AN ABSTRACT OF THE THESIS OF

Mahfuzur Rahman for the degree of Master of Science in Botany and PlantPathology presented on September 19, 2000.

Title: Sensitivity of Wheat Genotypes to a Toxic Fraction Produced byCephalosporium

ramineum and Correlation with DiseaseSusceptibility

Abstract approved: Redacted for Privacy

Christopher C. Mundt Thorhas J. Wolpert

Cephalosporium stripe, caused by the soil-borne ascomycete Cephalosporium gramineum, is becoming an increasingly important disease of winter wheat(Triticum aestivum) in several areas of the world, especially where stubble mulch is practiced to maintain soil moisture and prevent erosion. As cultural control of the disease is infeasible and no fungicides are registered, the development of resistant cultivarsoffers the best hope for disease control. Selection of resistant genotypes remainsproblematic due to the requirements of evaluating adult plants in variable field environments.The symptoms of cephalosporium stripe suggest the involvement ofpathogen-produced toxins, and the toxin called graminin A has previously been isolated from C. gramineum. The goals of this thesis were to determine if insensitivity of wheat

genotypes to a toxic fraction produced by C. gramineum is associatedwith resistance to

cephalosporium stripe, and to evaluate the potential use of this toxic fraction to screen

wheat genotypes for disease resistance. A method was developed to mass-produce a toxic fraction of C. gramineum by modifying the method of K. Kobayashi (Kobayashi and Ui 1977). Large volumes (9 L) of broth medium were inoculated with C. gramineum and incubated for 35 days. The culture filtrate was then extracted four times with ethyl acetate, which eliminated need for the most time-consuming step of rotary evaporation. Reversed phase fractionation of the crude extract was used to isolate the toxic fraction. Leaves from fourteen-day-old wheat plants were excised and placed in scintillation vials containing the toxic fraction.

Exposure to a concentration of(60d1ml) of the toxic fraction for 72 hours produced distinct wilting symptoms that allowed us to distinguish wheat genotypes in a repeatable manner. Degree of wilting was measured on a continuous 1-5 scale, where 1 = no wilting and 5 = a filly wilted leaf.

Twenty wheat genotypes, belonging to four distinct germplasm groups (common, club, durum, and synthetic) were included in three runs of the toxin assay. Wilting reactions for the 20 genotypes were very consistent among runs. Sensitivity of the individual genotypes to the toxic fraction did not differ significantly within germplasm groups, but all differences among the means of the four germplasm groups were highly significant (P < 0.001) based on linear contrasts.

Seventeen winter wheat genotypes representing the common, club, and durum germplasm groups were planted in infested fields at two locations. The percentage of tillers showing whitehead symptoms (early maturity and reduced grainfill) was significantly correlated with wilting symptoms measured by the toxin assay. The common and durum genotypes varied more for disease susceptibility than for toxin insensitivity. However, all pairwise linear contrasts between means of the whitehead percentages for the three germplasm groups were highly significant (P < 0.001). Failure of the toxin assay to identify some of the moderately resistant common wheat genotypes suggests that additional mechanisms of resistance are operative. As no toxin-insensitive genotype showed susceptibility to cephalosporium stripe in the field experiments, however, the toxin assay can be used as an initial screening procedure to reduce the

number of genotypes to be tested in the field. Copyright by Mahfuzur Rahman September 19, 2000 All Rights Reserved Sensitivity of Wheat Genotypes to a Toxic Fraction Produced by Cephalosporium gramineum and Correlation with Disease Susceptibility

by

Mahfuzur Rahman

A THESIS

Submitted to

Oregon State University

in partial fulfillment of the requirements for the degree of

Master of Science

Presented September 19, 2000 Commencement June 2001 Master of Science thesis of Mahfuzur Rahman presented on September 19, 2000

Approved:

Redacted for Privacy

Co-Major Professor, representing Botany and Plant Pathology

Redacted for Privacy Botany and Plant Pathology

Redacted for Privacy Char of Department of Botanythd Plant pathology

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Dean of

I understand that my thesis will become part of the permanent collection of Oregon State University libraries. My signature below authorizes release of my thesis to any reader upon request.

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Author ACKNOWLEDGEMENTS

I would like to thank the members of my committee, Christopher C. Mundt, Thomas J. Wolpert, and Oscar RieraLizarazu for their advice, support, and encouragement during this research. I am highly indebted to Christopher C. Mundt and Thomas J. Wolpert, my major professors, for shaping and guiding the research at every step. This specific project would not have been possible without the clear insight of Thomas J. Wolpert about toxin activity and practical assistance in extracting the toxic fraction in a simplified method for mass production. Christopher C. Mundt made it possible for me to conduct experiments in farmers' fields, which I hereby gratefully acknowledge. Oscar RieraLizarazu introduced me to the ITMI mapping population and actively helped in our search to identif' quantitative trait loci for resistance to cephalosporium stripe. Lynda M. Ciuffetti introduced me to laboratory techniques and allowed me to use her lab equipments throughout the study; her contributions to the project and generous help are also gratefully acknowledged. I would like to express my sincere thanks to my fellow lab member, Christina Cowger, from whom I received continuous inspiration about the project and exchanged a lot of views about cephalosporium stripe, which she worked with in her Master's project. My heartfelt appreciation also goes to Molly E. Hoffer, Senior Faculty Research Assistant, who helped me in various ways. Special thanks goes to Dan Coyle who was very friendly and willing to provide maintenance support for the growth chambers. I would like to express my gratitude to my authorities at the Bangladesh Rice Research Institute, who provided the opportunity and support for me to pursue higher study. Additional financial support for the project and myself came from USDA-CSREES-

SRG-STEEP, without which I could not have finished this study. I am deeply grateful to my wife, who consistently inspired and encouraged me towards completion of the study. I also appreciate the help of newly-joined Research

Assistants in Christopher C. Mundt's lab, Anne Bureau and LaRae Wallace. I would like to thank the staff and students of Lynda M. Ciuffetti and Tom Wolpert's lab for their friendship and support. CONTRIBUTION OF AUThOR

Dr. Oscar Riera-Lizarazu was involved in the analysis of ITMI mapping progeny evaluation data. He also assisted in the interpretation of data. TABLE OF CONTENTS

I. iNTRODUCTION .1

Importance of the disease .1 Pathogenbiology ...... 2

Influence of environment on disease development ...... 3

Role of inoculum on disease development ...... 4

Conditions affecting survival of inoculum ...... 5 InfectionProcess ...... 7

Resistance source and mechanism of resistance ...... 9 Potential control measures ...... 12

Role of toxin in cephalosporium stripe development ...... 15 Literaturecited ...... 17

II. SENSITiVITY OF WHEAT GENOTYPES TO A TOXIC FRACTION PRODUCED BY CEPHALOSPOPRIUM GRAM1NEUM AND CORRELATION WITH DISEASE SUSCEPTIBILITY ...... 25 Abstract ...... 26 Introduction ...... 27 Materials and Methods ...... 29 Laboratoryassays ...... 29 Extraction of toxic fraction ...... 30 Assaymethod ...... 32 Statistical analyses ...... 33 TABLE OF CONTENTS (Continued)

Fieldexperiments ...... 34

Locations and experimental design ...... 34 Data collection and statistical analysis ...... 35 Mapping progeny evaluation ...... 36 Results ...... 38

Reaction of genotypes to toxic fraction ...... 38 Fieldreactions ...... 38 Comparison of laboratory assay to field data ...... 40 Toxin sensitivity of the mapping progeny ...... 41 Discussion ...... 44 References ...... 47

III. CONCLUSIONS ...... 50 BIBLIOGRAPHY ...... 52 LIST OF FIGURES

Figure Pag

II. 1. Differential response of the mapping population parents at 24 hr. of exposure to the toxic fraction produced by Cephalosporium gramineum...... 33

II. 2. Scale used to assess the degree of wilting of wheat leaves after 72 hr. of exposure to a toxic fraction produced by Cephalosporium gramineum...... 34

II. 3. Correlation of leaf wilting in response to a toxic fraction produced by Cephalosporium gramineum with percent whiteheads produced by 17 wheat genotypes in a naturally infested field in Condon, OR...... 40

II. 4. Correlation of leaf wilting in response to a toxic fraction produced by Cephalosporium gramineum with percent whiteheads produced by 17 wheat genotypes in a naturally infested field in Dufur, OR...... 41

II. 5. Frequency distribution of wilting for 112 recombinant inbred line progeny resulting from a cross between the toxin sensitive parent Opata85 and toxin insensitive parent M6 after exposure to a toxic fraction produced by Cephalosporium gramineum...... 43 LIST OF TABLES

Table

II. 1. Variance components for analysis of sensitivity of 112 recombinant inbred wheat lines to a toxic fraction produced by Cephalosporium gramineum...... 37

II. 2. Mean wilting reaction of twenty winter and spring wheat genotypes to a toxic fraction produced by Cephalosporium. gramineum in the laboratory and Percentage of whiteheads expressed by seventeen winter wheat Genotypes in naturally infested fields at two locations...... 39

II. 3. Significance of linear contrasts between wheat germplasm Groups for wilting in reaction to a toxic fraction produced by Cephalosporium. gramineum in the laboratory and %whiteheads when grown in naturally infested fields...... 42

II. 4. Variance components analysis for the wilting reaction of the International Triticeae Mapping Initiative (ITMI) mapping progenies after exposure to a toxic fraction produced by Cephalosporium gramineum...... 43 Sensitivity of Wheat Genotypes to a Toxic Fraction Produced by Cephalosporium gramineumand Correlation With Disease Susceptibility

CHAPTER I INTRODUCTION

Importance of the disease

Cephalosporium stripe, caused by Cephalosporium gramineum Nisikado & Ikata

(Hymenula cerealis Eli. & Ev.), is an important vascular wilt disease of winter wheat

(Triticum aestivum) in many parts of the world including southern Africa, temperate

Europe, Japan, and several of northern U.S. states and adjoining Canadian provinces

(Commonwealth Mycological Institute, 1976; Nisikado et al., 1934; Bruehi, 1957; Grey

and Noble, 1960;). The causal organism is a soil-borne fungal pathogen that infects wheat

through the roots and colonizes the vascular system, causing stunting, leaf striping, and

blighted or prematurely ripening heads (whiteheads) with poor grain fill (Bruehl, 1957).

Under favorable conditions for disease, yield can be as low as 20% of a healthy plant

(Johnston and Mathre, 1972). In areas conducive to cephalosporium stripe, 50% yield

reduction from a generalized infection is very common (Mathre et al., 1970; Morton and

Mathre, 1980). Areas under monoculture of wheat with no residue destruction (to prevent

soil erosion) accumulate inoculum in the soil and plant debris, thus increasing the threat

of yield loss. Pathogen biology

Life cycle: Cephalosporium gramineum is an asexual ascomycete. In its parasitic phase, the organism enters wheat roots in the fall and winter. It ramifies throughout the root cortex and then enters the xylem either by active penetration or passively through

breaks in vascular tissue (Stiles and Murray, 1996). Once inside the xylem, the pathogen

colonizes the entire vascular system. Its ability to infest wheat from the base of the plant

to the tip of the racliis, creating a considerable reservoir of infected material upon death

of the host, is key to its survival and future spread (Bruehi and Lai, 1966).

C. gramineum lives in host debris as a saprophyte, and can survive for at least

three years between contacts with a living host (Wiese and Ravenscroft, 1975). Its

mycelium takes rapid and effective possession of the dead substrate, due to the high

degree of initial vascular colonization (Bruehi and Lai, 1968). C. gramineum produces

toxic metabolites which may serve to guarantee it sole access to the host substrate for

saprophytic growth (Bruehi et aL,1969). In the fall, with the onset of cooler, moister

weather, C. gramineum enters its sporodochial stage (Hymenula cerealis El.&Ev.), and

the infested superficial crop debris begins to support profuse sporulation (Bruehl, 1968;

Wiese and Ravenscroft, 1978). Sporodochia form on the outside of the wheat residue, and

large quantities of conidia are produced in a mucilaginous matrix (Wiese and

Ravenscroft, 1978). The conidia, which wash down into the root zone, are the infective

propagules for the next crop (Wiese and Ravenscroft, 1973; Mathre and Johnston, 1975).

As with many soil-borne pathogens, no sexual stage has been reported for C. gramineum.

It reproduces asexually by means of sporodochia that produce copious phialospores, phialides on mycelium, or blastogenously in host xylem vessels (Wiese and Ravenscroft,

1978).

Pathogen variability: Very little is known about the extent of genetic polymorphism in C. gramineum. Infonnation about the extent of variation in the genes controlling interactions of C. gramineum with its hosts could be helpful to improve the usefulness of resistance screening procedures. Van Wert et al. (1984) noted apparently different virulence patterns produced by two Michigan field isolates of C. gramineum in

15 wheat lines. However, Cowger (1998) inoculated winter wheat cultivars from the U.

S. Southern Plains and the Pacific Northwest with isolates from both regions, but no significant cultivar X inoculumsource interactions were found. Two aspects of the life cycle of C. gramineum suggest limited potential of variability. First, C. gramineum has no known sexual stage and is, therefore, likely to be highly clonalwith a limited spectrum of virulence within each lineage (Anderson and Kohn, 1995). Second, C. gramineum is a facultative parasite, and encounters living hosts only in alternate years in regions where summer fallowwheat is grown. Facultative parasites may develop less pathogenic variability than obligate parasites (Ellingboe, 1983).

Influence of environment on disease development Infection of wheat by C. gramineum and the development of disease is highly influenced by environmental factors (Bruehi, 1975; Pool and Sharpe, 1967; Martin et. Al,

1984). The organism survives between plantings of winter wheat by saprophytically

colonizing wheat debris. The survival and future spread of the fungus is greatly

deteimined by its ability to infest wheat through the xylem vessels from the base of the

plant to the tip of the rachis, creating a considerable reservoir of the infected material 4 upon death of the host. In the fall, with the onset of cooler, moister weather, thesurface crop debris begins to support profuse sporulation, and the conidia that wash into the root zone are the main infective propagules. The survival of the propagules, free in the soil, is temperature - dependent and limited. They have a half-life of 0.5 2.5 weeks at23°C in autumn-collected field soil, and a half-life of 17 weeks if the soil is dried at 7°C (Wiese and Ravenscroft, 1975).

Role of inoculum on disease development

The incidence of cephalosponum stripe is a function of inoculum density, although the relative importance of reducing soil inoculum levels in controlling cephalosponum stripe disease is unclear. Researchers have detected both linear and logarithmic relationships between inoculurn density and disease incidence (Mathre and

Johnston, 1975). For resistant genotypes, the response of winter wheat to varying C. gramineuminoculum levels, measured as either percent infection or grain yield reduction, was generally linear over a range of inoculum levels and curvilinear

(logarithmic) for the susceptible genotypes. Specht and Murray (1990) concluded from greenhouse studies that a logarithmic relationship exists between disease incidence and inoculum, implying that large reductions in inoculum are necessary in order to affect the prevalence of the disease. They described the role of inoculum as follows: "unless the amount of inoculum (i. e., naturally infested straw) in a field is reduced by rotation or other measures to a relatively low level, its influence on disease occurrence is likely to be

small. Rather the most important determinants will be environmental conditions (e.g. temperature, moisture, soil pH and other epidemiological factors, such as root wounding)".

Conditions affecting survival of inoculum

C. gramineum is capable of survival in quantities sufficient for substantial re-

infection for at least three growing seasons, which means that neither wheat in rotation

with another crop nor a wheat-fallow-wheat planting sequence can control the pathogen.

In an experiment in Washington, average survival of C. gramineum after 18 months at all

measured depths in the field was 32% at Pullman, 20% at Lind, and 11% at Vancouver.

When surface straw alone was considered for the same period of time, survival was 78%

at Pullman, 50% at Lind, and 38% at Vancouver (Lai and Bruehi, 1966). Michigan

experiments indicated that straw undisturbed on the soil surface decayed slowly enough

to support saprophytic growth during fall and winter for three consecutive years. Soil

populations of the fungus vary considerably during the year, and are usually highest

between October and February as the sporodochial stage, Hymenula cerealis, begins

profuse sporulation (Wiese and Ravenscroft, 1975). Survival in straw appears to vary

according to soil moisture and pH. Soil type and soil microflora type did not seem to

affect survival significantly (Lai and Bruehl, 1966).

pH: Low pH levels favor both sporulation and survival of C. gramineum conidia,

and, hence, the subsequent development of disease (Love and Bruehi, 1987; Murray

1987; Murray et al., 1991). Bruehl and Lai (1968) found that C. gramineum survived

longest in infested straw when soil pH was 3.9 to 5.5, and attributed this to the production

of a wide-spectrum antibiotic that is most active at low pH. Specht and Murray (1989)

concluded that sporulation on artificially colonized oat kernels on soil, and survival of 6 conidia in soil, was higher at soil pH 4.7 than in the pH range of 5.7-7.5. Further, they proposed that acid soil may favor pathogenicity by promoting increased host susceptibility to root infection, possibly as a result of greater root stress and damage and/or slower woundhealing in acid than in neutral or alkaline soil. Murray et al. (1992) suggested that soil pH has the greatest influence on cephalosporium stripe incidence under field conditions in years when root injury is relatively minor. They found that soil pH affects initial penetration and establishment of C. gramineum in the plant, but has no effect on subsequent colonization. Disease severity, in contrast to infection, is not significantly affected by soil pH. Instead, it may be more the result of temperature and rainfall in the autumn and winter, or cultural practices (Murray et al., 1992).

Soil moisture: Specht and Murray (1989) found that soil matric potential has a major effect on conidial longevity, with greatest survival in the driest (-0.O6MPa) soil tested.

They speculated that poor conidial survival in more moist (-0.03 and 0.01 MPa) soil was due to low oxygen concentrations. Sporodochia of H. cerealis are favored by wet, cool weather (Wiese and Ravenscroft, 1975).

Temperature: C. gramineum thrives at low temperatures. In laboratory studies, sporulation was especially profuse at 7°C; with rising temperatures it diminished and aerial hyphae flourished (Bruehi, 1957). Other laboratory trials conducted at 10, 15, and

20°C showed that C. gramineum persisted longer in straw at the lower temperatures, even though its maximum radial growth in pure culture is at 20°C (Lai and Bruehi, 1966). 7

Infection process

Alternative hypotheses have been proposed for the main mechanism of infection of host roots by C. gramineum. Mathre and Johnston (1975) asked how the pathogen travels from the straw in which it has survived saprophytically, and how it invades the host.

They concluded that while soil per se does not limit mycelial growth and sporulation, a fungistatic factor operating in natural soil inhibits both processes, especially sporulation.

Even the presence of exudates from crushed or severed roots of a highly susceptible wheat variety did not overcome this inhibitory effect. "We visualize the infection process," Mathre and Johnson wrote, " as being one where comdia are 'vacuumed' into xylem vessels exposed when roots are severed in the spring by frost heaving of soil."

In the same vein, Morton and Mathre (1980) examined the influence of soil

microflora on infection in environments in which mechanical root breakage did not

occur. Three red winter wheat cultivars with varying levels of resistance to C. gramineum

were grown in the greenhouse in field soil that was either heat-sterilized or left

unsterilized. Field seeding and inoculation rates were approximated. In neither treatment

was substantial infection observed. The same wheat varieties were also planted in the

field in spring, after danger of frost heaving had passed. Despite artificial inoculum of up

to 5 xi05conidia/gram of soil in the first month following planting, less then 2%

infection was evident. "Assuming that soil frost heaving causes root breakage," Morton

and Mathre concluded, "these data suggest that root wounding is necessary for successful

pathogenesis by C. gramineum." Morton et al. (1980) studied the relation between foliar

symptoms and systemic advance of C. gramineum during winter wheat development and concluded that the pathogen is incapable of penetrating living cells at any time during the disease cycle.

Specht and Murray (1990) agreed that broken roots provide a major avenue of infection for C. gramineum. In greenhouse studies using pot culture, they slashed roots with a knife 30-60 minutes before applying a 100-mi drench of conidia. They found that comdial inoculum densities needed to cause disease in unwounded plants were several fold of magnitude higher than those required to cause similar levels of disease in root- wounded plants.

Bailey et a! (1982) raised doubts about the passive infection theory. They observed that C. gramineum conidia were stimulated to germinate by freeze-induced root exudates and produce mycelium which penetrated the host roots. They found that exudates collected from wheat roots that had been frozen and thawed increased fungal spore germination, hyphal branching, and conidiogenesis more than did exudates from non- frozen roots. In fact, exudates from cut roots did not stimulate germination.

Noting that other soil-borne vascular pathogens, such as Fusarium and

Verticiiium, gain entry by active penetration, Bailey et al. questioned the importance of root-wounding in the natural infection process. They found that 62% of plants with cut roots developed infection when the roots were dipped in a conidial suspension and transplanted to soil, whereas only 0-3% of plant with cut roots became infected when transplanted into inoculated soil. This offered evidence, they said, that conidia are not readily transported in soil, at least in the clay loam soil used in the experiment. Bailey et al. agreed with other researchers that freeze stress is an important factor predisposing wheat plants to cephalosporium stripe. However, they suggested that the role of freeze- induced exudates in stimulating active penetration by C. gramineum may be greater than the role of wounds created by frost heaving in permitting passive ingress of conidia.

Buttressing their case, these workers noted that plant tissues subjected to freezing temperatures release more exudates than do non-frozen tissues. They also pointed out that an infection process relying on active transpiration to "vacuum" passive propagules into the vascular system would be hampered by the fact that, at least in Michigan, wheat fields are covered by snow and plant transpiration rates plummet during the December-January period when C. gramineum inoculum reaches peak density.

Resistance source and mechanisms of resistance

Various sources of resistance to cephalosporium stripe disease in winter wheat have been proposed. Nearly complete resistance is present in wild wheat relatives, including Agropyron elongatum (Mathre et al., 1985). Ultimately, a particular host's susceptibility may derive from a combination of several factors. Martin et al. (1986) identified three types of resistance among cultivars subjected to increasing inoculum levels. 1) Two moderately resistant varieties appeared to limit ingress to the host, based on a low score in total percentage of infected tillers. 2) Another moderately resistant variety experienced a relatively high infection level but only a smallgrainyield reduction, thus possibly possessing some kind of tolerance to C. gramineum. 3) A variety with intermediate infection and yield scores seemed to restrict pathogen movement after infection, as relatively few tillers were infected per plant. Among the proposed sources of resistance, the following are ordered in keeping with the development of the disease: 10

Root breakage and duration of wound susceptibility: Martin et al. (1989) found no differences among cultivars with respect to root tensile strength, which they took as a measure of resistance to root injury. The same researchers also found previously

(1975) that, in wheat cultivars with a range of overall resistance to C. gramineum, severed roots remained susceptible for at least 16 days after injury. They reported that the percent of plants infected, treated with either a high (5 x106 conidia/mi)or low (5 x iø comdia/ml) inoculum density was highest when plants were inoculated within 5 minutes after wounding. The infection percentage declined sharply until two days between wounding and inoculation, and then leveled off if the ends of roots broken during frost heaving are the main entry point for conidia, the time span over which the broken ends remain open may be an important determinant of variability among cultivars. Specht and

Murrray (1990) pointed out that wheat roots may be more prone to breakage in acid than in neutral soil, and wounds may heal more slowly at low pH. Hence, varying levels of tolerance among cultivars may partially explain differential resistance.

Crown or internal root barriers: Mathre and Johnston (1990) studied whether resistant wheat cultivars or wild wheat relatives present barriers to the movement of C. gramineum through roots or across the root-shoot interface. The researchers did fmd that movement of the fungus through root and especially through crown tissues was slowest in Elytrigea elongata and Thinopyron interinedium, followed by Agrotriticum (a fertile

cross between E. elongata and 7'. aestivum), and finally 7'. aestivum cultivars. Mathre and

Johnston compared this type of resistance to that observed in hosts challenged by other

vascular wilt diseases, such as Fusarium wilts of tomato, radish, and sweet potato. They

note that the Gramineae possess a particularly complex transition region between root 11

and culm tissues, including a scutellar nodal plate below the insertion of the scutellum, which they hypothesized could present a barrier to the pathogen. Unfortunately, they

noted, crown tissue characters are probably under complex genetic control and may be

difficult to transfer from wild relatives to cultivated wheat.

Host support to sporulation: Inoculum density is one factor affecting host

response (Mathre and Johnston, 1975; Specht and Murray, 1990). Mathre and Johnston

(1975), noting that a susceptible cultivar responded more strongly to increased inoculum

levels than did a moderately susceptible cultivar, suggested that the former supported

greater sporulation within the xylem vessels than the latter. Shefelbine and Bockus

(1990) investigated this thesis and concluded that 1) cultivars do infact differ with respect

to conidia produced per gram of straw, and 2) there is no evidence of correlation between

overall resistance to stripe disease and hospitality to sporulation.

Aluminum toxicity: It has been suggested (Love, 1994) that susceptibility to

cephalosponum stripe and to aluminum toxicity may be associated in winter wheat.

Among Pacific Northwest cultivars, Hill 81, Yamhill, and possibly Madsen possess

resistance to both cephalosporium stripe and aluminum damage (Lefever et al.,1977).

However, no formal investigation of such a possible link appears to have occurred to

date. Anderegg and Murray (1988) noted thatAl3and H, both present in soils they

studied, could injure wheat roots. They said that, "high soil moisture and low soil pH may

thus be important factors that predispose wheat plants to infection by C.gramineum."

Although the physiological effects of aluminum are still not well understood, it is

known to inhibit mitosis and elongation in roots of many plant species (Rengel, 1992). It

also affects membrane permeability for both electrolytes and non-electrolytes, and 12 reduces accumulation of divalent cations, especiallyCa2and Mg2, by interfering with membrane transport. Interference withCa2uptake and reduction of normal levels of cytosolicCa2may be a major component of aluminum toxicity (Huang et al., 1993).

Rhizosphere-dynamics, root camouflage, and microbial antagonism: For some crops, there is considerable evidence that the rhizospheres of resistant and susceptible cultivars differ in ways that affect the attractiveness of the host to soil-borne pathogens.

An important phenomenon that may distinguish potential hosts is the "rhizosphere effect": quantitative and/or qualitative differences between microbial populations on plant roots and in root-free soil (Gilbert et al. 1993). To date, investigation of possible rhizosphere effects in the winter wheat/C. gramineum disease system has been limited to the following: 1)the studies of Bailey et al. (1982) concerning freeze-induced exudates;

2) Mathre and Jonston's (1975) finding of fungistatic factors in natural soil; and 3)

Mortin and Mathre's (1980) conclusion that soils they tested did not appear to contain microorganisms capable of promoting infection by C. gramineum in the absence of mechanical root wounding. None of these findings is inconsistent with the idea that rhizosphere microorganisms may be affecting the pathogemcity of C. gramineum. The main area of disagreement concerns the relative importance of active and passive infection mechanisms.

Potential control measures

Cephalosporium stripe can be controlled by: 1) delaying seeding in the autumn; 2) liming the field 3) crop rotation 4) proper residue management! removing or destroying infested residue with deep plowing or burning, and 5) planting resistant cultivars. 13

Delayed seeding results in plants with smaller root systems that are less susceptible to winter root injury, and therefore disease incidence. Raymond and Bockus (1984), in a two year trial in Kansas, found that disease reduction in the first year due to late planting was significant but not in the second. Moreover, there was a 13.6% yield reduction for uninoculated plots with each week of delay beyond the optimum planting date and late planting also aggravates the problem of soil erosion.

Liming can reduce disease severity by elevating soil pH. Murray et al. (1992) found a

16.2% decrease of C.gramineuminfection by applying lime at 5000 lbs/acre in a

naturally infested field in Washington. However, liming is economically feasible only

where there is a ready and inexpensive source of lime.

Long crop rotations can be effective, but are not always economically feasible.Deep

plowing and burning of debris can reduce inoculum doses. Bockus et al. (1983)

concluded from their three-year field experiment that burning wheat stubble was the most

effective disposal method for reducing cepahalosporium stripe after a severe outbreak

under a continuous winter wheat production regime. After three years of plowing,

cephalosporium stripe incidence was the same as after three year of burning; therefore,

residue burning and! or continuous plowing is expected to effectively help maintain low

disease losses (Murray and Bruehi, 1983). Latin et al. (1982) also found that long rotation

and conventional plowing could decrease cephalosporium stripe incidence. However, in

most major wheat-growing regions, wheat debris on the soil surface is needed to hold

moisture in the soil and prevent soil erosion. 14

The development of host resistance currently offers the best hope for control of the disease, as there are no fungicides registered for the control of the disease. Shefelbine and

Bockus (1989) suggested that repeated cultivation of moderately resistant winter wheat varieties could reduce cephalosporium stripe incidence over time. In their three-year experiment, percentage reduction in disease measurements, even when normalized for environmental fluctuations by expressing disease severity as a percentage of that of the most susceptible cultivar, were substantially greater for moderately resistant cultivars than for susceptible cultivars. Though no soft white winter wheat cultivars show complete resistance to cephalosporium stripe, there is considerable variation in the degree of resistance among cultivars and the resistance is quantitative (Cai et al,. 1996; Bruehl et al., 1986; Mathre et al.,1977;Martin et. al 1983; Morton and Mathre, 1980). Levels of resistance currently available are adequate to reduce inoculum levels over time and control the disease in the long-term (Montana State Extension Service, 1978). However, identifing resistance in breeding programs remains problematic. Expression of resistance is incomplete and environmentally dependent (Martin et al., 1986) and the percentage of infection does not always correlate with actual resistance unless systemic symptoms are expressed (Bockus and Sim, 1982). Only one field trial can be completed per year and the ranking of cultivars can vary substantially from year to year depending on prevailing environmental conditions (Bruehi et al., 1986), rendering multiple trials both necessary and contradictory. Further, the disease tends to be aggregated within fields, thus requiring large plots to make useful comparisons, which is not possible in

early generations of cultivar development. Some other techniques such as root

inoculation proved to be laborious and difficult (Mathre and Johnston, 1983). Therefore, 15 rapid initial screening of cultivars under controlled conditions is desirable. Until now, most controlled environment work with cephalosporium stripe has been done with vernalized, adult wheat plants, requiring a total trial time of 5-6 months. Exceptions are the hydroponic seedling assay (Cowger and Mundt, 1998 ) for resistance to cephalosporiuin stripe and a method for screening winter wheat seedlings for cephalosporium stripe resistance in pots that distinguished highly susceptible from highly resistant lines, but was found less useful for varieties with intermediate susceptibility

(Van Wert et al., 1984). The hydroponic assay provides a turnaround time of only one month. However, variability associated with the technique requires significant replication, as well as production of large amounts of pathogen inoculum (Cowger and

Mundt 1998). The above problems suggest a need for evaluating resistance to cephalosporium stripe under more refined conditions.

Role of toxin in cephalosporium stripe development

Toxin production by pathogens, and the relationship of toxins to pathogenesis, have been studied for decades. However, disease severity depends on a succession of interactions between pathogen and the plant, and the specific role of toxin is not always clear. Application of toxin for resistance screening was first employed over 45 years ago by Wheeler and Luke (1955), by using relatively crude toxin preparations to screen large populations of host materials, e. g., whole seedlings, plant parts, protoplasts, or tissue cultures, for resistant individuals and this practice has continued (Bains and Tewari,

1987; Rines and Luke, 1985; Vidhyasekaran et al., 1986; Ling Ct al., 1985). The practice soon extended to the selection of useful plant genotypes or variants as a step in in vitro 16 breeding programs, before the regeneration of plants (Helgesen and Deverall, 1983;

Ingram, 1986). First applied using host-specific toxins (metabolites produced by pathogens causing plant diseases which selectively damage only varieties susceptible to the pathogens), the practice has now spread to non-host selective toxins as well

(Gengenbach and Rines, 1986; Hartman et al., 1986; Hess and Weber, 1988; Ling et al.,

1985; Nadel and Spiegel-Roy, 1987). The pertinent question is, whether in vitro selection is sufficiently effective. Simply showing that differential tolerance can be obtained is not adequate without some degree of assurance that these "resistant" individuals or their progeny will exhibit increased disease resistance under a variety of field conditions.

Although screening for disease resistance is a more straightforward task with host selective toxins, non-host selective toxins can also be used to identify resistant species, cultivars, or clones. For example Fusicoccin (FC) has been used to assess the sensitivity of stone fruit varieties to the toxic effects of Fusicoccin amygdali, the causal agent of bud canker (Turner and Graniti, 1976; Bottalico, 1976).

The symptoms of cephalosporium stripe suggest a pathogen-produced toxin or xylem-plugging compound and have led to investigation of the production of antibiotics, toxins, and extra-cellular polysaccharides by the pathogen. Extra-cellular polysaccharides have been implicated in wilt diseases (Spalding et al., 1961; Pool and Sharpe, 1969).

Symptoms of cephalosporium leaf stripe include wilting at higher disease severity, where wilting is observed with accompanying dwarfing of plants. Spalding et al. (1961)

concluded that extracellular polysaccharides produced by the fungus resulted in xylary

plugging; however, Weise (1972) reported that occlusions were due to fungal

proliferation that developed only after lateral extension of leaf striping. 17

Kobayashi and Ui (1977) isolated and characterized graminin A from culture filtrates of C. gramineum.This toxic compound was reported to cause yellowing at concentrations of 25 .tg mY' in excised leaves (Kobayashi and Ui, 1979). It was shown to be structurally similar to gregatin A, a compound isolated from culture filtrates of

Cephiosporium gregatum, which mimicked the symptoms of brown stem rot of soybeans; however graminin A was more selective for wheat. Graminin A possesses antimicrobial activity and has been shown to affect stomate function in the same manner as infection by

C. gramineum (Creatura et al., 1981), though Van wert et al. (1984) found that pathogenicity and virulence of C. gramineum is independent of in vitro production of extracellular polysaccharides and graminin A. If graminin A contributes to pathogenesis of wheat, it could be a useful compound for selecting resistant or tolerant wheat lines.

Hence, this study was undertaken as an attempt to evaluate the utility of in vitro produced toxic fraction(s), with or without 'graminin A', as a compound(s) to screen wheat genotypes for resistance by comparing the reactions obtained from field experiments and to understand its role in causing cephalosporium stripe of wheat.

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CHAPTER II

SENSITIVITY OF WHEAT GENOTYPES TO A TOXIC FRACTION PRODUCED BYCEPHALOSPORIUM GRAMINEUMAND CORRELATION WITH DISEASE SUSCEPTIBILITY

M. RAHMAN.', C. C MUNDT.', T. J WOLPERT.', AND 0.RIERA-LIZARAZU2

'Department of Botany and Plant Pathology.

2Department of Crop and Soil Science

Oregon State University

Corvallis, OR 973312902

Prepared for submission toPHYTOPATHOLOGY

American Phytopathological Society, St. Paul, MN

September 2000 26

Abstract

Cephalosporium stripe is an important disease of winter wheat (Triticum aestivum) in several areas of the world, especially where stubble mulch and early seeding are practiced to maintain soil moisture and prevent erosion. We developed a procedure to mass-produce a toxic fraction produced by Cephalosporium gramineum through a modification of the method of Kobayashi and Ui (1977). Exposure of excised wheat leaf to a concentration of(60p.1/ml) of the toxic fraction for 72 hours produced distinct wilting symptoms that allowed us to distinguish wheat genotypes on a continuous 1-5 scale in a repeatable manner. Twenty wheat genotypes, belonging to four distinct germplasm

groups (common, club, durum, and synthetic) were evaluated. Sensitivity of the

individual genotypes to the toxic fraction did not differ significantly within germplasm

groups, but all differences among the means of the four germplasm groups were highly

significant (P0.0001) based on linear contrasts. Seventeen winter wheat genotypes

representing the common, club, and durum germplasm groups were planted in C.

gramineum-infested fields at two locations. The percentage of tillers showing whitehead

symptoms significantly correlated with wilting symptoms measured by the toxin assay.

Failure of the toxin assay to identify some of the moderately resistant common wheat

genotypes suggests that additional mechanisms of resistance are operative as well. As no

toxin-insensitive genotype showed susceptibility to cephalosporium stripe in the field

experiments, however, the toxin assay can be used as an initial screening procedure to

reduce the number of genotypes to be tested in the field. 27

Introduction

Cephalosporium stripe of wheat, caused by Cephalosporium gramineum Nisikado

& ilcata (Hymenula cerealis Eli. & Ev.) is an important monocyclic, vascular diseaseof winter wheat in many parts of the world, including Europe, eastern Africa, Japan, several of the northern U.S. states and adjoining Canadian provinces (Commonwealth

Mycological Institute, 1976.; Nisikado et al., 1934; Bruehi, 1957; Grey and Noble, 1960).

The causal agent is a facultative ftingal parasite that enters wheat roots in the fall and

winter, and colonizes the vascular system. Symptoms can include general chiorosis and

diagnostic chlorotic leaf striping. Severe infections can cause wilting, leaf necrosis,

stunting, and prematurely ripening heads (whiteheads) in the spring and summer (Slope

1962). Under favorable conditions, the disease can cause as much as 80% yield loss

(Johnston and Mathre 1972). Colonized residue is returned to the soil after grain harvest

and serves as the source of primary inoculum. Conidia produced on the residue infect the

next crop to complete the disease cycle. In the absence of hosts, the fungus oversummers

as a saprophyte in wheat debris and can remain viablein superficial wheat straw for at

least three years (Bruehi et al., 1964; Lai and Bruehi, 1966).

Cultural controls for cephalosporium stripe (delayed planting, burning of crop

residue, and deep plowing) are either economically infeasible or increase soil erosion

(Latin et al., 1982; Bockus et al.,1983) and no chemicals are registered for control of the

disease. The development of host resistance currently offers the best hope for control of

cephalosporium stripe. Though no wheat cultivars show complete resistance to

cephalosporium stripe, there is considerable variation in the degree of resistance among

cultivars (Bruehi et al.,1986; Mathre and Johnston, 1975) and repeated planting of 28 moderately resistant cultivars can reduce both the incidence and severity of cephalosporiuin stripe (Shefelbine and Bockus, 1989). Levels of resistance currently available are adequate to reduce inoculum levels over time and control the disease in the long-term (Montana State Extension Service, 1989). However, current methods available to identify resistance in breeding programs are variable and prohibitively time-consuming

(Bruehi et al., 1986; Mathre and Johnston, 1983).The symptoms of cephalosporium stripe suggest the involvement of pathogen-produced toxins or xylem-plugging compounds and have led to investigations on the production of antibiotics, toxins, and extra-cellular polysaccharides by the pathogen (Pool and Sharpe, 1969). Spalding Ct al. (1961) found that extracellular polysaccharides produced by the fungus resulted in xylary plugging; however, Weise (1972) reported that occlusions were due to flingal proliferation that developed only after lateral extension of leaf striping. Kobayashi and Ui (1977) isolated and characterized graminin A from culture filtrates of C. gramineum . This toxic compound was reported to cause yellowing of excised leaves at concentrations of 25 ig mi'(Kobayashi and Ui, 1979). The toxin was shown to be structurally similar to gregatm

A, a compound isolated from culture filtrates of Cephalosporium gregatum, which mimicked the symptoms of brown stem rot of soybeans; however, graminin A was more selective for wheat. Graminin A possesses antimicrobial activity and has been shown to affect stomate function in the same manner as infection by C. gramineum (Creatura et al.,

1981). The goals of our study were to determine if resistance of wheat genotypes to toxin

is associated with resistance to cephalosporium stripe and to evaluate the utility of in-

vitro produced 'toxic activity' as a tool to screen wheat genotypes for resistance. 29

Materials and Methods

Laboratory assays

Biological materials.Twenty wheat genotypes were evaluated, representing four taxonomic groups: 1) Common wheat, Triticum aestivum L. (AABBDD). These included

9 soft white winter wheat cultivars and one soft white winter wheat breeding line that are adapted to the Pacific Northwest (PNW) region of the USA, and one hard red cultivar,

Opata 85. Opata 85 is a spring wheat cultivar developed at the International Maize and

Wheat Improvement Center (CIMMYT) and a parent of the International Triticeae

Mapping Initiative (ITMI) mapping population (Van Deynze et al., 1995). 2) Five winter club wheat cultivars adapted to the Pacific Northwest. Club wheats are hexaploid

(AABBDD) T. aestivum types with compact heads, and originate from a different and substantially smaller gene pool than the common wheats. 3) Three durum wheat genotypes, T. turgidum (AABB). These included two advanced winter durum lines from the Oregon State University (OSU) wheat-breeding program and one CIMMYT spring durum wheat cultivar (Altar 84) that is a "grandparent" of the ITMI mapping population

(see below). 4) The synthetic hexaploid wheat M6 (W-7984) is the second parent of the

ITMI population. M6 was produced by crossing Altar 84 with an accession of Aegilops tauschii (TA 2465,DD), followed by doubling chromosome number with colchicine to obtain a synthetic hexaploid.

We also evaluated a total of 112 Recombinant Inbred Lines (RILs) from the cross between M6 and Opata 85. Seed for the recombinant inbred line population, M6, and

Opata 85 were obtained from Dr.Calvin 0. Qualset (Genetic Resources Conservation 30 durum wheat breeding program advance lines were obtained from Dr. Karim Ammar

(Dept. of Crop and Soil Science, Oregon State University, Corvallis

Ten seeds of each genotype were planted in 10-cm plastic pots containing an organic soil mix and fertilized with the recommended rate of Miracle Gro (Stern's

Miracle Gro Product, Inc. Port Washington, NY 11050) at 7 days after planting. Pots were placed in a growth chamber in a completely randomized design at 21C with a 16- hr photoperiod.

C. gramineum-infected wheat stems were collected from a naturally infested wheat field near Dufur, Oregon. An isolate was obtained by placing a surface - sterilized stem piece on potato dextrose agar (PDA). The culture was maintained on PDA at 5-7C for use in subsequent laboratory assays.

Extraction of toxic fraction.

Culture conditions and extraction of the toxic fraction were first done according to the methods of Kobayashi (1978). The extracted substance produced yellowing of excised wheat leaves at low concentrations and wilting at higher concentrations. The substance produced dark blue spots under UV radiation, consistent with the properties of gramirnn A (Kobayashi and Ui, 1977). Extraction methodology was subsequently modified to further refine the toxic substance and increase its yield, as described below.

Nine liters of broth were prepared by mixing 50 g sucrose, 1.3 gK2HPO4,ig

KH2PO4, 0.5g(NH4)2SO4and 0.5g MgSO4.7H20 in each liter of distilled water. The medium was dispensed in 500 ml flasks (200 mi/flask) and autoclaved at 121°C and 15

PSI for 15 minutes. Fifty ml of sterile water was added to a PDA plate of C. 31 gramineumand then scraped with a sterile glass slide. The spore suspension from the plate was added to a 500 ml flask containing 100 ml potato dextrose broth, which was used as a seed-flask. This flask was then shaken on a Burrell Wrist-Action Shaker

(Burrell Corp., Pittsburgh, PA) for 72 hours to raise the spore concentration to approximately 1.3-1.5 X 1081m1. The spore suspension was then added to each flask to attain 2 XiC)5spores per ml. At 24 hr after inoculation, flasks were shaken well and then incubated in standing culture for 35 days at 25°C.

After 35 days of incubation, broth was sieved by a kitchen sieve and then filtered through 18.5 cm Whatman filter paper No. 3 (W. & R. Balston, Ltd.; England) followed by 47 mm Millipore (Prefilter AP 25, Millipore Corporation, Bedford, MA

01730). With our modified method, the most time-consuming step of rotary evaporation of the culture filtrate was avoided. Culture filtrate was directly extracted four times with ethyl acetate in a 2 L separating funnel. Ethyl acetate was added at one tenth of the volume of culture filtrate each time. Pooled ethyl acetate extracts were then subjected to rotary evaporation to give an oily residue. The oily residue was then solubilized in 18 ml chloroform and left under a hood overnight to dry.

After drying, the crude toxin was dissolved in 5.4 ml of ethanol. One third of the sample, 1.8 ml, was diluted to lOml with water and loaded onto a C18 reversed-phase

Sep Pak cartridge (Millipore Corp.,Millford, Mass), which had been equilibrated in water. The cartridge was then progressively eluted with 10 ml each of water, 25%, 50%,

75%, and 100% acetonitrile in water (v/v). This entire process was repeated two more times with the remainder of the ethanol-solubilized crude toxin and all equivalent fractions pooled. Each pooled fraction was subjected to rotary evaporation and the 32 fractions pooled. Each pooled fraction was subjected to rotary evaporation and the resulting oily residue dissolved in 6m1 of chloroform, transferred to a smaller vessel,

dried under a hood overnight, and finally dissolved in 900 tlethanol. Initial assays indicated that the majority of the toxic activity was contained in the75%acetonitriie/ water (v/v) fraction. Consequently, only this fraction was used for the genotype evaluations described below.

Assay method

Seven-mi Solvent Saver scintillation vials (Kimble, Vineland, NJ08360) were numbered and marked for each wheat genotype in three replications per run, plus one control vial. Each of the three treatment vials received 840 il water and 60 p1 ethanol- dissolved toxic fraction; control vials received 60 p1 ethanol. Leaves were excised from

14-day-old plants with scissors and placed in the marked vials, one leaf per vial. To maintain leaf uniformity for all genotypes,onlysecond leaves were chosen from robust plants in each pot, and leaves were cut to the same length. Vials containing the leaves were kept standing in a paperboard rack in a growth chamber at 21 C and a 16-hr photoperiod. After 24 hrs, 1 ml of water was added to proteót the leaves from desiccation.

The experiment was repeated twice, with three replications per genotype for each of the three runs. Leaves started showing wilting within 24 hour of exposure to the toxic fraction (Fig. 1) and sensitive genotypes were severely wilted within 72 hr of exposure when data were recorded. In contrast, control leaves did not change in appearance during this time. Wilting was evaluated on a continuous scale of 1 to5,with 1 indicating no change in leaf appearance and 5 indicating a fully wilted and dried leaf (Fig. 2). 33.

Statistical analyses

As none of the control leaves showed visible symptoms, theywere not included in

the statistical analyses. Wilting data for the toxin-treated leaves from allthree runs were

subjected to analysis of variance using the General Linear ModelsProcedure (PROC

GLM) of the Statistical Analysis System (SAS Institute Inc., Cary, NC).In the first

analysis, we considered genotype, run, and genotypex run as the experimental factors.

As the genotype x run interaction was found to be insignificant (P= 0.818), run and

genotype x run were dropped and the data reanalyzed. Fisher's protected least

significant difference test (P = 0.05) was calculated for thepurpose of comparing

individual genotype means. Differences among the four taxonomicgroups (common,

club, durum, and synthetic) were evaluated using six preplanned linearcontrasts.

Fig. 1. Differential response of the ITMI population parents at 24 hr ofexposure to a toxic fraction produced by Cephalosporium gramineum. 34

1' /

1.25 2 3 4 5

Fig. 2. Scale used to assess the degree of wilting of wheat leaves after 72 hr of exposure to a toxic fraction produced by Cephalosporium gramineum. Black bar on a leaf indicates the extent of wilting down the leaf.

Field experiments

Locations and experimental design

Experiments were conducted in commercial fieldsnear Dufur and Condon, Oregon.

Both fields had a history of severe cephalosporium stripe, andwere planted into stubble mulch and with mid-September planting dates to furtherencourage disease severity.

Elevation and mean annual precipitation are 900 m and 35.8cm for the Condon site, and 600 m and 31.8 cm for the Dufur site.

The experiments were arranged in a randomized complete block design withthree replications at Condon and four replications at Dufur. Allgenotypes evaluated in the laboratory/growth chamber experiment were included, except for the three spring 35 genotypes Opata 85, Altar 84, and M6. Plots were eight rows wide with 35.6 cm between rows, and were planted with a plot drill at 200 seeds per m2. Planting dates were 15 September 1999 at Condon and 16 September 1999 at Dufur. Due to limitations on available area, plot lengths varied from 7.5 to 30 m. Fertilization and other cultural practices were conducted by cooperating farmers and were standard for commercial production in the area.

Data collection and statistical analysis

Each plot was examined by two individuals, at the late milk/early dough stage, to estimate the percent whiteheads (heads showing early maturity and/or with poor or no grain fill). Whitehead data for each location were subjected to analysis of variance for a randomized complete block design using the General Linear Models Procedure (PROC

GLM) of the Statistical Analysis System (SAS Institute Inc., Cary, NC). To better satisfy the assumption of homogeneity of variance, data for Condon were first transformed to log10(Y + 0.5) and those of Dufur were transformed to log(Y + 0.5).

Fisher's protected least significant difference was calculated for comparison among individual genotypes. Germplasm (common, club, and durum) differences were evaluated using three pre-planned linear contrasts. In addition, whitehead percentages in field plots and wilting ratings from the toxin assays for all seventeen genotypes that were common to both the lab and field experiment were subjected to canonical correlation analysis using the Analyst application of the Statistical Analysis System

(SAS Institute Inc., Cary, NC). 36

Mapping progeny evaluation

Detached leaf assay described earlier was followed to obtain toxin sensitivity data from the individual mapping progeny. As the toxin extracted from the 9 L broth was standardized to evaluate 30 genotypes/progenies at a time, we evaluated the progenies in four different batches. Each time we included the resistant parent (M6) and susceptible parent (Opata 85) as a resistant and susceptible check, respectively.

Heritabifity estimate. Variance components were computed by equating mean squares to their expectations (Table 1) The genetic component of variance. (&g) was estimated as = (M3Ml)/r. Approximate heritability estimates were calculated on a plot basis ash2 =&gI(&g + &e/r) where &g and &e are the sample genotypic and error variances, respectively, and r is the number of replications The 90% confidence interval limits for estimates of heritability were calculated according to Knapp et al.

(1985). 37

Table 1. Variance components for analysis of sensitivity of 112 recombinant inbred wheat lines to a toxic fraction produced by Cephalosporium gramineum

Source of variation d.f. Mean squares Variance expectations Genotype g - 1 M3 +rog

Replications r-1 M2 a2g +g&r

Error (g1)(r-1) Ml O2gr = 38

Results

Reaction of genotypes to toxic fraction

The main effect of wheat genotype in the ANOVA was highly significant (P

0.0001). The common wheat genotypes were all highly sensitive to the toxic fraction

(wilting rating of 4 or greater). Fischer's protected LSD indicates that there is significant variation among genotypes within the common and club groups, but no overlap between these two groups. The three durum wheats showed highly similar reactions, and were intermediate between that of the common and the club groups. The synthetic wheat M6, which is a parent of the ITMI population, showed the highest level of resistance.

Fortuitously, the other parent of the ITMI population (Opata 85) was the most susceptible genotype evaluated, thus providing an opportunity for studying inheritance of resistance to the toxic fraction among progeny of this cross (see below). Linear contrasts were highly significant (P0.00 1) for the six pair wise combinations among the four germplasm groups (Table 3).

Field reactions Disease levels were lower (Table 2) and variation among plots was greater (data not shown) at the Condon than at the Dufur site. Nonetheless, rankings among genotypes were similar between sites (Tables 1), with a correlation coefficient of 0.90 (P0.0001) and the main effect of wheat genotype was highly significant in the analysis of variance for each site (P0.0001). On average, the two durum wheats were less susceptible than the common wheats (Table 2), though there was some overlap with the more resistant common wheat genotypes (Table 2). The club wheats were consistently more resistant than either the common or durum wheats (Tables 2 and 3). 39

Table 2. Mean wilting reaction of twenty winter and spring wheat genotypes to a toxic fraction produced by C.gramineumin the laboratory and percentage of whiteheads expressed by seventeen winter wheat genotypes in naturally infested fields at two locations

Germpl- Genotype Wiltinga Whiteheads asm COndonc Dufurd I group Untrans- Trans- Untrans- Trans- formed formed formed formed Common Opata85b Madsen 4.93 1.75 0.31 3.62 1.39 Macvicar 4.89 20.67 1.32 25.29 3.25 Malcolm 4.73 12.67 1.06 32.75 3.46 Stephens 4.68 14.17 1.13 46.75 3.83 Gene 4.52 8.67 0.96 33.00 3.50 Lambert 4.44 5.25 0.58 23.00 3.13 Rod 4.39 3.33 0.57 5.13 1.68 Weatherford 4.32 6.17 0.82 7.88 2.09 Hill 81 4.01 4.83 0.57 8.26 2.17 0R515 4.00 11.83 1.05 29.75 3.37 Mean 4.53 8.93 0.84 21.54 2.79 Durum 0R948927 2.99 4.00 0.65 6.75 1.96 0R971897 2.89 2.17 0.42 9.50 2.29 84b Altar 2.89 - - - - Mean 2.92 3.09 0.54 8.13 2.13 Club Temple 2.53 0.05 -0.25 1.44 0.61 Rohde 2.25 0.07 -0.25 1.13 0.49 Coda 2.09 0.62 0.05 1.38 0.55 Hyak 1.53 0.70 -0.04 0.35 -0.23 Tyee 1.47 0.68 0.03 0.05 -0.6 Mean 1.97 0.34 -0.09 0.87 0.16

Synthetic M6" 1.17 - - - -

Mean 1.17 - - -

LSD(0.05) 0.61 - 0.38 - 0.42 40

Table 2. (Continued) aCofltiflous scale with 1 = no effect of toxic fraction on leaf appearance and 5 = fully wilted and dried or necrotic leaf. bGenotype could not be included in the field experiments because it is a spring type. cDataare means of three replications, the transformed scale was log1o(%whiteheads + 0.5). dDataare means of four replications, the transformed scale was log(%whiteheads + 0.5).

Comparison of laboratory assay to field data There was more variation among genotypes within germplasm groups for the field data than for the toxin data (Table 2). Nonethless, rankings of mean disease levels for the three germplasm groups tested in the field were the same as for the toxin assay, with the common wheats most susceptible, the club wheats most resistant,and durum wheats intermediate. Wilting symptoms measured in growth chambers were significantly (P S 0.0001) correlated with percent whiteheads estimated at each site (Figure 3 and 4).

0 0.5 1 Log1o(%whiteheads + 0.5)

Fig. 3. Correlation of leaf wilting in response to a toxic fraction produced by Cephalosporium gramineum with percent whiteheads produced by 17 wheat genotypes in a naturally infested field in Condon, OR. 41

-

.-

U 1 2 3 Loge(%whiteheads + 0.5)

Fig. 4. Correlation of leaf wilting in response to a toxic fraction produced by Cephalosporium gramineum with percent whiteheads produced by 17 wheat genotypes in a naturally infested field in Dufi.zr, OR.

Toxin sensitivity of the mapping progeny

Frequency distribution of the wilting data (Fig. 5) suggests a continuous

distribution skewed towards insensitivity. Seventeen of the progenies were numerically

less sensitive than M6 and three were more sensitive than Opata 85, though none of these

differences were significant at P=0.05 based on either Duncan's Multiple Range or

Fisher's Least Significant Difference Test. Analysis of Variance (Table 4) indicates that

difference among the progenies are highly significant (P0.0001) with respect to toxin

sensitivity. Based on variance components analysis the genetic component of variance

was very large representing 88% of the phenotypic variance. Heritability of sensitivity to

the toxic fraction estimated on a genotype mean basis was 0.88, with a 90% confidence

interval of 0.85 0.91 42

Table 3. Significance of linear contrasts between wheat germplasm groups for wilting in reaction to a toxic fraction produced by C.gramineumin the laboratory and % whiteheads when grown in naturally-infested fields

Experiment Contrast Group mean 95% CI' P-value

Laboratorya Common vs. Club 4.53 vsl.97 1.46 to 2.03 <0.001

Common vs. Durum 4.53 vs. 2.92 2.28 to 2.76 <0.00 1

Common vs. Synthetic4.53 vs. 1.17 2.90 to 3.82 <0.001

Club vs. Durum 1.97 vs. 2.92 0.45 to 1.09 <0.001

Club vs. Synthetic 1.97 vs. 1.17 1.11 to 2.11 <0.001

Durum vs. Synthetic 2.92 vs.1.17 0.36 to 1.32 <0.001

Field: Condon"Common vs. Club 0.84 vs. -0.080.15 to 0.45 <0.001

Common vs. Durum 0.84 vs. 0.54 0.71 to 1.13 <0.001

Club vs. Durum -0.08 vs. 0.54 0.39 to 0.85 <0.001

Field: Dufurc Common vs. Club 2.79 vs. 0.16 0.53 to 0.85 <0.001

Common vs. Durum 2.79 vs. 2.10 2.39 to 2.87 <0.00 1

Club vs. Durum 0.16 vs. 2.10 1.68 to 2.20 <0.001 aData on wilting was taken on a continuous scale of 1-5 (1no wilting and 5 = maximum wilting) after 72 hr. of exposure to the toxic fraction. bPercent whitehead data on three replications were transformed to log1o(Y + 0.5).

CPercent whitehead data on four replications were transformed to lo&(Y + 0.5). 43

Table 4. Variance Components Analysis for the wilting reaction of the International Triticeae Mapping Initiative (ITMI) mapping progenies after exposure to a toxic fraction produced by Cephaloisporium gramineum

Source of d.f. Sum of Mean F value p>F Variation Squares Squares Genotype 111 472.01 4.2523 8.94 0.0001

Replications 2 2.0020 1.0010 2.10 0.1243

Error 222 105.57 0.4756

25

20

15 U>' C 0 a. 10 LL

U) Cl t C) U) U) U) C'4 Wilting rating (1-5 scale)

Fig. 5 Frequency distribution of wilting for 112 recombinant inbred line progeny resulting from a cross between the toxin-sensitive parent Opata 85 and toxin insensitive parent M6 after exposure to a toxic fraction produced by Cephaloisporium gramineum. 44

Discussion Our results are consistent with the hypothesis that toxin insensitivity is an important mechanism of resistance to cephalosporium stripe. Sensitivity to the toxic fraction was significantly correlated with percentage of whiteheads in two field trials.

Further, the common, durum, and club germplasm groups ranked the same for both toxin sensitivity and disease reaction in the field. M6, Altar 84, and Opata 85 could not be tested in the field because they are spring types, whereas cephalosporium stripe is mainly a disease of fall-sown wheat (Bruehl 1968; Bailey et al.,1982). However, preliminary studies with exposure of these genotypes to the pathogen in a liquid culture system in the greenhouse showed M6, Altar 84, and Opata 85 to be resistant, intermediate, and susceptible to cephalosporium stripe, respectively (unpublished), which is consistent with their reaction to the toxic fraction. The common wheat genotypes were all sensitive to the toxic fraction, but showed a substantial range of reactions in the field. It is important to note, however, that we found no case of an insensitive genotype being susceptible in the field. These results suggest that toxin insensitivity may be an important mechanism of resistance to cephalosporium stripe, but that other mechanisms are operative as well; similar results have been obtained for fusarium head blight of wheat (Lemmens et al., 1994). Differences in sensitivity to the toxic fraction were most notable among the four germplasm groups. Both common and club wheats are hexaploids that cany the A, B, and D genomes, but the club wheats originated from a different and substantially smaller gene pooi than the cormnon wheats. Durum wheat (T. turgidum) is a tetraploid that contains only the A and B genomes. The hexaploid synthetic wheat that we studied derived its D genome from Aegilops tauschii, which is a well known source of resistance to many wheat diseases (Cox et al., 1992; Lutz, 1994; Yildrim and Jones,1995; Thompson et al.,1997). This association of germplasm groups with toxin insensitivity should be 45

interpreted with caution, however, as only three durums and one synthetic wheat were evaluated in this study. Symptomology depended on concentration of the toxic fraction. We observed chiorosis of wheat leaves at low concentration (2OtI1ml), but this chiorosis did not reliably distinguish among wheat genotypes. At higher concentration (6Otl/ml), we

obtained distinct wilt symptoms that consistently distinguished wheat genotypes. Similarly, Kobayashi and Ui (1979) observed chiorosis after three days of exposure

of leaves to toxin and wilting in five days with a concentration of 25tI/ml graminin A. In the field, C. gramineum produces a wide array of symptoms depending on favorableness of conditions for disease. We have found the percentage of whiteheads, a severe symptom, to be a much more consistent and reproducible indication of resistance in the field than less severe symptoms such as leaf striping (unpublished). We used the 75% acetonitrile reversed phase fraction of the crude extract for our study. Though we detected evidence for the existence of graminin A in the crude toxin preparation using thin layer chromatography, we did not further analyze the fraction for the presence of other compounds. Thus, either graminin A alone, or in combination with other pathogen-produced compounds, may contribute to the toxic effect measured in this study. Further characterization of the toxic fraction may thus provide additional information regarding the role of toxin(s) in the pathogenesis of C.

gramineum. Most studies of plant pathogen toxins have involved highly host-selective interactions and qualitative inheritance of both susceptibility and virulence (Brown

and Hunger, 1993; Payne and Daly, 1980), though there are some exceptions (Lemmens et al.,1994). Studies of resistance to cephalosporium stripe by other workers (Mathre et al., 1977; Morton and Mathre, 1980; Martin et al.,1983) are all suggestive of incomplete expression of resistance and quantitative inheritance of disease reaction for cephalosporium stripe. These observations prompted us to study 46

toxin sensitivity in the ITMI mapping population of 112 progenies, derived from crossing the highly toxin-sensitive parent Opata 85 and the less sensitive synthetic parent M6. The resulting frequency distribution suggests quantitative inheritance, but with a very high heritability (0.88) that would allow for rapid response to selection. However, these results should be viewed with some caution, as we do not know if the

ITMI population is representative of wheat in general. Evaluation of the ITMI progenies for disease reaction would be useful in further elucidating the association between toxin insensitivity and disease resistance. This remains problematic, however, as the mapping population is in a spring wheat background. Efforts are underway to use the ITMI population to identify quantitative trait loci for toxin insensitivity and cephalosporium stripe resistance. Variability and environmental sensitivity in expression of cephalosporium stripe makes field screening difficult, especially in early generations of breeding material when limited seed stocks are available (Bruehi et al., 1986). Our study may thus provide options for more efficient screening of breeding material through direct assay with the toxic fraction in a controlled environment. Eliminating the most sensitive genotypes with a laboratory-based method could greatly reduce the number of genotypes to be tested in the field, thus reducing costs and allowing for more thorough evaluation of the genotypes retained. Subsequent selection of the more resistant types after exposure to the pathogen in the field would increase the probability that mechanisms in addition to toxin insensitivity were being selected. 47

References

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Bockus W. W., O'Connor, J. P., and Raymond, P. J. 1983. Effect of residue management method on incidence of Cephalosporium stripe under continuous winter wheat production. Plant Dis. 67:1323-1324.

Brown, C. A., and Hunger, R. M. 1993. Production of a chiorosis inducing,host-specific, low molecular weight toxin by isolates of Pyrenophora repentis, cause of tan spot of wheat. J. Phytopathol. 137:221-232

Bruehi G. W., Murray T. D., and Allan R.E. 1986. Resistance of winter wheat to cephalosporium stripe in the field. Plant Dis. 70:314-316.

Bruehl, G. W. 1957. Cephalosporium stripe disease of wheat. Phytopathology 47:64 1- 649.

Bruehi, G. W., Lai, P., and Huisman, 0. 1964. Isolation of Cephalosporium gramineum from buried naturally infested host debris. Phytopathology 54: 1035-1036.

Bruehi, G. W. 1968. Ecology of cephalosponum stripe disease of winter wheat in Washington. Plant Dis. Rep. 52:590-594.

Cox, T. S.,Raupp, W. J., Wilson, D. L., Gill, B. S., Leath, S., Bockus, W. W., and Brower, L. E. 1992. Resistance of foliar diseases in a collection of Triticum tauschii germplasm. Plant Dis. 76: 1061-1064.

Cowger, C., and Mundt, C. C.1998. A hydrophonic seedling assay for resistance to Cephalosporium stripe. Plant Dis. 82:1126-1131.

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Creatura, P. J., Safur, G. R., Scheffer, R. P., and Sharkey, T. D. 1981. Effects of Cephalosporium gramineum and a toxic metabolite on stomatal conductance of wheat. Physiol. Plant Pathol. 19:313-323.

Johnston, R. H., and Mathre, D. E. 1972. Effect of infection by Cephalosporium gramineum on winter wheat. Crop Sci. 12:817-819.

Knapp, S. J., Stroup, W. W., and Ross, W. M. 1985. Exact confidence intervals for heritability on a progeny mean basis. Crop Sci. 25:192-194. 48

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CONCLUSIONS

Cephalosporium stripe poses a substantial threat to the future of winter wheat production in areas where stubble mulch is required to preserve soil moisture and prevent erosion. Resistant cultivars can help to prevent yield losses and reduce inoculum concentration over time. Thus, a rapid and reliable screening method for identifying resistant sources will bring a significant benefit to wheat breeding programs. Strong environmental impacts on disease severity suggests the need for controlled environment screening techniques. Our findings open up the possibility of improved screening, as we were able to find easy methods for mass production of C.gramineumtoxin(s). However, the technique might need more calibration, e,g., with toxin concentration. This method is rapid and the symptoms are very distinct to read. The extent of wilting on a specific leaf is adequate to indicate quantitative variation with respect to toxin sensitivity among the lines/genotypes.

Our study also suggests that the toxin sensitivity trait is highly heritable. This information is the first of its type for cephalosporium stripe, and may encourage breeders to introgress cephalosporium stripe resistance to agronomically important cultivars.

Higher spore concentrations were assumed to be associated with the production of higher amounts of toxin. Thus, different concentrations of spore suspension for broth inoculation need to be investigated. It may be possible to find a concentration that more precisely reflects host resistance against cephalosporium stripe and the correlation with field reaction. The toxic fraction we used in our study needs to be subjected to High

Performance Liquid Chromatography to determine whether it contains only graminin A or if other compounds are responsible for the production of wilting symptoms. If more 51 than one compound is found, then all of them can be tried individually to find the best

one.

We used excised leaves to assay against toxin. For the same type of study,

workers have used other plant parts, such as coleoptiles, protoplasts, tissue cultures, and

roots. (Bains and Tewari 1987, Rines and Luke 1985). This toxin can also be used with

different plant parts of wheat to test for better correlations 52

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