characteristics for each of these are
listed in Table III-4. p-Coumaric acid (spot #6) was pre¬
sent in very small amounts in all plant sizes tested. It
could not be detected under UV light in any of the chromato¬
grams but was detected using either p-nitroaniline or sulphani-
lic acid reagent. Spot size and color intensity suggested
that caffeic acid (spot #1) was in the greatest concentra¬
tion in all samples.
Infected waterhyacinth leaves. The same six phenolic
acids that occurred in healthy leaves were also detected in
leaves infected with A. zonatum (Table III-3). p-Coumaric
acid was again in the smallest amounts while caffeic acid
was the mosT concentrated. The only qualitative change in
phenolic acids noted was the appearance of an unknown (spot
#7) in small- and medium-sized leaves after infection which
was not observed in healthy plants of the same size. This
unknown phenolic acid, along with the other two (spots #8 £
9), were also present in infected large leaves. Mean R-
values and color characteristics suggest that these
unknown acids are the same ones which are present in large
healthy leaves.
The mean values in each solvent and the color charac¬
teristics of each of the phenolic acids detected in healthy

110
and diseased waterhyacinth leaves are listed in Table III-4.
In each case, spots #1-6 were indistinguishable from the
authentic reference samples and consequently considered to
be the same.
Total Phenols
In healthy waterhyacinths, medium and large plants had
a significantly greater concentration of total phenols in
their leaves than did small plants (Fig. III-4). Data show
that leaves from small plants contained 92 yg phe/g fresh
weight while medium and large leaves contained 105 yg and
104 yg respectively. There was no significant difference
between the total phenols in medium and large healthy
leaves.
After infection with A. zonatum, the total phenol
concentration dropped significantly in both small- and
medium-sized plants, but remained constant in large plants.
Medium-sized diseased plants retained a significantly greater
concentration over small diseased plants (96 yg/g vs 80
yg/g) but was not significantly different from the concen¬
tration in large diseased plants (96 yg/g vs 105 yg/g).
Pholyphenoloxidase assay
In healthy plants, small leaves had a much greater PPO
activity than large leaves. Data show that the activity of
the enzyme in large leaves was 325% greater in small leaves
than in large leaves (Fig. III-5). Medium-sized leaves

Ill
exhibited a PPO activity between that of small and large
leaves. After infection with A. zonatum, the PPO rate in
small leaves decreased almost 40% (Fig. III-6). In large
leaves, the rate increased 300% to near the level found in
healthy small plants. Medium-sized infected leaves showed
little change in PPO activity.
Growth of A. zonatum on waterhyacinth-extract media
When A. zonatum was cultured on minimal media con¬
taining extracts from either small, medium, or large,
healthy waterhyacinth leaves it grew significantly better
than it did on minimal media without plant extracts (Table
III-5). When yeast extract was added to the medium as a
growth supplement, the fungus again grew significantly
better than it did on medium without healthy plant extracts.
Fungal growth on media containing extracts from diseased
leaves were similar (Table III-5). Growth was increased
on all media containing yeast and diseased-leaf extracts,
whether from small, medium, or large plants. However, growth
on the media which did not contain the yeast supplement
increased significantly only on plates containing extracts
from large diseased plants.
Comparisons of fungal growth rates between extracts
fren healthy and diseased plants of each morphotype revealed
no significant differences among any of the media which did
not contain yeast extract (Table III-5). However, each of

112
the cultures on diseased leaf-extracts grew significantly
better on media with yeast than their respective healthy
plant-extract counterparts.
Growth of A. zonatum on phenolic acid media
When A. zonatum was tested for its ability to grow on
media containing phenolic acids, it grew either as well or
better on each phenolic acid and concentration level tested
than it did on the controls (Table III-6). p-Coumaric acid
was the only phenolic compound tested which did not signifi¬
cantly increase growth of the fungus. In all other in¬
stances, the higher concentrations of phenolic acids (100 or
1000 ppm) Significantly stimulated the growth of A. zonatum.
In only one case did the lower concentrations increase
growth and that was with p-hydroxybenzoic acid at 10 ppm.
When yeast extract was added to the phenolic acid media
fungal growth was similar to that on phenolic acid media
without yeast extract with only a few noteable exceptions
(Table III-7). Most cultures grew better on the phenolic
acid media than they did on control plates, however, neither
ferulic or chlorogenic acids stimulated the growth of A.
zonatum. In addition, p-coumaric acid at 1000 ppm signifi¬
cantly decreased its growth compared to the controls. This
was the only acid tested which was inhibitory to fungal
growth in vitro.

113
Indoleacetic acid assay
Colorimetric detection. Cultures of A. zonatum grown
in Czapek broth with NaNO^ as the sole nitrogen source did
not synthesize auxin. When NaNO^ was replaced with trypto¬
phan as the sole nitrogen source, auxin was detected in the
filtrate after two days of growth (Fig. III-7). The auxin
concentration increased in the filtrates reaching a maximum
of 25 ppm after ten days and then decreased slightly after
14 days.
Chromatographic detection. Specificity of the Gordon-
Weber test for auxin, was verified by TLC, using filtrate
samples from ten-day-old cultures. Results showed two dis¬
tinct spots'from the filtrates containing tryptophan. Spot
#1, which was not identified, had an value of 0.71 and
turned brown after spraying with ferric chloride-perchloric
acid. A similar spot was observed from the filtrates con¬
taining NaiJO^ (R^ - 0.73). Spot #2 was ash colored under UV
light and turned pink when sprayed with the locating
reagent. It had an value of 0.87. The IAA standard
displayed identical colors under UV and after spraying as
spot #2, had a similar value (0.91), and was otherwise
indistinguishable from the authentic sample. It was con¬
cluded from this that spot #2 was most probably IAA. A
corresponding spot was not observed from filtrates con¬
taining NaN0~.

TABLE 111-1. Free phenolic acids detected in healthy and
A. ZONATUM - INFECTED WATERHYACINTHS BY THIN LAYER CHROMATOGRAPHY
Chlorogenic
ACID SOURCE
RF valuea
Color with locating reagents
(BuAW)
UVB
p-NAc p-NA/NaOHd
SAe
Standard
0.47
BLUE
GRAY-GREEN
yellow-brown
OLIVE-GREEN
Healthy water-
hyacinth leaves
0.45
BLUE
gray-green
yellow-brown
OLIVE-GREEN
Diseased water-
hyacinth LEAVES
0,43
BLUE
GRAY-GREEN
yellow-brown
OLIVE-GREEN
AN-BUTANOL:ACETIC AC ID¡WATER (40:10:20 v/v, ORGANIC PHASE)
BULTRAVIOLET LIGHT (253 NM)
CDIAZOTI ZED P-NITROANILINE
DP-NA OVERSPRAYED WITH 2N NaOH
EDIAZOTIZED SULPHANILIC ACID
114

TABLE 111-2, Phenolic acids detected in healthy waterhyacinth
LEAVES BY THIN LAYER CHROMATOGRAPHY AND VARIOUS LOCATING REAGENTS
AFTER ALKALINE HYDROLYSIS,
Plant Size
Phenolic
ACIDSA DETECTED WITH VARIOUS
REAGENTS
UVB
p-NAc
p-NA/NaOHd
SAe
Small
1,2,3,4,5
1,2,3,4,5,6
1,2,3,4,5,6
1,2,3,4,5,6
Medium
1,2^3,4,5
1,2,3,4,5,6
1,2,3,4,5,6
1,2,3,4,5,6
Large
1,2,3,4,5,
1,2,3,4,5,6,
1,2,3,4,5,6,
1,2,3,4,5,6,
and 8
7,8
7,8,9
7,8
Al= CAFFE ICj
2= PROTOCATECHUICj
3= P-HYDROXYBENZOIC! 4= VANILLIC!
5= FERULIC; 6= P-COUMARICJ 7= UNKNOWNJ 8= UNKNOWN! 9= UNKNOWN
bultraviolet light (253 nm)
CDIAZOTIZED P-NITROANILINE
°P-NA OVERSPRAYED WITH 2N NaOH
ediazotized SULPHANILIC acid
115

TABLE 111-3, Phenolic acids detected in A, zonatum - infected
WATERHYACINTH LEAVES BY THIN LAYER CHROMATOGRAPHY AND VARIOUS
LOCATING REAGENTS AFTER ALKALINE HYDROLYSIS
Plant Size
Phenolic
ACIDSA DETECTED WITH VARIOUS REAGENTS
UVB
p-NAc
p-NA/NaOHd
SAe
Small
1,2,3,A,5
1,2,3,A,5,6
1,2,3,A,5,6,7
1,2,3,A,5,6
Medium
1,2,3,A,5
1,2,3,A,5,6
1,2,3,A,5,6,7
1,2,3,A,5,6
Large
1,2,3,A,5,
1,2,3,A,5,6,
1,2,3,A,5,6,7
1,2,3,A,5,6,
AND 8
7,8,9
8,9
7,8
Al= CAFFEIC; 2= PROTOCATECHU IC; 3= P-HYDROXYBENZOIC; 4= VANILLIC;
5= FERULICj 6= P-COUMARIC; 7= UNKNOWN; 8= UNKNOWN; 9= UNKNOWN
bultraviolet light (253 nm)
CDIAZOTIZED P-NITROANILINE
dp-NA oversprayed with 2N NaOH
ediazotized sulphanilic acid

TABLE 111-A, RF values and color characteristics of the phenolic acids
DETECTED IN HEALTHY AND A, 70NATUM-INFECTED WATERHYACI NTH LEAVES AFTER
ALKALINE HYDROLYSIS
Spot
no,
Phenolic
acid
rf
VALUE
Color
CHARACTERISTICS
BzAWa
FAB
uvc
p-NAd
p-NA/Na0He
SAF
1
Caffeic
0,13
0,35
BLUE
OLIVE-BROWN
olive-brown
OLIVE-BROWN
2
Protocatechuic
0,11
0,59
PURPLE
YELLOW-BROWN
YELLOW-BROWN
YELLOW-BROWN
3
p-HBAg
0.46
0,60
PURPLE
PINK
PINK
YELLOW
A
Vanillic
0,67
0.5A
PURPLE
red-purple
PURPLE
GOLDEN-BROWN
5
Ferulic
0,65
0.37
BLUE
PINK
BLUE-GRAY
DEEP PINK
6
p-Coumaric
0.A7
0.A2
PURPLE
red-brown
PURPLE
GOLDEN-ORANGE
7
Unknown
0,59
0.69
NVH
PURPLE
PURPLE
YELLOW
8
Unknown
0,70
0.57
BLUE
PURPLE
PURPLE
YELLOW
9
Unknown
0.A1
0.73
NV
NV
PINK
NV
ABENZENE:ACETIC ACID:WATER (10:7:3 v/v, ORGANIC PHASE)j B2% FORMIC ACID
CULTRAVIOLET LIGHT (253 NM)j DDIAZOTIZED P"NITROANI LINEJ EP~NA OVERSPRAYED WITH 2N NaOHj
FDIAZOTIZED SULPHANILIC ACIDJ GP-HYDROXYBENZOIC ACIDJ HNOT VISIBLE

Figure III-4. Total phenol concentrations in healthy
and A. zonatum-infected waterhyacinth morphotypes.

3ZIS INVld
ai hi aw hw as hs
o
01
02
os
ot>
05
09
01
08
06
001
Oil
021
611
TOTAL PHENOLS (pg/g)

Figure III-5. Polyphenoloxidase activities in small,
medium, and large healthy waterhyacinth leaves. PPO rate
calculated as the change in optical density at 246 nm/time.

OPTICAL DENSITY (246 nm)
TIME (mins.)
121

Figure III-6. Polyphenoloxicase activities in small,
medium, and large diseased waterhyacinth leaves. PPO rate
calculated as the change in optical density at 246 nm/time

OPTICAL DENSITY (246 nm)
123

TABLE 111-5, Growth of A, zonatum on healthy and A. zonatum
INFECTED WATERHYACINTH LEAF-EXTRACT MEDIA
Plant
SIZE
Colony diameter (mm)
Without Yeast
With Yeast
Healthy
Diseased
Healthy
Diseased
Control-'-
25.6
25.6
30.2
30.2
Small
29,6a
28.2
34.2a
36.8ab
Medium
29,4a
27.2
32.8a
35.2ab
Large
30,4a
31.2a
33.4a
36.0AB
â– kzAPEK's AGAR
AC0L0NY DIAMETER SIGNIFICANTLY LARGER THAN CONTROL AT P= 0,05 LEVEL AS
DETERMINED BY ÃœUNCAN S MULTIPLE RANGE TEST.
BC0L0NY DIAMETER SIGNIFICANTLY LARGER THAN HELATHY COUNTERPART AT P= 0.05
LEVEL AS DETERMINED BY DUNCAN'S MULTIPLE RANGE TEST.
124

TABLE II1-6, Growth of A. zonatum on phenolic acid media
MFAN COI ONY niAMFTFR (MM) OF A. ZONATUM GROWN ON
CZAPEK'S AGAR WITH PHENOLIC ACIDS ADDED.
CONC.(PPM)
p-HBA1
PROTOCAT,2
VANILLIC
FERULIC
p-COUMARIC
CAFFEIC
CHLOROGENIC
0
25,2
25,2
25,2
25,2
25,2
25.2
25.2
10
27,6a
26,2
25,8
27,4
25,0
25.2
24,8
100
28.0a
27, A
27.8a
28,2a
26.0
26,4
29.6a
1000
28,6a
29.2A
30.0A
27,8a
25.2
31.2A
26.0
1p-HBA: p-HYDROXYBENZOIC ACID
2PR0T0CAT.: PROTOCATECHU IC ACID
ACoLONY DIAMETER SIGNIFICANTLY LARGER THAN CONTROL AT P= 0.05 LEVEL AS DETERMINED BY DUNCAN'S
MULTIPLE RANGE TEST,
r-O
cn

TABLE 111-7, Growth of A. zonatum on phenolic acid
MEDIA WITH YEAST EXTRACT
MEAN COLONY DIAMETER (MM) OF A, 70NAT1IM GROWN
ON CZAPEK'S AGAR W/YEAST EXTRACT AND PHENOLIC ACIDS ADDED
CONC, (PPM)
p-HBA1
PROTOCAT.2
VANILLIC
FERULIC
p-COUMARIC
CAFFEIC
CHL0R0GENIC
0
30,A
30,4
30,4
30,4
30,4
30.4
30.4
10
34,2a
30,4
32,8
31,2
32.6
33,0A
32,2
100
32.4
33.0a
31,4
32.6
34,4a
34.6a
32.6
1000
33.0a
32.0
35.4a
32.0
27,6b
32,4
32,2
1p-HBA: p-HYDROXYBENZOIC ACID
2PROTOCAT.: PROTOCATECHU IC ACID
ACOLONY DIAMETER SIGNIFICANTLY LARGER THAN CONTROL AT P= 0,05 LEVEL AS DETERMINED BY DUNCAN'S
MULTIPLE RANGE TEST,
BCOLONY DIAMETER SIGNIFICANTLY SMALLER THAN CONTROL AT P= 0,05 LEVEL AS DETERMINED BY DUNCAN'S
MULTIPLE RANGE TEST,
ro
cn

Figure III-7 .
from tryptophan by
In vitro synthesis
Acremonium zonatum•
of indoleacetic acid


129
Discussion
Varietal resistance of plants to pathogens has been
i: : led to polyphenol content on numerous occasions. The
oV/.sic works of Walker (183,184) and Walker and Link (185)
were perhaps the first positive evidence for a role of
phenols in disease resistance. Working with the onion
pathogen, Colletotrichum circinans, they demonstrated that
the presence of certain phenolic compounds, mainly proto-
catechuic acid and catechol, in the scales of red onions
imparted resistance to the smudge pathogen. Since then
numerous investigators have correlated host phenolic com¬
pounds to disease resistance (12,35,50,145,159,179,186).
Condensed and hydrolyzable tannins have been implicated
in resistance of cotton, strawberry, and apricot to Verticil-
lium wilt and the resistance of chestnut to Endothia para¬
sitica (12). Other compounds such as 3,4-dihydroxypheny-
lalanine (DOPA) and benzoic acid have been implicated in the
resistance of banana to Fusarium wilt (102) and apples to
Nectria rot (17) respectively. Chlorogenic acid content has
been correlated with resistance to such disease as coffee
canker (^4), potato scab (82), and Verticillium wilt of
potatoes (93).
The idea that phenolic compounds may be involved in
disease resistance stems from the many articles which appear
in the literature demonstrating a biostatic or biocidal

130
effect on microorganisms. Microorganisms, however, very
considerably in their sensitivity to different phenolic
acids. Bell (12) states that the order of activity of
fungal pathogens to polyphenols is obligate parasites >
facultative saprophytes > facultative parasites > saprophytes.
Thus certain pathogens such as the mildew and rust fungi are
frequently inhibited by polyphenols while many saprophytes
are stimulated by moderate concentrations of polyphenols as
â–  carbon source.
The toxicity of phenols is generally believed to be due
to their oxidation products, the quiñones, brought about
either by enzymatic action or autooxidation (12). Thus,
Schaal and Johnson (161) demonstrated that chlorogenic acid,
caffeic acid, catechol, and tetrahydroxybenzoin were toxic
to Streptomyces scabies in vitro. p-Hydroxybenzoic acid was
not as an effective inhibitor as were the other four. They
concluded that the inhibition effect of these phenolic
compounds was due to autooxidation to their respective
quinor.es, and although p-hydroxybenzoic acid also autooxi-
dizes, it does so much more slowly since it is a monophenol
whereas the others are polyphenols.
The effectiveness of polyphenols in disease resistance
depends on many factors (12): (i) the quantity and type of
polyphenols present in healthy tissue, (ii) the speed and
quantity of polyphenol synthesis induced by infection, (iii)

131
the quantity and type of oxidases present in healthy tissue,
Civ) the speed and quantity of oxidase synthesis induced by
infection, (v) location of the polyphenols and oxidases in
the host, (vi) the sensitivity of the pathogen to these
compounds, and (vii) the cellular environment in which these
occur. The studies presented in this chapter were designed
to evaluate most of these criteria as they relate to water-
hyacinth .
The only phenolic acid detected in the free state
was chlorogenic, which is an ester of caffeic and quinic
acids. Alkaline hydrolysis liberates the phenols which are
bound in plants as esters while acid hydrolysis, on the
other hand, liberates those phenols which are bound as
glycosides (152). The six phenolic acids identified in
waterhyacinth after alkaline hydrolysis and the three uniden¬
tified ones, are therefore most probably present in the
plant as esters, either with quinic acid (152) or with
sugars (70,71). Liberation of the aglycones in vivo from
their respective linkages, either by host-mediated or
pathogen-mediated reactions could then make them available
to the host during pathogenic attack and possibly serve in
defensive reactions. The mechanism by which this is accom¬
plished, has not been fully investigated.
It has been demonstrated that polyphenols increase,
either qualitatively or quantitatively, or both, in many
plants following infection (159). The decrease in

132
concentration of total phenols in small- and medium-sized
plants after infection was therefore, unexpected. However,
this decrease in total phenols may be indicative of a
decrease in resistance of small plants.
Results similar to those obtained with waterhyacinths
were obtained by McLean et al. (115) working with potatoes
and Verticillium wilt. They observed that wilt developed
more rapidly and more severly in susceptible varieties,
coincident with or following the decrease in phenol com¬
pounds in the vascular system. Patil et al. (136,137) also
showed that young potato roots which are partically resis¬
tant to infection by Verticillium have a relatively high
level of phenols until five weeks after sprouting. From the
time of sprouting, the chlorogenic acid content decreased
continuously and was correlated with an increase in suscep¬
tibility to infection.
The decrease in total phenols in small diseased water-
hyacinths also correlates with the decrease in PPO activity
after infection. It would be logical to assume that reduced
levels of substrate would lead to reduced levels of enzyme
activity. In large waterhyacinths, however, the total
phenol content did not decrease after infection, but re¬
mained at the level found in healthy plants. In this case,
increased resistance can best be correlated with the greatly
accelerated PPO rate. The concentrations of polyphenols in

133
large healthy plants is significantly higher than that in
small healthy plants, but the PPO rate is much lower. This
would account for a reduced rate of enzymatic oxidation.
After infection, however, PPO rates increase, thereby in¬
creasing the oxidation of phenols to the quiñones. In
contrast to small plants, large plants are. initally more
susceptible to fungal attack by virtue of their lower PPO
rate but gradually build up resistance as the PPO activity
increases over 300%. Thus, disease severity in each plant
morphotype balances at about 40% diseased leaf area, small
plants by an increasing susceptibility and large plants by
an increasing resistance.
On first inspection, the qualitative data on the pheno¬
lic acids present in small waterhyacinth might not appear to
coincide with the total phenol content. An additional
phenolic acid was detected on chromatograms from diseased
small plants which was not detected on chromatograms from
the same morphotypes of healthy plants. This could result
from two possibilities. First, the compound is in healthy
tissues but in concentrations too low to detect and after
infection, synthesis is increased, or second, it is net in
healthy tissues, but is synthesised de_ novo during disease.
In either case, Its concentration should increase, resulting
in an increase in total phenols. It is believed, however,
that even though this compound is increasing, the remaining

134
phenolic acids are decreasing, thereby reducing the total
phenolic content. Preliminary results with caffeic acid
support this hypothesis (R. Kartyn, unpublished). Caffeic
acid content, which is in the highest concentrations of all
the phenolic acids in waterhyacinth, decreased greatly in
small diseased plants from that present in healthy plants.
The higher content of total phenols in large healthy plants,
as opposed to small healthy plants, is most probably due to
increased concentrations of each acid plus the presence of
three additional ones not found in smaller plants.
One of the most striking results of this study was the
high tolerance of A. zonatum to phenolic acids, in vitro.
Acremonium zonatum is a facultative parasite, and according
to Bell's sensitivity ranking of pathogens, it should be
relatively tolerant of polyphenols. Results presented in
this chapter support this concept. Its tolerance was
evidenced by its ability to grow significantly better on
minimal media with various concentrations of phenolic acids
incorporated Into them than it did on minimal media alone.
This fact could explain why A. zonatum is able to cause rela¬
tively severe damage on waterhyacinth.
The above finding is in agreement with the results of
fungal growth on plant-extract media. Healthy waterhyacinths
of each morphotype have a relatively high phenol content in
their leaves and fungal growth was stimulated on media

135
containing such plant extracts. The significance of this can
be noted in the growth of A. zonatum on media containing
extracts from diseased plant morphotypes. An increase in
growth was only observed on media containing extracts from
large diseased leaves. Extracts from both small and medium
diseased leaves failed to stimulate fungal growth which may
be indicative of their reduced phenol content after infection.
Oddly, p-coumaric was the only phenolic acid which did
not stimulate fungal growth in vitro on minimal media. When
additional growth supplements were added to the media, p-
counaric was the only phenol found zo be inhibitory. This
suggests that in the absence of additional nutrients, A.
zonatum is capable of metabolizing several different pheno¬
lic acids as a carbon source. In the presence of an en¬
riched medium, however, these same phenolic acids lose some
of their simulatory effect, noticeably ferulic and chlcro-
genic acids, while one (p-coumaric) becomes inhibitory.
The reason for the inhibitory effects of p-coumaric
acid is not known but a recent study by Elstner and Heupel
(46) nay add some information as to its mode of action.
Working with isolated cell walls from horseradish, they
demonstrated that hydrogen peroxide production was inhibited
by dihydroxyphenols but stimulated by monohydroxyphenols such
as p-coumaric acid. If p-coumaric acid stimulated hydrogen
peroxide production in A. zonatum as it does in horseradish,

136
it may well account for the inhibitory effect. Studies on
the peroxidase system of this fungus should be done
to test this hypothesis.
Data from electron microscopy studies during pathogensis
(see Chapter IV) revealed that the penetration of a phenol
cell by A. zonatum resulted in death of the invading hyphae.
It is presumed that either the concentrations of phenols
in those cells are such that they are no longer stimulatory
but toxic to the fungus or that some other factor of meta¬
bolism, such as PPO activity, is increased to the point where
the fungus can no longer tolerate it. Unfortunately, data
from this study do not permit a conclusion on either
possibility.
In the present discussion, it has been suggested that
host phenols play a major role in the defense against poten¬
tial pathogens of waterhyacinths by being biocidal or bio¬
static. This concept seems plausible when dealing with an
organism that is susceptible to the toxic properties of
phenols. However, this is not the case with A. zonatum, and
it appears that some waterhyacinth plants have an additional
defensive mechanism which is indirectly linked tc phenol
metabolism. It was observed that after infection, large
plants displayed almost a three-fold increase in new leaf
production over that of either small or medium plants.
Increased meristematic activity in plants Is not uncommon

137
during disease (61). It has been demonstrated many times
that auxin content markedly increases in diseased tissue
(16,173). The increased levels of auxin in diseased tissue
may result from either (i) increased synthesis of IAA by
either host or pathogen or (ii) a decreased rate of auxin
degradation. There is no direct evidence that reveals
whether the host plant or the pathogen is the actual source
of increased IAA levels in infected tissues (61), but a
number of studies indicate that pathogens are capable of
synthesizing auxin in_ vitro (103,106,171,193, 194). Acre-
monium zonatum was also able to synthesize high concentra¬
tions of IAA in vitro which is a possible source of extra
growth hormone necessary to promote accelerated rates of
leaf regeneration. If the fungal-IAA was responsible for
the accelerated leaf production, then higher growth
rates would be expected to occur In all infected plant
morphotypes but it did not; only large plants increased leaf
production.
A second possible means of increased levels of auxin in
the host suggested was a decrease in the rate of auxin
degradation. This may be correlated with an inhibition of
IAA oxidase activity in the plant. IAA oxidase is a peroxi¬
dase mediated system responsible for keeping a balanced
level of auxin in the plant and phenolic compounds are known
inhibitors of this system (61). In large diseased

138
waterhyacinths, phenolic compounds do not decrease as they
do in small and medium, but remain at their preinfection
level. It is highly possible that these phenols are inhi¬
biting the normal "checks and balances" system of auxin
regulation in large plants consequently allowing it to build
up to abnormal levels resulting in a faster growth.
The waterhyacinth's primary disease defense system
appears to lie in its unique phenol cell-PPO complex and
operates effectively against numerous potential pathogens.
However, this system breaks down somewhat when the plant is
attacked by an organism such as A. zonatum which can utilize
the plant's phenolics for its own growth. In such event,
the role of the phenols is to block auxin degradation al¬
lowing the plant to "outgrow" the infection.

CHAPTER IV
AN ULTRASTRUCTURAL STUDY OF PENETRATION AND COLONIZATION
OF WATERHYACINTH BY ACREMONIUM ZONATUM AND CYTOLOGICAL
CHANGES ASSOCIATED WITH INFECTION
Introduction
The process of pathogenesis can be viewed as a battle
between a plant and a pathogen refereed by the environment
(191). A small change in a single environmental factor,
such as temperature or moisture, often can mean the dif¬
ference between crop success and crop failure. When stu¬
dying plant diseases, one should always be conscious of the
climatic conditions under which the disease is evident.
In nature, the environmental variables are numerous,
and as Matta (109) has suggested, plant-pathogen inter¬
actions can sometimes be best studied in the laboratory
where some of the variables can be controlled.
It is axiomatic that before disease can ensue, a viru¬
lent pathogen must come into contact with a susceptible
host. Although some pathogens are brought to their hosts
through a vector relationship, most fungal pathogens make
contact with their hosts fortuitously in the form of wind¬
blown or water-borne spores. Merely establishing contact is
not enough in most cases to ensure a parasitic relationship.
139

140
Entry into the host by the parasite must usually be accom-
*
plished. The manner in which this is achieved has been
debated for many years. In general, there are two mecha¬
nisms through which a pathogen can enter its host: (i)
direct penetration through natural openings, wounds, or
through unbroken surfaces by mechanical pressure and (ii)
penetration facilitated by enzymatic degradation. Although
there is little argument against direct penetration through
openings in the plant surfaces, it is generally believed
that penetration through unbroken surfaces involve a combi¬
nation of both mechanisms (61). Pressure for direct pene¬
tration is presumably supplied by the appresorium of the
fungus which serves as an anchoring device and from which
the infection peg emerges. Cuticle and cell wall degrading
enzymes are secreted from the hyphal tip, facilitating
entrance of the fungus into the host.
After penetration fungal pathogens may spread from the
site of infection throughout the host. Only if the fungus
enters into a parasitic relationship with its host is colo¬
nization successful, and hence pathogenesis is initiated.
Unlike bacteria, most fungi invade their hosts Intracellu-
lary and obtain nutrients from those cells. This may be
accomplished by distinctive fungal structures termed hausto-
ria which penetrate the cell wall and absorb nutrients
through the host cells' plasmalemma, or by secretion of

141
toxins which may act on the cell membrane casing its dis¬
ruption and electrolyte leakage. Those fungi that do not
form haustoria or produce toxins may penetrate within the
host cell and gain nutrients directly from the hosts'
cytoplasm. This appears to be the case with A. zonatum. In
general, after penetration has occurred, the protoplasm of
the host cell becomes granular and the nucleus migrates
towards the Infecting hyphae. Later the nucleus increases
in size considerably (61). In advanced stages of disease
the nucleus, as well as other cell organelles, begins to
collapse ppd degenerate.
This chapter» presents the results of an ultrastructural
study of the penetration and colonization of waterhyacinth
leaves by A. zonatum. Special attention was given to the
possible method(s) by which the fungus was able to gain
entrance into its host cells and the cytological features
which changed as a result of the infection.
Materials and Methods
Electron qjfcroscopy
Leaves, displaying characteristic symptoms of
disease, from both SUall and large waterhyacinth plants were
excised and used for the ultrastructural study. The leaves
were placed in petri dishes fitted with filter paper and
moisten with approximately 5 ml of Karnovsky's fixative.

142
Sections (2-4 mm) were cut from three areas on each leaf:
(i) center of the lesion, (ii) periphary of the lesion and,
(iii) asymptomatic tissue several centimeters (2-5) from the
lesion. Each section was the! fixed, dehydrated, and em¬
bedded as previously described (see Chapter II). Thin
sections were cut using a diamond knife, placed on single
hole grids, poststained in uranyl acetate and lead citrate,
and observed on an Hitachi HU-11E electron microscope.
Production of pectinases.
The procedure used for the detection of extracellular
pectinases produced by A. zonatum follows closely that de¬
scribed by English (47). Five hundred grams of peeled sweet
potatoes were boiled in one liter of distilled water until
soft (approximately 20 minutes). The solution was filtered
through successive layers of cheese cloth and 100 ml of the
filtrate placed into each of ten 250 ml sterile Erlenmeyer
flasks. Each flask was seeded with a 7 mm plug of A.
zonatum and incubated for ten days at room temperature.
After incubation the media were filter-sterilized through
0.45 p Killipore filters and each filtrate divided Into two
fractions. One fraction was then boiled for ten minutes
while the other fraction was left unboiled. Fresh sweet
potato discs (0.5 x 7 mm) were placed Into sterile petri
dishes fitted with moisten filter paper and 1.0 ml of either
the boiled or unboiled filtrate pipetted onto each disc.

14 3
Five replicates of each plate were made. In addition, small
pieces of the mycelial growth were also placed onto addi¬
tional sweet potato discs. All plates were incubated at
room temperature and examined after 4, 8, 12, 16, and 24
hours for signs of tissue maceration. Cultures of Rhizopus
stolonifer, supplied by J.W. Kimbrough, were grown and
treated in an identical manner and were used alp positive
controls.
Production of cellulase
Carboxymethylcellulose (CMC) medium (63, and R.E.
Stall, personal communication) was prepared in the following
manner. The CMC gum (Hercúlea Chemical Co.) was autoclaved
for ten minutes at 15 psi in 80% ethanol and washed twice in
fresh 80% ethanol for 15 minutes each. The resin was dried
at room temperature overnight and the dried powder added to
hot distilled water to give a final concentration of 2.0%.
Bacto-agar (Difco) was added to a concentration of 0.5% and
yeast extract added to 0.3%. The entire mixture was then
autoclaved and poured into standard size (100 x 14 mm)
sterile petri dishes. The CMC agar was allowed to solidify
and seeded with a 7 mm plug of A. zonatum. The plates were
incubated Ipoom temperature and monitored daily for fungal
growth and liquefaction of the gel around the colonies (pit
formation).

144
Results
Production of pectinases and cellulases
Acremonium zonatum did not produce detectable amounts
of extracellular pectinases when tested with the potato
assay method (Fig. IV-1). The fungus grew well in the sweet
potato broth and on the sweet potato discs, but it failed to
cause any noticeable signs of tissue maceration. Similarly,
the unboiled filtrates failed to cause any detectable break¬
down of the discs. After 24 hours the potato discs became
dehydrated and were discarded. Fotato pieces treated with
R. stolonifer filtrate showed tissue breakdown after eight
hours which increased over the next 16 hours.
Electron micrographs of infected waterhyacinth leaves
revealed that A. zonatum penetrated the middle lamellae of
cells (Fig. IV-2c £ d) but did so without destroying the
integrity of the surrounding portions. If extracellular
pectinases were produced by A. zonatum as an aid d_n host¬
cell penetration, they were localized at the tip of the
invading hypha and hence were produced in concentrations too
low to be detected in_ vitro by the sweet potato disc mace¬
ration test.
On the contrary, A. zonatum produced detectable amounts
of an extracellular cellulase in_ vitro as evidenced by the
pit formation or liquefaction when grown on CMC medium (Fig.
IV-1). This is supported by electron micrographs which

revealed the hyphae within the cell walls and by large areas
of lysis and wall breakdown well away from the advancing
mycelia (Fig. IV-3b and c).
Ultrastructure of penetration and colonization
Penetration of waterhyacinth leaves by A. zonatum
occurred either through the stomata (Fig. IV-2a) or directly
through the cuticle (Fig. IV-2b). In most cases the hyphae
firmly cemented themselves to the leaf surface by secretion
of a mucilaginous substance (fig. IV-2a). In other cases,
however, there did not appear to be a cementing matrix (Fig.
IV-2b).
Acremonium zonatum did not appear to produce appreso-
ria. However, direct penetration of the cuticle by mecha¬
nical force did apparently occur, at least in part, as
evidenced by the inward displacement of the cuticle at the
of penetration (Fig. IV-2b). Similar results were
. Gained with Colletotrichum graminicola on maize (141).
Additional evidence for direct penetration by A.
zonatum was the formation of papillae (Fig. IV-2b). Papil¬
lae were seen to form immediately opposite the infection peg
in the epidermal cell wall. There is good evidence (3) that
papillae are formed in response to mechanical pressure
during penetration.
Penetration through either the epidernal walls (Fig.
IV-2b and c) or xylem walls (Fig. IV-2d) was apparently

146
accomplished with equal ease. There was no sign of wall
tearing at the penetration site or disturbance of adjacent
wall material which indicated that penetration was enhanced
by localized secretions of cellulases and other wall de¬
grading enzymes. The resolution of the micrographs did not
permit a clear interpretation as to whether a haustorium was
formed, or not since it was not possible to distinguish
between fungal cell wall and host-cell plasmalemma.
Figure IV-4a is a cross-section of a collection of
hyphal strands which pulled away from the epidermis but
illustrates the extent to which this fungus secretes a
mucilaginous matrix to cement itself to the host cell sur¬
face. Micrograph IV-4b also illustrates this matrix
ana perhaps the first stage of penetration through the
epidermis. In the area of immediate contact between the
host cell wall and the fungus there was an area of wall
material which appeared to be undergoing degradation. This
may be the point of localized enzyme secretion.
Penetration of the phenol cell by A. zonatum is shown
in Figs. IV-5 and IV-6. The hyphae were cemented to the
cell surface and penetration through the wall occurred as it
did in other cells. Once inside the phenol cell, however,
the hyphae appeared extremely vesiculated, granulated, and
distorted. It appeared that the hyphae were killed,
by the high concentrations of polyphenolic compounds within
the cell.

147
An additional observation made from electron micro¬
graphs of diseased waterhyacinths was the presence of hyphae
within the xylem (Fig. IV-3a). Disease symptoms incited by
zcnatum do not include those indicative of vascular
■ issue infection such as wilting and it was therefore unex¬
pected when hyphae were consistently observed within the
tracheary elements.
vtolcgical alterations induced by infection
One of the most striking changes in waterhyacinth cells
infected with A. zonatum was the disappearance of starch
granules in the chlcroplasts. The chlcroplasts in palisade
cells of large healthy waterhyacinth leaves had an abundance
of starch granules in them (Fig. IV-7a). After infection,
there was a noticeable absence of starch in the chloroplasts
(Fig. IV-7b ) .
A second noticeable change after infection was the
build up of plastoglobuli within the chloroplasts. Chloro¬
plasts of healthy waterhyacinth leaves consistently
had several plastoglobuli (Fig. IV-8a). After infec-
ton, however, the number and size of these plastoglobuli in¬
creased greatly (Fig IV-8b).
A third cytological change after infection was a
build up of microbodies within the cells. Healthy water¬
hyacinth leaf cells had only a few microbodies but after
infection they increased in number (Fig. IV-9a). These

148
generally appeared during the later stages of disease after
starch disappeared and the plastoglobuli started to in¬
crease .
Associated with the appearance of microbodies in the
cell, the chloroplasts began to distort and the thylakoids
started to distend (Fig. IV-9a). During the final stages of
disease, the chlorplasts, as well as other organelles, lost
their integrity completely, plastoglobuli fused forming
large, irregular complexes, and the cytoplasm took on a very
granular appearance (Fig. IV-9b). Eventually, the entire
cell became convoluted and filled with dark, electron-dense
material which was indicative of necrosis and cell death
(Fig. IV-10).

Figure IV-1.
degrading enzymes
Flow diagram for testing of carbohydrate
produced by Acremonium zonatum.

A. zonatum
PECTINASES
grow in sweet
potato broth
CELLULASES
filter
inoculate on
waterhyacinths
I
examine microscopically
tissue maceration
destruction of
middle lamellae
destruction of
cel
wall
negative
positive
negative
grow on CMC1
medium
observe for pits
positive
Test for Production of Carbohydrate Degrading Enzymes by
A. zonatum
150

Figure IV-2 (a - d). Penetration of waterhyacinth leaf
by Acremonium zonatum.
a. Hyphae (h) penetrating through an open stoma.
sc= substomatal cavity, gc= guard cell. Arrow
points to mucilaginous matrix secreted by the
fungus (6,200 X).
Hyphae (h) penetrating the cuticle (cu) and epi¬
dermal cell wall (cw). ip= infection peg, p=
papillae (14,400 X).
c Hyphae (h) penetrating the epidermis of a water-
hyacinth leaf. cw = cell wall (22,400 X).
d. H- :e penetrating the cell wall (cw) of a xylary
ele - '30,000 X) .

152
h Jr

Figure IV-3a. Cross-section of Acremonium zonatum
observed in xylem tissue of diseased waterhyacinth leaf.
h= hyphae, x- xylem. (5,2C0 X).
Figure IV-3 (b - c). Degradation of wall material in
•terhyacinth by Acremonium zonatum.
b. Growth of A. zonatum within the cell wall. h=
hyphae, cw= cell wall, ml= middle lamellae
(10,000 X).
o Growth of A. zonatum within the cell wall showing
large areas of lysis (arrows). Middle lamellae
(ml) remains intact (27,500 X).

154

Figure
the cuticle,
icuci] aginous
IV-4a. Attachment of Acremonium zonatum to
h= hyphae, E= epidermis. Arrow points to
cementing substance (9,200 X).
Figure IV-4b. Attachment of Acremonium zonatum to
eDidermis and the possible area of localized enzyme secretion.
h= hyphae, cw= cell wall. Fart of the cell wall immediately
beneath the hyphae is eroded and suggestive of enzymatic de¬
gradation (32, C'OO X).

156

Figure IV-5 Ca - c). Penetration of phenol cell
by Acrenoniun zonatun•
Hyphae on the external cell surface appear viable
while those inside the phenol cell are highly vesiculat
and appear nonviable. h=hyphae, phc= phenol cell, pb =
nhenol body, sp= septum. a= 6,200 X; b= 27,000 X;
c= 40J100 X.

158

Figure IV-6. Phenol cell invaded by Acremonium
zonatum. liyphae (h) appear granulated ar.d nonviable.
phc= phenol cell, pb= phenol body. C 9 ,200 X).

16o

Figure IV-7 (a - b). Breakdown of starch reserves in
chloroplasts during disease.
a. Chloroplasts in healthy palisade cell (pc) showing
an abundance of starch (s) (6,800 X).
b. Choroplasts in diseased palisade cell (pc) showing
the absence of starch. c= chloroplast, n= nucleus,
pl= plastoglobuli (6,200 X).


Figure IV-8 (a - b). Increase in the number of plasto-
globuli in chloroplasts during disease.
a. Chloroplast of a healthy cell depicting only a
few plastoglobuli (pi). (48,000 X).
b. Chloroplasts in an infected cell showing the in¬
crease ir. plastoglobuli (pi). m= mitochondrion.
Arrow points to area of thylalcoid disruption
(2.500 X).


Figure IV-9a. Increase in the number of microbodies
in cytosol as a result of infection with Acremonlum
zonatum. mb = microbody, m= mitochondria, th= thylakoids
(16,000 X).
Figure IV-9b. Destruction of chloroplast integrity
during later stages of disease. c= chloroplast, cw = cell
wall (18,000 X).

166
V

Figure IV-10. Diseased palisade cell showing
of necrosis and cellular breakdown. E- epidermis,
wall, h= hyphae. (9,200 X).
extent
cw= cell

168

169
Discussion
When a fungal cell contacts a susceptible host prior to
penetration, an amorphous, electron-dense layer is often
found between the walls of the host and the pathogen (15).
This material is probably a fungal secretion and is assumed
to aid adhesion of the pathogen to the host cell. Electron
micrographs of infected waterhyacinth leaf tissue indicate
that A. zonatum produces abundant secretions which serve as
a cementing matrix. It is quite common at almost any inter¬
face between the pathogen and host cell but is apparently in
greatest amounts on the surface of epidermal cells and
stomata. Since the outer surface of a leaf would be ex¬
pected to be subjected to greater physical stresses than the
internal surface, it is quite logical that a fungus would
require greater adhension forces externally than it would
internally. It is also probable that the mucilaginous
matrix serves as a buffer or insulator from the external
environment during the course of penetration, thereby
protecting the pathogen from dessication or other delete¬
rious conditions.
Entry of A. zonatum Into waterhyacinth leaves was
through the stomata or by direct penetration of the unbroken
cuticle. Under natural conditions it is presumed that entry
is by way of the stomata since conditions of high humidity
favor disease development and the substomatal cavities would

170
conceivably provide this needed moisture. Similar results
were obtained by Rintz (152). However, when the relative
humidity was maintained at 99-100% after inoculation, A.
zonatum was capable of penetrating the cuticle directly.
The method by which fungi penetrate their hosts has
been a topic of considerable interest. For many years,
direct penetration of plant cuticles and cell walls by
pathogens was thought to be primarily a mechanical process
(61). More recently, this idea has been modified to include
enzymatic activity in addition or in combination with mecha¬
nical force (191).
The idea that enzymes may be involved in plant tissue
breakdown during pathogenesis was initiated by DeBary in
1886 [see Bateman and Millar, (8)]. Working with the fungus
Sclerotinia libertiana, DeBary demonstrated that a thermo-
labile substance from the fungus brought about disorgani¬
zation of the host tissue. Brown (18) subsequently described
a similar substance from Botrytis cinereaâ–  Tissue macerating
enzymes have since been described from numerous micro-
â– cganisms. For a detailed discussion of these enzymes see
reviews by Brown (19), Albersheim et al. (4), and
&. 'Leman and Millar (8).
Pectic enzymes are implicated almost routinely as a
feature of host-pathogen interactions. Their involvement in
the degradation of pectic constitutents of cell walls and of
the middle lamella in plant tissues has been reported for

171
such diverse types of diseases as soft rots, dry rots,
wilts, blights, and leaf spots and for many types of patho¬
genic agents such as fungi, bacteria, and nematodes (8). It
is perhaps significant that many of the pathogenic fungi and
bacteria examined have been found capable of producing
pectic enzymes. Cn the other hand, that a pathogen has this
property does not explain why the organism is pathogenic
(8). Rather, pectic enzyme production is likely to be but
one of several properties of the pathogen, all of which are
acting in concert to determine the pathogenecity of the
organism.
In regard to A. zonatum, pectic enzymes were not de¬
tected in_ vitro; however, micrographs of this fungus
penetrating cell walls Indicated that they were perhaps
produced, at least in small concentrations, and most pro¬
bably were localized near the hyphal tip. Although some
fungi are capable of excerting enough force to cause loca¬
lized indentations in films of cured resins (117) there is
little evidence that such pressure alone is responsible for
penetration of plant cell walls. Mechanical pressure alone
.ould be expected to show evidence of tearing and distortion
of the cell wall around the penetration site and these signs
were not observed. Instead, penetration was accomplished
t: i'O'igh smooth-bordered holes in the cell wall which indi-
cal.^d a presoftening of the wall constitutents prior to

172
penetration. Mechanical pressure was involved to some
extent, however, during penetration of the cuticle. Elec¬
tron micrographs showed an inward bending of the cuticle
which can best be explained by the application of force.
In addition, there was no excessive damage to the
middle lamella, which is composed chiefly of pectin sub¬
stances, while extensive damage occurred to the more cellu-
losic walls. In this regard, A. zonatum did produce exten¬
sive amounts of a cellulolytic enzyme in vitro as evidenced
by its ability to cause pit formation on CMC medium. Like¬
wise, in_ vivo cellulolytic activity of the fungus was demon¬
strated by the extensive degradation and lysis of the cell
walls.
It is not possible from, this study to determine which
cellulolytic enzyme(s) is involved but Reese et al. (147)
have reported that the ability of microorganisms to develop
an enzyme capable of hydrolyzing the 6, 1-4 glucosidic
linkage found in cellulose and its derivatives is widespread
aim-ng microorganisms. The degradation of native cellulose,
hoe-ver, is less common. Thus, microorganisms which are
c.; 'd as non-cellulolytic may develop the C^ enzyme, the
activi.Ly of which is measured by its capacity to degrade
carboxymethylcellulose, but are unable to breakdown native
cellulose .
The earliest and most consistent morphological response

173
to pathogens which penetrate directly is the formation of
structures called papillae (191). Several lines of evidence
suggest that mechanical forces applied by the pathogen
during penetration may provide the stimulus for papilla
formation (3). First, papillae are restricted to areas
beneath or immediately adjacent to the point of penetration.
Second, papilla-like structures can be induced by gentle
pin-pricks. Third, when no evidence of localized mechanical
force is found during penetration, palillae are not formed.
Finally, papilla-like structures are not formed in response
to treatment with the pathotoxin victorin which causes other
morphological changes typical of disease.
The proposed function of papillae is to impede or block
penetration by some pathogens but evidence for such a role
is far from conclusive (191).
Papillae are formed in epidermal cells of waterhya-
cinths during penetration by A. zonatum but are not formed
in other cell types. This is additional evidence to support
mechanical force as the most probable means of initial entry
; io its host.
.In summary it can be stated that entry of A. zonatum
through the cuticle was mechanical; penetration through cell
walls involved production of cellulases and to a lesser
extent pectinases; and penetration from cell to cell was
also enhanced by localized pectinase production.

174
One of the proposed functions of the phenol cell, is
defense against invading pathogens. Electron micrographs of
invaded phenol cells support this theory. The hyphae pene¬
trate the phenol cells, perhaps as they would any other
cell, but once inside are confronted with a collection of
phenolic compounds in concentrations that are toxic. The
hyphae appear less electron-dense than those in other cell
types, highly vesiculated, and cytoplasmic organelles such
as mitochondria are not evident. In other words, the hyphae
are nonviable. It would appear that the concentration of
phenols in those cells is such that they are no longer
stimulatory to the fungus' growth but are toxic to it. In
this case the phenol cell is able to stop the advancing
mycelia at various points within the leaf, thereby limiting
the infection. This could also help explain why disease
severity in this plant is limited to only 40% of the leaf.
The presence of hyphae in the tracheary elements was
unexpected. However, A. zonatum is closely related to
members of the genus Cephalosporium, which has many species
that are vascular pathogens of other plants. It is postu¬
lated from the results of this study, that A. zonatum is
i anslocated through its host in the xylem without causing
tissue destruction or blockage. On the other hand, it does
cause extensive wall degradation of the surrounding meso-
phyll cells. This suggests that this fungus lacks the

175
necessary enzymes to degrade the lignified materials present
in the secondary wall thickenings of xylary cells.
Several cytological changes due to infection were
observed in the cells of waterhyacinth leaves. The most
noticeable was the disappearance of starch granules from the
chloroplasts of palisade cells. Changes in starch content
following infection have been observed in many foliar di¬
seases (191). The general pattern is an initial decrease
followed by a marked increase with heavy accumulations
around the margins of the lesions. This presumably is
brought about by the increased respiration which occurs soon
after infection and serves to increase anabolic pathways
(61). The accumulation of starch shortly after infection
has been attributed to an increase on CC^ fixation in the
dark by plants (198). Similar results were obtained by Luke
and Freeman (99) in victorin-treated oat leaves and by Wang
(137) in Uromyces phaseoli infected bean leaves. Later in
infection, however, starch content declines drastically
until most or all of it is gone.
A second noticeable cytological change during pathoge¬
nesis was the build up of plastoglobuli in chloroplasts. A
similar increase in plastoglobuli was reported in spinach
plants infected with Albugo occidentales (60). The function
of plastoglobuli is unknown but their occurrence in the
stroma is a characteristic feature of chloroplasts fixed

176
with osmium tetraoxide (176). Several lines of evidence
indicate that they are not artifacts (66). Studies also
suggest that they may be a product of senesence since they
increase in size and number during aging (43). The exact
chemical nature of plastoglobuli is not clear although it is
generally believed that they represent a reservoir of excess
lipid (65). An interesting observation has been made by
Adams and Strain (1) in a study of the drought-deciduous
desert plant Cercidium. They found that chloroplasts in the
rather ephemeral leaves which appear after a heavy rainfall
contained starch and are ultrastructurally similar to those
of higher plants. The chloroplasts in the green bark
tissue, which evidently provides a major source of
photosynthetically fixed carbon to the plant, resemble those
in the ephemeral leaves of other plants, except they lacked
starch and have numerous large plastoglobuli. They sug¬
gested that the plastoglobuli may represent a form in which
photosynthetically fixed carbon is stored in these chloro¬
plasts. Thus, plastoglobuli may have the same general role
as starch in most other plants. Thomson (176) considers
this an attractive hypothesis, particularly in regard to
pi 'nts where limited water may be available for metabolic
pri :esses and from the point of view of effecient energy
conservation and utilization. It Is unlikely that a free-
floatir.g aquatic plant such as the waterhyacinth would be

177
subjected to limited water supplies under natural growing
conditions. Therefore, plastoglobuli in this plant must
have a different function.
The last major cytological change observed in infected
waterhyacinths was the increase in microbodies within the
cells. The term "microbody" was introduced into the litera¬
ture by P.hodin in 1954 [see DeDuve (41)] to designate a
special type of cytoplasmic body present in the convoluted
tubule cells of the mouse kidney, characterized by a single
membrane and finely granular matrix. Similar structures
have since been found in yeast cells (57), other fungi
(111,116) green algae (168) and higher plants (20,67,84).
These structures have been called by various names
(peroxisomes, glyoxysomes and crystalloids) depending on the
type of function proposed for them (178). The biological
significance of microbodies has been attributed to several
things, including serving as mitochondrial precursors (158).
The main functions appear to be involved with gluconeogensis
(178). In germinating fatty seeds, glyoxysomes are involved
in the conversion of fats to sugars via 8-oxidation and the
i’lyoxylate cycle (178). Leaf peroxisomes, however, are
â–  volved in the conversion of nonphosphorylated compounds
d-: ■ ‘ved from photosynthesis to glycine, serine, and C-^
cc pounds via the glycolate pathway (178). From glycolate,
glycine, or serine, the pathway is gluconeogenic in the

178
light since it leads to phosphoglycerate and then to sugars
(178). For a detailed discussion of microbodies the reader
is referred to the excellent review by Tolbert (178).
The appearance of microbodies in infected waterhyacinth
leaves at about the time starch is lost in the chloroplasts
represents, perhaps, an alternate method of producing
needed carbohydrates for energy production.
During the final stages of pathogenesis, chloroplasts
were observed to break down along with other cell orga¬
nelles. The grana distend, plastoglobuli may coalesce, and
organelle structure is lost completely. Plasmolysis occurs,
oxidation of cellular components takes place, and the end
result is the formation of dark brown melanin pigments
typical of necrosis and death.

SUMMARY AND CONCLUSIONS
The waterhyacinth [Eichhornia crassipes (Mart.) Solms.]
is a free-floating vascular hydrophyte that has colonized
much of Florida's inland waters. In 1970, a program was
initiated at the University of Florida to study biological
control of this noxious plant with phytopathogens. One of
the pathogens currently being studied is the fungus Acre-
monium zonatum (Sawada) Gams. It causes severe spotting on
both leaves and petioles of this plant under conditions of
high humidity.
During field trials with this fungus, it was observed
than small, young, plants displayed fewer symptoms after
infection than did larger plants in the same plots. It also
appeared that large plants infected with A. zonatum exhi¬
bited a faster rate of leaf regeneration than did smaller
plants. The present study was initiated to determine if
small plants were in fact more resistant to A. zonatum than
large plants; if meristematic activity in the plants was
altered after infection; and, if so, to what extent host
phenolic compounds and their oxidizing enzymes, namely
polyphenoloxidase (PPO), were responsible.
Waterhyacinths displayed various degrees of resistance
to A. zonatum depending on their morphotypic state of deve¬
lopment. Results of this study indicate that these dif¬
ferences in resistance are due to the variations in phenol
179

180
chemistry among plants of different sizes and to subsequent
changes induced by infection (Table S-l).
Small plants are more resistant to fungal attack than
are medium or large plants, based upon the number of lesions/
leaf after infection. It appears that the presence of high
concentrations of phenolic compounds does not itself impart
resistance to the pathogen. Rather it is the oxidation of
these compounds by enzymes, such as polyphenoloxidase (PPO),
which is responsible for the resistance. This view is
supported by qualitative and quantitative data on the
phenols in plant morphotypes and is coincident with the
observed differences in resistance.
2
Small plants, by virtue of having fewer phenol cells/mm
leaf area, have less total phenol content/leaf than larger
plants. If phenol content alone, was responsible for di¬
sease resistance, then small plants would be more suscep¬
tible than large plants but they were not. In this case PPO
activity is apparently the mediating factor. The rate of
enzyme activity in small plants is three-fold that in large
plants; presumably therefore, oxidation of polyphenols to
quiñones is much greater in small plants. Thus, small
plants are initially more resistant to pathogenic attack
than are larger plants.
After the disease has progressed for several weeks the
differences in resistance among the morphotypes is no

TABLE S-l, Differences and similarities among healthy and
A. ZONATUM-INFECTED WATERHYACI NTH MORPHOTYPES.
Assessment criteria
Morphotype
“Small
Medium
Large
Mean ft lesions/leaf
3,7
12.8
18.3
% TOTAL DISEASE
41.3
37.0
39.5
Mean ft phenol cells/mm^
33,6
41.8
48,7
PPO RATE (healthy)
1.53
0.80
0.47
PPO RATE (DISEASED)
0,90
0.70
1,36
PPO localization (healthy)
3 cell typesa
3 CELL TYPESA
3 CELL TYPESA
PPO localization (diseased)
ALL CELLS®
ALL CELLS®
ALL CELLS®
Type of phenolic acids (healthy)
6
6
9
Type of phenolic acids (diseased)
7
7
9
Total phenols (healthy)
92 ug/g
106 ug/g
104 ug/g
Total phenols (diseased)
80 ug/g
96 ug/g
105 ug/g
Fungal growth (healthy)
stimulative0
stimulative0
STIMULATIVE0
Fungal growth (diseased)
stimulative0
STIMULATIVE0
STIMULATIVE0
Leaf regeneration (healthy)
27,3%
28,5%
46.1%
Leaf regeneration (diseased)
21.6%
33,9%
93.3%
AVASCULAR PARENCHYMA, BUNDLE SHEATH, AND PHENOL CELLSj BALL CELLS WHICH CONTAIN CHLOROPLASTS
CP= 0.05J GROWTH INCREASED OVER CONTROLS! °P= 0.05J GROWTH INCREASED OVER HEALTHY COUNTERPART
181

182
longer evident. Each plant size exhibits a percent-
total-diseased leaf area which is statistically the same
(approximately 40%). It is believed that this equalization
of disease severity results from a gradual loss in resis¬
tance by small plants while at the same time there is a
gradual increase in resistance by large plants. Again,
quantitative data of the phenol metabolism can be correlated
with this change.
The total phenol content decreased significantly after
infection in small- and medium-sized plants. This is coin¬
cident with a reduction in PPO activity. The coupling of
these two phenomena may account for the decrease in resis¬
tance of small plants. Large plants, on the other hand,
retain their total phenol content and at the same time
exhibit a three-fold increase in PPO activity. Therefore,
an increase in polyphenol oxidation would be expected to
occur and could account for the increase in resistance in
large plants.
In essence, then, the point being made is: if infected
â– small plants retained the phenol content and PPO activity of
’■■■-jlthy plants, then disease severity would probably be
lif.ihed to much less than 40%. Similarly, if infected large
pi ml’ retained the PPO activity of healthy plants, disease
would progress to a much higher percentage, perhaps 60-70%.
Howev- , because each morphotype responds to infection

183
differently (in most cases in contrast to each other) di¬
sease severity balances among the plant sizes at approxi¬
mately 40% of the leaf-surface area.
If disease severity is viewed, not from a percentage of
leaf-area infected, but as a reduction in plant growth, then
data on leaf regeneration rates among the morphotypes be¬
comes of prime importance. It has been observed that in¬
fected large plants regenerate two to three times as many new
leaves as do infected small plants. This too, is correlated
with the plant's phenol chemistry.
It has been shown that A. zonatum is capable of synthe¬
sizing indoleacetic acid in vitro and that this is one
explanation for the increased growth observed in large
plants. More important, however, is the fact that phenols
are known inhibitors of IAA oxidase, the enzyme responsible
for controlling the IAA level in the plant. It has already
been pointed out that the different waterhyacinth morpho¬
types vary in phenol content, both prior to and after infec¬
tion. The higher phenol content in large plants could
account for the increased growth observed in large plants by
inhibition of the IAA oxidase system.
Perhaps the most significant data supporting a positive
na for phenols in disease resistance comes from the loca¬
tion studies of PPO in healthy and diseased plants.
' ..ne activity is localized in the thylakoids of

184
chloroplasts in only three cell types in healthy plants.
After infection there is a "turn on" in PPO activity in all
cells which contain chloroplasts. This turn on in PPO
activity is highly suggestive of a vital role for enzymatic
oxidation of polyphenols during disease.
Before disease can ensue , the pathogen must come into
contact with and penetrate its host. In this regard, A.
zonatum can enter the waterhyacinth by either of two ways:
through open stomata or by directly penetrating the unbroken
cuticle of the leaf. Intracellular colonization is enhanced
by the diffuse secretion of cellulolytic enzymes and perhaps
by the localized secretion of pectolytic enzymes.
Growth of A. zonatum was either unaffected or stimu¬
lated by seven different phenolic acids in concentrations up
to 1000 ppm in minimal media. When yeast extract was added
to the media as a growth supplement, one phenolic acid, p-
coumaric, was found to be inhibitory. In addition, fungal
growth was enhanced on media containing yeast extract and
extracts from diseased leaves over that on media containing
healthy leaf extracts.
Several cytological changes were observed in the cells
cm infected waterhyacinth leaves. First, chloroplasts in
c Lis of healthy leaves have an abundance of starch granules
which disappear after infection. Second, there are only a
few plastoglobuli in chloroplasts in healthy cells, but

185
after infection, they increase both in size and in number.
Third, there is a noticeable increase in microbodies in the
cytosol of infected cells. It is believed that each of
these cytological changes is the result of a shift in host
metabolism induced by infection.
It is concluded that waterhyacinths have at least two
distinct biochemical defense mechanisms that are related to
phenol metabolism and plant size. The first is the presence
of high concentrations of polyphenols in specialized phenol-
cells which, under the proper conditions, can serve as
toxicants to potential pathogens. The second proposed
defense mechanism of waterhyacinths is an acceleration of
its growth rate brought about by the inhibition of IAA
oxidase by the phenolic compounds.
Which of the above mechanisms is operational is depen-
-nt upon the plant's morphotypic stage of development. It
â–  u believed that initially small plants defend against
p, ¡-hogenic attack by virture of their high PPO activity
whereas large plants respond by increased leaf production.
Mi Hum-sized plants appear to have a combination of both
mti. nisms.
In consideration of A. zonatuni as a potential biocon¬
trol agent for waterhyacinths, it is concluded that best
control would be achieved with small, young, plants rather
than with larger, more mature plants. In this regard,

186
control procedures should be initiated early in the spring
when new plants start to grow and colonize the body of
water.

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BIOGRAPHICAL SKETCH
Raymond D. Martyn, Jr. was born in Washington, D.C. on
December 15, 1946. He was graduated from Pompano Beach
Senior High School in June, 1964, and obtained an Associate
of Arts degree in Engineering from Palm Beach Junior College
in April, 1966. He was awarded the degrees of Bachelor of
Science in Biology in June, 1969 and Master of Science in
Microbiology in June, 1971, from Florida Atlantic Univer¬
sity .
From September, 1971 to August, 1973, he served as
laboratory supervisor at the Biological Control Laboratory
at Florida Atlantic University. From September, 1973 to
May, 1974, he was a biology instructor at Palm Cove Beach
High School.
In June, 1974, he entered the Plant Pathology Depart¬
ment at the University of Florida to pursue graduate studies
towards the degree Doctor of Philosophy and is presently a
candidate for that degree in June, 1977.
He is married to Jane D. Brooks and has a six-year-old
daughter, Susan, by a former marriage.
204

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.
Thomas E. Freeman, Chairman
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.
«■¿i P■ 1.. C
Daniel A. Roberts
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.
Ronert E.Stall
Professor of Plant Pathology
â– I certify that I have read this study and that in my
â– pinion 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.
: „. ■' ; • - ivtxbr
Herbert K. Luke , , -
Plant Pathologist, U.S.D.A.

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.
Professor of Botany
This dissertation was submitted to the Dean of the College
of Agriculture and to the Graduate Council, and was accepted
as partial fulfillment of the requirements for the degree of
Doctor of Philosophy.
June, 1977
Dean, Graduate School