Will

I
\

ill

  

§§§ IIHNIWHWIWWWIHUHIJIWIIWINN

THESE

\{3

(\

ICHIGAN STATE WWE 8 TY LIBRARIES L

J Ill/1W”!!!IllIll!l/llI/llllll/lllill/[Wml

3 1293 02058 9994

 

 

This is to certify that the

thesis entitled

Ascospore Discharge and Perithecium
Development in Gibberella zeae.

 

presented by
Corrie L. Andries

has been accepted towards fulfillment
of the requirements for

Masters Science

degree in

 

 

\ Major professor

Date 5,1“ loo

0-7639 MS U i: an Affirmative Action/Equal. Opportunity Institution

 
 

ttBnAnY
M’Chigan State
University

    
    

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

 

DATE DUE DATE DUE DATE DUE

 

afij
'0 I! n 0 2: ‘ I,
._.' b ‘ “

U ‘1'”-

 

MAY 2 5 2007

 

 

 

 

 

 

 

 

 

 

 

 

 

moo chlRC/DneOmpss-p.“

 

ASCOSPORE DISCHARGE AND PERITHECIUM DEVELOPMENT IN
GIBBERELLA ZEAE

By

Corrie L. Andries

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

2000

ABSTRACT

Fusarium Head Blight (FHB), caused by Gibberella zeae, has incurred billions of
dollars in yield losses to wheat and barley growers from 1993 to 1999. The fungus
overwinters in crop debris and undergoes sexual reproduction to form perithecia that
contain ascospores. These ascospores are forcibly discharged and deposited on the
developing grain head during flowering. I conducted two studies: a field study to
pinpoint the timing of perithecium formation in the field on crop debris, and a random
mutagenesis project to understand the mechanism of forcible discharge of ascospores.

A total of 2186 samples of infected corn and wheat stubble were collected monthly
from 1997 to 2000. Of these, 1532 samples were com debris and 654 samples were
wheat debris. A total of 241 com debris samples (15%) and 31 wheat debris samples
(3%) had perithecia of G. zeae. The number of perithecia present was highest in May,
June, and July. Average temperatures at 4 days and at two-weeks prior to sample
collection were correlated with the presence of perithecia.

In a separate study, 5005 transformants were generated with random insertional
mutagenesis of a plasmid harboring the hygromycin resistance gene. Transformants were
screened for a loss of the ability to forcibly discharge ascospores. Southern blot analysis
indicated the insertion of the vector was random and remained stable through meiosis and
mitosis. Four mutants were found that were unable to forcibly discharge their
ascospores. One of these mutants (designated 729G-112) appeared morphologically
normal and the other three had mutations associated with the development of mature

perithecia. Isolate 729G-112 also exuded irregularly shaped cirrhi.

ACKNOWLEDGEMENTS
I would like to thank the following people for their guidance and assistance while
completing my masters project and thesis: Dr. Frances Trail, Ifi‘a Gaffoor, Dr. John Linz,
Dr. L. Patrick Hart, Dr. Andy Jarosz, David Johnson, Luis Velasquez, Cheryl Calhoun,
Rachel Loranger, Dr. Haixin Xu, Dr. Seanna Annis, Dr. John Halloin, Jennifer Schaupp,
Dr. Gerry Adams, Dr. John Scott-Craig, and Kerry Pedley.
I would also like to thank my husband, Erik Andries and my parents, Kathy and John
Jacomino, my nana, Kathryn Kiser, and my sister, Kimberly Jean Platt for their support

and encouragement throughout my education.

iii

TABLE OF CONTENTS

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

LIST OF TABLES -- v
LIST OF FIGURES - vi
ABBREVIATIONS . ...... 1
INTRODUCTION 1
CHAPTER 1
PERITHECIA SURVEY OF GIBBERELLA ZEAE - 11
INTRODUCTION 11
MATERIALS AND METHODS 13
RESULTS 17
DISCUSSION -21
CHAPTER 2
USE OF RANDOM INSERTIONAL MUTAGENESIS TO STUDY THE
MECHANISM OF ASCOSPORE DISCHARGE IN GIBBERELIA ZEAE ............. 23
INTRODUCTION 2?
MATERIALS AND METHODS 26
RESULTS 10
DISCUSSION 18
APPENDICES 40
APPENDIX A
Plasmid Maps: pHA1.3 and pUCH2-8 - .41
APPENDIX B
Protocol for Random Insertional Mutagenesis ---.-44
APPENDIX C
Protocol for Large Scale Random Insertional Mutagenesis ........ 48
APPENDIX D
Protocol for Screening of Discharge Minus / Perithecia Mutants 52
BIBLIOGRAPHY ..... -- ...... 54

iv

LIST OF TABLES
Table 1. Variables for regression analysis.
Table 2. Field data summary.
Table 3. Effect of temperature on development of perithecia.

Table 4. Segregation analysis of mutations.

LIST OF FIGURES
Figure 1. Disease cycle of Gibberella zeae (F usarium graminearum)
Figure 2. Sampling pattern for crop debris collection
Figure 3: Timing of perithecium production on corn debris samples 1997 — 1999
Figure 4. Proportion of perithecia in relation to average 14-day temperature.
Figure 5. Southern hybridization analysis G. zeae transformants.
Figure 6. Southern hybridization analysis of pHAl .3 transformants.
Figure 7. Southern hybridization analysis of pUCH2-8 transformants.
Figure 8. Perithecia of mutants 729G-112, 720R-2 and wild type PH-l X 400.
Figure 9. Perithecia and cirrhi of 720R-2, 729G-112 and wild type PH-l.
Figure 10. Map of plasmid DNA pHA1.3.

Figure 11. Map of plasmid DNA pUCH2-8.

vi

ABBREVIATIONS

CA: carrot agar

CMC: carboxymethylcellulose medium

Dis': discharge minus

Dish wild type for discharge

DON: deoxynivalenol

FHB: Fusarium Head Blight

Hng: hygromycin resistant

Hy : hygromycin sensitive

Kb: kilobase

MMTS: minimal medium with Tergitol Type NP- 1 0
Mt: nitrate non-utilizing, auxotrophic

Nit“: nitrate utilizing

PEG: polyethylene glycol

REM]: restriction enzyme mediated integration
RH: relative humidity

RIM: random insertional mutagenesis

RM: regeneration medium

RT: room temperature

Tx: transformants

VM: V8 juice medium

WA: water with 2% agar

YEPD: yeast extract / peptone / dextrose broth
YES: yeast extract/ sucrose broth

INTRODUCTION
Disease overview. F usarium Head Blight (FHB) or scab of wheat (Triticum aestivum
L.) has reached epidemic proportions in the United States over the last ten years. The
causal agent, Gibberella zeae (Schwein.) Petch (anamorph: F usarium graminearum
Schwabe), also causes seedling blight, ear rot of maize and affects other small grain
cereals such as barley and rye. The US. Wheat and Barley Scab Initiative reports that
FHB has incurred over 4 billion dollars in yield losses to wheat and barley growers

between 1993 and 1999 (McMullen, 1999). Yield losses in wheat occur primarily as a

result of sterile florets and diminished, low weight kernels (McMullen, 1999). Infested

wheat kernels that are harvested, may produce a poor flour quality with a lowered protein
content (Parry et 01., 1995). Barley infected with G. zeae can result in uncontrolled
“gushing” during beer brewing which has led the brewing companies to establish “zero
tolerance” levels for F HB (Parry et al., 1995). Gibberella zeae also produces mycotoxins
including deoxynivalenol (DON) and zearalenone. DON or vomitoxin, is a trichothecene
produced in diseased kernels that causes vomiting and feed refusal in livestock. The
Food and Drug Administration (FDA) has limited levels of DON in grains to l, 5, and 10
ppm for human, swine, and cattle consumption, respectively (McMullen, 1999).
Zearalenone, an estrogenic compound, has been shown to cause reproductive disorders
when fed to young female pigs (Long et al., 1982). There is also evidence it can
accumulate in milk from dairy cattle (Kuper—Goodman, 1987). The National Veterinary
Services Laboratories at United States Department of Agriculture, Animal and Plant
Health Inspection Service (USDA, APHIS) has advised that detectable levels of
zearalenone in grains remain below 1 and 10 ppm, for swine and cattle consumption,
respectively (Stanton, 1995).

The disease cycle of F HB (Figure 1) has been well described. Gibberella zeae
reportedly overwinters as saprophytic mycelia on crop debris of maize, wheat, and
barley. The fungus undergoes sexual reproduction and produces ascospores in a fruiting
body known as a perithecium. Ascospores are forcibly discharged from perithecia and
become airborne. The spores are deposited on the wheat head sometime during flowering
(Paulitz, 1996; Sutton and Procter, 1982) when susceptibility is highest (Fernando et al.,
1997). Ascospores serve as the primary source of inoculum, although the asexual

macroconidia may also produce some localized infection due to wind and splash dispersal

(Fernando et al., 1997; Parry et al., 1995; Sutton, 1982). The hyphae then penetrate the
glume stomata (Pritsch, 2000), and spread throughout the wheat head resulting in
chlorosis of infected kernels (Parry er al., 1995; Sutton, 1982). Favorable conditions for
disease outbreaks appear to coincide with extended periods of high relative humidity
(RH) and warm temperatures (24 - 29°C). (McMullen et al., 1997a). If warm, wet
conditions are prolonged, secondary infection can occur from production of ascospores
and macroconidia on infested wheat plants (McMullen et al., 1997b).

Control of FHB has been problematic. Currently, there are no known wheat or barley
cultivars entirely resistant to FHB and most are highly susceptible (Parry et al., 1995).
Durum wheat and barley are the most susceptible, but all classes of wheat, including
spring and winter varieties, are affected (McMullen et al., 1997a). Four types of
resistance have been described: inhibition of initial infection (Type I), limiting the spread
of infection throughout the host (Type II) (Schroeder and Christensen, 1963), ability to
degrade DON (Type III), and tolerance of high DON concentrations (Type IV) (Parry et
aL,1995)

Chemical seed treatments can inhibit seedling blight, but do not prevent F HB
(McMullen, 1999). Currently, an effective fungicide or biocontrol agent that can
consistently control FHB outbreaks has not been established. Delivery of fungicides is
also difficult because the disease develops inside the grain head (Parry et al., 1995) and
food safety and economic considerations limit the frequency of application (McMullen et
al., 1999). Even when fungicides achieve some reduction of disease severity, high levels

of DON may still be present (Moss and Frank, 1985). A disease forecasting system is

under development but its efficacy in predicting disease outbreaks has not been
established (McMullen et al., 1997a).

Crop rotation and tillage practices have an impact on the severity of F HB. Incidence
of F HB increases when wheat is planted immediately after maize or cereals (wheat,
barley, oats) as opposed to soybean or other non-host crops (McMullen, 1999; Teich and
Hamilton, 1985; Teich and Nelson, 1984; Tusa et al., 1981). Tillage practices that bury
crop debris have been shown to be effective in reducing levels of primary inoculum
(Teich, 1989). However, the effectiveness of these cultural controls is variable and is it
not always possible due to the practice of soil conservation tillage and grower
commitments to the USDA to produce specific crops (Bai and Shaner, 1994; McMullen
et al., 1997a; Miller et al., 1998; Parry et al., 1995). We believe study of the causal
organism, specifically, the mechanism of forcible discharge of ascospores and the timing
of perithecium formation in field conditions, may lead to a better understanding of the
spread of this disease and lead to novel methods for control.

Timing of perithecium formation in the field. Fungi in the Phylum Ascomycota
are characterized in Ainsworth and Bisby ’3 Dictionary of the Fungi (Hawksworth et a1 .,
1996) by the production of ascospores following meiosis. Ascospores are contained
within sac-like cells known as asci. Certain ascomycetes bear asci in flask-shaped
fruiting bodies called perithecia. Gibberella zeae is a member of Hypocreaceae De. Not,
a family whose members are characterized by brightly colored perithecia (Alexopoulos et
al., 1996). Members of Hypocreaceae, along with those of Nectriaceae and
Clavicipitaceae are all able to forcibly discharge their ascospores when mature. The

morphological development of perithecia of G. zeae has been described in detail (Trail

and Common, 2000). First, the vegetative hyphae start to enlarge and develop into a
perithecium body. Inside the perithecia, sterile hyphae, termed paraphyses begin to grow
from the top and later attach to the base. Another type of sterile hyphae, called
periphyses, form inside the developing ostiole. Once these periphyses and paraphyses
mature, they will collapse, creating open space in the perithecia. The collapse of the
periphyses creates the opening in the ostiole and the shrunken paraphyses leave an area
for the maturing asci. The nucleus within the developing ascus undergoes meiosis,
followed by one sequence of mitosis, which creates eight nuclei. These nuclei then
become delimited by the formation of ascospore walls (Trail and Common, 2000). The
mature perithecia are bright blue-black, globose in shape, with asci that contain eight, one
to three septate ascospores in a biseriate arrangement.

Gibberella zeae is subdivided into two groups, Group 1, which does not form
perithecia in culture and seldom forms them in the field, and Group 2, which easily forms
perithecia under both conditions (Burgess et al., 1975; Francis and Burgess, 1977).
Khonga and Sutton have found that G. zeae can form perithecia and ascospores for up to
three years in the field on infected wheat and corn debris; two years longer than
macroconidia formation. This may be related to the nutrient content (high C:N ratio) at
later stages of decay being more conducive to perithecium than macroconidium
production (Khonga and Sutton, 1988). Formation of perithecia appear to be primarily
limited by exhaustion of the host substrate and competition with other organisms (Pereyra
et al., 1999). Although these long-term studies have helped to understand the longevity
of perithecium formation, little is known about what triggers perithecia] formation in the

field.

Perithecium formation in G. zeae and other fungi appear to favor surfaces that are
exposed to direct light (El-Gholl et al., 1979; Halama and Lacoste, 1992; Khonga and
Sutton, 1988; Reis, 1990b). It has also been suggested that near ultra-violet (UV) light
and RH may be a significant factor in triggering perithecium development in G. zeae and
other fungi (Halama and Lacoste, 1992; Paulitz, 1996). Under laboratory conditions, a
perithecium typically forms and discharges its ascospores within seven days of induction
(Trail and Common, 2000). As part of this study, we collected wheat and corn stubble
over the course of year, examined it for the presence of mature perithecia, and compared
it to local weather data to determine if any correlation exists between perithecium
formation and environmental parameters.

Forcible discharge of ascospores. Ingold has described the physiological events
of ascospore discharge in detail. First, the mature ascus extends its tip through the ostiole
and then rapidly ejects the ascospores. Once an ascus has been emptied, it collapses back
into the perithecium body and the next ascus moves into place and fires its spores. This
is repeated until all asci have been emptied (Ingold, 1971). Three mechanisms have been
proposed to explain how an ascus builds up the necessary force to eject its ascospores:
force induced by explosion of a gas-bubble inside the ascus, a spring mechanism
involving microtubules, and the bursting of turgid ascus which forces ascospores through
its narrow opening.

It was first proposed that a gas-filled bubble might build up enough pressure to
forcibly discharge ascospores (Olive, 1964). However, it was later found that ascospore
discharge in a species of Sordaria was not influenced by changes in air pressure but wase

primarily affected by moisture (Ingold and Dann, 1968).

In a transmission electron microscopy study, Czymmek and Klomparens found an
extensive microtubule network in the epiplasm of mature ascospores of Thelebolus
crustaceus. Since microtubules have been implicated in active movement of cells and
their organelles, they suggested these microtubules could contract, drawing the
ascospores together, and eject the ascospores through a spring mechanism (Czymmek and
Klomparens, 1992). It has also been found that benomyl, which inhibits microtubule
formation, reduces the discharge of ascospores in Venturia inaequalis (Miller, 1970).

The idea that turgor pressure is behind forcible discharge was proposed by CT
Ingold (1968). He observed a stretching of the ascus wall in Loramyces sp. as it built up
osmotic pressure under moist conditions. This stretching eventually caused the ascus to
burst and subsequently contract, which forced the ascospores out of a narrow slit at the
top of each ascus (Ingold, 1968). In a field study of Venturia inaequalis, a relationship
was found between leaf surface wetness and the mean distance ascospores were fired. A
correlation between increased ascospore discharge and high RH or rainfall has been
observed in several fungi Venturia inaequalis (Gadoury et al., 1998) (Stensvand et al.,
1998), Anisogramma anomala (Pinkerton et al., 1998), and Pleospora allii (Prados
Ligero et al., 1998). Some early studies showed a similar correlation in G. zeae (Chen
and Yuan, 1984; Reis, 1990a). However, a later study by Paulitz (1996) did not find a
definitive correlation between ascospore discharge by the perithecia of G. zeae and
moisture in the field. During periods of heavy rainfall, ascospores were not forcibly
discharged, but instead oozed out of the ostiole in a mucilaginous spore-tendril, or cirrus.
However, since microclimates on the surface of the debris were difficult to evaluate, he

concluded that high moisture levels were still important for perithecium and ascospore

formation, and RH may play a role in discharge in addition to cirrhi production. (Paulitz,
1996). Under laboratory conditions, it has also been observed that although discharge
occurred at low RH levels, the maximum number of spores were released in 100% RH
conditions (Trail et al., 1998).

Since many fungi are known to actively discharge their spores, determining the
mechanism behind this phenomenon would lead to an increased understanding of the
physiology of spore dissemination. The role of environmental parameters is difficult to
determine in fungi, due in part, to microclimates that exist around a perithecium on its
host. Physiological study of the thin-walled, delicate asci are also problematic. Instead,
we have chosen to study this mechanism by disabling ascospore discharge through a
random gene disruption process, and study phenotypic changes in vitro. Based on current
studies, some mutant phenotypes that might affect discharge would be the following:
inability to build up osmotic pressure in the ascus, limited microtubule formation,
changes in composition of the epiplasm of ascospores, and morphological mutations of
the paraphyses, ostiole, or asci.

Molecular study of ascospore discharge. Random insertional mutagenesis can be
useful for a large-scale search for genes involved in a specific mechanism. A large
number of tagged transformants can be generated and then screened for a change in the
phenotype of interest. In this study, a screening method was developed that quickly
distinguished isolates that had lost their ability to forcibly eject their spores. Mutants
showing a change in this phenotype could then be studied in further detail.

Random mutagenesis had been traditionally performed by exposure of the organism

to UV light or a mutagenic chemical. Ultra-violet (UV) mutagenesis has been used for

G. zeae as an effective method of generating large numbers of mutants (Leslie, 1983).
However, UV and chemical mutagenesis do not leave any molecular tag indicating the
location of the affected gene(s). Recombinant DNA technology has currently become a
powerful tool in molecular research. This technology allows for integration of foreign
DNA into the host genome, leaving a molecular marker at a specific gene. Mechanical
methods of fungal cell wall disruption such as the gene gun or biolistics (Armaleo et al.,
1990; Lorito et al., 1993) (Durand et al., 1997) (Yu Jieh and Cole Garry, 1998), and
electroporation (Chakraborty et al., 1991; Redman and Rodriguez, 1994), have been
successfully used to allow incorporation of foreign DNA. Polyethylene-glycol (PEG)
mediated transformation of protoplasts has also been shown to achieve an increased rate
of transformation, particularly when coupled with restriction enzyme mediated
integration (REMI) in some fungi (Brown et al., 1998; Redman and Rodriguez, 1994).
Dissolution of the cell wall occurs during enzymatic digestion followed by the uptake of
foreign DNA in a PEG solution (transformation). Foreign DNA is linearized with
restriction enzymes prior to transformation. REMI transformation, originally developed
for Dictyostelium discoidium (Kuspa and Loomis, 1992), uses additional restriction
enzymes to nick the host DNA. This is thought to facilitate insertion of the linearized
plasmid. The REMI system has been successfully used to isolate desirable mutants in
many fungi including Saccharomyces cerevisiae (Schiestl and Petes, 1991), Ustilago
maydis (Bolker et al, 1995), Cochliobolus heterostrophus (Lu et al., 1994), Magnaporthe
grisea (Shi et al., 1995; Sweigard et al., 1998), Aspergillus nidulans (Sanchez et al.,

1998), and Penicillium paxilli (Itoh and Scott, 1997).

     
   

AEaEmESEM 53.2335 moon SESAEG mo 298 0335 ; oSwE

omega 3 8:383

”can. 8 83.5. 528.22 .8... _

2.8: 03.20»?
a we 8338.00

 

10

CHAPTER 1
PERITHECIA SURVEY OF G. ZEAE
INTRODUCTION

Fusarium Head Blight (FHB) or scab of wheat (Triticum aestivum L.), caused by
Gibberella zeae (Schwein.) Petch (anamorph: F usarium graminearum Schwabe), has
reached epidemic proportions in the United States over the past decade. Fusarium Head
Blight has incurred over 4 billion dollars in yield losses to wheat and barley growers in
the US. between 1993 and 1999 (McMullen, 1999). Yield losses in wheat occur
primarily as a result of sterile florets and diminished, low weight kernels (McMullen,
1999). Gibberella zeae also produces the mycotoxins deoxynivalenol (DON) and
zearalenone that cause vomiting, feed refusal, and reproductive disorders when
contaminated grain is fed to livestock (McMullen, 1999; Stanton, 1995).

The species is subdivided into two groups: Group 1, seldom forms perithecia in the
field and does not form them in culture; Group 2, easily forms perithecia under both
conditions (Burgess et al., 1975; Francis and Burgess, 1977). The disease cycle of FHB
has been well described (Parry et al., 1995; Sutton, 1982). Gibberella zeae overwinters
as saprophytic mycelia on the crop debris of maize, wheat, and barley. The fungus
undergoes sexual reproduction and produces ascospores in a fruiting body known as a
perithecium. Ascospores are forcibly discharged from perithecia and become airborne.
The spores are deposited on the wheat head sometime during flowering (Paulitz, 1996;
Sutton and Procter, 1982) when susceptibility is highest (Fernando et al., 1997).
Ascospores serve as the primary source of inoculum, although the asexual macroconidia

may also produce some localized infection due to wind and splash dispersal (Fernando et

11

al., 1997; Parry et al., 1995; Sutton, 1982). The hyphae then penetrate the glume stomata
(Pritsch, 2000), and spread throughout the wheat head resulting in chlorosis of infected
kernels (Parry et al., 1995; Sutton, 1982). Favorable conditions for disease outbreaks
appear to coincide with extended periods of high relative humidity (RH) and warm
temperatures (24 - 29°C). (McMullen eta1., 1997a).

An effective and consistent method to control FHB has not yet been established
(Parry et al., 1995). Crop rotation and tillage, when employable, can reduce the severity
of FHB. Incidence of F HB increases when wheat is planted immediately after maize or
cereals (wheat, barley, oats) (McMullen, 1999; Teich and Hamilton, 1985; Teich and
Nelson, 1984; Tusa et al., 1981) as compared to a non-host crop. Deep tillage that can
bury crop debris has been shown to be effective in reducing levels of primary inoculum
(Teich, 1989).

Gibberella zeae can form perithecia and ascospores for up to three years in the field
on infected wheat and corn debris; two years longer than macroconidia formation
(Khonga and Sutton, 1988). Formation of perithecia appear to be primarily limited by
exhaustion of the host substrate and competition with other organisms (Pereyra et al.,
1999). Although these long-term studies have helped to understand the longevity of
perithecium formation, little is known about what environmental triggers may induce
formation. Perithecium formation in G. zeae and other fungi appears to favor the light-
exposed surfaces of plant debris (El-Gholl et al., 1979; Halama and Lacoste, 1992;
Khonga and Sutton, 1988; Reis, 1990b). It has also been suggested that day length, light
intensity, wavelength and relative humidity (RH) may all be significant factors in

triggering perithecium development in G. zeae and other fungi (Halama and Lacoste,

12

1992; Paulitz, 1996). Under laboratory conditions, perithecia typically form and
discharge ascospores within 7-14 days of induction (Trail and Common, 2000).

Knowledge of the timing of formation of perithecia in the field with respect to
flowering of wheat would be important in designing novel control methods. To evaluate
the timing of perithecium formation in the field, we collected wheat and corn stubble
from commercial fields throughout the year from 1997 to 2000. Additionally, we
analyzed the timing of perithecium formation to determine if it positively correlated to
local temperature and rainfall. Based on previous research, our hypothesis was that the
timing of perithecium formation coincides with the cool, wet weather conditions that that
typically occur near the time of wheat flowering.

MATERIALS AND METHODS

Field collection. Between June 1997 to March 2000, thirteen commercial fields were
surveyed that had experienced previous outbreaks of F usarium Head Blight (FHB) in
lngham County, Michigan. The traditional crop rotation scheme is com, soybean, and
wheat, however rotation patterns often varied. Fields containing wheat or corn stubble
from the previous year’s harvest were sampled (Table 2). A decreasing number of fields
were sampled in 1999 and 2000 due to progressive replacement of wheat by soybean in
the crop rotation scheme.

Debris samples that had visible perithecia were collected monthly unless impeded by
snow cover. During the period of wheat flowering, sampling was increased to two-week
intervals. Eight locations were sampled along transects in each field (Al-D2) in a
diamond pattern (Figure 2). The length of transects were scaled to the acreage of each

field. Two to four samples exhibiting symptoms of infestation were collected from each

13

location. Samples were stored in plastic bags at —20° C. Daily weather data, from the
nearest National Weather Service (N WS) station at the Lansing Capitol Airport, was
obtained from the National Oceanic and Atmospheric Administration National Climatic
Data Center (NOAA-NCDC) database (N OAA-NNDC, 2000). All fields were within a
60-mile radius of the NWS station.

Identification. Corn and wheat stubble were microscopically examined at 70X
magnification for the presence or absence of perithecia. Species identification was
confirmed by perithecium wall structure, color and ascospore morphology at 400X
magnification as described by Nelson et al. (1983). Samples harboring one or more
developing perithecia of G. zeae were scored as positive.

Data Analysis. Data was expressed as a proportion of samples with perithecia per
total samples collected. The proportion of samples with perithecia was compared to
calendar days. Wheat stubble samples were not included in analyses due to the low
number of mature perithecia.

The distribution of proportion data was normalized with the arcsine square root
transformation. All analyses were performed using proc REG of SAS version 7.0 (SAS
Institute, Cary, NC.) Single factor regression analysis was used to evaluate the effect of
rainfall and temperature. A stepwise regression analysis for all variables was used to
determine if combined variables increased correlation. Variables used are defined in

Table 1.

14

Table 1. Variables for regression analysis.

 

Label Time period included

 

T4 Average daily temperature 1 — 7 days preceding sampling
T7 Average daily temperature 4 — 10 days preceding sampling
T14 Average daily temperature 11 — 17 days preceding sampling
R4 Average daily rainfall 1 — 7 days preceding sampling

R7 Average daily rainfall 4 — 10 days preceding sampling

R14 Average daily rainfall 11 — 17 days preceding sampling

 

15

 

 

 

 

Figure 2. Sampling pattern for crop debris collection. Outer square indicates field
perimeter, diamond indicates sampling transect. Letter and number notations locate
sampling locations along the transect.

16

 

RESULTS

Of the 2186 samples collected, 272 were found to have perithecia of G. zeae (Table

1). Approximately 83% of the samples with perithecia were found on corn debris [(241

of the 1532 corn debris samples or 15%) (31 of the 654 wheat debris samples or 3% had

perithecia)]. Over the three years sampled, the highest proportion of samples with

perithecia were found during the summer months, with the highest number in May, 1997

and 1998 and July, 1999 (Figure 3).

Table 2. Field data summary.

 

 

 

 

 

Sampling Average No. No. Total No. Total No. Corn Wheat
year No. of Com Wheat corn wheat samples samples
fields debris debris samples samples w/ w/
fields fields perithecia perithecia
Spring 1997 10 8 2 320 80 31 l
- Fall 1997
Fall 1997- 11 7 4 368 312 40 24
Fall 1998
Fall 1998- 9 7 2 684 200 161 6
Fall 1999
Fall 1999- 8 5 2 160 62 9 0
Winter 2000
TOTAL 38 27 10 1532 654 241 31

 

 

 

 

 

 

 

 

 

 

 

In single variable regression, variables T4 and T14 were found to have the highest
correlation with the proportion of corn debris samples with perithecia (Table 2).
Temperatures below 9° C appear to inhibit perithecium formation (Figure 4). If data
points of T14 below 9° C are removed from the analysis, the R2 of T14 increases to 0.75.

Table 3. Effect of temperature on development of perithecia.

 

 

Condition at days prior F value P8 > F R‘ Adjusted R‘Z
to collection

T4 12.290 0.0017 0.2949 0.2949
T14 13.168 0.0012 0.3107 0.3107
T4 and T14 days 7.42 0.0030 0.3725 -

T14 using 9° C 50.866 0.0001 0.7495 0.7348
threshold

 

‘ Regressions were significant at P < 0.05

17

Stepwise regression analysis showed that inclusion of all other variables (T7, R4, R7,
R14) in the T4 and T14 model improved the R2 (including data below the 9° C threshold)
to 0.4811, but no individual variable had a large effect on the predictivity of the model.
Combination of T4 and T14 with a stepwise regression analysis showed that R2 was only

slightly improved, indicating that the variables may covary.

18

mag - 32 SEES £526 88 no 832603 Saoofitom mo wEEE. ”m oSwE

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

   

 

 

 

 

 

 

 

 

 

 

 

€333 $33 32030
W. m m. m m m m. m m m.
E illluql' I ooo.o
8Q
5- -11 - oo_.o
- | - 5 SS
snow
l 1 - 1 1 . ,. .. oomd
.82
1 11 111... 151i- .>. s .- 2:3
m -. 0:3. «2
oamfifl 0:3.
702- .3... 8.5 g 82
‘ 3.3 I mutate:— .3.—>9
1 - 1 -1- 1 1 -- 11 l- -- .1 11 11. coed
I l- 11 1|| l ;1- 1.83
.22

 

 

 

oowd

egoaqrpad qrtm saIdures go uoruodoxd

l9

om

883389 amp 3 owmuozw 8 notice 5 floofitom mo comtomoi .v 23.5

ADV EBEoQEoH b5 E 0353..

mm om 2 2 m

 

 

 

Sod

oomo

oomd

_..__. .. ___*f . 4

 

oofio

oovd

_' __ l _ I—

- cemd

oocd

 

oond

-__~ __Y.
1

D. cows

(1210150 uotuodord) eroaqruad qitm saldumg

20

DISCUSSION

In the study area, corn stubble was the predominant substrate for perithecium
formation of G. zeae (83% of total debris samples with perithecia). However, the disease
outbreak of FHB in wheat fields was minimal in 1998 and 1999 due to hot, dry
conditions during the spring (Hart, 1998; 1999). Weather conditions that decreased
colonization of the wheat plants would have resulted in lower numbers of perithecia on
wheat stubble.

Average daily temperatures below 9° C appeared to limit perithecium formation at
T14, and temperatures above 9° C at T4 and T14 were found to have a linear correlation
to the proportion of perithecia present. Combining these two factors in a stepwise
regression only increased the R2 slightly, which indicates a covariance. The covariance
of these two factors indicates they may both be necessary for perithecium formation.
Mature perithecia of G. zeae form within 7-14 days following induction under laboratory
conditions (Trail et al., 1998). The data did not show a limiting high temperature for
perithecium formation.

There was a peak in the proportion of samples with perithecia in 1997 and 1998 just
prior to wheat flowering. In 1999, the peak occurred in July afler heavy rainfall in late
June and early July (approximately 7 in). Rainfall in May (near the time of wheat
flowering) was minimal (approximately 1.7 in.) and may have delayed perithecium
formation until conditions became more favorable in June and July. Although it is likely
moisture also plays a role (Paulitz, 1996), the rainfall data used in this study was
collected 25 - 60 miles away from the field sites and may not adequately represent field

conditions.

21

Perithecium formation may also be linked to day length and solarization (El-Gholl et
al., 1979; Halama and Lacoste, 1992). Although microclimates on the surface of the
debris were not monitored they would be a major factor in perithecium development
(Paulitz, 1996). Further studies that monitor on-site weather conditions and controls for
factors such as plant variety, pesticide and fertilizer applications, and initial inoculum are

needed to construct a predictive model for G. zeae perithecium development.

22

CHAPTER 2
USE OF RANDOM INSERTIONAL MUTAGENESIS TO STUDY THE
MECHANISM OF ASCOSPORE DISCHARGE IN GIBBERELLA ZEAE
INTRODUCTION

Fusarium Head Blight (FHB) or scab of wheat (Triticum aestivum L.), caused by
Gibberella zeae (Schwein.) Petch (anamorph: F usarium graminearum Schwabe), has
reached epidemic proportions in the United States over the last ten years. F usarium Head
Blight has incurred over 4 billion dollars in yield losses to wheat and barley growers in
the US. between 1993 and 1999 (McMullen, 1999). Yield losses in wheat occur
primarily as a result of sterile florets and diminished, low weight kernels (McMullen,
1999). G. zeae also produces the mycotoxins deoxynivalenol (DON) and zearalenone
that cause vomiting, feed refusal, and reproductive disorders when contaminated grain is
fed to livestock (McMullen, 1999; Stanton, 1995).

The disease cycle of FHB has been well described (Parry et al., 1995; Sutton, 1982).
Gibberella zeae overwinters as saprophytic mycelia on crop debris of maize, wheat, and
barley. In the spring, the fungus undergoes sexual reproduction and produces ascospores
in a fruiting body known as a perithecium. The forcible discharge of ascospores, possibly
along with wind and splash dispersal of the asexual macroconidia, serves to distribute the
primary inoculum. Although both spore types may be deposited on the wheat head
sometime during flowering, (Paulitz, 1996; Sutton and Procter, 1982) the forcible release
of ascospores likely plays a major role in spreading the disease over large distances

(Fernando et al., 1997).

23

The process of ascospore discharge has been described by Ingold (1971). A mature
ascus swells and then rapidly ejects its ascospores through the ostiole. Once an ascus has
been emptied, it collapses back into the perithecium body and the next ascus moves into
place and fires its spores. This is repeated until all asci have been emptied. The
mechanism for pressure build-up inside the ascus is not well understood in any fungus.

The idea that turgor pressure is behind forcible discharge was proposed by CT.
Ingold (1968). He observed a stretching of the ascus wall in Loramyces sp. as it built up
osmotic pressure under moist conditions. This stretching eventually caused the ascus to
burst and subsequently contract, which forced the ascospores out of a narrow slit at the
top of each ascus. The role of moisture in ascospore discharge has been described in
field studies of several fungi, G. zeae (Chen and Yuan, 1984; Reis, 1990a), Venturia
inaequalis (Gadoury et al., 1998) (Stensvand et al., 1998), Anisogramma anomala
(Pinkerton et al., 1998), and Pleospora allii (Prados Ligero et al., 1998). Under
laboratory conditions, it has also been observed G. zeae discharged the maximum number
of spores in 100% RH conditions (Trail et al., 1998).

In addition, a spring mechanism has been proposed based on the presence of an
extensive microtubule network in the epiplasm of mature asci of Thelebolus crustaceus
(Czymmek and Klomparens, 1992). It has also been found that benomyl, which inhibits
microtubule formation, reduces the discharge of ascospores in Venturia inaequalis
(Miller, 1970).

Random insertional mutagenesis can be useful for a large-scale search for genes
involved in a specific mechanism. A large number of tagged transformants can be

generated and then screened for a change in the phenotype of interest. In this study, a

24

screening method was developed that quickly distinguished isolates that had lost their
ability to forcibly eject their spores. Mutants showing a change in this phenotype could
then be studied in further detail.

Polyethylene-glycol (PEG) mediated transformation of protoplasts has also been
shown to achieve an increased rate of transformation, particularly when coupled with
restriction enzyme mediated integration (REMI) in some fungi (Brown et al., 1998;
Redman and Rodriguez, 1994). REMI transformation, originally developed for
Dictyostelium discoidium (Kuspa and Loomis, 1992), uses additional restriction enzymes
to nick the host DNA which is thought to facilitate insertion of the linearized plasmid.
The REMI system has been successfully used to isolate desirable mutants in many fungi
Saccharomyces cerevisiae (Schiestl and Petes, 1991), Ustilago maydis (Bolker et a1,
1995), Cochliobolus heterostrophus (Lu et al., 1994), Magnaporthe grisea (Shi et al.,
1995; Sweigard et al., 1998), Aspergillus nidulans (Sanchez et al., 1998), and G.
pulicaris (Salch and Beremand, 1993).

We have chosen to elucidate the mechanism of discharge in G. zeae through a
random gene disruption strategy. We believed random disruption of a large number of
isolates would produce a mutant(s) that was unable to discharge their spores. Subsequent
characterization of the disrupted genes may lead to a better understanding of the
processes involved in forcible ascospore discharge. Since many fungi are known to
actively discharge their spores, determining the mechanism behind this phenomenon
would lead to an increased understanding of the physiology of spore dissemination and

novel methods for control of these fungi.

25

MATERIALS AND METHODS

Strains, plasmids, and culture conditions. Two strains of G. zeae (F usarium
graminearum) were used for transformation. Strain W-8 (U -5373) (Adams et al., 1987)
was provided by G. Adams (Michigan State University). Strain PH-l is a field isolate
from Michigan provided by L.P. Hart (Michigan State University). Nitrate-nonutilizing
mutants 4232 and 4233, provided by R. Bowden (Kansas State University) were used for
meiotic stability tests.

Two plasmids were used for insertional mutagenesis, both harboring the coding
region on E. coli for the hygromycin B phosphotransferase gene (hph) (Appendix A).
Plasmid pHA1.3 has the hph coding region linked to the Aspergillus parasiticus trpC
promoter and terminator sequences (Redman and Rodriguez, 1994). Plasmid pUCH2-8
(Hohn, 1997) has the hph coding region fused to promoter 1 from Cochliobolus
heterostrophus (Turgeon et al., 1987).

Spore suspensions of fungal strains were prepared by flooding 4 day old cultures on
carrot agar (CA) medium (Bowden and Leslie, 1999) with approximately 0.01% Tween
20. All strains were maintained as soil stocks at —20°C (Nelson et al., 1983).

Transformation. Transformations were performed on germinated conidia using a
previously published protocol (Hohn, 1997) with the following modifications.
Approximately 0.3 g soil stock was used to inoculate carboxymethylcellulose (CMC)
medium and incubated for 72 hr at 25°C at 250 rpm (Cappellini and Peterson, 1965)..
Conidia were harvested by centrifugation and germinated in YEPD (0.3% yeast extract,
1% bactopeptone, and 2% D-glucose) broth for 12 - 14 hours at room temperature (RT) at

175 rpm. Isolation of protoplasts occurred in 25 mg/ml driselase, 0.05 mg/ml chitinase

26

(Sigma Chemical Co., St. Louis), and either 0.5 mg/ml mureinase (USB-Amersham
Pharrnacia Biotech Inc., Piscataway, NJ) or 5 mg/ml lysing enzyme (Sigma Chemical
Co., St. Louis) in a 1.2 M KC1 buffer. Protoplasts were collected by filtration through a
30 um Nitex nylon membrane (Tetko Inc., Kansas City, MO) and washed three times in
STC buffer (1.2 M sorbitol; 10 mM Tris-HCl, 50 mM CaClz, pH 8.0). Transformation
took place in the presence of 30% polyethylene glycol solution (10 mM Tris-HCl, 50 mM
CaClz, pH 8.0), STC buffer and linearized plasmid. Restriction enzyme mediated
integration (REMI) transformations included an additional 10, 50, or 100 units of NdeI or
HindIlI. Protoplasts recovered on Regeneration Medium (RM): 0.1% yeast extract, 0.1%
casein enzyme hydrosylate, 0.8 M sucrose , 1% agarose, for 15 hr and then were overlaid
with 10 ml of RM amended with 150 ug/ml hygromycin B (HygB) (Calbiochem-
Novabiochem Corp., San Diego, CA). Putative transformants were selected within 4 - 7
days and selected again on V8 juice (V M) medium amended with HygB. Unless

otherwise noted, hygromycin transformants were screened on 450 ug/ml for strain PH-l

and 300 ug/ml for strain W-8. Hygromycin resistant (Hng) colonies were transferred to
a 2% water agar (WA) medium and hyphal-tipped to obtain genetically pure isolates.
Detailed protocols for small and large-scale transformations are presented in Appendix B
and C.

Screening for mutants. To induce perithecium formation, transformants were placed on
CA (Klittich and Leslie, 1988) in 24 cell well microplates [17 mm well diameters
(Evergreen Inc., Los Angeles, CA)] or 60 mm Petri plates. Cultures were maintained
under continuous 15-watt cool white and black light fluorescent lighting. After 2 - 4 days

of growth, perithecia were induced from vegetative hyphae with 200 pl (microplates) or 1

27

ml (Petri plates) of 2.5% aqueous Tween 60 solution (Bowden and Leslie, 1999).
Approximately 7 - 14 days after perithecium induction, the plate lid was examined under
a dissecting microscope at 50X magnification for discharged ascospores. Cultures with
plate lids that had no visible accumulation of ascospores were screened more vigorously
as follows. Agar blocks containing approximately 20 - 40 perithecia were removed from
mature cultures and placed on a glass slide so that spores would be discharge in a
trajectory that ran parallel to the slide. Blocks were placed in a high humidity chamber.
The perithecia were kept in the chamber for 1 - 2 days away from direct light and re-
examined microscopically. A detailed screening protocol is presented in Appendix D.
Discharge mutants were embedded and sectioned as described in Trail and Common
(2000) and examined microscopically.

Screening for auxotrophic mutants. Transformants were tested for nitrate
utilization by growing on Minimal Medium (Corell, 1987) amended with 0.05% Tergitol
Type NP-10 (Sigma Chemical Co., St. Louis, MO) and 2% sorbose (MMTS) (Bowden
and Leslie, 1999). Nitrate non-utilizing (N it') and nitrate utilizing (N it+) phenotypes have
distinct morphologies on MMTS and are easily distinguished. Transformants unable to
grow on MMTS were then rescreened on Czapeks medium (Difco Laboratories, Detroit,
MI) which would not support the growth of Nit' mutants.

Molecular analyses of transformants and progeny. Transformants were grown in
a 2% yeast extract, 6% sucrose (YES) broth for 4 - 6 days. Mycelium was collected by
filtration and stored at -80° C and subsequently ground to a fine powder using liquid

nitrogen. DNA was isolated either according to the manufacturer’s instructions using

28

DNeasy Plant Maxi Kit (Qiagen Inc., Valencia, CA) or by the Autogen DNA isolation
system.

Plasmid DNA isolation and Southern hybridization analyses were performed using
published procedures (Ausubel et al., 1987). Digested genomic DNA was separated on an
0.8% agarose gel in TBA buffer (0.44 M Tris-HCl, 0.01 M EDTA, 0.44 M boric acid, pH
8.0) and subsequently transferred to a nylon Hybond-N+ membrane (U SB-Amersham
Pharmacia Biotech Inc., Piscataway, NJ). Restriction enzymes were used as instructed by
supplier (Gibco BRL, Grand Island, NY, or Boehringer Mannheim Biochemicals,
Indianapolis, IN). Linearized probe DNA (pHA1.3 or pUCH2-8) was labeled with [01-
32P]dCTP using Gibco Random Primers DNA Labeling System (Gibco BRL, Grand
Island, NY).

Mitotic stability test. Twelve transformants that were Hng (six containing pHAl .3
or six containing pUCH2-8) were placed on VM amended with HygB, and incubated at
25° C for 7 days. Mycelia were transferred to VM without HygB and incubated under
the same conditions. This procedure was repeated three times.

Meiotic stability test. Transformants showing deviations in perithecium formation
or ascospore discharge were selfed and rescreened for phenotype. Transformants were
also mated with nitrate-non-utilizing (N it') hygromycin sensitive (Hygs) mutants 4232
and 4233. Ascospores were selected from eight perithecia and germinated on MMTS
medium. Colonies were then placed on VM amended with HygB. Inheritance of

mutations was compared to expected Mendelian ratios.

29

RESULTS

Two fungal strains of G. zeae were used to generate and screen 5005 transformants
(139 with W-8 and 4866 with PH-l). Of these, four mutants (all PH-l) were identified
that failed to forcibly discharge ascospores in our assays. Three of the isolates were
developmental mutants that did not form mature asci and ascospores. One transforrnant,
designated 729G-112, appeared to develop normal perithecia and ascospores and was
studied in further detail.

Transformation. Transformation of protoplasts of G. zeae was conducted by
incubation of protoplasts of linearized plasmid DNA, with and without the presence of
restriction enzymes (10, 50, or 100 units of NdeI or HindIII). Rates of transformation
were higher without REMI (0.2 - 2.37 transformants (Tx)/ug plasmid DNA using
linearized pHA1.3 only and 0.08 Tx/ug plasmid DNA from pHA1.3 using REMI). The
transformation rate of protoplasts incubated with linearized pUCH2-8 was 0.2 - 1.2
Tx/ug plasmid DNA. No transformants were recovered from control transformations
without vector DNA. Strain PH-l was weakly resistant to low concentrations of HygB
(100 — 400 ug/ml) so only transformants able to grow vigorously on VM containing 450
ug/ml HygB were selected. Strain W-8 was more sensitive to HygB and was selected at
300 ug/ml.

Molecular analysis of transformants. Examination of 20 putative transformants (10
from pHAl .3 and 10 from pUCH2-8) by Southern analysis indicated that both plasmids
had successfully integrated into the genome by single or multiple-copy insertion (Figure

5).

30

Analysis of pUCH2-8 transformants digested with Clal, which cuts outside, indicated
single, multiple, and possibly partial copy insertions. Lanes 2 and 7 — 10 are considered
to be single-copy insertions with the additional bands attributed to partial digestion of
high concentrations of genomic DNA. The presence of three or more bands in lanes 1, 3,
and 4, indicates multiple insertion of pUCH2-8. Lanes 1 and 6 may contain partial copy
insertions as each contain a band(s) approximately 1 — 2 Kb smaller than the size the
whole plasmid.

Analysis of pHAl .3 transformants digested with Clal, which cuts once within the
plasmid, revealed the majority of transformants had a single-copy insertion (lanes 11 -
18, 20). One isolate, shown in lane 19, has four bands, which indicates multiple
insertions of pHAl .3. This isolate was transformed with pHA1.3 DNA that had been cut
by NdeI, which cuts twice within the plasmid. This may have resulted in multiple
insertions of truncated copies of pHAl .3.

Isolates in lanes 15 - 18 and 20 were further analyzed by digestion with Xhol which
does not out within the plasmid. The presence of a single band in each sample, ranging
from 4 — 6 kilobases (Kb) in size (lanes 15a — 18a, and 20a), suggests these isolates do
not result from a tandem repeat insertion. A tandem repeat would be indicated by the
presence of a fragment at least twice the size of the plasmid (>12 Kb) when digested with
a restriction enzyme that cuts outside the plasmid.

Undigested samples separated on 0.8% agarose gel did not show migration of low
molecular weight DNA that would indicate an autonomous plasmid (data not shown).

Southern analysis did not reveal any hybridization to bands that were similar in size to an

31

autonomous plasmid (data not shown). Both DNA probes, linearized labeled pHA1.3

and pUCH2-8, did not hybridize to unnansformed PH-l (data not shown).

    

            

12345678910 11121314151617181920
5 fl 5 5 -23.l ' 5 5 -23.1
-9.6 ,. . ;r -9-6
-6.6 ‘ ;_'., ‘ i. ‘
' . -6.6
4.3 '
-2.3
.20 I .-' -4.3
:f;.1_ .2.0
3;
.1:
-23.1
-9.6
-6.6
4.3
-2.3
-2.0

 

Figure 5. Southern-hybridization analysis G. zeae transformants. DNA from G. zeae
isolates transformed with pUCH2-8 and digested with restriction enzyme ClaI
(recognizes no restriction sites in the vector) (lanes 1 — 10). DNA from G. zeae isolates
transformed with pHAl .3 and digested with restriction enzyme ClaI (recognizes one
restriction site in the vector) (lanes 11 - 20) and XhoI (recognizes no restriction site in the
vector) (lanes 15a-18a and 20a). Lane 11, discharge minus mutant. Probed with 32F
labeled with HindIII cut pUCH2-8 (lanes 1-10) or pHA1.3 (lanes 11 — 20).

32

Mitotic stability and insertional pattern. Twelve isolates (six transformed with
each vector) were subjected to a mitotic stability test by transferring three successive
times on and off VM amended with HygB. Southern hybridization indicated the
integrated plasmid typically remained stable throughout these transfers (Figures 6 and 7).
However, a distinct heavy band appeared in DNA at week 7 of the isolate shown in lanes
2a and b in Figure 6. Differences in the size of large fragments in lanes 3a and b, 4a and
b, and 5a and b are attributed to partial digests of genomic DNA.

1b Za 2b

 

3b

           

4a 5a

   

Figure 6. Southernahybridization analysis of pHAl .3 transféfinants. DNAfim G. zeae
isolates transformed with pHA1.3 and digested with restriction enzyme ClaI (recognizes
one restriction site). DNA isolated from transformants from week 0 (lanes a) DNA
isolated from transformants after 7 weeks of transfer on/off medium amended with HygB
(lanes b). Probed with 32P labeled with HindIII cut pHA1.3.

33

 

155%
Figure 7. Southern hybridization analysis of pUCH2- 8 transformants. DNA from G. zeae
isolates transformed with pUCH2- -8 and digested with restriction enzyme CIaI
(recognizes one restriction site). DNA isolated from transformants from week 0 (lanes a)
DNA isolated from transformants after 7 weeks of transfer on/off medium amended with
HygB (lanes b). Probed with 32P labeled with HindIII cut pUCH2-8.

Meiotic Stability. The discharge mutant 7290-112, was selfed and tested for
stability of hygromycin resistance and retention of the mutant phenotype. Progeny were
resistant to HygB concentrations of 150 - 300 ug/ml HygB (compared to parental
resistance at 450 ug/ml HygB) and did not recover the ability to discharge ascospores.

The genetic inheritance of mutations was tested by mating selected mutants with
isolates unable to utilize nitrate (Bowden and Leslie, 1999). Progeny were tested on
MMTS for nitrate utilization, screened for perithecia production and ascospore discharge
on CA, and then screened on VM amended with 150 - 300 ug/ml HygB. Cosegregation
of the phenotype and hygromycin resistance was observed in 729G-112 and 729R-57 for
the majority of progeny. Transformants 720R-2 and 610A—173 produced few viable
ascospores after mating with Nit' isolates (Table 4). The Nit' phenotype segregated

independently.

34

Table 4. Segregation analyses of mutations.

 

 

Mutants

7290-1 12“” 720R-2W 729R-57W 61 OA- 1 73"”
Vector“ PHA1.3 PUCH2-8 PUCH2-8 PHAl.3
Enzfiymbedfinn Rm Hinldlll 1111311111 H1131!” Hindi]!
Nit ° is yg 0
Nif'Dis'Hygs 1 o o 1
Nit‘Dis’Hyg" 15 0 3 o
Nimisilygs 17 o 4 1
Nit’Dis’Hng 9 o 3 o
Nimismygs 14 3 4 3
Nit'DisW-Iyg“ o o o o
Nit'Dis‘HygS 2 O 0 0

 

‘ Discharge minus mutant

b Perithecium developmental mutant

° Plasmid used in transformation

d Restriction enzyme used to linearize plasmid prior to transformation
' Nitrate utilizing (N it') or nitrate non-utilizing (Nit')

' Ability to forcibly discharge ascospores

3 Hygromycin resistant (R) or Hygromycin sensitive (S)

Specific mutants. Of the 5005 transformants generated, four appeared to have lost

the ability to forcibly discharge ascospores. After microscopic examination, it was

confirmed three of the mutants were arrested during perithecium formation. Cross-

sections of perithecia are presented in Figure 8. Perithecia of 7296-1 12 (Figure 8A) have

mature ascospores, and wild type morphology of paraphyses, periphyses, and ostiole.

This mutant also produced amorphous cirri instead of the distinct tube formation of the

wild type cirrhi (Figure 9). Mutant 720R-2 (Figure 8B) lacked mature asci and was

arrested at an early stage of perithecia] development. Mutants 729R-57 and 610A-173

were also arrested in early stages of development (data not shown).

35

 

A) Discharge minus mutant 729G-112 at earlier stage maturity B) Developmental mutant
720R—2 C) Wild type perithecia at full maturity.

36

 

Figure 9. Perithecia and cirrhi of 720R-2, 729G-112 and wild type PH-l. A) Perithecium
at 50X of: D. developmental mutant 720R-2 W. wild type. B) Ascospore discharge
mutant 7296-112 at 70X (arrow) cirrus C) Wild type perithecia at 70X (arrow) cirrus.

37

DISCUSSION

Random insertional mutagenesis is an effective method to disrupt and locate a gene
sequence associated with a change in phenotype. The ease of isolation of surrounding
genes relies on vector insertion in a single location per genome and that the insert
remains stable. Examination of 20 transformants of G. zeae, has shown insertions are
random and remain stable in the majority of transformants. Transformants with pHA1.3
had predominantly single-copy insertions and transformants of pUCH2-8 had single and
multiple-copy insertions. Multi-COpy and unstable insertional events have been
documented in Gibberella pulicaris, (Salch and Beremand, 1993), Gibberellafitjikuroi
(Leslie and Dickman, 1991), Alternaria alternata (T anaka et al., 1999), Gibberella
pulicaris (Salch and Beremand, 1993) and Colletotrichum sp. (Redman and Rodriguez,
1994). We do not know the cause of the higher number of multiple copy inserts in
transformants of pUCH2-8.

Although the transformation rates for G. zeae were low, we have modified an existing
transformation protocol to make the generation of large numbers of mutants more
efficient. The majority of our transformation experiments did not use REMI mutagenesis
because we found it decreased the transformation rate. This was also observed in
F usarium mom'loforme (Shim and Woloshuk, 1998). The reason for this is unknown,
however it is possible that the additional restriction enzyme may have led to
chromosomal breakage of the host genome and a loss of plasmid insertion. Although
vector pHA1.3 has a telomeric sequence that was reported to cause chromosomal

breakage and lead to a loss of plasmid integration (Redman and Rodriguez, 1994), we

38

observed it to yield a higher transformation rate, greater stability, and more single copy
inserts than pUCH2-8.

In selflngs and outcrossings with another strain, progeny of matings were more
sensitive to hygromycin than the parental generation. We were not able to determine the
pattern of genetic inheritance for two of the four mutants due to low ascospore viability.
Ascospore abortion was also seen during a REMI mutagenesis project in G. pulicaris
(Salch, 1993) and may be caused by the “spore-killer” gene complex similar to that found
in Neurospora crassa (Hasunuma, 1984; Turner and Perkins, 1991 ).

Of the four mutants isolated during this study, the phenotypes appeared to either be
normal morphologically with a loss or defect in the mechanism of ascospore discharge or,
arrested early in perithecium development. The discharge mutant also has a distinct
cirrhus morphology that may be related to the mechanism of forcible discharge. We have
promising candidates to study the gene(s) associated with each phenotype and plan to

pursue gene isolation and sequencing.

39

APPENDICES

40

APPENDIX A

Plasmid Maps: pHA1.3 and pUCH2-8

41

Nar 0.24
Ndet 0.18 M11031 .
38p a” Hindlll 0.0
Pstl 0.40
Salt 0.41

0181 0.77
. 50081 1.@

   
   
 

Pstl1.11
Pvut 1.14
Baal-I1 1.82
PM 4.37
Boom 4.14
83:11 4.13
Sim/Diana Xbal 2.53
Baird-l1 2.54
8pm 2.72
2.83 Xbal 2.81

12.82

Figure 10. Map of plasmid DNA pHA1.3.

42

l.SacI (666)

'1 Xbal (686)

,' //BamHl(674)
,l // Pstl(714)

,1' .1/ /, EcoRl (716)

I

I! 1" .” ’l/
1 1"! ,4/ fl / ECO RV (724)
.I f l, (I I
’ / ,{i/ / _,,, H 1nd111 (728)

l .

l . .

. . ,

5 I J .’

. V. ." “I,

1 1 ;' "z ,"
. . u . ,
'1, y r ,’

) [,1],

 
  
 

/ EcoRV (886)

 
   

\

\ Pstl (2056)

Promoter l
\
\ EcoRI (2142)

\_

)/// / 1
// ,l/ {1’ 1
Kpnl(324l // / , Pstl(2745)

Apal(3235)/,/ .
Xhol(3222)/ /

Sall(3216) l

Pstl(3125)’

Figure 11. Map of plasmid DNA pUCH2-8.

43

APPENDIX B

Protocol for Random Insertional Mutagenesis

44

10.

11.

12.

13.

14.

. Inoculate 100 ml of Carboxymethylcellulose Medium (in a 250 ml Erlenmeyer flask)

with 0.3 g soil suspension. Incubate for 72 hr on a rotary shaker table at 25° C at 250
rpm (Cappellini and Peterson, 1965).

Filter culture through sterile miracloth in a Buchner funnel under vacuum. Place into
one 250 ml oakridge bottle. Spin at room temperature (RT) at 5,000 rpm (4000 g) in
a GSA rotor for 10 minutes.

Discard all but 4—5 mls of the supernatant and resuspend solution to a concentration
of 108 conidia/ml in the remaining solution.

Place conidia suspension in 100 ml of YEPD Broth (in a 250 ml Erlenmeyer flask)
and grow on a rotary shaker table for 12-14 hours at 25° C at 175 rpm.

Filter culture through sterile miracloth in a Buchner funnel under vacuum and collect
mycelia] mat. Place mat in sterile 250 Erlenmeyer flask. Add Protoplasting Buffer at
rate of 30 ml solution/ 100 ml initial YEPD culture

Digest for 1 hr on a rotary shaker table at 30° C at 80 rpm.

Filter digestion mixture through a 30 um Nitex nylon membrane (Tetko Inc., Kansas
City, MO) into a sterile beaker. Filtered solution should be turbulent due to the
presence of protoplasts.

Distribute solution evenly in 30 ml oakridge tubes and spin at room temperature (RT)
at 3,000 rpm (1500 g) for 5 minutes in a GSA rotor.

Discard supernatant and gently resuspend protoplasts in 10 ml of STC Buffer using
wide orifice glass pipettes (Fisher Scientific, Pittsburgh, PA). Spin solution in 88-34
rotor at 5,000 rpm (4000 g).

Discard supernatant and gently resuspend protoplasts in 1 ml STC Buffer using wide
orifice pipet tips. Transfer to a 2 ml eppendorf tube. Spin in a microcentrifuge at RT
at 3,500 rpm (3500 g) for six minutes. Repeat once.

Resuspend protoplasts in a final volume of 300 - 600 1.11 (106 - 108 protoplasts /ml)
and suspend in the following mixture: 100 111 - Protoplast Buffer, 100 111 -STC Buffer,
50 pl 30% PEG Solution, and 10 ul of linearized plasmid. Incubate at room
temperature for 20 minutes.

Add 2 ml 30% PEG Solution and incubate 5 minutes

Add 4 ml STC Buffer and gently mix by inversion.

Place 600 111 of suspension in 20 ml Regeneration Medium (RM) at 47—50°C.

45

15. Allow protoplasts to regenerate for 15 hours and then overlay with 10 ml of RM
amended with 150 ug/ml HygB (Calbiochem-Novabiochem Corp., San Diego, CA).

16. When transformants emerge (generally within 4 - 7 days), screen putative
transformants on V8 Medium containing 450 ug/ml HygB (strain PH-l) or 300 ug/ml
HygB (strain W-8).

CMC Medium

15 g carboxymethylcellulose (low viscosity) Sigma

1.0 g NH4NO3

1.0 g KH2P04

0.5 g MgSO4-7H20

1.0 g Yeast Extract

in l L H20

*note dissolve CMC in warm H20 before adding remaining ingredients
dispense 100 mls into ten 250 ml Erlenmeyer flasks

 

YEPD Broth

3.0 g Yeast Extract (Difco Laboratories, Detroit, MI)
10.0 g Bactompeptone ((Difco Laboratories, Detroit, MI)
20.0 g Dextrose (D-glucose)

dispense 100 mls into ten 250 ml Erlenmeyer flasks

 

Protoplasting Buffer

20 ml - 1.2 M KCl

500 mg - driselase (Sigma Chemical Co., St. Louis)

1 mg - chitinase (Sigma Chemical Co., St. Louis)

either:

10 mg - mureinase (USB-Amersham Pharmacia Biotech Inc., Piscataway, NJ)
or:

100 mg - lysing enzyme (Sigma Chemical Co., St. Louis)

 

STC Buffer

For 100 mls

60 ml - 1.2 M sorbitol

1 ml - 1M Tris - HCl pH 8.0
5 ml - 1 M CaClz

autoclave 20 minutes

30% PEG Solution

For 10 mls

7.5 ml - 40% PEG 8000

0.1 ml - l M Tris-HCI, pH 8.0

0.5 ml - l M CaClz,

1.9 ml - H20

Filter sterilize in a 0.45 pm Millex-HA filter (Millipore, Bedford, MA).

 

46

 

Regeneration Medium

For 1 L (500 ml each solution)

2X solution

1.0 g - yeast extract

1.0 g - casein enzyme hydrosylate
10 g - agarose

bring to 500 mls

Sucrose solution

547 g - sucrose

bring to 500 mls

autoclave each solution 20 minutes and then combine

 

V8 Medium

163 ml V8 Juice
g CaCO3

15 g agar

in l L H20

47

 

APPENDIX C

Protocol for Large Scale Random Insertional Mutagenesis

48

10.

11.

12.

13.

. Inoculate thirty-two flasks of 100 ml CMC Media (in 250 m1 Erlenmeyer flasks) with

9.6 g soil suspension (0.3 g each flask). Incubate for 72 hours on a rotary shaker table
at 25°C at 250 rpm (Cappellini and Peterson, 1965).

Filter cultures through sterile miracloth in a Buchner funnel under vacuum. Divide
into sixteen 250 ml oakridge bottles. Spin at room temperature (RT) at 5,000 rpm
(4000 g) in a GSA rotor for 10 minutes.

Discard all but 10 mls of the supernatant and resuspend solution to a concentration of
108 conidia/ml in the remaining solution in each bottle.

Distribute conidia suspension (from the sixteen 250 ml oakridge bottles) evenly into
thirty-two 100 ml cultures of YEPD Broth and grow for 12-14 hours on a rotary
shaker table at 25°C at 175 rpm.

Filter cultures through sterile miracloth in a Buchner funnel under vacuum. Collect
mycelia] mats and evenly divide into eight sterile 250 Erlenmeyer flasks. Add
Protoplasting Buffer at rate of 30 ml solution/100 ml initial YEPD culture

Digest for 1 hour at on a rotary shaker table 30° C at 80 rpm.

Filter digestion mixture through a 30 um Nitex nylon membrane (Tetko Inc., Kansas
City, MO) into a sterile beaker. Filtered solution should be turbulent due to the
presence of protoplasts.

Distribute solution evenly in four 250 ml oakridge bottles and spin at room
temperature (RT) at 3,000 rpm (1500 g) for 5 minutes in a GSA rotor.

Discard supernatant and gently resuspend protoplasts in 10 ml of STC Buffer using
wide orifice glass pipettes (Fisher Scientific, Pittsburgh, PA). Transfer to four 30 ml
oakridge bottles.

Discard supernatant and gently resuspend protoplasts in 1 ml STC Buffer. Transfer to
sixteen 2 ml eppendorf tubes. Spin in a microcentrifuge at RT at (3 500 g) for six
minutes. Repeat once.

Resuspend protoplasts in 2 ml tubes 3 final volume of 600 pl (1 X 106 - 1 X 108
protoplasts /ml). Combine two tubes for a total of eight tubes. Prepare eight 50 ml
sterile conical polypropylene centrifuge tubes with the following mixture: 600 pl -
protoplast solution, 600 pl -STC Buffer, 300 1,11 30% PEG Solution, and 60 1.11 of
linearized plasmid. Incubate at room temperature for 20 minutes.

Add 12 ml 30% PEG Solution and incubate 5 minutes

Add 24 m1 STC Buffer and gently mix by inversion.

49

14. Gently pour each conical tube into 120 ml Regeneration Medium (RM) at 47—50°C.
Aliquot 20 mls into each 60 x lS-mm Petri plate dish.

15. Allow protoplasts to regenerate for 15 hours and then overlay with 10 ml of RM
amended with 150 ug/ml HygB (Calbiochem—Novabiochem Corp., San Diego, CA).

16. When transformants emerge (generally within 4 - 7 days), screen putative
transformants on V8 Medium containing 450 ug/ml HygB (strain PH-l) or 300 ug/ml
HygB (strain W-8).

CMC Medium
15 g - carboxymethylcellulose (low viscosity) Sigma F
1.0 g - NH4NO3

1.0 g - KH2PO4

0.5 g - MgS04-7H20

1.0 g - yeast extract (Difco Laboratories, Detroit, MI)

in 1 L H2O

*note dissolve CMC in warm H2O before adding remaining ingredients
dispense 100 mls into ten 250 ml Erlenmeyer flasks 1"

 

 

YEPD Broth

3.0 g - yeast extract (Difco Laboratories, Detroit, MI)

10.0 g - Bactompeptone ((Difco Laboratories, Detroit, MI)
20.0 g - dextrose (D-glucose)

dispense 100 mls into ten 250 ml Erlenmeyer flasks

 

Protoplasting Buffer

960 ml - 1.2 M KCl

24 g - driselase (Sigma Chemical Co., St. Louis)

48 mg - chitinase (Sigma Chemical Co., St. Louis)

480 mg - mureinase (USB-Amersham Pharmacia Biotech Inc., Piscataway, NJ)

stir for 30 minutes and filter sterilized in a 0.45 pm Millex-HA filter (Millipore, Bedford,
MA).

 

STC Buffer

For 100 mls

60 ml - 1.2 M sorbitol

1 ml - 1M Tris-HCl, pH 8.0
5 ml - l M CaCl2
autoclave 20 minutes

50

30% PEG Solution

For 10 mls

7.5 m1 - 40% PEG 8000

0.1 ml - 1 M Tris-HCl, pH 8.0

0.5 ml - 1 M CaCl2,

1.9 ml - H2O

Filter sterilize in a 0.45 pm Millex-HA filter (Millipore, Bedford, MA).

 

Regeneration Medium

For 1 L (500 ml each solution)
2X solution

1.0 g - yeast extract

1.0 g - casein enzyme hydrosylate
10 g - agarose

bring to 500 mls

Sucrose solution

547 g — sucrose

bring to 500 mls

autoclave each solution 20 minutes and then combine

 

 

V8 Medium

163 ml V8 Juice
1.0 g CaCO3

15 g agar

in 1 L H20

51

APPENDIX D

Protocol for Screening of Discharge Minus / Perithecia Mutants

52

Transformants were placed on carrot agar (CA) medium (35% carrots, 2% agar)
(Klittich and Leslie, 1988) in 24 cell well microplates with 17 mm well diameters
(Evergreen, Los Angeles, CA) or 60 mm Petri plates. Plates were maintained under
continuous 15-watt cool white and black light fluorescent lighting. Atmospheric
moisture was intensified by covering incubation area with plastic sheeting. During dry
periods (winter months) a standard room humidifier was used to further supplement air
moisture. Prior to induction, conidia were harvested in sterile water containing a few
drops of Tween 20. After 2 - 4 days of growth, vegetative hyphae was covered with 200
pl or 1000 pl of 2.5% aqueous Tween 60 solution (Bowden and Leslie, 1999) in 17 mm
cell well plates and 60 mm Petri plates, respectively. Hyphae was flattened using a glass
rod or hockey stick in 17 mm cell well plates and 60 mm Petri plates, respectively.
Approximately 5 - 7 days after perithecium formation, the cell well microplate lid was
examined under a microscope at 5X magnification for discharged ascospores.

Cultures that had no visible signs of forcible discharge after two trials were
rescreened more vigorously as follows. Agar blocks containing approximately 20 - 40
perithecia were removed from mature cultures and placed on a glass slide so that spores
would be parallel to the slide in a high humidity chamber. The perithecia were kept in

the chamber for 1 - 2 days away from direct light and re-examined microscopically.

53

 

BIBLIOGRAPHY
National Climatic Data Center National Oceanic and Atmospheric Administration.
[Online] Available
http://www.ncdc.noaa.gov/ol/climate/climatedata.html#DAILY, 2000.

Adams, G., Johnson, N., Leslie, J. F., and Hart, L. P., 1987. Heterokaryons of Gibberella
zeae formed following hyphal anastomosis or protoplast fusion. Exp. Mycol.
11(4): 339-353.

Alexopoulos, C. J ., Mims, C. W., and Blackwell, M., 1996, Introductory Mycology, John
Wiley & Sons, Inc., New York, pp. 869.

Armaleo, D., Ye, G. N., Klein, T. M., Shark, K. B., Sanford, J. C., and Johnston, S. A.,
1990. Biolistic nuclear transformation of Saccharomyces cerevisiae and other
fungi. Curr. Genet. 17(2):97-103.

Ausubel, F. M., Brent, R., Kingston, R. E., Moore, D. D., Siedman, J. G., Smith, J. A.,
and Struhl, K., 1987. Current protocols in molecular biology. John Wiley & Sons,
New York.

Bai, G. H., and Shaner, G., 1994. Scab of Wheat - Prospects For Control. Plant Dis.
78(8):760-766.

Boelker, M., Boehnert, H. U., Braun, K. H., Goerl, J., and Kahmann, R., 1995. Tagging
pathogenicity genes in Ustilago maydis by restriction enzyme-mediated
integration (REMI). Mol. Gen. Gen. 248(5):547-552.

Bowden, R. L., and Leslie, J ., F., 1999. Sexual recombination in Gibberella zeae.
Phytopathology 89(2): 182-1 88.

Brown, J. S., Aufauvre Brown, A., and Holden, D. W., 1998. Insertional mutagenesis of
Aspergillusfumigatus. Mol. Gen. Gen. 259(3):327-335.

Burgess, L. W., Wearing, A. H., and Toussoun, T. A., 1975. Surveys of Fusaria
Associated with Crown Rot of Wheat in Eastern Australia. Aust. J. A gric. Res.
26:791-799.

Cappellini, R. A., and Peterson, J. L., 1965. Macroconidium Formation in Submerged
Cultures by a Non-Sporulating Strain of Gibberella zeae. Mycologia 57:962-966.

Chakraborty, B. N., Patterson, N. A., and Kapoor, M., 1991. An electroporation-based
system for high-efficiency transformation of germinated conidia of filamentous
fungi. Can. J. Microbiol. 37(11):858-63.

Chen, X. M., and Yuan, C., 1984, Application of microcomputer in studying wheat scan
epidemiology and forecasting, Zhejiang Agric. Sci. 2:55-60.

54

Corell, J. C., Klittich, C.J.R., Leslie, J .F, 1987. Nitrate Nonutilizing Mutants of F usarium

oxysporum and Their Use in Vegetative Compatibility Tests. Phytopatholog»
77: 1640-1646.

Czymmek, K. J ., and Klomparens, K. L., 1992. The ultrastructure of ascosporogenesis in
freeze-substituted T helebolus crustaceus: enveloping membrane system and
ascospore initial development. Can. J. Bot. 70(8):1669-1683.

Durand, R., Rascle, C., Fischer, M., and Fevre, M., 1997. Transient expression of the
beta-glucuronidase gene after biolistic transformation of the anaerobic fungus
Neocallimastixfiontalis. Curr. Genet. 31(2):158-61.

El-Gholl, N. E., Schoulties, C. L., and Ridings, W. H., 1979. Factors affecting perithecia]
production in Gibberella (F usarium) tricincta. Can. J. Bot. 57:2497-2500.

Fernando, W. G. D., Paulitz, T. C., Seaman, W. L., Dutilleul, P., and Miller, J. D., 1997.
Head blight gradients caused by Gibberella zeae from area sources of inoculum in
wheat field plots. Phytopathology 87(4):414-421.

Francis, R. G., and Burgess, L. W., 1977. Characteristics of F usarium roseum
'Graminearum' in Eastern Australia. Trans. Brit. Myc. Soc. 68:421-427.

Gadoury, D. M., Stensvand, A., and Seem, R. C., 1998. Influence of light, relative
humidity, and maturity of populations on discharge of ascospores of Venturia
inaequalis. Phytopathology 88(9):902-909.

Halama, P., and Lacoste, L., 1992. Detenninism of sexual reproduction of Phaeosphaeria

nodorum agent of wheat septoriosis: II. Effect of temperature and light. Can. J.
Bot. 70(8):1563-1569.

Hart, L. P., 1998. NCR-184 Regional committee scab of wheat progress report 1998, in:
The 1998 National F usarium Head Blight Forum (L. P. Hart, Ward, R., Bafus, R.,
Bedford, K., ed.). University Printing, East Lansing, MI, Michigan State
University, East Lansing Michigan, pp. 111-113.

Hart, L. P., 1999. 1999 NCR-l 84 State report management of head scab of small grains,
in: 1999 National Head Blight Forum (J. A. Wagester, R. Ward, L. P. Hart, S. P.
Hazen, J. Lewis, and H. Borden, eds.). University Printing, Michigan State
University, East Lansing, MI, Sioux Falls, SD, pp. 201-203.

Hasunuma, K., 1984. Meiosis in Neurospora crassa: 2. Isolation and partial
characterization of dominant meiotic mutants constituting meiotic gene cluster, V-
2 or LG V. Jap. J. Gen. 59(5):517-526.

55

Hawksworth, D. L., Kirk, P. M., Sutton, B. C., and Pegler, D. N., 1996. Ainsworth and
Bisby's dictionary of the fungi (1. M. Institute, ed.). CAB International, New
York.

Hohn, T., 1997. Transformation Protocol for Gibberella zeae. USDA, Peoria, IL.
Ingold, C. T., 1968. Spore liberation in Loramyces. Transaction of British Mycological
Society 51(2):323-341 .

Ingold, C. T., 1971. Fungal spores: their liberation and dispersal. Clarendon Press,
Oxford, UK.

Ingold, C. T., and Dann, V., 1968. Spore discharge in fungi under very high surrounding
air-pressure, and the bubble-theory of ballistospore release. Mycologia 60:285-
289.

Itoh, Y., and Scott, B., 1997. Effect of de-phosphorylation of linearized pAN7-l and of
addition of restriction enzyme on plasmid integration in Penicillium paxilli. Curr.
Genet. 32:147-151.

Leslie, J .F. and Dickman, M.B., 1991. Fate of DNA encoding hygromycin resistance
after meiosis in transformed strains of Gibberellafitjikuroi (F usarium
monoliforme). Appl. Env. Microb. 57(5): 1423-1429.

Khonga, E. B., and Sutton, J. C., 1988. Inoculum production and survival of Gibberella
zeae in maize and wheat residues. Can J Plant Pathol Rev Can Phytopathol
10(3):232-239.

Klittich, C. J. R., and Leslie, J. F., 1988. Nitrate reduction mutants of F usarium
moniliforme (Giberellafiljikuroi). Genetics 118:417-423.

Kuper-Goodman, T., Scott, P.M., Watanabe, H., 1987. Risk assessment of the mycotoxin
zearalenone. Regulatory toxicology and pharmacology 7:253-306.

Kuspa, A., and Loomis, W. F., 1992. Tagging Developmental Genes in Dictyostelium By
Restriction Enzyme-Mediated Integration of Plasmid DNA. Proc. Natl. Acad. Sci.
USA 89(18):8803-8807.

Leslie, J. F ., 1983. Some genetic techniques for Gibberella zeae. Phytopathology
73(7): 1005-1008.

Long, G. G., Dickman, M., Tuite, J ., Shannon, G. M., and Vesonder, R. F., 1982. Effects

of F usarium roseum corn culture containing zearalenone on early pregnancy in
swine. Am. J. Vet. Res. 43:1599-1603.

56

 

Lorito, M., Hayes, C. K., Dipietro, A., and Harman, G. E., 1993. Biolistic Transformation

of T richoderma-Harzianum and GIiocladium— Virens Using Plasmid and Genomic
DNA. Cur. Gen. 24(4):349-356.

Lu, 8., Lyngholm, L., Yang, G., Bronson, C., Yoder, O. C., and Turgeon, B. G., 1994.
Tagged mutations at the Toxl locus of Cochliobolus heterostrophus by restriction
enzyme-mediated integration. Proc. Natl. Acad. Sci. U. S. A. 91(26):12649-53.

McMullen, M., Jones, R., and Gallenberg, D., 1997a. Scab of wheat and barley: a re-
emerging disease of devasting impact. Plant Dis. 81(12):]340-1348.

McMullen, M., Milus, G., and Prom, L., 1999. 1999 Uniform fungicide trials to identify
products effective against F usarium head blight in wheat. in: I 999 National Head
Blight Forum (J. A. Wagester, R. Ward, L. P. Hart, S. P. Hazen, J. Lewis, and H.
Borden, eds.), University Printing, Michigan State University, East Lansing, MI,
Sioux Falls, SD, pp. 64-68.

McMullen, M. P., Stack, R.W., 1999. US Wheat and Barley Scab Initiative. North
Dakota State University Extension Service. [Online] Available
http://www.scabusa.org.

McMullen, M. P., Enz, J ., Lukach, J ., and Stover, R., 1997b. Environmental conditions
associated with F usarium head blight epidemics of wheat and barley in the
northern Great Plains, North America. Cer. Res. Com. 25(3 Part 2):777-778.

Miller, J. D., Culley, J ., Fraser, K., Hubbard, S., Meloche, F., Ouellet, T., Seaman, W. L.,
Seifert Keith, A., Turkington, K., and Voldeng, H., 1998. Effect of tillage practice
on fusarium head blight of wheat. Can J Plant Pathol. March 20(1):95-103.

Miller, P. M., 1970. Reducing discharge of ascospores of Venturia inaequalis with a
spring application of benomyl, thiabendazole, or urea. Plant Dis. 54(1):27.

Moss, M. 0., and F rank, J. M., 1985. Influence of the fungicide tridemorph on T-2 toxin
production by Fusarium sporotrichoides. Trans. Brit. Myc. Soc. 84:585-590.

Nelson, P. E., Toussoun, T. A., and Marasas, W. F. 0., 1983. Fusarium species : an
illustrated manual for identification. Pennsylvania State University Press,
University Park.

Olive, L. S., 1964. Spore discharge mechanism in Basidiomycetes. Science 146:542.

Parry, D. W., Jenkinson, P., and McLeod, L., 1995. Fusarium ear blight (scab) in small
grain cereals-a review. Plant-Pathology (United Kingdom) v. 44(2):p. 207-238.

Paulitz, T. C., 1996. Diurnal release of ascospores by Gibberella zeae in inoculated wheat
plots. Plant Dis. 80(6):674-678.

57

Pereyra, S. A., Dill-Macky, R., and Sims, A. L., 1999. Survival and Incolum potential of
F usarium graminearum in wheat residues. in: 1999 National Head Blight Forum
(J. A. Wagester, R. Ward, L. P. Hart, S. P. Hazen, J. Lewis, and H. Borden, eds.),
University Printing, Michigan State University, East Lansing, MI, Sioux Falls,
South Dakota, Pp. 96-98.

Pinkerton, J. N., Johnson, K. B., Stone, J. K., and Ivors, K. L., 1998. Maturation and
seasonal discharge pattern of ascospores of Anisogramma anomala.
Phytopathology 88(1 1): 1 165-1 173.

Prados Ligero, A. M., Gonzalez Andujar, J. L., Melero Vara, J. M., and Basallote Ureba,
M. J ., 1998. Development of Pleospora allii on garlic debris infected by
Stemphylium vesicarium, Eur. J. Plant Pathol. 104(9):861-870.

Pritsch, C., Muehlbauer, G.J., Bushnell, W.R., Somers, D.A., and Vance, C.P., 2000.
Fungal development and induction of defense response genes during early

infection of wheat spikes by F usarium graminearum. M01. Plant-Microbe
Interact. 13(2): 159-169.

Redman, R. S., and Rodriguez, R. J ., 1994. Factors affecting the efficient transformation
of Colletotrichum species. Exp. Myc. l8(3):230-246.

Reis, E. M., 1990a. Effects of rain and relative humidity on the release of ascospores and
on the infection of wheat heads by Gibberella zeae. F itopatologia-Brasileira
(Brazil) 15(4):p. 339-343.

Reis, E. M., 1990b. Survival of perithecia of Gibberella zeae on naturally infected wheat
kernels under field conditions, F itopatologia-Brasileira (Brazil) 15(3):254-255.

Salch, Y. P., and Beremand, M. N., 1993. Gibberella pulicaris transformants: state of
transforming DNA during asexual and sexual growth, Curr. Gen. 23:343-3 50.

Sanchez, 0., Navarro, R. E., and Aguirre, J ., 1998. Increased transformation frequency
and tagging of developmental genes in Aspergillus nidulans by restriction
enzyme-mediated integration (REMI), Mol. Gen. Gen. 258(1-2):89-94.

Schiestel, RH, and Petes, T.D., 1991. Integration of DNA fragments by illegitimate
recombination in Saccharomyces cerevisiae, Proc. Natl. Acad. Sci. 88: 7585-7589

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

Shi, Z., Christian, D., and Leung, H., 1995. Enhanced Transformation in Magnaporthe-

Grisea By Restriction Enzyme-Mediated Integration of Plasmid DNA.
Phytopathology 85(3):329-333.

58

Shim, W. B., and Woloshuk, C. P., 1998. Use of restriction enzyme mediated integration
for F usarium moniliforme. in: American Phytopathology Society, Las Vegas, NV.

Stanton, N., 1995. Mycotoxin levels in the 1995 midwest preharvest corn crop. The
National Veterinary Services Laboratories [Online] Available
http://www.usda.org. USDA, APHIS. Ames, Iowa 50010.

Stensvand, A., Amudsen, T., Semb, L., Gadoury, D. M., and Seem, R. C., 1998.
Discharge and dissemination of ascospores by Venturia inaequalis during dew.
Plant Dis. 82(7):761-764.

Sutton, J. C., 1982. Epidemiology of wheat head blight and maize ear rot caused by
F usarium graminearum. Can. J. Plant Path. 4:195-209.

Sutton, J. C., and Procter, R., 1982. Dispersion analysis of gibberella ear rot in maize
Gibberella zeae. Can. J. Plant. Pathol. 4(3):254-258.

Sweigard, J. A., Carroll, A. M., Farrall, L., Chumley, F. G., and Valent, B., 1998.
Magnaporthe grisea pathogenicity genes obtained through insertional
mutagenesis. Mol. Plant. Microbe. Interact. ll(5):404-12.

Tanaka, A., Shiotani, H., Yamamoto, M., and Tsuge, T., 1999. Insertional mutagenesis
and cloning of the genes required for biosynthesis of the host-specific AK-toxin in

the Japanese pear pathotype of Alternaria alternata. Mol. Plant. Microbe.
Interact. 12(8):691-702.

Teich, A. H., 1989. Epidemiology of wheat (T riticum aestivum L.) scab caused by
F usarium spp. in: F usarium Mycotoxins, Taxonomy and Pathogenicity (J.
Chelkowski, ed.), Elsevier, Amsterdam, pp. 269-282.

Teich, A. H., and Hamilton, J. R., 1985. Effect of cultural practices, soil phosphorus,
potassium and pH on the incidence of F usarium head blight and deoxynivalenol
levels in wheat (Triticum aestivum). Appl. Environ. Microbiol. 49(6): 1429-143 1.

Teich, A. H., and Nelson, K., 1984. Survey of F usarium head blight and possible effects

of cultural practices in wheat fields in Larnbton County in 1983. Can. Plant Dis.
Sur. 64:11-13.

Trail, F., and Common, R., 2000. Perithecia] development of Gibberella zeae: A light
microscopy study. Mycologia 92(1): 130-138.

Trail, F., Gadoury, D., and Loranger, R., 1998. Environmental parameters of ascospore
discharge in Gibberella zeae, in: The 1998 National F usarium Head Blight
Forum. University Printing, Michigan State University, East Lansing Mi, pp. 11-
13.

59

Turgeon, B. G., Garber, R. C., and Yoder, O. C., 1987. Development of a fungal
transformation system based on selection of sequences with promoter activity.
Mol. Cell Bio. 7(9):3297-3305.

Turner, B. C., and Perkins, D. D., 1991. Meiotic drive in Neurospora and other fungi.
American Naturalist 137(3):416-429.

Tusa, C., Munteanu, I., Capetti, E., Pirvu, T., Bunescu, 8., Sin, G. L., Nicolae, H., Tianu,
A., Caea, D., Romascanu, I., and Stoika, V., 1981. Aspects of the F usarium
attacks on wheat in Romania, Probleme de Protectia Plantelor 9: 15-3 1.

Yu Jieh, J ., and Cole Garry, T., 1998. Biolistic transformation of the human pathogenic
fungus Coccidioides immitis. J Microbial. Meth. July 33(2): 129-141.

60

"I11111111111111“