Title: Disease resistance mechanisms in waterhyacinths and their significance in biocontrol programs with phytopathogens /
Full Citation
Permanent Link: http://ufdc.ufl.edu/UF00099392/00001
 Material Information
Title: Disease resistance mechanisms in waterhyacinths and their significance in biocontrol programs with phytopathogens /
Physical Description: xvi, 204 leaves : ill. ; 28 cm.
Language: English
Creator: Martyn, Raymond DeWint, 1946-
Publication Date: 1977
Copyright Date: 1977
Subject: Water hyacinth -- Control   ( lcsh )
Plant Pathology thesis Ph. D
Dissertations, Academic -- Plant Pathology -- UF
Genre: bibliography   ( marcgt )
non-fiction   ( marcgt )
Thesis: Thesis--University of Florida.
Bibliography: Bibliography: leaves 187-203.
Statement of Responsibility: by Raymond DeWint Martyn, Jr.
General Note: Typescript.
General Note: Vita.
 Record Information
Bibliographic ID: UF00099392
Volume ID: VID00001
Source Institution: University of Florida
Holding Location: University of Florida
Rights Management: All rights reserved by the source institution and holding location.
Resource Identifier: alephbibnum - 000011390
oclc - 03386351
notis - AAB3883


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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


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.


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


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


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

LEAVES . ... . . . . . . 90

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



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

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

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

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


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





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


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


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.


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


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 . . . . . . .


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


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



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


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


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.


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.)


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


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

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-


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


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.



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


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-
40 cm and (iii) large plants, with leaves greater than 40
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


maintained under identical conditions. Two weeks post-

inoculation, leaves were excised and used for subsequent


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.


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


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

a. Waterhyacinth with zonate lesions on leaves and

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

c. Waterhyacinth leaf showing coalescence of leaf

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





Lu 75








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.




0 0 0 0 0 0

HIMOd9 .IV3-1



0 0
CM -





F,71? .0





PvcII-F!- 7-x---- J


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/


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



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


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)].


\ a
\ 0



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

0 a

om o
0 Z

S I I o

0 -
U) U ,


/ 4 0

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


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)

wash in buffer (4x)

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

embed in epoxy resin

post-stain w/ UrAc (10min.)

post- stain w/ PbCi (5min.)

Flow Diagram for Electron


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)

treat w/ DDC3

wash in buffer(5x)
I -

boil sections
(10 mir)

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

t t

post-stain w/

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.


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
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


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).


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).



cU 40


E 30






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).


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

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