DISEASE RESISTANCE MECHANISMS IN WATERHYACINTHS AND THEIR
SIGNIFICANCE IN BIOCONTROL PROGRAMS WITH PHYTOPATHOGENS
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
This above all: to thine own self be True
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!"
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
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
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 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
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
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
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
LIST OF FIGURES
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
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
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 .. ...
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 ...
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 . . . . . .
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
Raymond DeWint Martyn, Jr.
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
(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
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
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
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
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
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.
RESPONSES OF WATERHYACINTHS TO INFECTION WITH ACREMONIUM
ZONATUM AND ITS IMPLICATIONS IN BIOLOGICAL CONTROL
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)
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
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
SMALL MEDIUM LARGE
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
I I I I I I
0 0 0 0 0 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
A CYTOCHEMICAL AND ULTRASTRUCTURAL STUDY OF THE PHENOL
CELLS AND POLYPHENOLOXIDASE ACTIVITIES IN HEALTHY AND
DISEASED WATERHYACINTH LEAVES
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
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)].
S I I o
U) U ,
/ 4 0
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
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)
fix in Karnovsky's fixative'
(2hr- 22 C)
wash in buffer (4x)
post-fix in 1% OsO4
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
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.
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)
wash in d.w.-sucrose (5x)
post-fix w/ 2% Os04
dehydrate in EtOH
embed in epoxy resin
Flow Diagram for C
ytochemical Localization of
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
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).
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).
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
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
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.
Figure II-12 (a b). Localization of polyphenol-
oxidase in chloroplasts of phenol cells in healthy water-
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
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
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
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