Q7603
5756‘”? /

  

  

leifinv
Michigan State

 

L

University

 

i‘

 

This is to certify that the
thesis entitled

THE COLONIZATION OF WHEAT STEMS AND
SUSEQUENT PERITHECIUM DEVELOPMENT BY
GIBBERELLA ZEAE (ANAMORPH FUSARIUM
GRAMINEARUM)

presented by
JOHN C. GUENTHER

has been accepted towards fulfillment
of the requirements for the

M. S. degree in Botany and Plant Pathology

 

 

Eff/15x AMOA Mat mo 19

MajoF’Professof's Signature
_ 8/020/0 3

Date

MSU is an Afiinnafive Action/Equal Opportunity Institution

PMCE IN RETURN BOX to remove this checkout from your record.
To AVOID FINES return on or before date due.
MAY BE RECALLED with earlier due date if requested.

DATE DUE DATE DUE DATE DUE

 

6/0! c:/CIRC/DateDue.p65~pt15

 

THE COLONIZATION OF WHEAT STEMS AND SUBSEQUENT
PERITHECIUM DEVELOPMENT BY GIBBERELLA ZEAE (ANAMORPH
FUSA RIUM GRA MINEAR UM)

By

John C. Guenther

A THESIS

Submitted to
Michigan State University
In partial fulfillment of the requirements
For the degree of

MASTER OF SCIENCE
Department of Botany and Plant Pathology

2003

ABSTRACT
THE COLONIZATION OF WHEAT STEMS AND SUBSEQUENT
PERITHECIUM DEVELOPMENT BY GIBBERELLA ZEAE (ANAMORPH
FUSARI UM GRAMHVEA R UM)
By
John C. Guenther
A devastating pathogen of wheat, Triticum aestivum, is the head blight
fungus, Gibberella zeae. Sexual spores of this fungus develop on cereal debris
remaining in the field following harvest and provide the primary inoculum for
infection of the next cereal crop. The colonization of vegetative wheat tissue by
G. zeae in the green house and perithecium development on stem debris was
investigated. To observe the patterns and extent of colonization within wheat
stems, plants were inoculated with a fungal strain expressing the endow-green
fluorescent protein. For observations of perithecium development, colonized
wheat stems from an infected field were returned to a new outdoor plot in mesh
bags and collected as perithecia formed. The results of this study showed that
stems were systemically colonized following head infections and that this
colonization was extensive. Descriptive observations implicated G. zeae as a
pathogen that invades the vascular system and indicated that host anatomy was
important to the development of disease. Pen'thecium formation occurred on
segments of all stems placed in the field and was in close association with specific
cell types of the wheat stem epidermis. The characterization of possible resistance

mechanisms, systemic colonization and role of host anatomy in association Wlth

potential inoculum development are discussed.

ACKNOWLEDGEMENTS

I would like to thank the following people for their guidance and assistance while
completing my Master’s degree: Dr. Frances Trail, Dr. Karen Klomparens, Dr.
Annemiek Schilder, Luis Valesquez, Iffa Gaffoor, Hiaxin Xu, Chil Kwon, Dr.

Shirley Owens and Dr. Joanne Whalen.

iii

TABLE OF CONTENTS

LIST OF FIGURES ........................................................................ iv

CHAPTER ONE

LITERATURE REVIEW .................................................................. 1
INTRODUCTION .................................................................. l
GEOGRAPHIC DISTRIBUTION AND EPIDEMIC LOSS ................. l
MYCOTOXINS ..................................................................... 3
DISEASE CYCLE AND INOCULUM .......................................... 4
SYMPTOMS AND ANATOMY OF INFECTED WHEAT HEADS ...... 6
CONTROL OF F HB ............................................................... 7
SUMMARY ......................................................................... 9

CHAPTER TWO

COLONIZATION OF SENESCENT WHEAT STALKS BY THE HEAD
BLIGHT F UNGUS, GIBBERELLA ZEAE (ANAMORPH F USARI UM
GRAMINEAR UM), AND SUBSEQUENT DEVELOPMENT OF

PERITHECIA .............................................................................. l I
INTRODUCTION ................................................................ 1 1
MATERIALS AND METHODS ................................................ 13
RESULTS ........................................................................... 1 7
DISCUSSION ..................................................................... 33

LIST OF FIGURES

Figure 1. Wheat stem anatomy.

Figure 2. Progression of stem colonization.

Figure 3. Cross section of an infected vascular bundle within a wheat culm.
Figure 4. Cross section of an infected wheat culm showing horizontal
colonization.

Figure 5. Direct penetration of parenchyma cell wall by hyphae.

Figure 6. Longitudinal section of wheat stem node showing vertical colonization.
Figure 7. Primary and secondary hyphae within pith cavity of a wheat culm.
Figure 8. Secondary hyphae and stroma initials within chlorenchyma of a wheat
culrn.

Figure 9. Perithecia developing on culm surface.

Figure 10. Perithecia developing on stem node surface.

Figure 11. Longitudinal section of wheat culm on which perithecia have formed
above stomates.

Figure 12. Longitudinal section of wheat culm on which perithecia have formed
showing hyphal masses below perithecia.

Figure 13. Cross section of wheat culm and leaf sheath with perithecia forming on
the leaf sheath.

Figure 14. Longitudinal section of leaf base with perithecia forming on surface.

CHAPTER ONE
LITERATURE REVIEW
Introduction- Fusarium head blight (F HB), sometimes known as wheat
scab or ear blight, is a worldwide disease of wheat and other small grains.
Epidemics during the last century have been documented throughout the world
and losses to the grain industry have been significant. The epidemiology of F HB
and the losses attributed to this disease have been well reviewed by several
authors (Atanasoff, 1920; Sutton, 1982; Parry, 1995). These authors emphasize
the complexity of this disease and how this may increase loss and decrease
effective measures of control. This disease afflicts a variety of hosts, including
wheat, maize, barley, oats and rye. The fungi that cause this disease have also
been isolated from other grasses and soybean and while not causing disease on
these plants, these hosts could serve as an alternate means of survival. In addition
to the confusion of multiple hosts, multiple fungi have been associated with the
disease. Most documentation of F HB associates the disease with five causal
organisms: F usarium graminearum Schwabe (Gibberella zeae (Schwein.) Petch),
F. culmorum (Smith) Sacc., F. avenacum (Corda ex Fr.) Sacc., F. pane and
Microdochium nivale (FL) Ces. Most of these organisms are also capable of
causing stem rot, root rot and seedling blight. The complex interaction between
multiple hosts, causal organisms and diseases is poorly understood and confusing
in literature. This review will focus on FHB of wheat and G. zeae, the primary

causal agent in North America.

Geographic distribution and epidemic loss- FHB has been documented
in all regions in North America where wheat or other small cereals are grown, but
epidemics do not present a significant threat to all regions. Within the last century
epidemics have been sporadic and at times, particularly in the early 1900’s, not
well documented. This sporadic nature makes it difficult to study and therefore
accurately estimate total loss. The geographic distribution of epidemics within the
United States and Canada does suggest that some regions are predisposed to
greater disease occurrence (Sutton, 1982; McMullen et al., 1997). These regions
within the US. include: the Red River Valley (Minnesota, North Dakota and
South Dakota), the Midwest (Michigan, Ohio, Indiana, Illinois and Wisconsin),
and the Mid—Atlantic (Pennsylvania and New York). In Canada these regions are:
the Ottawa Valley and areas of Ontario, Manitoba and Quebec. Losses within
these regions are due to the combination of intense production of cereals that are
host to this disease and climatic conditions that are favorable for disease
development. Economic losses can be severe and have been documented to be as
high as $1 billion for a tri-state area within a single growing year (McMullen et
al., 1997). Through the 19903, the total losses to wheat and barley farmers have
been estimated at $3 billion. Economic loss is usually associated with damage that
is directly caused by the fungal pathogen, but losses may also be due to indirect
effects (Bai and Shaner, 1994; Parry et al., 1995). The direct effects of this disease
are loss of yield and reduced grain quality. Diseased plants exhibit reduced grain
filling, lowered test weights and grain discounts at the elevator due to fungal

contamination all at a cost to the producer. Indirect effects are more difficult to

estimate, but include: increased grain costs to bakers and brewers, losses to
seedling blight from contaminated seed, reduced quality of products made with
contaminated grain (gushing of beer and non-rising bread) and adverse reaction of
livestock fed with grain contaminated by mycotoxins.

Mycotoxins- Mycotoxins are secondary metabolites produced by fungi
that are toxic to animals and humans (Seaman, 1982). There has been a growing
interest in this area of research, as more is understood about the prevalence of
mycotoxins and the damage they cause. G. zeae produces several mycotoxins.
These may be produced in association with the infection process in the field or
within contaminated grains in storage (Mills, 1982). The most harmful class of
toxins found in grains following FHB epidemics is the tricothecenes (V esonder
and Hesseltine, 1981; Snijders, 1990). These mycotoxins are considered to be the
most potent inhibitors of protein synthesis for eukaryotic cells (Martins and
Martins, 2002). The tricothecenes produced by G. zeae are deoxynivalenol,
acetyldeoxynivalenol (isomers 3-ADON and 15-ADON) and nivalenol. Of these
toxins, nivalenol is the most toxic, followed by acetyldeoxynivalenol and finally
deoxynivalenol is the least toxic (Joffe, 1986). Deoxynivalenol is the most
commonly isolated tricothecene from contaminated grains and while less toxic
than the other tricothecenes, it can result in adverse reactions if consumed. Feed
grains contaminated with deoxynivalenol can cause vomiting and feed refiisal in
swine but has a lesser effect on chickens and cattle (F orsyth et al., 1977). It can
also induce emesis, headaches, convulsions and cause gastrointestinal illness in

humans if consumed in high enough concentrations. With regard to the infection

of wheat plants, it is a known virulence factor that affects severity of disease
following infection by the firngus (Bai et al., 2001). This toxin is carried within
the xylem and phloem of wheat heads away from the point of infection and
exhibits a close relationship with pathogenic changes at the cellular level (Kang
and Buchenauer, 1999). Zearalenone is another mycotoxin produced by G. zeae
that is well studied. It is a stable estrogenic compound that elicits the same
reaction as estrogen in humans and especially swine. It can cause estrogenic
syndrome, including feminization of young male swine, when consumed with
infected grain (Caldwell and Tuite, 1970). The impact of mycotoxins on humans
and livestock significantly contributes to losses suffered due to F HB.

Disease cycle and inoculum- The disease cycle of F HB on wheat is well
studied and understood (Sutton, 1982 and Parry et al., 1995). A crucial component
of the disease cycle of F HB is the presence of infested host debris that serves as
the principal source of inoculum. G. zeae survives within old stems and ears of
corn, as well as the stubble and pieces of wheat and other cereals left in the field
following harvest. The pathogen survives the winter as mycelia within the debris
and then forms sporulating structures in association with periods of prolonged
wetness throughout the year. The fungus may survive within debris on the soil
surface for as long as three years within spikelet and grain residues (Khonga and
Sutton, 1988). Two types of spores are formed on field residues and serve as the
inoculum that perpetuates field infection; macroconidia and ascospores.

Macroconidia are asexual spores that form from globose phialides (Booth, 1971).

The phialides may aggregate into sporodochia forming slimy, salmon-colored

masses on upright plant and debris surfaces. The spores within sporodochia are
splash dispersed by rain and can be deposited short distances vertically and
horizontally fiom a debris source by water droplets (J enkinson and Parry, 1994).
This inoculum source can effectively spread disease, however it is generally not
regarded as the initial or primary source of inoculum (Francl et al., 1999).
Ascospores are a more likely source of primary inoculum based on their ability to
travel long distances, daily periodicity in release and large numbers of ascospores
collected in the field when disease occurs (Paulitz, 1996; Fernando et al., 1997;
Fernando et al., 2000). Ascospores are formed within asci (eight per ascus) that
are densely packed with paraphyses within somatic perithecia (Trail and
Common, 2000). Perithecia form on debris surfaces throughout the year following
periods of increased moisture or rainfall events. The initiation of perithecium
development requires low-intensity ultra-violet light (wavelength below 320 nm)
and the rate of perithecium formation increases with temperature up to 29 °C
(Tschanz et al., 1976). It is worth noting that requirements for the development of
perithecia and spore release differ. Ascospore release is by forcible discharge into
the air or by “oozing” to form a cirrhus. Ascospore release events have been
correlated with increases in relative humidity and require light (Trail et al., 2002)
however, temperature requirements are lower (26 °C maximum) than those
recorded for perithecium development. When released in the field ascospores, by
the force of discharge, are wind dispersed and can be carried long distances on the
magnitude of kilometers. For this reason perithecia and the ascopores they

produce are an important component of this disease in epidemiological terms.

Both macroconidia and ascospores have similar potential to germinate and cause
disease (Stack, 1989). However, airborne ascospores can easily reach wheat heads
in the field where infested debris is present as well as wheat heads in adjacent
fields. Spores that are delivered to wheat heads are capable of germinating,
infecting and colonizing wheat heads at the time of anthesis; this is the stage of
host development that is most susceptible to F HB. Infested plants can then
potentially become field debris and therefore a source of inoculum in following
seasons, thereby perpetuating the disease.

Symptoms and anatomy of infected wheat heads- Wheat heads, and
more specifically wheat spikelets, are the major sites of initial infection by G.
zeae. Head infections in the field initiate the onset of F HB and cause the majority
of damage suffered to crops. Arthur (1891) was the first to report F HB as a floral
disease. Later observations by Atanasofi' (1920) confirmed that the spikelets of
the wheat head are the site of initial penetration by G. zeae. The histological
studies of Pugh et al. (1933) and Tu (1953) later identified specific floral
components of the spikelets as favoring infection. Pugh et al. (1933) identified the
infection of anthers as being a first crucial step and observed that all subsequent
infections arose from this initial infection. Tu (1953) found that in addition to
anthers, the stigma, ovaries and inner walls of the palea and lemma were
susceptible to infection when the fungus was present in the floral cavity. Studies
using transmission electron microscopy by Wanyoike et al. (2002) confirmed that
all of these floral parts are likely involved in the infection of wheat spikes. In an

additional study by Pritsch et al. (2000) it was shown that the outer surfaces of the

glumes are also susceptible to direct penetration by the fungus by means of
stomatal openings. Microscopy studies that followed the colonization of wheat
heads after initial penetration (Pugh etal., 1933; Tu, 1953; Ribichich et al., 2000;
Wanyoike et al., 2002) are all in agreement that the fungus proceeds to colonized
the rachis through the vascular bundles of the infected spike. Subsequent
colonization up and down the rachis to adjacent spikes was also through the
vascular bundles. G. zeae may penetrate wheat plants at the stem base, however,
these infections do not directly result in head symptoms (Clement and Parry,
1998) and therefore the major site of infection for F H3 is the wheat head. The
colonization of the wheat head and the damage caused by colonization is directly
observable by eye in the form of distinct symptoms associated with F HB.

The symptoms associated with F HB are easily identifiable following
infection of plants in the field (Atanasofl', 1920; Wiese, 1987). The first symptom
of infection within immature heads is the appearance of a slightly brown water-
soaked spot on the surface of the glume. This spot spreads to cover the whole
spikelet and then adjacent spikelets. Shortly after, the point of infection begins to
lose its water-soaked appearance and dries up taking on the appearance of a
mature spikelet. This premature bleaching closely follows the infection downward
and also occurs in all tissues above the point of infection. Varying degrees of
infection may occur within a head depending on field conditions and wheat
cultivar. This can vary from a single spikelet to the entire head showing
symptoms. Colonization may continue down the culm (below the head) exhibiting

the same symptoms. Small purplish fruiting bodies (perithecia) or salmon to rose-

colored spore masses may eventually form on the outer surfaces of the head,
particularly at the spikelet bases. In fields that are particularly wet at the time of
infection, heads may become covered with white or pink mycelia and take on a
cottony or fuzzy appearance. Heavily infected fields typically have a pink or red
appearance from a distance.

Control of FHB- The damage sustained to the wheat industry by F HB is
extensive and therefore the search for an effective means of control is warranted.
As stated earlier, F H8 is a complex disease; the causal organisms can afflict
multiple hosts in multiple ways. For this reason cr0p rotations are not an effective
means of control when used alone (Martin and Johnston, 1982). In a similar way,
tilling debris into soil can reduce disease incidence in following seasons (Miller et
al., 1998; Dill-Macky and Jones, 2000). Tilling can also compromise measures to
control soil erosion. Seed treatments with fungicides can reduce seed-bome
inoculum sources and increase seedling vigor however this does not prevent
inoculum from infecting older wheat plants. Some safe fungicides are available
for the late treatment of wheat plants but difficulties in application to wheat heads
and small reductions in mycotoxin contamination can make fungicide applications
less efficacious (Shaner and Buechley, 1992). Alone, each of these measures is
only in part effective for the control of FHB. Martin and Johnston (1982) suggest
that the integration of tilling, crop rotation and fungicide application for seed and
foliage may potentially control this disease. The integration of control measures is
not always an attractive prospect to the grower; the time spent and expense are

greater and do not guarantee a better yield. Currently, breeding for resistant

cultivars of wheat holds the greatest potential for control of this disease
(McMullen et al., 1997).

Resistance to FHB within wheat cultivars has shown promise but as yet
there is not a completely resistant wheat line available. Resistance to F HB can be
regarded as passive or active in nature. Passive mechanisms involve phenotypic
traits such as plant height, presence of awns, spikelet density and timing of
flowering (Mesterhazy, 1995). Active resistance can be divided into five
components: Type I (resistance to initial infection), Type II (resistance to spread
of infection), Type III (resistance to kernel infection), Type IV (tolerance) and
Type V (resistance to mycotoxin accumulation) (Schroeder and Christensen,
1963; Mesterhazy, 1995; Pritsch et al., 2000). Type I and Type II resistance are
very widely accepted and screened for in wheat breeding programs. There tends
to be less interest in the other types of active resistance, with the possible
exception of Type V where resistant cultivars of wheat have been observed to
have greater resistance to tricothecene accumulation (Wang and Miller, 1988).
The genetics of FHB resistance is better understood than the physical mechanisms
conferring resistance. Several defense response genes are expressed in direct
correlation with head infections by G. zeae. Transcripts for genes encoding
peroxidase, chitinase and thaumatin-like proteins all accumulated quickly
following infection (Pritsch et al., 2000). The peroxidase genes were also
expressed systemically; indicating that direct contact by the fungus is not required
for induction of defense response genes (Pritsch et al., 2001 ). In addition to the

study of the expression of genes involved in plant defense, the inheritance of these

genes in wheat lines has also been investigated. Resistance can be variable in
susceptible lines of wheat, but is generally stable in wheat lines that are
considered to be resistant lines (Bai and Shaner, 1996). The mechanism of
resistance has not been elucidated, however, microscopy studies by Ribichich et
al. (2000) and Siranidou et al. (2002) show that extensive cell wall alterations
occur as the fungus comes in contact with wheat cells. Ribichich et al. (2000) also
noted vascular occlusions that corresponded to infection.

Summary- F HB is a devastating disease of wheat throughout most of the
wheat growing regions of the US. and Canada. Wheat growers suffered billions
of dollars in losses through the 19903 and countless more in losses prior to that.
The majority of losses occurred in sporadic epidemic years and is mostly
attributable to loss of yield and reduced grain quality due to fungal contamination.
The disease cycle is well understood. However, the complexity of multiple
pathogens, lack of host specificity and several sites of infection makes this disease
difficult to manage. Efforts to control F HB are marginal at best. The integration
of cultural practices, crop rotations and fungicide applications has been
unsuccessful in completely limiting this disease. The search for resistant wheat
lines shows promise but as yet, complete resistance to FHB has not been found.
Further understanding of resistance mechanisms and means of controlling initial
inoculum will yield better control methods for this disease. It is understood that
the formation of perithecia and the discharge of ascospores is a major factor in the
FHB complex, but little is understood about the formation of perithecia on debris

or the colonization of wheat leading up to the production of inoculum. An

10

understanding of the process of colonization of wheat and its impact on inoculum
formation is required if initial inoculum is to be controlled in the field.
Investigating the colonization process will also result in the discovery of potential
resistance mechanisms present in the host. Currently, sporadic epidemics continue
to damage the wheat production industry. A focus on resistant wheat lines and

limitation of inoculum in the field will increase control of F HB.

11

CHAPTER TWO
COLONIZATION OF SENESCENT WHEAT STALKS BY THE HEAD
BLIGHT FUNGUS, GIBBERELLA ZEAE (ANAMORPH FUSARIUM
GRAMINEARUM), AND SUBSEQUENT DEVELOPMENT OF PERITHECIA
INTRODUCTION
F usariurn head blight (F HE) is a destructive disease of wheat, T riticum
aestivum L., and other cereal crops worldwide. Loss of yield, contamination of
seed with mycotoxins and reduced seed quality all contribute to the impact of this
disease (McMullen et al., 1997). The trichothecene deoxynivalenol, a potent
protein biosynthesis inhibitor and, zearalenone, an estrogenic toxin, are found in
grains following FHB epidemics (Neish et al., 1982; Seaman, 1982). Zearalenone
and deoxynivalenol have been linked to feed refusal and toxicoses in livestock
and also present a threat to human safety (Forsyth et al., 197 7; Vesonder and
Hesseltine, 1981). Gibberella zeae (Schwein.) Petch (anamorph F usarium
graminearum Schwabe) is the predominant species, among several, to cause FHB
on wheat or barley in North America. Limited control of this disease can be
achieved through fungicide applications, resistance breeding and crop rotation in
conjunction with proper tillage (Bai and Shaner, 1994). To date, resistant cultivars
of wheat impart only partial protection from FHB. Fungicide applications, crop
rotations and tillage are not always economically feasible or efficacious.
The epidemiology and development of FHB has been best studied in
wheat (Sutton, 1982; Parry et al., 1995). An important component of the

development of this disease is inoculum in the form of ascospores or

macroconidia. While both ascospores and macroconidia have an impact, the

12

relative contribution of each spore type to FHB epidemics remains unresolved
(Stack, 1989). It is known that airborne spores are the primary inoculum and
infect wheat spikelets during anthesis when wheat heads are the most susceptible.
Symptoms begin to appear at the point of infection in the form of water-soaked
brown spots and then spread in all directions. Eventually this discoloration
spreads to the rachis and then to additional spikelets within the head. Blighting or
bleaching is also a common symptom of this disease and can be clearly seen on
emerged immature wheat heads (Parry et al., 1995; personal observations). The
fungus over-winters within crop debris in the form of mycelia and forms
ephemeral perithecia in the fall and spring as temperature and moisture levels rise
(Khonga and Sutton, 1988; Fernandez and F emandes, 1990; Reis, 1990). An
investigation into the development of perithecia in planta will further describe the
role of perithecia in the disease cycle of F HB.

The colonization of plant tissue is important to pathogen survival and
several studies have addressed this process. The infection and colonization of
stem-base tissue were investigated microsc0pically by Clement and Parry (1998).
This study showed that while G. zeae does colonize wheat stems upward,
infections do not reach the head and therefore this type of infection does not
directly contribute to FHB. Microscopic studies on the mode of infection, initial
colonization and development of symptoms within wheat heads show that the
penetration and infection of wheat spikelets directly impacts FHB (Tu, 1953;
Pugh etal., 1933; Pritsch et al., 2000; Wanyoike et al., 2002; Bushnell et al.,

2003). In general, these studies observed the penetration of ovaries, glumes and

13

inner walls of the palea and lemma by G. zeae and the head blight symptoms that
followed, however, stern colonization was not addressed. Strausbaugh and Maloy
(1986) documented symptoms in the stem below heads after head infections, but
did not describe the process of stem colonization. If head infections contribute to
the colonization of wheat stems, in addition to stem-base infections, both the
inoculum production and survival of G. zeae would depend on the extent of stem
colonization.

Our objective in the present study is to understand the process of
colonization and the development of perithecia on wheat stems following head
infections. Specifically, we investigated the process of colonization of wheat
stems by G. zeae and the development of perithecia on the surface of wheat stems.
We present our findings that the downward colonization of wheat stems is
extensive and that specific host tissues are preferential for colonization and
perithecium development by G. zeae.

MATERIALS AND METHODS

Wheat and fungal stock- Seeds of spring wheat (Triticum aestivum L.)
cv. Norm were planted into 9 cm. clay pots under greenhouse conditions with
constant supplemental light. Two fungal strains of G. zeae were used in this study,
wild-type PH-l (NRRL 31084) and ZTE-2a carrying the green fluorescent protein
as a reporter gene (provided by Robert Proctor at USDA-Peoria, IL). Fungal
stocks were maintained at —20 0C in sterile soil. For the production of
macroconidia as an inoculum source, potato dextrose agar plates were point

inoculated by transferring hyphae of the fungal strains and allowed to grow under

14

cool white light (Phillips F 34CW) at room temperature until mycelia reached the
edge of the petri dish. Plates were then washed with sterile distilled water and the
resulting solution was filtered using Miracloth (Calbiochem, La Jolla, CA) to
remove hyphae and debris (Pat Hart, personal communication). Spore
concentrations were measured using a hemacytometer and adjusted to a final
concentration of 5 X 105 spores/m1 in sterile distilled water.

Inoculation of wheat plants- Plants were inoculated 3-4 days prior to
anthesis by pippetting 10 pl of conidial suspension into a spikelet at the midpoint
of the rachis. Plants were then placed in a mist chamber at approximately 24 °C
for 4 days. Symptoms were observed daily following mist chamber treatments and
plants were removed from the greenhouse for observations of stem colonization
starting at 12 days after the plants were inoculated.

Determination of infection front- Greenhouse-inoculated plants were
prepared as follows: Heads were cut from each plant immediately below the
lowest spikelet and the remaining stem was cut at 5 cm intervals. At each cut
point thin sections were cut using a razor blade. These were mounted in distilled
water, coverslipped and observed on the microscope immediately. As a cut point
exhibited no fungal infection, the preceding 5 cm segment was cut into 1 cm
segments and similarly sectioned to determine the infection front. In total 65
stems were analyzed. Regression analysis was conducted using Si grnaplot 2000
for Windows Version 6.10 software.

Treatment of wheat residue- Wheat plants (cultivar Norm) grown under

field conditions in Ramsey County, Minnesota, were tagged for collection when

15

symptoms were visible. Tagged samples were then removed from the field at
grain harvest. Each sample was collected without root material or heads and
consisted of a single wheat stem with leaves but did not include tillers. To
stimulate perithecium deve10pment under field conditions, samples were placed in
vinyl mesh bags (l-mm mesh, 43 X 20 cm), two samples to a bag, and returned to
the soil surface in a field in Ingham County, Michigan on 6 September 2001. The
contents of one mesh bag were processed immediately without field treatment.

Microscopy- Sections fiom greenhouse-inoculated plants were observed
using either a Zeiss Standard epifluorescence microscope or a Zeiss Pascal 5 laser
scanning confocal microscope with a 488 nm krypton laser (Jena, Germany). For
both microscopes, a 488 nm excitation filter was used to illuminate samples. For
the Zeiss standard microscope, along pass 520 nm barrier filter allowed for the
observation of host autofluorescence and the GFP simultaneously. Images
collected with the Zeiss Pascal 5 were collected in two channels. The first channel
had a long pass 560 nm barrier filter and the second channel had a band pass
barrier filter with a range of 505-530 nm. Images from the Zeiss Standard
microscope were collected using a Nikon Coolpix 995 (Tokyo, Japan). Images
from the Zeiss Pascal were reconstructed using Laser Scanning Microscope LSM
5 Pascal software, version 3.0 SP3. All images were transferred to Adobe
Photoshop v. 6.0. Images in this thesis are presented in color.

The surfaces of field-inoculated wheat stem samples were inspected for
perithecia and collected as perithecia formed. The vegetative portion of two plants

was collected at each time point. Sub-samples (intemode, leaf or node pieces 0.5

16

to 1 cm in length) were cut from each stem. Each stem that was processed
included a minimum of: three node sub-samples, eight intemode sub-samples,
four sheath sub-samples and three leaf blade sub-samples. The sub-samples were
placed in FAA (3.5% formaldehyde, 5% glacial acetic acid and 47.5% ethanol in
water) and vacuum infiltrated. After a 24 h fixation, sub-samples were placed in a
solution of glycerol and ethanol (1:1) for an additional 24 h. Sub-samples were
dehydrated through a tert-butanol series (O’Brien and McCully, 1981), embedded
in Tissue Prep paraffin wax (Fisher Scientific, Pittsburg, PA) and stored at 4°C
until sectioned. Embedded samples were sectioned to a thickness of 15 um using
a rotary microtome. Microscopic inspection of serial sections was conducted
using a Zeiss Standard microscope (model: GFL 654-633) with phase contrast
optics.

Host anatomy- For the purposes of this study, it was necessary to
consider each of the parts of wheat stems separately. For observations of stem
colonization, vegetation was divided in terms of culm, leaf sheath, leaf blade and
node tissue as shown in figure 1 (Leaf blade not shown). The cellular arrangement
differed or varied in each of these tissues as is well defined by Esau (1953). For
observations of perithecium development, stems were divided into node and
intemode tissue as defined by figure 1. The division of vegetation into node and
intemode was based on differences within the epidermis of wheat stems; namely,
intemode epidermis contains all examples of short cells including stomatal guard
cells whereas node epidermis contains only silica and cork cells. Both regions

contain long epidermal cells.

17

 

 

A B

Figure 1. Wheat stem anatomy. A, Outer surface of a wheat stem showing
intemode, node and sheath. B, Longitudinal section of wheat stem.

RESULTS

Colonization of wheat culms following head infections in the
greenhouse- The colonization of wheat stems following head inoculations was
extensive. While 97% of all inoculated heads became infected and exhibited
symptoms, these symptoms varied among the plants. In 72% of the inoculated
plants, the fungus moved down the rachis and the colonized culms below. In
general, the colonization progressed down the wheat culms throughout the time of
seed development and slowed as plants senesced and became completely dry (Fig.
2). All symptoms corresponded to the presence of the molecular marker GFP in
the fungal hyphae and this confirmed that the GFP strain, ZTE-Za, and not
contaminating fungi caused the infections. The GFP marker was detected in
hyphae growing down from the infected head , remaining at the infection front,
indicating that colonized stem tissue resulted only from head infections. Wheat
heads inoculated by PH—l (positive control) developed symptoms equivalent to
those inoculated by the ZTE-Za strain. Wheat heads inoculated with distilled

water (negative control) exhibited no symptoms.

19

Progression of Stem Colonization

 

 

 

 

 

70
R=.75

60 - y=yo+ax+bx2 O
S 50 -
“o
a)
13 40 -
C
o
6
O 30 -
.C
‘3’
0 20 —
_r
E
2 10 -
(D

0 -
‘10 l r T T I l
U 10 20 30 40 50 60 70

Days After Inoculation

Figure 2. Length of stem (cm) that had been colonized a given number of days
after susceptible wheat heads were inoculated with G. zeae. At time=0 days
wheat heads are flowering and at time=65 days wheat stems have completely
senesced.

20

It is interesting that in 25% of inoculated wheat plants symptoms remained
in the head. In these plants, the culm was not colonized and the fimgus either
remained in the inoculated spikelet or colonized the rachis downward infecting an
additional 1-2 spikelets. Upon examination, all plants in which culms were not
colonized contained vascular occlusions within the rachis below the inoculated

spikelet and the culm directly below the rachis.

Anatomy of colonization within wheat stems- The mode of colonization
of wheat stems, following head inoculations, was consistent throughout this study
for all plants observed and is summarized as follows. A single floret near the
middle of the wheat head was inoculated and head blight symptoms began to
appear within 4-5 days. Colonization continued down the rachis until the fimgus
entered the culm 10-12 days after inoculation. Hyphae penetrated culms through
xylem vessels within the large vascular bundles. At 12-14 days after inoculation
the diaphragm subtending the head was penetrated by hyphae within the rachis.
Colonization down the culm (vertical colonization) continued within the vessels
(Fig. 3) and pith cavity as hyphae extended away from the rachis. This pattern
was maintained at the infection front as the stem was colonized, making the
vessels and pith cavity the two major means of systemic colonization. At 14-16
days after inoculation older, established hyphae within the vessels and pith cavity
began to branch and colonize horizontally within culms. Horizontal colonization
was centrifugal away from vessels and pith cavity and began with the penetration

and infection of inter-cellular spaces within the xylem and fibers of the vascular

bundles and parenchyma near the pith cavity (Fig. 4). At 15-17 days after

21

inoculation hyphae began to penetrate parenchyma, xylem and phloem cells and
intra-cellular growth was initiated. Hyphae then began to grow from cell to cell
through pits and by direct penetration of adjacent cells (Fig. 5). Fungal
colonization continued outward toward the stem surface and the chlorenchyma,
just below the stem surface, was reached at 16-18 days after inoculation. Hyphae
filled sub-stomatal cavities of exposed culms but did not grow through stomates.
Heavily lignified fibers and cutinized epidermis were the last cell types to be

colonized and penetrated and then only occasionally.

22

 

Figure 3. Cross section of a large vascular bundle within a wheat culm. Primary
hyphae within a vessel (arrow) are colonizing the culm vertically. Scale bar = 20
um.

23

 

Figure 4. Cross section of a wheat culm. Horizontal colonization away
from vessels by primary hyphae. Scale bar = 50 um.

24

 

Figure 5. Horizontal colonization of parenchyma cells within the culm.
Primary hyphae penetrating adjacent cells (arrow) through pits and by
direct penetration. Scale bar = 20 um

25

Two distinct symptoms were observed on stem surfaces as stems were
colonized. Blighting or bleaching was first observed near the rachis at 14-16 days
after inoculation and were directly correlated with the initiation of horizontal
colonization. At 16-18 days after inoculation, brown streaks appeared on stem
surfaces that were simultaneous with the colonization of chlorenchymatous tissue.
This pattern of colonization was continued down the stems until the top-most
node was reached.

Nodes were first reached by hyphae vertically colonizing the pith cavity
and shortly after by hyphae vertically colonizing the space between the culm and
sheath. Hyphae between the culm and sheath were critical to the horizontal
colonization of the parenchyma and collenchyma cells within the leaf sheath base,
but did not continue to colonize vertically. Hyphae colonizing the pith cavity were
important to vertical colonization of nodes. The pith above the nodes was filled
with parenchyma cells and vertical colonization of stem nodes was initiated by
inter-cellular growth within these parenchyma cells. Inter—cellular growth
continued downward through parenchyma cells until the vascular traces were met
at the leaf joint. The xylem and phloem were both penetrated and vertical
colonization continued downward within the vascular bundles into the culm
below the node (Fig. 6). Horizontal colonization of parenchyma cells just below
the leaf joint lead to the eventual penetration of the pith cavity subtending the

node. The pattern of colonization within culms below nodes was very similar to

26

that below the rachis. All of the nodes observed in this study were colonized in

this manner.

27

 

Figure 6. Longitudinal section of node region. Primary hyphae are colonizing
downward from pith cavity into vascular tissues within the node.

28

The colonization of leaf sheaths was initiated as hyphae protruded through
culm stomates and filled the space between culm and sheath. Hyphae within this
space colonized vertically downward similarly to hyphae in the pith cavity.
Hyphae penetrated the adaxial surface of the leaf sheath and began inter-cellular
growth through epidermal cells and fibers. Horizontal colonization through sheath
mesophyll was both inter-cellular and intra-cellular in nature. Hyphae continued
to grow outward toward the sheath surface colonizing vascular bundles and filling
stomatal cavities. Vessels within the vascular bundles of the leaf sheath were
penetrated, however vertical colonization did not progress. Hyphae grew out of
stomates on the abaxial surface of the leaf sheath and were observed growing on
the surface.

Anatomy of colonizing hyphae— Hyphae colonizing wheat stems
following head infections were observed to have three distinct morphological
states. Primary hyphae were thin, regularly septate and uninucleate. These hyphae
occurred at the infection front in vessels and the pith cavity. Growth by primary
hyphae was both intra-cellular and inter-cellular throughout stem tissues and
remained in this state unless specialization occurred. Established primary hyphae
within the vessels, pith cavity and chlorenchyma developed into wider secondary
hyphae as they matured (Fig. 7). These hyphae were composed of short cells and
dikaryotic with paired nuclei. Secondary hyphae within the vascular bundles and
pith cavity branched to form the primary hyphae that initiated horizontal
colonization. Established secondary hyphae growing within the chlorenchyma

began to form ascogeneous initials below stomates (Fig. 8). The initials

29

specialized into stromatic hyphae that filled the stomatal cavities and grew into

chlorenchyma cells.

30

 

Figures 7—8. Hyphal morphologies observed within wheat stems. 7. Hyphae
colonizing the pith cavity. Both primary and secondary hyphae have developed.
8. Chlorenchyma tissue of the culm directly below a stomatal row. Stroma initials
(arrows) have formed below stomates and are connected by a secondary hyphum.
Scale bar = 20 um.

31

Development of perithecia in field-incubated tissue- Strain ZTE-Za
colonized stems readily however, this strain did not produce perithecia under
laboratory conditions and, due to its recombinant nature, could not be placed in
the field for perithecium development. Therefore, field-inoculated wheat stems
were collected and placed in the field for the development of perithecia.
Colonization patterns within field-inoculated stems were visually assessed before
and after perithecium development. While the process of colonization was not
observed for field-inoculated samples, the pattern ofcolonization in mature plants
was identical to that observed in greenhouse-inoculated plants as described above.
In stems, where perithecia formed, intracellular and inter-cellular colonization by
primary hyphae was observed throughout stem tissues as described previously.
Secondary hyphae had developed within the pith cavity, vascular bundles and
chlorenchymatous tissue. Stromatic hyphae formed within stomatal cavities
extending into chlorenchyma and also formed within parenchyma cells directly
underlying silica cells in the node regions. Infection fronts were not observed
within field-inoculated wheat stems, as the stems were fully colonized at
collection.

Perithecia developed on nodes and intemodes of all wheat stems recovered
after field treatment. On culms and leaf sheaths, perithecia developed only above
stomatal Openings (Figs. 9 and 11-12). Perithecia formed in association with silica
cells on nodes (Figs. 10 and 14). On all stem samples stroma formed prior to
perithecium formation. Leaf blades were not observed to support perithecium

development, but were colonized(not shown). Primary hyphae colonized

32

epidermal cells and mesophyll of the leaf blade. Secondary hyphae were not
observed and sub-stomatal cavities did not contain stroma. Genninating conidia
were observed on the leaf blade surface but the penetration process was not

obvious.

33

   

{i rill g: f
.4 r

‘5 .v

u.

‘

 

Figures 9—10. Development of perithecia on stem surfaces. 9. Outer surface of a
culm infected by G. zeae. Small black structures are perithecia erupting fiom
stomates. Perithecia emerge only through stomates, as seen here by their
development in rows. 10. Outer surface of a node region infected by G. zeae.
Perithecia were most dense on the upper ridge of the leaf base (arrow).

34

Perithecium development on sheathes was directly associated with
colonization and development of hyphae from underlying intemode tissue (Fig.
13). Where sheath tissue covered culm stomates, stromatic hyphae formed in the
culm sub-stomatal cavities, but hyphae extended through the stomatal pores to the
abaxial sheath surface. Hyphae then grew through the chlorenchyma cells of the
sheath and formed stroma in the sub-stomatal cavities of the adaxial sheath
surface. Perithecium development in intemode stern tissue occurred where
stomates were exposed and chlorenchyma cells were colonized.

Perithecium development on node tissue was dense in comparison to
development on intemode tissue. Perithecia formed from epidermal cells adjacent
to silica cells, but did not colonize silica cells (Fig. 14). Perithecia were most
dense on the ridges of the node, where silica cells were prevalent. Minimal
perithecium development occurred in the center of the thickened leaf base. Early
development of perithecia in nodal regions was initiated by the growth of
secondary hyphae within the epidermal and parenchyma cells. However the nodes
did not have stomates except at their boundaries. In the absence of stomates,
stroma formed within epidermal cells. Stroma within the epidermal cells were
much smaller relative to those formed in intemode tissue; however, the "knotted"
appearance was similar. Perithecia erupted from the epidermal cells or,
infrequently, formed from within a stroma that formed on the node surface.
Despite the presence of silica cells among the stomates on the intemode surface
perithecia were not observed emerging from epidermal cells adjacent to silica

cells.

35

 

Figures 1 1-12. 11. Arrows indicate stomates in the epidermis of the culm.
Mature perithecia have formed above the stomates; Scale bar = 100 um. 12.
Hyphal masses (arrows) within chlorenchyma tissue. Figs. 11 and 12 are
serial sections showing the same perithecia. The hyphal masses shown in
Fig. 12 developed below the stomates in Fig. 11 Scale bar = 100 um.

36

 

' x .
x ,\ .,
"f \f“; is) A?'.' xxk‘: -‘.
0‘ '

Figure 13.1ntemode and sheath supporting perithecium development by G.
zeae. Arrows indicate stomatal openings. Note that hyphae erupting from
intemode stomates subtending sheath, penetrate the adaxial surface of the

sheath. Perithecia develop through exposed stomates on the abaxial surface of
the sheath; Scale bar = 100 um.

37

i..." 3": ~v-rfir’r’ /Mf’v

I

 

Figure 14. Perithecium formation on the leaf- base ridge 1s dense and compact
and affected by placement of silica cells (arrows) within the epidermis.
Perithecia did not formed on regions adjacent to the ridge (Not shown); Scale
bar = 100 um.

38

DISCUSSION

The experiments conducted in this study provide the first documentation
of the colonization of wheat culms from anthesis through plant senescence.
Colonization of culms is extensive following head infections. This provides an
adequate explanation of how the vegetative portions of wheat plants are colonized
in the field prior to becoming debris. Straughsbaugh and Maloy (1986)
demonstrated that F. graminearum could be isolated from wheat culm segments
an average of 19 cm below infected heads in field studies. Unfortunately, they
failed to record the growth stage at which the segments were collected, making it
difficult to determine if the distance of stem colonized was representative of the
total amount of colonization possible prior to becoming debris. The extent of
culm colonization is important. Increased culm colonization directly affects the
potential for culms to become debris that may serve as a source of inoculum. It is
probable that field conditions would provide an environment that is suitable for
culm colonization, much the same as the greenhouse. A significant difference
between greenhouse and field conditions is rainfall events. In this study it was not
determined if wetting and drying events increase or decrease the extent of culm
colonized, but it is likely that it would increase sporulation.

The tissues colonized within the culm are also colonized within the head.
This shows that the process of colonization following head infection is continuous
from the spikelet and rachis through vegetative tissues of the wheat plant. The
vascular system plays an important role in this process. Within wheat culms

colonizing hyphae move through xylem vessels before colonizing other tissues.

39

Tu (1953), Ribichich et al. (2000) and Wanyoike et al. (2002) observed that the
vascular bundles leading from the spikelet to the rachis and the bundles of the
rachis were well colonized and hypothesized that the vascular system was
important to the colonization process. Involvement of the vascular system
throughout the colonization process indicates that this pathogen is mostly a
vascular pathogen of wheat plants. The phloem and chlorenchyma of the culm
were especially damaged as a result of colonization by G. zeae. Both tissues were
discolored and degenerated following fungal infection. Pugh et al. (1933) and Tu
(1953) reported similar damage in the same tissues of the rachis and glumes
within wheat heads. Sporodochia formed above stomates on cuhns prior to
treatment for perithecium formation. Pugh et al. (1933) reported sporodochium
formation at the base of spikelets and above stomates of the glumes. They also
observed that as mycelia massed within these tissues, sporodochia erupted
through the epidermis elsewhere. This was not observed at any point in this study.
In this study, 25% of all plants inoculated did not develop
symptoms below the infected head. Observations under the microscope confirmed
that the fungus had not colonized the culm but was present in the inoculated
spikelet. The inability of the fungus to colonize culms in these plants is likely due
to an unstable Type II (resistance to spread of infection) resistance mechanism
within the wheat cultivar Norm. Large variations in F HB severity have been
reported for cultivars with moderate resistance (Bai and Shaner, 1996). For each
of the plants in which culms were not colonized, vessel occlusions were present in

the rachis and the culm. Colonized culms did not contain occlusions. The

40

 

occlusions were clear and gel-like in appearance; similar to occlusions reported
for other vascular diseases (V anderMolen et al., 1983; Bishop and Cooper, 1984).
Ribichich et al. (2000) reported that vascular occlusions were present in the
highly resistant wheat cultivar Sumai-3 following inoculation, but were not
present in susceptible cultivars of wheat. Vascular occlusions are an important
component of resistance for many plants that are host to vascular pathogens
(Beckrnan and Roberts, 1995) and may also be an important component of
resistance to F HB for wheat.

The presence of secondary hyphae in chlorenchymatous tissue was an
expected result. Trail and Common (2000) observed secondary hyphae near the
surface of culture plates in regions where perithecia were forming. Similarly,
chlorenchyma tissue of the wheat culm is near the outer surface of the stem and in
regions where perithecia form. In the findings of this study and that of Trail and
Common (2000) secondary hyphae were uninucleate or binucleate and wider than
primary hyphae. The presence of secondary hyphae within the vascular system
and pith cavity of wheat culms was not an expected result as it was thought that
these hyphae were associated with perithecium development. It is more likely that
secondary hyphae are survival structures. The characteristic wide morphology of
these hyphae could imply storage in preparation of survival, and in fact Trail and
Common (2000) found that secondary hyphae contained many refractive drops
that stained positive for lipids.

The formation of perithecia is selective with regard to specific host cells.

However, the selection for stomatal openings, cork cells or silica cells must begin

41

 

with the associated formation of stroma; the developmental step preceding
perithecium formation. Factors inducing the initiation of stroma and perithecium
development are understood only on a gross environmental level that does not
easily explain an association with specific host cells. Light is required for the
formation of perithecia (Tschanz et al., 1976) and could be a factor that induces
the development of stroma and perithecia through stomatal openings and light
transmitting silica cells. Stomatal Openings are selected over silica cells when
both are present as in the case of culrn and sheath where perithecia only form over
stomates. This could be the easiest course or it could present the greatest light
stimulus. Perithecia did not form on leaf sheath blades where stomates and silica
cells are both present. A possible explanation for this lack of development could
be that the tissues of the leaf sheath blade are not colonized or only superficially
colonized. This would limit possibilities for survival of the fungal pathogen. The
tissues of the leaf sheath blade also break down quickly and therefore might not
support perithecium development.

The colonization of wheat vegetation prior to grain harvest in the field
increases the potential for infested debris that may act as a source of initial
inoculum in following seasons. Vegetation may become colonized by means of
stem-base infections and head infections, allowing the head blight pathogen to
establish itself prior to saprophytic invasion by other organisms. Strategies to
control or eliminate inoculum in the field should focus on slowing or reducing the
colonization of wheat vegetation or reducing sporulating structures on debris

surfaces. An obvious target for the reduction of vegetative colonization is Type II

42

resistance mechanisms. Vascular pathogen-induced resistance exists in wheat
cultivars and could be selectable in wheat breeding programs. Reducing
sporulation on debris will require a fitrther understanding of the factors that
initiate sporulation. Studies are currently underway to elucidate the molecular
basis for light intiation of perithecium development in planta. Further research is
being conducted to determine the source and timing of lipid acquisition by G.
zeae within the wheat host. An understanding of these processes will aid in
measures to reduce potential inoculum production and survival by this fungal

pathogen.

43

LITERATURE CITED

Arthur, J.C. 1891. Wheat scab. Indiana Agricultural Experimental Station Bulletin.
36: 129-132.

Atanasoff, D. 1920. Fusarium blight (scab) of wheat and other cereal crops. Journal
of Agricultural Research. 20: 1-32.

Bai, G-H. and G. Shaner. 1994. Scab of wheat: prospects for control. Plant Disease.
78:760-766.

Bai, G-H. and G. Shaner. 1996. Variation in Fusarium graminearum and cultivar
resistance to wheat scab. Plant Disease. 80:975-979.

Bai, G-H., R. Plattner, A. Desjardins, F. Kolb. 2001. Resistance to Fusarium head
blight and deoxynivalenol accumulation in wheat. Plant Breeding. 120:1-6.

Beckrnan, OH. and EM. Roberts. 1995. On the nature and genetic basis for
resistance and tolerance to fungal wilt diseases of plants. In Advances in
Botanical Research incorporating Advances in Plant Pathology, J .A. Callow, ed.
APS Press, St. Paul MN.

Bishop, CD. and RM. Cooper. 1984. Ultrastructure of vascular colonization by
firngal wilt pathogens. II. Invasion of resistant cultivars. Physiological Plant
Pathology 24:277-289.

Bushnell, W. R., B. E. Hazen, and C. Pritsch. 2003. Histology and physiology of
Fusarium head blight. In Fusarium Head Blight of Wheat and Barley, K. Leonard
and W. R. Bushnell, eds. APS Press. St. Paul MN.

Booth, C. 1971. The Genus Fusarium. Farnham, Commonwealth Agricultural
Bureaux. 23 7pp.

Caldwell, R.W. and J. Tuite. 1970. Zearalenone production in field corn in Indiana.
Phympathology. 60: 1 696- 1697.

Clement, J .A. and D.W. Parry. 1998. Stem-base disease and fungal colonization of
winter wheat grown in compost inoculated with Fusarium culmorurn, F.
graminearum and Microdochium nivale. European Journal of Plant Pathology.
104:323-330.

Dill-Macky, R. and R.K. Jones. 2000. The effect of previous crop residues and tillage
on F usarium head blight of wheat. Plant Disease. 84:71-76.

Esau, K. 1953. Plant Anatomy. Wiley & Sons, New York. 399pp.

Fernandez, M.R., and J .M.C. Femandes. 1990. Survival of wheat pathogens in wheat
andn soybean residues under conservation tillage systems in southern and central
Brazil. Canadian Journal of Plant Pathology 12:289-294.

Fernando, W.G.D., T.C. Paulitz, W.L. Seaman, P. Dutilleul, J.D. Miller. 1997. Head
blight gradients caused by Gibberella zeae from area sources of inoculum in
wheat field plots. Phytopathology. 87 :414-421.

Fernando, W.G.D., J .D. Miller, W.L. Seaman, K. Seifert, T.C. Paulitz. 2000. Daily
and seasonal dynamics of airborne spores of Fusarium graminearum and other

Fusarium species sampled over wheat plots. Canadian Journal of Botany. 78:497-
505.

Forsyth, D.M., T. Yoshizawa, N. Morooka, J. Tuite. 1977. Emetic and refusal activity
of deoxynivalenol to swine. Applied and Environmental Microbiology. 34:547-
552.

Francl, L., G. Shaner, G. Bergstrom, J. Gilbert, W. Pedersen, R. Dill-Macky, L.
Sweets, B. Corwin, Y. J in, D. Gallenberg, J. Wiersma. 1999. Daily inoculum
levels of Gibberella zeae on wheat spikes. Plant Disease. 83:662-666.

Jenkinson, P. and D.W. Parry. 1994. Splash dispersal of conidia of Fusarium
culmorum and F usarium avenaceum. Mycological Research. 98:506-510.

Joffe, AZ. 1986. F usarium species: their biology and toxicology. Wiley & Sons, New
York. 588 pp.

Kang, Z. and H. Buchenauer. 1999. Irnmunocytochemical localization of fuasriurn
toxins in infected wheat spikes by Fusarium culmorum. Physiological and
Molecular Plant Pathology. 55:275-288.

Khonga, EB. and J .C. Sutton. 1988. Inoculurn production and survival of Gibberella
zeae in maize and wheat residues. Canadian Journal of Plant Pathology. 10:232-
239.

Martin, R.A. and H.W. Johnston. 1982. Effects and control of Fusarium diseases of
cereal grains in the Atlantic Provinces. Canadian Journal of Plant Pathology.
4:210-216

Martins, ML. and HM. Martins. 2002. Influence of water activity, temperature and
incubation time on the simultaneous production of deoxynivalenol and
zearalenone in corn (Zea mays) by Fusarium graminearum. Food Chemistry.
79:3 1 5-3 1 8.

McMullen, M., R. Jones, D. Gallenberg. 1997. Scab of wheat and barley: A re-
emerging disease of devastating impact. Plant Disease. 81:1340-1348.

45

Mesterhazy, A. 1995. Types and components of resistance to Fusarium head blight of
wheat. Plant Breeding. 1 14:377-3 86.

Miller, J .D., J. Culley, K. Fraser, S. Hubbard, F. Meloche, T. Ouellet, W.L. Seaman,
K.A. Seifert, K. Turkington, H. Voldeng. 1998. Effect of tillage practice on
fusarium head blight of wheat. Canadian Journal of Plant Pathology. 20:95-103.

Mills, J .T. 1982. Development of fusaria and fusariotoxins on cereal grains in storage.
Canadian Journal of Plant Pathology. 4:217-218.

Neish, G. A., E. R. F arnworth, and H. Cohen. 1982. Zearalenone and trichothecene
production by some Fusarium species associated with Canadian grains. Canadian
Journal of Plant Pathology 4: 191-194

O’Brien, T. P. and M. E. McCully. 1981. The study of plant structure: principles and
selected methods. Terrnarcarphi Pty. Ltd., Melbourne.

Parry, D.W., P. Jenkinson, L. McLeod. 1995. Fusarium ear blight (scab) in small
grain cereals-a review. Plant Pathology. 44:207-238.

Paulitz, TC. 1996. Diurnal release of ascospores by Gibberella zeae in inoculated
wheat plots. Plant Disease. 80:674-678.

Pritsch, C., G]. Muehlbauer, W.R. Bushnnell, D.A. Somers, C.P. Vance. 2000.
Fungal development and induction of defense response genes during early
infection of wheat spikes by Fusarium graminearum. Molecular Plant-Microbe
Interactions. 13: 159-169.

Pritsch, C., C.P. Vance, W.R. Bushnell, D.A. Somers, T.M. Hohn, G.J. Muehlbauer.
2001. Systemic expression of defense response genes in wheat spikes as a
response to F usarium graminearum infection. Physiological and Molecular Plant
Pathology. 58: 1 ~12.

Pugh, G.W., H. Johann, J .G. Dickson. 1933. Factors affecting infection of wheat
heads by Gibberella saubinetii. Journal of Agricultural Research. 46:771-797.

Reis, EM. 1990. Survival of perithecia of Gibberella zeae on naturally infected
wheat kernels under field conditions. F itopatologia Brasilia 15:254-255.

Ribichich, K.F., S.E. Lopez, A.C. Vegetti. 2000. Histopathological spikelet changes
produced by Fusarium graminearum in susceptible and resistant wheat cultivars.
Plant Disease. 84:794-802.

Schroeder, H.W. and U. Christensen. 1963. Factors affecting the resistance of wheat
to scab caused by Gibberella zeae. Phytopathology. 53:831-838.

Seaman, W.L. 1982. Epidemiology and control of mycotoxigenic Fusaria on cereal
grains. Canadian Journal of Plant Pathology. 4: 187-190.

46

Shaner, G. and G. Buechley. 1992. Effect of foliar fungicides on wheat, 1991.
F ungicide and Nematicide Tests. 47 :206-207.

Siranidou, E., Z. Kang, H. Buchenauer. 2002. Studies on symptom development,
phenolic compounds and morphological defence responses in wheat cultivars
differing in resistance to Fusarium head blight. Journal of Phytopathology.
150:200-208.

Snijders, C.H.A. 1990. F usarium head blight and mycotoxin contamination of wheat,
a review. Netherlands Journal of Plant Pathology. 96: 1 87-198.

Stack, R.W. 1989. A comparison of the inoculum potential of ascospores and conidia
of Gibberella zeae. Canadian Journal of Plant Pathology. 11:137-142.

Strausbaugh, C. A. and O. C. Maloy. 1986. Fusarium scab of irrigated wheat in
central Washington. Plant Disease 70: 1104-1106.

Sutton, J .C. 1982. Epidemiology of wheat head blight and maize ear rot caused by
Fusarium graminearum. Canadian Journal of Plant Pathology. 4: 195-209.

Trail, F. and R. Common. 2000. Perithecial development by Gibberella zeae: a light
microscopy study. Mycologia. 92:130-138.

Trail, F., H. Xu, R. Loranger, D. Gadoury. 2002. Physiological and environmental
aspects of ascospore discharge in Gibberella zeae (anamorph F usarium
graminearum). Mycologia. 94:1 81 -1 89.

Tschanz, A.T., R.K. Horst, P.E. Nelson. 1976. The effect of environment on sexual
reproduction of Gibberella zeae. Mycologia. 68:327-340.

Tu, D.S. 1953. Factors affecting the reaction of wheat blight infection caused by
Gibberella zeae. Ph.D. Thesis, The Ohio State University, Columbus.

VanderMolen, G.E., J .M. Labavitch, L.L. Strand, and J .E. DeVay. 1983. Pathogen-
induced vascular gels: Ethylene as a host intermediate. Physiological Plant
Pathology 59:573-580.

Vesonder, RF. and CW. Hesseltine. 1981. Vomitoxin: natural occurrence on cereal
grains and significance as a refusal and emetic factor to swine. Process
Biochemistry. Dec/Jan. 1980/81 :12-15, 44.

Wang, Y.Z. and JD. Miller. 1988. Effects of Fusarium graminearum metabolites on
wheat tissue in relation to F usarium head blight resistance. Journal of
phytopathology. 122: 1 18-125.

Wanyoike, M.W., Z. Kang, H. Buchenauer. 2002. Importance of cell wall degrading
enzymes produced by F usarium graminearum during the infection of wheat heads.
European Journal of Plant Pathology. 108:803-810.

47

 

Wiese, M.V. 1987. Compendium of wheat diseases. The American Phytopathological
Society press, St. Paul: 112pp.

48

 

   

i111i;giiiiijiijiiuri