ABSTRACT

TAYLOR, TANYA VALERA. Characterization of a Xanthomonas campestris pv. zinniae Oxidoreductase Involved in the Biodegradation of Cercosporin. (Under the direction of Dr. Margaret E. Daub.) The fungal photoactivated toxin cercosporin plays a key role in pathogenesis by

Cercospora spp. The bacterium Xanthomonas campestris pv. zinnia (XCZ) is able to rapidly

degrade this toxin. Growth of XCZ strains in cercosporin-containing medium leads to the

breakdown of cercosporin and to the formation of a non-toxic breakdown product called

xanosporic acid. EMS mutagenesis was used to generate five non-degrading mutants of a

rapid-degradation strain (XCZ-3). Mutants were complemented using a wild-type genomic

library. All five mutants were complemented with the same fragment, which encoded a

putative transcriptional regulator and an oxidoreductase. Simultaneous expression of the two

genes was necessary to complement the mutant phenotype. Sequence analysis of the mutants

showed that all five had point mutations in the oxidoreductase sequence and no mutations in

the regulator.

Quantitative RT-PCR showed that expression of both of these genes in the wild-type

strain is upregulated after exposure to cercosporin. Both the oxidoreductase and

transcriptional regulator genes were transformed into three cercosporin non-degrading

bacteria to determine if they are sufficient for cercosporin degradation. Quantitative RT-

PCR analysis confirmed that the oxidoreductase was expressed in all transconjugants.

However, none of the transconjugants were able to degrade cercosporin, suggesting that other

factors are involved in this process.

In an effort to determine if the oxidoreductase gene has a role in pathogenesis,

quantitative RT-PCR was used to assay for oxidoreductase regulation in XCZ-3 grown in the

presence of zinnia and tobacco extracts. The oxidoreductase gene was upregulated in the

presence of both plant extracts, but the response was greater with the tobacco extract. The

five XCZ-3 oxidoreductase mutants were assayed for pathogenicity on zinnia. The mutants

yielded disease symptoms equal to those of the wild-type. Furthermore, previously identified

bacteria (Xanthomonas axonopodis pv. pruni, strains XAP-50 and XAP-75, and

Pseudomonas syringae pv. tomato, PST DC3000) with oxidoreductase homologues were not

able to infect zinnia, suggesting that the presence of a functional oxidoreductase is not

correlated with pathogenesis of zinnia.

Characterization of a Xanthomonas campestris pv. zinniae Oxidoreductase Involved in the

Biodegradation of Cercosporin

By

TANYA VALERA TAYLOR

A dissertation submitted to the Graduate Faculty of

North Carolina State University

In partial fulfillment of the

Requirements for the degree of

Doctor of Philosophy

In

PLANT PATHOLOGY

Raleigh, NC

2005

Approved by:

______________________ ______________________ Margaret E. Daub Gary A. Payne Dissertation Committee Chair ______________________ ______________________ David F. Ritchie R. Gregory Upchurch

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PERSONAL BIOGRAPHY

I was born to Thomas and Virginia Taylor in 1973 and raised in Hillsborough, NC

along with my brother Harlan. After graduating from Orange High School in 1991, I enlisted

in the Navy for four years. Through the Navy, I was able to travel to various parts of the

United States, as well as to Spain. After completing my tour of duty, I attended the

University of North Carolina at Chapel Hill and earned a Bachelors of Science degree in

Biology in 2000. I began working on my Ph.D. at North Carolina State University in the

Department of Plant Pathology in the fall of 2000. My project, which focused on

characterization of bacterial genes responsible for degradation of the fungal toxin

cercosporin, was completed under the direction of Dr. Margo Daub in the Department of

Plant Pathology.

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TABLE OF CONTENTS

Page

LIST OF FIGURES ………………………………………………………………….…...... vi

LIST OF TABLES …………………………………………………………………..……. viii

Chapter 1: REVIEW OF RELEVANT LITERATURE .………………………………...…. 1 The fungal genus Cercospora …………………………………………………….…. 2

The toxin cercosporin …………………………………………………………..…….3 Cercospora resistance mechanisms ……………………………….……………...…..9 Control of Cercospora diseases ………………………………………………….….13 Cercosporin degradation ………………………………………………………….…16 Cytochrome P450s ………………………………………………….. ………….......19 Disease control through transgenic resistance to toxins ……….………………....…24 Summary …………………………………………………………………….…....…27 References cited ………………………………………………………………….….29

Chapter 2: A PUTATIVE OXIDOREDUCTASE IS INVOLVED IN CERCOSPORIN DEGRADATION BY THE BACTERIUM XANTHOMONAS CAMPESTRIS PV. ZINNIAE ..……….………………………………………………………………………….42 Abstract ………………………………………………………………………….…..43 Introduction ……………………………………………………………………….…44 Materials and Methods …………………………………………………………..…..48 Bacterial cultures …………………………………………………..………..48 Cercosporin stocks and chemicals ………………………………………..…48 Chemical mutagenesis of Xanthomonas campestris pv. zinniae and

screening for mutants unable to degrade cercosporin ..………………….…..48 Box PCR …………………………………………………………………….49 Xanthomonas campestris pv. zinniae genomic library construction .…….…49

Complementation assays ……………………………………………………50 Subcloning of complementing library clone ………………………………...51 Sequencing of subclone …………………………………………………..…51 Sequencing of mutant genes ………………………………………………...51 Gene complementation ……………………………………………………...52 Quantitative RT-PCR of oxidoreductase and regulator in wild-type XCZ-3 …………………………………………………………….………....52 Southern analysis …………………………………………………………....53 Transformation of the regulator and oxidoreductase genes into non-degrading bacteria ………………………………………………………54 Time course of cercosporin degradation by transconjugants ……………......54

Results …………………………………………………………………………….…56 Isolation of Xanthomonas campestris pv. zinniae mutants unable to

degrade cercosporin …………………………………………………………56

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Identification of complementing genes ……………………………………...56 Search for additional genes ………………………………………………….58 Sequencing of mutants ……………………………………………………....59 Identification of sequences necessary for mutant complementation .……….59 Presence of cercosporin upregulates gene expression ………………………60 Southern analysis of oxidoreductase homologues in degrading and

non-degrading species ……………………………………………………….60 The transcriptional regulator and oxidoreductase are not sufficient for cercosporin degradation in non-degrading species ………………………….61

Discussion …………………………………………………………………………...63 References cited ……………………………………………………………………..68 Chapter 3: THE XANTHOMONAS CAMPESTRIS PV. ZINNIAE OXIDOREDUCTASE GENE IS REGULATED BY PLANT EXTRACTS, BUT IS NOT NECESSARY FOR PATHOGENESIS .……………………………………………………………………....…..88 Abstract ……………………………………………………………………………...89 Introduction ………………………………………………………………………….90 Materials and Methods …………………………………………………………...….93 Bacterial Cultures ……………………………………………………………93 Plant Extract ……………………………………………………………..…..93 Quantitative RT-PCR of Oxidoreductase Expression in Wild-Type

XCZ-3 ………………………………………………………………….…....94 Zinnia Inoculation …………………………………………………………...94 Results ……………………………………………………………………………….96 Presence of Plant Extract Upregulates Oxidoreductase Gene Expression …..96 Presence of the Oxidoreductase Not Correlated with Pathogenicity ………..96

Discussion …………………………………………………………………………...98 References cited ……………………………………………………………………100 APPENDIX - SEARCH FOR PROTEINS DIFFERENTIALLY EXPRESSED IN XCZ-3 UPON EXPOSURE TO CERCOSPORIN ………………………………….……..105 Goal …………………………………………………………………………….…..105 Methods ……………………………………………………………………….…....105 Results …………………………………………………………………………...…107 Conclusion …………………………………………………………………………107 References cited …………………………………………………………………....108 APPENDIX – XANTHOMONAS AXONOPODIS PV. CITRI DOES NOT DEGRADE CERCOSPORIN …………………………………………………………………………..110 Goal …………………………………………………………………………….….110 Methods …………………………………………………………………….….…..111 Results ………………………………………………………………………….….111 Conclusion …………………………………………………………………….…...112 References cited ………………………………………………………….…….…..113

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APPENDIX – TRANSFORMATION OF TOBACCO WITH THE XANTHOMONAS CAMPESTRIS PV. ZINNIAE OXIDOREDUCTASE GENE INVOLVED IN CERCOSPORIN DEGRADATION ………………………………………………….…...117

Goal ………………………………………………………………………………..117 Methods ……………………………………………………………………………118 Cloning of Genes ………………………………………………….….……118 Plant Tissue Transformation ……………………………………….….…...118 Recovery of Plants and Callus ……………………………………...……...119 Screening of Callus ………………………………………………….……..120 Results ……………………………………………………………………….……..120 Conclusion …………………………………………………………………….…...121 References cited ……………………………………………………………..……..122

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LIST OF FIGURES Page

A PUTATIVE OXIDOREDUCTASE IS INVOLVED IN CERCOSPORIN DEGRADATION BY THE BACTERIUM XANTHOMONAS CAMPESTRIS PV. ZINNIAE

Figure 1 Chemical Structure of cercosporin and the breakdown product (xanosporic acid) produced by X. campestris pv. zinniae, isolate XCZ-3 ………………….…………………………………….76

Figure 2 Cercosporin non-degrading XCZ-3 mutants ..……………………….77 Figure 3 Agarose gel of Box PCR products ………………………………..…78 Figure 4 Complementing fragment derived from sequencing the

complementing 7.5 kb subclone …………………………………….79 Figure 5 Sequence containing the putative transcriptional regulator and oxidoreductase genes of XCZ-3 (GenBank Accession #DQ087176) ………………………………….80 Figure 6 Alignment of amino acid sequences of oxidoreductase genes from

Xanthomonas campestris pv. zinniae (XCZ-3) and Xanthomonas axonopodis pv. citri (XAC-306) ………………………………….…82

Figure 7 Alignment of amino acid sequences of transcriptional regulator genes from Xanthomonas campestris pv. zinniae (XCZ-3) and Xanthomonas axonopodis pv. citri (XAC-306) ……………………..83

Figure 8 Quantitative RT-PCR analysis of expression of the transcriptional regulator and oxidoreductase genes in XCZ-3 following exposure of the bacterium to cercosporin ……………………………………..84

Figure 9 Southern hybridization of total DNA isolated from cercosporin-degrading (XCZ-3, XCP-50, XCP-75, XCP-1, XCP-39, XCP-49, XCP-34, XCP-31, and XCP-79) and non-degrading (XCP-76, XCV-79, XCV-126, PST DC3000, and PSP 209-21-3R) isolates using a PCR-amplified digoxigenin- labeled oxidoreductase gene as a probe ………………………….….85 Figure 10 Quantitative RT-PCR analysis of expression of the oxidoreductase

in transconjugants of the non-degrading bacteria XCV-79, XCV-126, and PST DC3000 as compared to the XCZ-3 degrading strain …………………………………………………..…86

Figure 11 Degradation of cercosporin in liquid culture by cercosporin-degrading and non-degrading isolates and oxidoreductase-expressing transconjugants …………………………87

THE XANTHOMONAS CAMPESTRIS PV. ZINNIAE OXIDOREDUCTASE GENE IS REGULATED BY PLANT EXTRACTS, BUT IS NOT NECESSARY FOR PATHOGENESIS Figure 1 Bacterial isolates used to inoculate zinnia ………………………....102

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Figure 2 Quantitative RT-PCR analysis of expression of the oxidoreductase gene in XCZ-3 following exposure of the bacterium to plant extracts ……………………………………....103

Figure 3 Zinnia leaves inoculated with the XCZ-3 wild-type and the cercosporin non-degrading mutants 120-2, MutA, MutB, MutC, and MutD ……………………………………….………..104

APPENDIX - SEARCH FOR PROTEINS DIFFERENTIALLY EXPRESSED IN XCZ-3 UPON EXPOSURE TO CERCOSPORIN Figure 1 One-dimensional polyacrylamide gels displaying the

banding patterns of XCZ-3 exposed to cercosporin or to 0.5 % acetone as a control ……………………………………….109

APPENDIX – XANTHOMONAS AXONOPODIS PV. CITRI DOES NOT DEGRADE CERCOSPORIN Figure 1 Alignment of amino acid sequences of oxidoreductase

genes from Xanthomonas campestris pv. zinniae (XCZ-3) and Xanthomonas axonopodis pv. citri (XAC-306) ………….…114

Figure 2 Alignment of amino acid sequences of transcriptional regulator genes from Xanthomonas campestris pv. zinniae (XCZ-3) and Xanthomonas axonopodis pv. citri (XAC-306) …..115

Figure 3 Xanthomonas axonopodis pv. citri isolate xc62 does not degrade cercosporin when grown on cercosporin-containing medium ……………………………………………………..…...116

APPENDIX – TRANSFORMATION OF TOBACCO WITH THE XANTHOMONAS CAMPESTRIS PV. ZINNIAE OXIDOREDUCTASE GENE INVOLVED IN CERCOSPORIN DEGRADATION Figure 1 Tobacco callus on cercosporin-containing medium……………..124

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LIST OF TABLES Page

A PUTATIVE OXIDOREDUCTASE IS INVOLVED IN CERCOSPORIN DEGRADATION BY THE BACTERIUM XANTHOMONAS CAMPESTRIS PV. ZINNIAE Table 1 Location of each mutation in the oxidoreductase gene

of the five cercosporin non-degrading mutants ……………………..75 APPENDIX – TRANSFORMATION OF TOBACCO WITH THE XANTHOMONAS CAMPESTRIS PV. ZINNIAE OXIDOREDUCTASE GENE INVOLVED IN CERCOSPORIN DEGRADATION

Table 1 Callus tissue transformed with the oxidoreductase gene was unable to degrade cercosporin after 21 days incubation on cercosporin-containing medium.…………………………..…...…..123

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CHAPTER 1

Review of Relevant Literature

2

Review of Relevant Literature

The Fungal Genus Cercospora

Members of the fungal genus Cercospora are pathogenic on plants, causing damaging

leaf spot and blight diseases. The host range for this genus is large, with individual species

showing specificity to certain host plants (17). Cercospora spp. can infect such hosts as

tobacco, corn, soybean, rice, sugar beet, and banana, resulting in extreme crop damage and

loss (5, 23, 30, 65, 91). Blossoms, fruits, pods, succulent petioles, seed coats, and young

stems are affected, but Cercospora spp. are mainly responsible for leafspot diseases (64, 97).

Symptoms include tan-colored, necrotic lesions surrounded by a slightly raised, reddish-

brown region (17, 64, 72). The spots may appear zonate and can eventually coalesce. There

may also be a chlorotic halo around the lesion.

The genus Cercospora has traditionally been grouped in the Deuteromycota, order

Hyphomycetales (17). However, modern science groups many Cercospora species in the

Ascomycota based on their known sexual stages, which are in the genus Mycosphaerella

(17).

Although some Cercospora spp. have a known Mycosphaerella sexual stage, the life

cycle of Cercospora species is usually asexual. Conidia are the principal signs of the

presence of these fungi, and are produced in the leaf spots. Conidia are multiseptate and

hyaline, with thick walls (17). Transmission of the conidia is usually passive, such as by

wind or rain, but can also be active, such as by humans and farm equipment (64). Once

disseminated, they land on a plant host and germinate. Disease development is dependent on

extended periods of high humidity (1). With conducive conditions, such as wet weather and

3

temperatures between 20 and 30 °C, the conidia can germinate within 24 hours (6).

However, it is common for Cercospora spp. to have an extended latent period, such as that

reported for C. musae, which can extend from 16 to 120 days (6). An extended latent period

is possible because many Cercospora spp. are able to grow epiphytically on leaf surfaces and

tolerate dry conditions which are not conducive to host penetration (1). For example, C.

zeae-maydis was shown to survive at least one week on corn leaf surfaces prior to penetration

(6).

Cercospora pathogens most commonly enter the host through wounds or natural

openings, such as stomata (1, 6, 64). Depending on the species, there may also be production

of an appressorium (6). Growth of Cercospora spp. is limited to the intercellular spaces (89).

Once the pathogen has penetrated the host, its growth and reproduction leads to the

characteristic leafspots. This cycle will continue until winter arrives, or until conducive

weather conditions deteriorate. The fungus then overwinters in plant debris, on which it

produces new conidia the following spring (64, 78). For sugar beet leaf spot, primary

inoculum can also originate from common weed hosts surrounding crop production fields

(78, 103).

The Toxin Cercosporin

The genus Cercospora is known for the production of the non-host-specific toxin

cercosporin. Cercosporin, deep red in color, was first isolated from the dried mycelium of

Cercospora kikuchii, a soybean pathogen, in 1957 by Kuyama and Tamura (58). It has the

chemical name 4,9-dihydroxyperylene-3,10-quinone [1,12-bis(2-hydroxypropyl)-2,11-

dimethoxy-6,7-methylenedioxy-4,9-dihydroxyperylene-3,10-quinone] and the molecular

4

formula C29H26O10, as shown independently by both Yamazaki, et al. (104) and Lousberg, et

al. (63, 104). It is a seven-membered ring molecule that is bilaterally symmetrical (104).

Cercosporin is produced by most members of the Cercospora genus and has been

isolated from many Cercospora species, as well as from infected host plants (2, 3, 30, 41, 58,

65, 73, 98). Interestingly, production of cercosporin in lab-grown cultures was shown to be

dependent upon the medium used, with the ratio of carbon to nitrogen being particularly

important (53). Production of cercosporin is sometimes strain- or isolate-specific and

producing isolates may react differently to the same set of growth conditions (45). For

example, cercosporin production can be affected by temperature and light availability,

depending on the Cercospora isolate (53). An example of the importance of light in

cercosporin production has been revealed in wild-type cultures of Cercospora kikuchii.

Growth of C. kikuchii in the light yielded high levels of cercosporin, whereas cultures grown

in the dark produced 100-fold less (83).

Phylogenetic evidence suggests that because only species within the Cercospora

genus can produce cercosporin, the ability to produce cercosporin must have a single

evolutionary origin (45). Furthermore, Fajola, et al. have suggested that Cercospora spp. that

do not produce cercosporin may actually belong to a related genus (45). Alternatively, non-

producing Cercospora spp. may have produced cercosporin in the past, but simply lost the

ability to do so over time (45). Interestingly, not all red pigments found in Cercospora

cultures are cercosporin (43, 53). Four isolates of C. arachidicola were shown to produce a

red pigment, but were unable to produce cercosporin (43).

Cercosporin is a polyketide secondary metabolite and a nonspecific toxin, showing

toxicity to plants, animals, bacteria and fungi (3, 20, 41, 66, 105). It belongs to a group of

5

toxins known as the photosensitizing perylenequinones; these compounds must absorb light

energy to become active (20, 105). In general, photosensitizers enter the excited triplet state

after exposure to visible wavelengths of light. Once in the triplet state, photosensitizers are

able to interact with molecular oxygen to form the activated oxygen species superoxide and

singlet oxygen (22, 24).

Activated oxygen species are generated by photosensitizers in two ways: through type

I and type II reactions (23, 87). In type I reactions, electron transfer from the triplet

photosensitizer via a reducing substrate yields reduced and radical forms of oxygen, i.e.

superoxide, hydrogen peroxide, or the hydroxyl radical. In type II reactions, direct energy

transfer between the triplet photosensitizer and molecular oxygen yields singlet oxygen.

With cercosporin, superoxide is more readily produced when a reducing substrate is present.

Otherwise, singlet oxygen is the more common product (50).

Although cercosporin has been shown to produce both activated oxygen species and

both have damaging effects, singlet oxygen appears to cause the most damage (20, 24, 50,

60). Cercosporin is lipid-soluble and localizes in the lipid bilayer of host cell membranes.

When activated oxygen species contact cell membranes, they react with the double bonds of

the lipids in the membranes. Presence of singlet oxygen, in particular, can lead to lipid

peroxidation and the breakdown of cell membranes (21, 22), which may occur both in vitro

and in vivo (11, 21). Studies in tobacco have elucidated the changes that occur in cell

membranes in the presence of cercosporin (21, 22). Upon exposure to light, the cell

membranes exhibit increased rigidity due to the destruction of unsaturated acyl chains of the

fatty acids. This change increases the ratio of saturated to unsaturated fatty acids in the

membrane and reduces membrane fluidity. At this point, as shown in sugar beet, the

6

endoplasmic reticulum and mitochondrial membranes begin to swell and thicken (88, 89).

Finally, there is a disruption of plasmalemma, tonoplast, and chloroplast membranes,

resulting in a leakage of cell nutrients which the fungus can then use (22).

In tobacco leaf disks, light intensity and cercosporin concentration have been shown

to affect the rate of electrolyte leakage from the cells (21). Tobacco leaf disks were

infiltrated with cercosporin and assayed for electrolyte leakage by measuring the change in

conductivity of the bathing solution. Some treated disks were exposed to light, leading to a

rapid increase in electrolyte leakage. Untreated disks and treated disks that were kept in the

dark did not show an increase in electrolyte leakage. The intensity of the light used to

irradiate the disks affected the rate of electrolyte leakage. Decreasing the light intensity

increased the amount of time it took for the electrolyte leakage to occur. Decreasing

cercosporin concentration also delayed the initiation of electrolyte leakage.

Several lines of evidence suggest that the production of the toxin cercosporin is

associated with parasitism and development of disease in host plants (23). First, it is

produced by most Cercospora spp. and has also been isolated from infected tissues, such as

soybean and groundnut (41, 98), confirming production during disease development. In

addition, studies by Steinkamp, et al. showed that ultrastructural changes caused by

Cercospora beticola on infected sugar beet are almost identical to the changes caused by

cercosporin on sugar beet (22, 88, 89). In both cases, lesions showed a loss of cell

membranes and the accumulation of large amounts of granular, electron-dense material in the

intercellular spaces.

Upchurch, et al. (94) have shown a more direct role for cercosporin in disease. Stable

mutants of Cercospora kikuchii were induced using UV mutagenesis. These mutants were

7

affected in their ability to produce cercosporin, synthesizing no more than 2% of wild-type

cercosporin levels. When these mutants were inoculated on soybean, they yielded only a

few, small flecks, in contrast to the wild-type, which yields large, coalescing lesions (94).

The authors concluded that production of cercosporin by C. kikuchii is necessary to induce

disease on the host plant.

Research by Chung, et al. (15) and Choquer, et al. (12, 15) also demonstrated the

importance of cercosporin in disease-causing ability. Mutants of C. nicotianae that were

deficient in the synthesis of cercosporin were created through the use of plasmid insertion

and REMI (restriction enzyme-mediated integration) (15). These isolates were termed ctb

(cercosporin toxin biosynthesis) mutants. In one of these ctb mutants, the sequence flanking

the insert site was analyzed to reveal homology to fungal type I polyketide synthases. Fungal

polyketide synthase genes typically encode several catalytic domains, resulting in a large

protein (15). The polyketide synthase gene from C. nicotianae (CTB1) was found to encode

five catalytic domains: a keto synthase, an acyltransferase, a thioesterase/claisen cyclase, and

two acyl carrier protein domains. The CTBI gene was then cloned and disrupted to yield a

disruption construct, which was used to replace the wild-type CTBI gene with the disrupted

version in C. nicotianae (12). When the resulting disruption mutants were inoculated onto

tobacco, lesions were fewer and smaller compared to wild-type. Complementation of the

disrupted CTBI gene restored wild-type levels of cercosporin synthesis and virulence.

Regulation studies of CTBI have also shown that gene expression is induced by both light

and growth on PDA medium, consistent with the initiating factors for cercosporin

biosynthesis already established (36, 53).

8

As further evidence for the role of cercosporin in pathogenesis, disease intensity has

been shown to be directly related to light intensity and amount of light exposure. For

example, symptoms caused by Cercospora coffeicola on coffee were reduced when plants

were grown close together, an effect attributed to shading of the leaves (23).

Calpouzos, et al. studied the effect of light intensity on Cercospora-infected sugar

beets (10). High light intensities led to an earlier onset of disease. Leaf spots were round,

with defined borders, small and not expanding in size, and surrounded by a purple halo.

These symptoms are characteristic of those observed under field conditions. However,

exposing infected sugar beets to low light intensities altered the symptoms. Leaf spots were

larger, irregularly shaped, and expanded over time. In addition, the purple halo was absent

and disease severity was reduced.

The disease severity of Cercospora leaf spot disease has also been shown to be

directly related to light intensity in banana (9, 91). Disease caused by Cercospora musae had

been observed to be reduced on stands of bananas when the amount of sunlight was reduced,

such as by shading from the canopies of taller trees (91). Also, the onset of disease

symptoms occurred earlier on banana plants exposed to higher light intensities, whereas low

light intensities caused a delay in disease symptoms (91). Calpouzos, et al. further studied

this observation and found that under greenhouse conditions, banana plants inoculated with

C. musae and exposed to high daylight conditions developed abundant disease after seven

days, whereas inoculated plants exposed to low light conditions did not develop disease (9).

This result was shown to be true for both natural and artificial light.

9

Cercospora Resistance Mechanisms Several studies have been conducted in an effort to elucidate the mechanisms used by

Cercospora spp. to protect themselves from their own toxin. Carotenoids have been shown

to play at least a small role in resistance of fungi to cercosporin (27). Carotenoids are

excellent quenchers of singlet oxygen and of photosensitizers in the triplet (activated) state

(57). Assaying of carotenoid-deficient mutants of Neurospora crassa and Phycomyces

blakesleeanus demonstrated that carotenoid content was correlated with cercosporin

resistance in these fungi (27). However, a carotenoid over-producing mutant of P.

blakesleeanus (producing 30 times more carotenoids than C. nicotianae) was still more

sensitive to cercosporin than C. nicotianae. In addition, a wild-type isolate of N. crassa that

produces equivalent amounts of carotenoids as C. nicotianae shows a sensitivity to

cercosporin that C. nicotianae lacks (27). Further, Ehrenshaft, et al. used targeted gene

disruption of the carotenoid biosynthetic phytoene dehydrogenase gene to create mutants of

C. nicotianae defective in carotenoid biosynthesis (35). The mutants did not show increased

sensitivity to cercosporin, demonstrating that carotenoids do not play a major role in

protection of Cercospora spp. from cercosporin. Interestingly, the carotenoid-minus mutants

were not affected in their ability to infect tobacco plants, suggesting that carotenoids are also

not required for pathogenesis.

Cell biology studies identified a role for cell surface reducing ability as a major

source of self-protection (25, 26, 60, 86). When cercosporin is reduced, it becomes bright

yellow and produces a green fluorescence (59). In the reduced state, the molecule has been

shown to yield less singlet oxygen and to lose its toxicity (25, 54, 86). Reduction of

cercosporin is reversible and will spontaneously revert when the reducing agents have been

10

removed (25, 59, 86). Confocal microscopy revealed that hyphal cells of Cercospora treated

with cercosporin maintain the internal cercosporin located in the cytoplasm in the reduced

state (25, 26). Cercosporin that was exported out of the cell was not reduced, presumably

due to spontaneous reoxidation. Wild-type Cercospora spp. resistant to cercosporin were

shown to have a higher cell surface reducing capability than cercosporin-sensitive fungi (86).

It was hypothesized that maintenance of a reducing environment ensures that the hyphae are

in contact with only the non-toxic (reduced) form of cercosporin.

Genetic studies demonstrated that Cercospora spp. could achieve partial resistance to

cercosporin by transporting the toxin out of the fungus. Work by Upchurch, et al. revealed

the role of the cercosporin facilitator protein (CFP) in C. kikuchii as a means of exporting

cercosporin from the fungus. The CFP gene was initially identified in a subtractive library

representing genes whose transcript accumulation was upregulated by light (36). The

corresponding CFP transcript was shown to be light-regulated, increasing as much as 20-fold

in the C. kikuchii strain PR when exposed to light. CFP also shows a temporal expression

pattern that matches cercosporin accumulation. Expression of the CFP gene in Cochliobolus

heterostrophus, which does not produce cercosporin and is cercosporin-sensitive, resulted in

reduced accumulation of cercosporin in mycelium treated with the toxin as well as an

approximately 20% increase in cercosporin resistance (93).

Work by Callahan, et al. showed that CFP is required for cercosporin production,

resistance, and virulence by C. kikuchii strain PR (8). Targeted disruption of the CFP gene

yielded mutants that produced only 5% of the cercosporin produced by wild type strains. In

addition, the mutants showed reduced virulence on soybean. Growth of disruptants was also

attenuated on cercosporin-containing medium. Over-expressing the CFP gene in C. kikuchii

11

resulted in fungal strains that exhibited increased production and secretion of cercosporin

(92). These results suggested that CFP plays a role both in cercosporin resistance and in

production.

Sequence analysis of the CFP gene suggested that it is an MFS (Major Facilitator

Superfamily) transporter protein, having highest similarity to the MFS drug resistance

transporter group (8). MFS proteins actively export toxic molecules from the membrane

and/or cytoplasm by means of the proton motive force (92). Callahan, et al. proposed that

CFP plays a role in cercosporin resistance by actively exporting cercosporin out of the

fungus, effectively maintaining low concentrations of the toxin inside the cells (8).

In an attempt to identify additional cercosporin-resistance genes, a total of six

mutants of C. nicotianae were generated that showed sensitivity to the presence of

cercosporin (54). Five of the mutants, CS2, CS6, CS7, CS8, and CS9, demonstrated

inhibited growth when cercosporin was present, but their sensitivity depended on cercosporin

concentration, as well as on light intensity. These mutants retained the ability to produce

cercosporin, but at slightly reduced amounts (52). Increased light intensity led to increased

cercosporin sensitivity, although there was no light sensitivity in the absence of cercosporin.

Growth of the sixth mutant, CS10, was only partially inhibited by the presence of

cercosporin. It was also able to synthesize cercosporin at somewhat reduced levels, but did

not exhibit increased sensitivity to cercosporin with increasing light intensity. The mutants

were unaltered in endogenous levels of reducing agents known to protect against cercosporin

toxicity (52). Production of the carotenoid β-carotene in the mutants also did not differ from

that of the wild type. When tested for pathogenicity on tobacco, CS2, CS8 and CS10 yielded

fewer lesions than wild-type.

12

Complementation of the cercosporin-sensitive mutants identified three genes that are

involved in cercosporin resistance. Complementation of the CS10 mutant led to

identification of the gene CRG1, which encodes a zinc cluster transcription factor (14, 16).

Targeted disruption of the CRG1 gene in C. nicotianae showed that CRG1 is needed for

resistance against cercosporin as well as for toxin production (14). The genes regulated by

CRG1 are currently being investigated as possible genes involved in both cercosporin

resistance and biosynthesis. Complementation of the remaining mutants led to the discovery

of a gene originally termed SOR1, which is required for resistance to cercosporin and to

several other singlet oxygen-generating photosensitizers (34). SOR1, determined to be

present in a single copy in C. nicotianae, was later shown to be essential for the synthesis of

pyridoxine (vitamin B6) in C. nicotianae and Aspergillus flavus, and the name was changed

to PDX1 (31, 32). Subsequent work identified a second gene, PDX2, also required for

pyridoxine biosynthesis (33). These discoveries led to the identification of a novel pathway

for pyridoxine synthesis, distinct from the previously defined pathway in E. coli. Ehrenshaft,

et al. showed that organisms can encode the genes for production of pyridoxine by the PDX1

pathway, or by the traditionally known method utilized by E. coli, but not by both (31).

Subsequent work determined that pyridoxine and its vitamers can quench singlet oxygen,

with quenching constants similar to those of vitamins C and E, two of the best known cellular

antioxidants, suggesting the mechanism by which pyridoxine may play a role in cercosporin

resistance (31). Unfortunately, constitutive expression of the C. nicotianae PDX1 and PDX2

genes in tobacco did not result in increased cellular levels of vitamin B6, as well as no

increase in resistance to cercosporin (Herrero and Daub, unpublished).

13

Control of Cercospora Diseases

Presently, control of diseases caused by Cercospora spp. relies primarily on

fungicides and development of resistant cultivars (42). Disease scouting and monitoring of

weather conditions can allow for use of prediction models for disease forecasting (103),

which may help increase the efficacy of fungicide applications.

Some research has suggested that the age of a plant can affect the susceptibility to

infection, making the planting date an important factor. In sugar beet, it has been shown that

leaf spot disease caused by C. beticola Sacc. first appears on older leaves and later appears

on younger leaves (103). Leaf disk experiments with corn have shown that younger corn

plants are more sensitive to the toxin cercosporin (48), but older corn plants are more

sensitive to Cercospora infection (99). However, greenhouse studies carried out by

Beckman, et al. showed that neither corn plant age or leaf age affected susceptibility to C.

zeae-maydis and that disease development was more dependent on conditions of high

humidity, such as that created in an older plant canopy (6).

Some cultural practices may be quite effective at controlling Cercospora diseases,

such as removal or plowing under of plant debris and crop rotation (18, 78). The common

use of conservation tillage practices in corn-producing areas in the 1980s and 1990s has led

to a resurgence of the gray leaf spot disease caused by C. zeae-maydis (6, 18, 61, 62, 78).

When infected corn debris is left on the soil surface, this allows the primary inoculum to

form and infect the following year’s crops. Plowing under would be an effective means of

limiting inoculum and controlling the disease. But, avoiding tillage helps prevent erosion

and preserve the nutrients that are present in the soil (17). Therefore, crop rotation would aid

14

in disease avoidance for the infested no-till fields. However, inoculum can still carry over

from neighboring fields and cause disease.

Resistance to Cercospora diseases is uncommon, making the development of

resistant cultivars difficult. However, resistant cultivars of rice, corn, and sugar beet have

been reported (5, 18, 46, 68, 69, 76, 77, 85). Rice has been shown to have unusually high

levels of resistance to the pathogen C. oryzae (5). Louisiana red rice, an annual weed, shows

widespread resistance to C. oryzae races. Other rice cultivars are more specific, showing

resistance only to certain C. oryzae races. This resistance has been shown to involve

carotenoids (5).

Several lines of corn resistant to gray leaf spot (GLS) have also been found (18, 46,

68, 69). Coates, et al. worked with three resistant lines and showed that breeding for

resistance to GLS could be a successful strategy, but that expression of resistance may

depend on the environment (18). Resistance was shown to be a dominant trait for some loci.

However, they found that resistance was primarily the result of multiple genes, which could

be inherited, and that resistance was additive (18). Gordon, et al. studied marker-assisted

selection as a means of selecting for resistance and found that this was a more reliable and

efficient method than determining resistance based on phenotype after pathogen infection

each growing season (46). They also demonstrated that inherited resistance to GLS could be

effective in diverse environments. Recently, Menkir, et al. reported 20 lines of maize

displaying resistance to GLS (68). Further work with these lines showed that cytoplasmic

genes played a significant role in GLS resistance in hybrid offspring (69). Therefore, they

suggested that when crossing resistant and susceptible maize lines, the resistant lines should

be used as the female.

15

Breeding for sugar beet varieties resistant to Cercospora leaf spot has been very

successful (76, 85). However, achieving the desired resistance often results in the

appearance of other traits that are undesirable, such as reduced sugar content. In addition,

Orth, et al. (77) showed that the presence of resistance to foliar infection by Cercospora was

not related to resistance to seed infection. Therefore, development of cultivars resistant to

one form of infection will not necessarily protect the plants from another form of infection.

Infection rate and the progress rate of epidemics, however, may be significantly reduced (84).

Work by Nilsson, et al. revealed five quantitative trait loci that were responsible for

Cercospora leaf spot resistance in sugar beet (76). They also found that resistance was

primarily additive, but sometimes dominant, as well. Based on their results, they suggested

that high-performing sugar beet lines resistant to Cercospora infection may best be achieved

through use of marker-assisted breeding.

In the past, breeding plants for resistance has mainly focused on the acquisition of

defense and resistance genes. Maher, et al. suggested that an alternate approach to breeding

for resistance would be to focus on increasing preformed plant protectants (67). They

showed that susceptibility of tobacco plants to C. nicotianae was increased with suppression

of plant phenylpropanoid biosynthesis. Biosynthesis of phenylpropanoids usually increases

in response to wounding or inoculation with avirulent pathogens and leads to activation of

the plant defense genes. Therefore, increasing the production of phenylpropanoids may

result in an increased resistance to pathogen invasion and a reduction of disease symptoms.

Production and use of transgenic lines is another possible source for disease

resistance. Experimental studies on transgenic resistance to Cercospora spp. could focus on

resistance to fungal invasion. Given the importance of cercosporin in disease, studies could

16

also address strategies that would target cercosporin. For example, the cercosporin-

resistance genes already identified in Cercospora spp. may be useful in developing resistant

plant lines. Additionally, genes that would decrease production or accumulation of the toxin

may also be beneficial.

Cercosporin Degradation

A patent filed in 1993 by Robeson, et al. revealed a method for identifying microbes

that could degrade cercosporin (82). Mixed bacterial populations were harvested from the

soil, leaf surfaces and leaf tissue of Cercospora beticola-infected sugar beet plants. The

bacteria were then inoculated onto cercosporin-containing medium and incubated in the dark.

Addition of cercosporin turns the medium red. Bacteria that are able to degrade cercosporin

can be identified by a cleared halo surrounding the colony on the red-colored plates. As

further support for degradation, agar plugs were collected from both red-colored and cleared

areas near the tested bacterial colonies and extracted with acetone. The samples were

assessed spectrophotometrically and by qualitative thin layer chromatography. Four isolates

were discovered that could degrade cercosporin: Bacillus thuringiensis, Pseudomonas

fluorescens biovar V, Mycobacterium sp., and Bacillus subtilis. Interestingly, the bacteria

were only able to degrade the cercosporin in the dark.

An additional study using 14C-labelled cercosporin incorporated into the medium also

supported the degradation data (82). Only Bacillus thuringiensis was used in this particular

experiment. After growth of the bacterium on the medium revealed the cleared halo, the agar

was extracted and assayed for radioactivity. The level of radioactivity was reduced in the

cleared zones, suggesting degradation of the 14C-labelled cercosporin. The bacteria were also

17

assayed for radioactivity to show that the cercosporin had not disappeared around the colony

due to absorption into the cells.

Mitchell, et al. followed up on this work by screening 244 bacterial isolates,

representing 12 different genera and 23 different species, for the ability to degrade

cercosporin (71). The bacteria were incubated on cercosporin-containing medium and

assayed for a cleared halo. The organisms that were found to degrade cercosporin were

Xanthomonas arboricola (pseudonym campestris) pv. pruni (80, 96), Xanthomonas

campestris pv. zinniae, Pseudomonas syringae pv. pisi, Mycobacterium smegmatis, Bacillus

subtilis, Bacillus cereus, and two unidentified propane users. Interestingly, of the 32 isolates

of Xanthomonas campestris pv. zinniae (XCZ) that were tested, all 32 were able to degrade

cercosporin.

Mitchell, et al. further studied two degrading isolates, Xanthomonas arboricola

(pseudonym campestris) pv. pruni, strain 77 (XAP-77) and Xanthomonas campestris pv.

zinniae, strain 3 (XCZ-3), in order to determine if the degraded cercosporin could be used as

a carbon source (71). The results showed that neither bacterium was able to derive carbon

from cercosporin. Also, the ability to degrade cercosporin provided only a minimal benefit

to the bacteria when exposed to light. While the isolate XCZ-3 easily survived

concentrations of 50 μM cercosporin in the dark, exposure to light reduces the level of

resistance to only 1.25 μM cercosporin (71).

To characterize the degradation process, Mitchell, et al. assayed for the breakdown of

cercosporin using the degrading isolates XCZ-3 and XAP-77, and the non-degrading isolate

XAP-76 (71). The isolates were incubated in the dark in liquid medium containing

cercosporin and cercosporin concentration was determined spectrophotometrically. The

18

uninoculated control and the non-degrading isolate XAP-76 showed that cercosporin was

stable in the dark, as the concentration did not decrease over the course of the experiment

(71). Both of the degrading isolates showed a decrease in cercosporin concentration, such

that it was no longer detectable after 108 hours. Although degradation rates of the two

isolates were equivalent, XCZ-3 began degrading cercosporin at an earlier time point and

was considered to be the best isolate to use for further studies.

After noticing that the appearance of a green compound in the culture medium was

inversely related to the decreasing concentration of cercosporin, Mitchell, et al. performed a

time-course assay to compare the concentrations of the two compounds (71). Results showed

that cercosporin concentration began rapidly decreasing within the first 12 hours. The green

breakdown product was present as early as 24 hours after inoculation of the bacterium and

continued to accumulate for 60 hours, after which it gradually decreased. The appearance of

the breakdown product did not directly follow the loss of cercosporin, but this was suggested

to be due to a sequestration of cercosporin in the capsular polysaccharide (which was not

extractable), instead of being due to an immediate degradation of cercosporin (71). Thin

layer chromatography (TLC) was used to separate the breakdown product after extraction

from culture medium (71). TLC confirmed the results of the time course assays. As

cercosporin was degraded, the presence of the green breakdown product increased.

The breakdown product was extracted from culture medium and separated by TLC.

Spectral analysis of the purified product using HRMS and NMR revealed the chemical

formula C28H22O10 and the product was named xanosporolactone (70, 71). However, this

lactonized version of the breakdown product was a result of the purification process. The

19

true breakdown product formed by XCZ-3 in culture was actually found to have the chemical

formula C28H24O11 and was named xanosporic acid (70, 71).

Mitchell, et al. proposed that the conversion of cercosporin to xanosporic acid was

due to a single enzymatic step; insertion of an oxygen atom into one of the cercosporin

quinoid rings by a cytochrome P450 monooxygenase (70). Oxygen insertion would yield an

unstable seven-membered ring, possibly resulting in spontaneous reorganization and loss of a

methoxyl group.

Both xanosporic acid and xanosporolactone were tested to determine possible toxicity

to cercosporin-sensitive organisms (71). Xanosporic acid was tested for toxicity to the

cercosporin-sensitive mutant CS-8 of C. nicotianae. To generate xanosporic acid,

cercosporin-containing medium was pre-incubated with the XCZ-3 degrading isolate or the

non-degrading isolate XAP-76 as a control. Culture filtrates were then tested for toxicity

against CS-8. Cercosporin degraded to xanosporic acid by XCZ-3 showed no toxicity to CS-

8. In addition, purified xanosporolactone was tested for toxicity to tobacco by infiltrating

concentrations of 1, 5, 10 and 50 μM into leaves. After eight days, no necrotic lesions had

appeared on the leaves, whereas lesions appeared after only four days when infiltrated with

cercosporin at 0.9 μM. Mitchell, et al. concluded that both xanosporic acid and

xanosporolactone were non-toxic (71).

Cytochrome P450s

As indicated above, Mitchell, et al. suggested that the conversion of cercosporin to

xanosporic acid may be due to the action of a cytochrome P450, which would catalyze the

reaction by insertion of a single oxygen atom (71). Cytochrome P450s were initially

20

characterized by discovery of a reduced pigment from rat liver microsomes that had an

absorbance maximum of 450 nm after binding with carbon monoxide (37, 44). The pigment

was later characterized as a b-type cytochrome containing the heme group iron-

protoporphyrin IX (7, 44). P450s are monooxygenase enzymes which typically bind and

split apart molecular oxygen, O2, and introduce a single oxygen atom into a lipophilic

substrate (95, 101). In order to carry out this function, P450 monooxygenases require an

electron donor to reduce the heme iron and activate the molecule. In eukaryotic systems,

NAD(P)H is needed as the source of electrons. In bacterial systems, this donor is NADH.

In general, the entire P450 reaction is carried out in several steps (7). First, the

substrate is bound to the P450. Then the first electron is transferred from an iron-sulfur

ferredoxin to the P450 to yield a ferrous P450. Third, the P450 complex binds molecular

oxygen. Next, the second electron is transferred from the ferredoxin to the P450, fully

reducing the P450 and activating it. This is followed by splitting of the oxygen-oxygen bond

in molecular oxygen to yield activated oxygen. One of the activated oxygen atoms can then

be inserted into the substrate, while the other is released in the form of water. Finally, the

newly oxidized substrate can be released from the P450. Presence of substrate increases the

rate of P450-ferredoxin binding and the rate of transfer of the first reducing electron (40).

Protein-protein interaction is required for the electron to be shuttled from the ferredoxin to

the P450 (39). However, this interaction is short-lived (38). The reducing molecule may be

responsible for activating 10 to 25 different P450 molecules, and so must move rapidly from

P450 to P450, reducing (activating) each in turn (79).

Typically, P450 oxygen insertions occur at terminal portions of molecules, often

resulting in alcohols. P450 monooxygenases can function at allylic positions, double bonds,

21

and non-activated C-H bonds (95). Oxygen atoms can also be inserted into internal sites of a

molecule, such as into quinone rings, as an example of the Baeyer-Villiger reaction (102).

P450s frequently function in biosynthetic pathways, such as in alkaloid, lipid, or

antioxidant synthesis (13, 100), or in metabolization of foreign compounds, such as toxins

and drugs (7, 49). P450s that function in biosynthetic pathways have a higher degree of

substrate specificity than those that function in xenobiotic degradation (49). Harmful

xenobiotics, such as drugs and poly-brominated biphenyls are usually metabolized to a form

that is no longer toxic (7). However, xenobiotics are not always detoxified. Sometimes

alteration of the substrate can yield a harmful product, such as with the conversion of the

harmless benzo[a]pyrene to the mutagenic diol-epoxide derivative (7).

P450 gene expression is often inducible based on presence of the substrate (44). It

was recognized that when animals were treated with different compounds, certain

monooxygenase activities were induced (7). It has also been shown that treatment of rats

with certain xenobiotics specifically induced metabolization of the xenobiotics (44). P450

genes may be induced by various substances, such as steroids, phenobarbital and

hypolipidemic drugs. In addition to inducible expression, P450 genes may be expressed

constitutively, expressed only during certain developmental stages, expressed only in certain

tissues, expressed only in males or females, or specifically suppressed under certain

conditions (44). Some P450s are regulated posttranscriptionally, such as the P450 IA2 from

rats. The specific form of P450 regulation that is used depends on the organism and on the

P450 subfamily.

The structures of P450 enzymes are all very similar, having the shape of a triangular

prism, although the amino acid sequences are usually less than 20% identical (49, 95). P450s

22

classified in the same family are at least 40% identical at the amino acid level, whereas

subfamilies are at least 55% identical (100). Structural homology between amino acid

sequences of five different cytochromes (from eukaryotic and prokaryotic systems) has been

shown to be conserved in three regions (7). Two of the three regions are positioned around a

conserved cysteine residue, one around amino acid 152 and the other around position 438.

The third region of conservation occurs around position 390. The presence of conserved

regions in cytochromes from different organisms suggests an important role for each region

in enzyme function.

Cytochrome P450s are classified based on their redox partners (49, 95). In class I,

typically found in prokaryotes, two molecules are required to activate the P450: a

flavoprotein, which binds a flavin adenine dinucleotide (FAD) and an iron sulfur ferredoxin,

which binds a flavin mononucleotide (FMN). FAD and FMN are both essential cofactors for

transferring electrons to the cytochrome P450. The model system for this P450 class is

P450cam from Pseudomonas putida (95). Class II P450s interact with a cytochrome P450

reductase, a single molecule that binds both FAD and FMN. Enzymes in class II are found

in eukaryotes and are usually membrane-bound, such as to the endoplasmic reticulum (95).

Typical class II P450s are those found in mammalian liver cells, which are responsible for

steroid metabolism and detoxification (81). Class III P450s also use FAD and FMN as

cofactors, but these cofactors are bound to the P450, yielding a single, self-sufficient enzyme.

Class III P450s act on substrates that already contain oxygen and rearrange the molecule

(95). An example of a prokaryotic class III P450 is the P450 BM-3 of Bacillus megaterium,

which has functional and structural similarity to the class II enzymes (49). However, P450

BM-3 is self-sufficient, having no need of an additional reductase molecule. In P450 BM-3,

23

the P450 monooxygenase and the FAD/FMN domain occur as a single molecule (95).

Finally, class IV P450s are described as needing no other protein components other than the

P450 itself. The only known example of this class of P450s is P450nor, from Fusarium

oxysporum (75, 95). P450nor obtains its electrons directly from NAD(P)H, without the need

for reducing molecules. In addition, FAD and FMN are not needed as cofactors.

Most bacterial P450s are soluble and belong to the class I monooxygenases (74). As

indicated above, class I monooxygenases require two component molecules: a FAD-binding

reductase and an FMN-binding iron-sulfur ferredoxin. The FAD-binding reductase accepts

electrons from NADH and transfers them to the ferredoxin. The ferredoxin then shuttles the

electrons to the substrate-bound P450 to activate the complex. Bacterial P450s are usually

substrate specific, with the reactions they carry out being regioselective and stereoselective

(7).

The most well-studied prokaryotic P450 system is that of P450cam from

Pseudomonas putida (74). P450cam belongs to the monooxygenase class I and catalyzes the

5-exo hydroxylation of camphor (49, 74). It was the first cytochrome P450 to have its three-

dimensional structure determined, which has the shape of a flattened, triangular prism (7).

The heme group is buried within the interior of the structure. The genes camA, camB, and

camD code for the FAD-binding putidaredoxin reductase, the iron-sulfur protein

putidaredoxin, and the 5-exo-hydroxy-camphor reductase, respectively, and occur in an

operon along with camC (74). This operon, known as camDCAB, is negatively regulated by

the product of camR. When the camphor substrate is bound to P450cam, hydrophobic

interactions and hydrogen bonding hold the molecule in place (7). The rate-limiting step in

P450cam performance has been determined to be the electron transfer to the heme group of

24

the P450 (74). Much of what is known about cytochrome P450s, including those of

eukaryotes, has been determined through study of the P450cam system.

Disease Control Through Transgenic Resistance to Toxins The use of transgenes that can provide resistance to pathogen-produced toxins has

proven to be a successful means of disease control under laboratory conditions. One

example of transgenic resistance allowed for resistance in tobacco against phaseolotoxin

(28). The bacterium Pseudomonas syringae pv. phaseolicola causes halo blight disease on

bean. It produces the non host-specific toxin phaseolotoxin, which inhibits the enzyme

ornithine carbamoyltransferase (OCTase) needed for amino acid biosynthesis. OCTase,

found in plant chloroplasts, converts ornithine and carbamoyl phosphate to citrulline, which

is necessary for the production of arginine. The presence of phaseolotoxin interrupts this

reaction and leads to an accumulation of ornithine and to the production of chlorotic

symptoms. P. syringae pv. phaseolicola is resistant to its own toxin due to the production of

a phaseolotoxin-insensitive OCTase. An experiment in which the bacterial phaseolotoxin-

insensitive OCTase was transformed into tobacco and targeted to the chloroplast resulted in

transgenic tobacco plants that did not show the typical chlorotic symptoms in response to

phaseolotoxin (28). Although tobacco is not the normal host for P. syringae pv.

phaseolicola, the OCTase transgene provided resistance to the toxin.

Another example of using transgenes to achieve toxin resistance is the use of the

tabtoxin resistance protein against tabtoxin (4). Tabtoxin is produced by Pseudomonas

syringae pv. tabaci, a pathogen of tobacco that causes wildfire disease. When tabtoxin is

introduced into the host plant, it is cleaved to yield tabtoxin-ß-lactam. Tabtoxin-ß-lactam

25

inhibits glutamine synthetase, which leads to ammonia accumulation and the death of plant

cells. It also results in the production of chlorotic symptoms on tobacco leaves. P. syringae

pv. tabaci naturally carries a gene, ttr, which makes it resistant to its own toxin. The product

of the tabtoxin resistant gene, TTR, catalyzes the acetylation of tabtoxin, rendering the toxin

harmless (29). Transformation of tobacco plants with the ttr gene resulted in transgenic

plants that were resistant to tabtoxin, as well as to wildfire disease caused by P. syringae pv.

tabaci (4). In addition, the presence of the gene and the resulting resistance was shown to be

hereditary.

Resistance in sugarcane to Xanthomonas albilineans has also been accomplished

through transgenic approaches (107). Sugarcane is susceptible to X. albilineans, which

invades the xylem and causes leaf scald disease. X. albilineans produces toxins known as

albicidins, which are low molecular weight molecules that can block prokaryotic DNA

replication, such as that occurring in chloroplasts. The resulting under-development of the

chloroplasts leads to chlorosis. Albicidins have been indicated as playing a key role in the

systemic invasion by the pathogen. Expression of the gene albD, found naturally in

Pantoeae dispersa, codes for an esterase and provides for albicidin detoxification (106).

Expression of the albD gene in transgenic sugarcane not only provided resistance against the

usual chlorotic symptoms during X. albilineans infection, but also hampered bacterial

multiplication and systemic invasion (107). Plant lines with high expression of albD showed

little to no disease, whereas plant lines with low AlbD activity showed a high level of disease

severity.

Successful transgenic resistance has also been accomplished with fungal toxins. The

fungus Eutypa lata can produce up to six different toxins, although individual strains may

26

produce various combinations of the toxins (55). One of the toxins, eutypine, has been

shown to play an important role in causing dying arm disease of grapevines by E. lata (47).

Eutypine has been shown to accumulate in the cell cytoplasm due to an ion trapping

mechanism, and to uncouple mitochondrial oxidative phosphorylation. Presence of the toxin

leads to dwarfed and withered shoots and inflorescences, necrosis of leaf margins, and

eventual death of entire branches. Eutypine can be reduced by an enzyme derived from

mung bean (Vigna radiata), a non-host plant of E. lata. This enzyme is a NADPH-dependent

reductase identified as eutypine-reducing enzyme (VR-ERE). The reduced form of eutypine

has been designated as eutypinol. Although eutypinol has been shown to have inhibitory

effects on mitochondrial respiration in yeast, it is non-toxic to grapevine tissue (19). The

gene for VR-ERE was transgenically over-expressed in grapevine cells cultured in vitro (47).

When treated with purified eutypine, the transgenic cells were shown to be resistant to the

toxin. As of yet, there are no reports as to whether transgenic plants expressing the VR-ERE

gene have been recovered and tested for resistance to E. lata or eutypine.

Trichothecenes are fungal toxins produced by cereal pathogens in the genus Fusarium

(51). Trichothecenes inhibit protein synthesis in eukaryotes and prevent polypeptide chain

initiation or elongation (56). They act as virulence factors during pathogen infection, causing

damage to the infected plants. In addition, they can yield serious problems in the organisms

that eat the infected plants. Trichothecene-producing fungi protect themselves from the

toxins through expression of the gene Tri101, a gene for 3-O-acetyltransferase. TRI101 acts

specifically on the 3-hydroxyl group located at the C-3 position on tri-oxygenated

trichotriols. Higa, et al. targeted the Tri101 gene for transgenic expression in the model plant

tobacco (51). High-expressing transgenic lines were protected from the toxic effects of

27

trichothecene exposure. Although tobacco is not the natural host for Fusarium spp., the

results are promising for field application.

Recently, transgenic lines of sugar beet were developed that express superoxide

dismutase genes from tomato (90). Superoxide dismutase is responsible for converting

harmful superoxide radicals into hydrogen peroxide. Previous research had shown that

increased levels of superoxide dismutase activity in plants resulted in increased tolerance to

stress. Tertivanidis, et al. transformed sugar beet with two superoxide dismutase genes from

tomato for expression in the chloroplast and the cytosol (90). The transgenic plants

displayed a higher tolerance to oxidative stress induced by purified cercosporin, continuing

to grow and appear healthy, whereas the control plants did not display continued growth and

appeared unhealthy. The transgenic plants were also more resistant to disease caused by C.

beticola infection, displaying fewer leaf spots when compared to the wild-type control. In

addition, there were no developmental differences observed between the transgenic and

control plants. Field trials of these transgenic plants have not yet been reported.

Summary Cercospora spp. pose a serious threat to crops throughout the world. Production of

the non-specific toxin cercosporin by most Cercospora spp. has been shown to play a major

role in causing disease. Unfortunately, present cultural controls often do not work, and

fungicide applications are impractical due to the high expense. Breeding for resistance has

the potential to produce resistance crop varieties, but this method is slow and often results in

undesired phenotypic traits when resistance is achieved. However, using transgenes that can

provide resistance to pathogen-produced toxins can be a useful strategy for disease control.

28

Although successful examples of this are still limited to laboratory trials, the potential for

success in the field exists.

Xanthomonas campestris pv. zinniae is a potent degrader of cercosporin, rapidly

converting cercosporin into a non-toxic breakdown product. The goal of this research will be

to identify and characterize the gene(s) in XCZ responsible for cercosporin degradation.

These genes will then be cloned into tobacco using Agrobacterium-mediated transformation.

The resulting transgenic plants can then be tested in order to determine if the degradation

genes are sufficient to yield resistance to cercosporin and to Cercospora- infection.

29

References Cited

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3. Balis, C., and M. G. Payne. 1971. Triglycerides and cercosporin from Cercospora

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42

CHAPTER 2

A putative oxidoreductase is involved in cercosporin degradation by the bacterium

Xanthomonas campestris pv. zinniae

43

Abstract:

The fungal toxin cercosporin plays a key role in pathogenesis by Cercospora spp.

The bacterium Xanthomonas campestris pv. zinniae (XCZ) is able to rapidly degrade this

toxin. Growth of XCZ strains in cercosporin-containing medium leads to the breakdown of

cercosporin and to the formation of a non-toxic breakdown product called xanosporic acid.

Five non-degrading mutants of a rapid-degradation strain (XCZ-3) were generated by EMS

mutagenesis. Mutants were complemented using a genomic library of the wild type strain.

All five mutants were complemented with the same fragment, which encoded a putative

transcriptional regulator and an oxidoreductase. Simultaneous expression of the two genes

was necessary to complement the mutant phenotype. Sequence analysis of the mutants

showed that all five had mutations in the oxidoreductase sequence and no mutations in the

regulator. Quantitative RT-PCR showed that expression of both of these genes in the wild

type strain is upregulated after exposure to cercosporin. Both the oxidoreductase and

transcriptional regulator genes were transformed into three cercosporin non-degrading

bacteria to determine if they are sufficient for cercosporin degradation. Q-RT-PCR analysis

confirmed that the oxidoreductase gene was expressed in all transconjugants. However, none

of the transconjugants were able to degrade cercosporin, suggesting that other factors are

involved in this process. Further study of cercosporin degradation in XCZ may allow for the

engineering of Cercospora-resistant plants.

44

Introduction:

Members of the fungal genus Cercospora are pathogenic on plants, causing damaging

leaf spot and blight diseases. The genus collectively has a wide host range, with individual

species parasitizing specific hosts. Cercospora spp. can infect such hosts as tobacco, corn,

soybean, rice, sugar beet, and banana (4, 14, 17, 28, 40). Crop damage and losses can be

very high.

Cercospora fungi are known for the production of the non-host-specific toxin

cercosporin. Cercosporin is deep red in color and was first isolated from the dried mycelium

of Cercospora kikuchii, a soybean pathogen, in 1957 by Kuyama and Tamura (23). It has

the molecular formula C29H26O10 (Fig. 1), as shown independently by both Yamazaki, et al.

(46) and Lousberg, et al. (26). Cercosporin is produced by most members of the Cercospora

genus and has since been isolated from many Cercospora species, as well as from infected

host plants (1, 3, 17, 18, 23, 28, 31, 43). It is a polyketide secondary metabolite and a

nonspecific toxin, showing toxicity to plants, animals, bacteria and fungi (14). Cercosporin

is a photosensitizer, and uses light to become toxic to cells (11, 47). In general,

photosensitizers absorb light energy and enter the excited triplet state. Once in the triplet

state, photosensitizers are able to interact with molecular oxygen to form the activated

oxygen species superoxide and singlet oxygen. Although cercosporin has been shown to

produce both activated oxygen species, singlet oxygen appears to cause the most damage (11,

15, 21, 24). Presence of singlet oxygen in a plant can lead to lipid peroxidation and the

breakdown of cell membranes (12, 13).

Production of the toxin cercosporin has been associated with parasitism and

development of disease in host plants (14). A role for cercosporin in causing disease has

45

been shown by Upchurch et al. (41). UV-induced mutants of Cercospora kikuchii that were

able to synthesize no more than 2% of wild-type cercosporin levels yielded a few, small

flecks when inoculated on soybean, whereas the wild-type yields large, coalescing lesions

(41). Research by Chung et al., and Choquer, et al. (7, 8) also showed the importance of

cercosporin in disease-causing ability. The polyketide synthase responsible for the synthesis

of cercosporin was isolated, and then disrupted, to yield cercosporin-deficient mutants in

Cercospora nicotianae. When these mutants were inoculated onto tobacco, lesions were

fewer and smaller compared to wild-type (7). Complementation of the mutated polyketide

synthase restored wild-type levels of cercosporin and virulence. As further evidence, disease

severity of Cercospora leaf spot diseases has been shown to be directly related to light

intensity in both sugar beet and banana (6, 40). The onset of disease symptoms occurs earlier

on plants exposed to higher light intensities, whereas low light intensities cause a delay in

disease symptoms. In addition, studies by Steinkamp, et al. showed that ultrastructural

changes caused by Cercospora beticola on infected sugar beet are almost identical to the

changes caused by cercosporin treatment (13, 37, 38). In both cases, inoculated or treated

tissue showed a loss of cell membranes and the accumulation of large amounts of granular,

electron-dense material in the intercellular spaces.

Presently, control of diseases caused by Cercospora species relies on the use of

fungicides and appropriate cultural practices, such as tillage and crop rotation (44, 45).

Development of resistant cultivars has been attempted, but high levels of resistance to

Cercospora diseases are uncommon. Although limited disease resistance has been found in

corn, sugar beet, and soybean (9, 33, 36, 44, 45), alternate methods of control would be

46

desirable. One such approach would be to target the toxin cercosporin through engineering

of plants to express genes encoding degradation enzymes.

A patent was filed by Robeson et al. in 1993 on a method to identify bacteria which

had the ability to degrade cercosporin (35). Mixed bacterial populations were harvested from

the soil, leaf surfaces and leaf tissue of Cercospora beticola-infected sugar beet plants and

inoculated onto cercosporin-containing medium. After incubation in the dark, bacteria that

are able to degrade cercosporin were identified by a cleared halo surrounding the colony on

the red-colored plates. Four isolates were discovered that could degrade cercosporin:

Bacillus thuringiensis, Pseudomonas fluorescens biovar V, Mycobacterium sp., and Bacillus

subtilis.

In previous work in our laboratory, we followed up on the findings of Robeson, et al.

and screened bacteria for the ability to degrade cercosporin. A total of 244 isolates,

representing 12 different genera and 23 different species, were screened. Of these, 43

isolates were identified that were able to degrade cercosporin based on the production of a

cleared halo when bacteria were cultured on cercosporin-containing medium (30). Of the

isolates tested, Xanthomonas species had the largest proportions of cercosporin-degraders

and among those, isolates of Xanthomonas campestris pv. zinniae (XCZ), in particular, had a

strong ability to degrade cercosporin, with all 32 isolates tested having this ability. Time

course assays of cercosporin degradation carried out with one particular isolate, XCZ-3,

revealed that cercosporin degradation began as early as 12 hours after exposure to

cercosporin.

Mitchell, et al. subsequently showed that cercosporin degradation leads to the

production of a green breakdown product identified as xanosporic acid (29) (Fig.1).

47

Xanosporic acid was shown to be nontoxic to a cercosporin-sensitive mutant of Cercospora

nicotianae (30). The lactonized derivative of xanosporic acid, xanosporolactone, was used in

a plant infiltration assay of tobacco and was also shown to be nontoxic (30).

The goal of the work reported here is to isolate and characterize the gene(s)

responsible for cercosporin degradation by the wild-type strain XCZ-3. We show that the

ability of XCZ-3 to degrade cercosporin relies on the presence of a putative operon,

containing genes encoding a putative oxidoreductase and transcriptional regulator.

48

Materials and Methods:

Bacterial Cultures: The XCZ-3 strain (30) and XCZ-3 mutants were maintained on Luria-

Bertani (LB) medium (10 g of tryptone, 10 g of NaCl, and 5 g of yeast extract per liter; pH

7.2) containing rifampicin at 100 μg/mL. Isolates Xanthomonas arboricola (pseudonym

campestris) pv. pruni (34, 42) strains XAP-76, XAP-50, XAP-75, XAP-1, XAP-39, XAP-49,

XAP-34, XAP-31, and XAP-79 were obtained from Dr. David Ritchie (North Carolina State

University) and were maintained on LB medium without antibiotics. Isolates of

Xanthomonas axonopodis (pseudonym campestris) pv. vesicatoria (34, 42) (XAV-79 and

XAV-126), also obtained from David Ritchie, were maintained on LB containing 200 μg/mL

copper-sulfate. Isolates Pseudomonas syringae pv. tomato (PST) DC3000 and P.s. pisi (PSP)

209-21-3R were obtained from Dr. Peter Lindgren (North Carolina State University) and

were maintained on LB containing rifampicin (as above). All of the bacterial isolates were

maintained at 28 ºC in the dark.

Cercosporin Stocks and Chemicals: Cercosporin was isolated and purified as previously

described (11). Dried cercosporin crystals were dissolved in acetone for use in media. All

other chemicals were obtained from Sigma-Aldrich Co. (St. Louis, MO) unless noted

otherwise. All restriction enzymes were obtained from Promega Corporation (Madison, WI).

Chemical Mutagenesis of Xanthomonas campestris pv. zinniae and Screening for Mutants

Unable to Degrade Cercosporin: Three of the non cercosporin-degrading mutants (MutA,

MutB and MutC) were created through ethyl methanesulfonate (EMS) mutagenesis following

the protocol of Thorne et al. (39). The other two non-degrading mutants (MutD and MutE)

49

were obtained by first transforming the wild-type with the multicopy vector pLAFR6

(obtained from Dr. Brian Staskawicz, University of California, Berkeley) containing both the

transcriptional regulator and the oxidoreductase. EMS mutagenesis was then performed to

yield the mutants. Screening for cercosporin non-degrading mutants (Fig. 2) was carried out

as previously described (30). For MutD and MutE, the pLAFR6 vector was maintained

through antibiotic selection until the mutants were discovered, then removed through plasmid

curing. Mutant colonies were grown on LB medium containing rifampicin at 100 μg/mL to

ensure that the mutants retained the RifR marker.

Box PCR: The whole cell method of Box PCR was performed according to the protocol

established by Louws et al. (27) in order to confirm that the non-degrading mutants were

derived from the isolate XCZ-3. The following cycle conditions were used: 95 °C for 2 min.,

followed by 35 cycles of 94 °C for 3 sec., 92 °C for 30 sec., 50 °C for 1 min., 65 °C for 8

min. Ten microliters of each PCR product were run on a 1.5% agarose gel at 36 volts for

approximately 15 hours at 4 °C. The banding patterns on the ethidium bromide stained gel

were viewed under UV light (Fig. 3).

Xanthomonas campestris pv. zinniae Genomic Library Construction: Genomic DNA was

purified from XCZ-3 as described (2). A genomic library was created based on the protocol

of Thorne et al. (39). The genomic DNA was digested with Sau3A and fragments of

approximately 21 kb were selected. The fragments were cloned into the BamHI restriction

site of the expression vector pLAFR6, which carries tetracycline resistance as the selectable

marker. Plasmids were then transformed into Epicurian Coli JM109 competent cells

50

(Stratagene, La Jolla, CA). Transformants were selected for tetracycline resistance. A total

of 2016 clonal colonies were selected in order to form the library. Colonies were grown in

96-well plates and stored at –80 °C.

Complementation Assays: Conjugation of the non-degrading mutants with the library clones

was carried out using a tri-parental mating procedure with the helper cell line pRK2013 (16).

The library clones were cultured in LB liquid medium containing tetracycline at 20 μg/ml

and incubated at 37 ºC. The helper cell line pRK2013 was cultured in LB liquid medium at

37 ºC. The non-degrading mutants were cultured in LB liquid medium containing 100 μg/ml

rifampicin at 28 ºC in the dark. For conjugation, 133 μL of each culture was mixed together

and plated onto 0.45 µm nitrocellulose membrane on LB solid medium using a 96-well

replicator. The bacteria were allowed to dry on the membrane for 15 minutes, and then were

incubated for 48 hours at 28 ºC in the dark. Colonies were then moved to LB solid medium

containing 100 μg/ml rifampicin and 20 μg/ml tetracycline. For screening, a minimum of

three colonies per conjugation were collected and transferred to nutrient agar (NA)

containing 50 μM cercosporin. As a control, the colonies were also grown on NA containing

0.5 % acetone (used to solubilize cercosporin). A density of 12 colonies per plate was used

for the screening process. Two controls, the cercosporin-degrading XCZ-3 isolate and the

non-degrading XAP-76 isolate, were also inoculated onto each plate. The colonies were

incubated in the dark at 28 ºC for 12 days and screened for a cleared halo surrounding the

colony. Colonies were also screened on M9 minimal medium to identify auxotrophs.

51

Subcloning of Complementing Library Clone: Subcloning was carried out through use of

restriction enzyme digests with EcoRI and HindIII. The digested clone was run on a 1.0%

agarose gel, and the individual bands were excised and purified using the Qiagen Gel

Purification kit (Qiagen, Valencia, CA). Subclones were ligated into the expression vector

pLAFR6 (linearized with EcoRI), and transformed into DH5α chemically competent cells

(Invitrogen, Carlsbad, CA) following the manufacturer’s protocol. Complementation assays

were performed with each subclone as described above.

Sequencing of Subclone: The single complementing subclone (a 7.5 kb fragment obtained

from the EcoRI digest) was sheared using a Hydroshear. Sheared fragments were run on a

1.0 % agarose gel, and DNA fragments sized between 1 – 2 kb were excised and purified

using the Qiagen Gel Purification kit (Qiagen, Valencia, CA). The End-Repair kit

(Epicentre, Madison, WI) was used to blunt-end the fragments. Fragments were ligated to

pBluescript II SK+ (Stratagene, La Jolla, CA) and the ligation was transformed into DH10β

electrocompetent cells (Invitrogen, Carlsbad, CA). Colonies were chosen using blue/white

screening and ampicillin resistance, and then cultured overnight. Plasmids were purified

from each colony and were amplified by PCR using the standard primers T3 and T7. The

PCR products were sequenced at the North Carolina State University Genome Research Lab.

Sequencing of Mutant Genes: The transcriptional regulator and oxidoreductase from each

non-degrading mutant were sequenced to identify the mutations. Sequences were amplified

by high fidelity PCR using a combination of both Taq DNA Polymerase (Promega, Madison,

WI) and PfuUltraTM High-Fidelity DNA Polymerase (Stratagene, La Jolla, CA). PCR

52

products were loaded on a 1.0 % agarose gel and the desired bands were excised. The

Promega Wizard SV Gel and PCR Clean-Up Kit was used to purify the DNA from the

agarose gel. The bands were then cloned into the pGEM-T Easy vector system (Promega,

Madison, WI) according to the manufacturer’s protocol. The genes were sequenced using

the pGEM-T Easy sequencing primers T7 and SP6.

Gene Complementation: The oxidoreductase gene was PCR-amplified from XCZ-3 using

the primers F8kb1 (5’-TCCTGCTCGATGTCAATGC-3’) and R8kb1 (5’-

GAAGTTCTAGGGGCGACCAG-3’). The PCR product was cloned into the pGEM-T Easy

vector, excised by restriction digest with EcoRI, and cloned into pLAFR6. A second

construct, containing both the transcriptional regulator and the oxidoreductase gene, was

made in the same fashion using primers newsense2 (5’-AGCACCATTGACGCAAGCACC-

3’) and R8kb1. A third construct, containing 330 bp at the 3’ end of the transcriptional

regulator, and all of the oxidoreductase was made by amplification with primers F915 (5’-

TGATCGGGCCATTACTGCTGTT-3’) and R8kb1. All three pLAFR6 constructs were

tested for the ability to complement the cercosporin non-degrading mutants.

Quantitative RT-PCR of Oxidoreductase and Regulator in Wild-Type XCZ-3: XCZ-3 was

grown in LB liquid medium or LB medium containing 50 μM cercosporin or an equal

volume of acetone (0.5%). RNA was isolated at 24 hours using the Promega SV Total RNA

Isolation kit. A total of 1 μg RNA for each sample was used to make cDNA using Applied

Biosystems (Foster City, CA) TaqMan® Reverse Transcription Reagents kit. Primers for the

oxidoreductase and transcriptional regulator genes were designed using Primer Express

53

(Applied Biosystems, Foster City, CA) software and acquired from Sigma Genosys (The

Woodlands, TX). For the oxidoreductase, the primers F1 (5’-TGCGCTACGAACGGATCA-

3’) and R1 (5’-GGCACGAGCTTGGGAATGT-3’) were used. For the transcriptional

regulator, primers F2 (5’-TCTTCAATCGCTTGGTCGAT-3’) and R2 (5’-

ATCGCTTCAAGCTTGTCCAG-3’) were used. As an internal control, the internal

transcribed sequence (ITS) that resides between the 16S and 23S rRNAs was amplified using

primers F-ITS (5’-GTTCCCGGGCCTTGTACACAC-3’) and R-ITS (5’-

GGTTCTTTTCACCTTTCCCTC-3’) (20). Quantitative RT-PCR was carried out in the MJ

Research DNA Engine Opticon® 2 thermal cycler using 2X SYBR Green Master Mix

(Applied Biosystems, Foster City, CA) with the following conditions: 95 °C for 10 min.,

followed by 40 cycles of 95 °C for 15 sec., 60 °C for 1 min., 72 °C for 20 sec. The sample

plate was read after every cycle. Analysis of the qRT-PCR results was carried out according

to Livak, et al. (25).

Southern Analysis: A labeled probe was created using Digoxigenin (Roche, Indianapolis,

IN) and the primers F8kb1 (5’-TCCTGCTCGATGTCAATGC-3’) and R8kb1 (5’-

GAAGTTCTAGGGGCGACCAG-3’) as per the manufacturer’s protocol. Genomic DNA

was isolated (2) for Southern analysis from XCZ-3, XAP-76, XAV-79, XAV-126, PST

DC3000, PSP 209-21-3R, XAP-50, XAP-75, XAP-1, XAP-39, XAP-49, XAP-34, XAP-31,

and XAP-79. A total of 5 μg of genomic DNA was digested with 10 μL EcoRI from

Promega (Madison, WI) in a total volume of 100 μL following the manufacturer’s protocol.

The digest was carried out at 37 °C for 18.5 hours. The EcoRI enzyme was heat-inactivated

for 15 minutes at 65 °C. The DNA was cleaned using the Promega Wizard® DNA Clean-Up

54

kit and the entire digest was loaded on a 0.7% agarose gel for Southern analysis. Treatment

and transfer of the Southern gel was carried out according to Ausubel, et al. (2), with the

exception of soaking the gel in 0.25 N HCl for eight minutes instead of 30 minutes. After

transfer, the membrane was exposed to hybridization buffer for four hours at 65 °C, and then

incubated overnight at 65 °C with hybridization buffer containing the Dig-labeled probe.

The Southern blot was developed according to the manufacturer’s protocol for Dig-labeled

probes (Roche, Indianapolis, IN).

Transformation of the Regulator and Oxidoreductase Genes into Non-Degrading Bacteria:

The original complementary 23 kb library clone contained in the pLAFR6 vector (p6-23kb)

was conjugated into cercosporin non-degrading bacteria (XAV-79, XAV-126 and PST

DC3000) following the tri-parental mating protocol previously described. Transconjugant

bacteria, as well as the corresponding wild-type strains, were grown in LB liquid medium

containing 50 μM cercosporin or 0.5% acetone. RNA was isolated at 24 hours and used to

make cDNA as previously described. Quantitative RT-PCR was carried out using the F1 and

R1 primers mentioned above in order to assess expression of the oxidoreductase gene. The

F-ITS and R-ITS primers were used to amplify the internal transcribed spacer region as an

internal control.

Time Course of Cercosporin Degradation by Transconjugants: Cercosporin degradation was

assessed by growing the transconjugants and the corresponding wild-type strains in 50 mL

LB liquid medium containing 60 μM cercosporin. 5 mL aliquots were taken at 0, 24, and 48

hours and stored at –20 °C until ready for use. Each sample was extracted twice with 2 mL

55

chloroform (30), followed by partitioning of the cercosporin into the aqueous phase using 1

N NaOH. Mixtures were vortexed, centrifuged, and the supernatant collected. The

supernatant was treated with 1 N HCl (to partition cercosporin into the organic phase) and

extracted with an additional 1 mL of chloroform. The chloroform extractions for the

individual samples were combined to yield a total of 5 mL for each culture. Cercosporin

degradation was quantified by measuring absorbance at 472 nm using a Beckman DU 650

spectrophotometer. The experiment was carried out twice.

56

Results:

Isolation of Xanthomonas campestris pv. zinniae Mutants Unable to Degrade Cercosporin:

Wild-type XCZ-3, when grown on cercosporin, produces a cleared halo surrounding the

colony (Fig. 2). By contrast, non-degrading strains do not produce a halo and turn dark in

color due to uptake of cercosporin. XCZ-3 cultures were mutagenized with EMS and

screened by transferring individual colonies onto cercosporin-containing medium. Presence

or absence of halo formation was determined at 12 days. Two initial screens resulted in the

isolation of three non-degrading XCZ-3 mutants. These were termed MutA, MutB and MutC

(Fig. 2). When tested on M9 minimal medium, none of the mutants showed auxotrophy. All

three of the mutants also retained the rifampicin resistance marker. Mutants were screened

by Box PCR to confirm their identity (Fig. 3).

Identification of Complementing Genes: Complementation of the cercosporin non -

degrading mutants with the genomic library was carried out using the tri-parental mating

procedure as outlined in the Materials and Methods. The transconjugant colonies were

screened for cercosporin degradation on cercosporin-containing medium. A single library

clone of 23 kb was found that complemented all the mutants. The 23 kb clone was

subcloned, and a 7.5 kb subclone was identified that complemented all the mutants. This

subclone was sequenced, the sequences aligned, and the resulting contigs blasted against the

NCBI database. A total of six genes were identified. These had sequence similarity to: a

pectate lyase, a soluble lytic murein transglycosylase, a TonB-dependent receptor, an

oxidoreductase, a transcriptional regulator, and a choline dehydrogenase (Fig. 4).

57

Initial efforts were focused on the transcriptional regulator and oxidoreductase genes.

Subcloning indicated that digestion of the complementing 23 kb library clone with HindIII

eliminated complementation ability, and the 7.5 kb subclone has two HindIII digestion sites,

one each in the transcriptional regulator and oxidoreductase genes. In addition, the

oxidoreductase was a likely candidate based on the proposed reaction for the formation of

xanosporic acid from cercosporin, which involves an oxygen insertion followed by

spontaneous rearrangement of the molecule (29). The region spanning the transcriptional

regulator and oxidoreductase was amplified by PCR and tested for the ability to complement

mutants MutA, MutB and MutC. This putative operon was able to complement all three

mutants.

The nucleotide sequence spanning the oxidoreductase and regulator is shown in Fig. 5

(GenBank Accession #DQ087176). The transcriptional regulator and oxidoreductase genes

appear to form an operon. The genes are transcribed in the same direction and are located

only 61 bp apart. A search done using the Basic Local Alignment Search Tool (BLAST),

version 2.2.11, showed that the translated form of the oxidoreductase gene shows highest

similarity to an oxidoreductase from Xanthomonas axonopodis pv. citri (XAC) strain 306

(Accession #AAM36534; E = 2e -114) (10), a flavoprotein monooxygenase from

Pseudomonas syringae pv. syringae strain B728a (Accession #YP_233205, E = 9e -105) (19),

and a monooxygenase from Pseudomonas syringae pv. tomato strain DC3000 (Accession

#NP_790149; E = 1e -85) (5). An alignment between the two xanthomonad oxidoreductase

sequences, carried out using the programs Vector NTI 8 (Informax, Inc., Frederick, MD) and

Clustal W 1.82 (EMBL-EBI, Heidelberg, Germany), showed that the two genes are 63%

similar at the amino acid level (Fig. 6). Despite the similarity, the XCZ-3 oxidoreductase

58

gene does not have any defining domains or motifs known to be associated with

monooxygenases according to an ExPASy PROSITE motif scan.

The translated XCZ-3 transcriptional regulator shows highest similarity to a

transcriptional regulator for cryptic hemolysin from XAC-306 (Accession #AAM36535)

(10), with an E value of 1e-37. An alignment between these two genes using Vector NTI and

Clustal W shows 70.1% similarity at the amino acid level (Fig. 7). Interestingly, the

homologous transcriptional regulator from XAC-306 lies directly upstream of the

homologous oxidoreductase. As in XCZ-3, these two genes appear to occur as an operon.

They are transcribed in the same direction and are only 30 bp apart. An alignment of these

two putative operons from XCZ-3 and XAC-306 shows 55.2% similarity at the nucleotide

level (data not shown).

The second most similar match to the transcriptional regulator is a MarR

transcriptional regulatory protein from Gloeobacter violaceus strain PCC 7421 (Accession

#NP_926807) that shows 53% similarity, with an E value of 5e-12 (32). An ExPASy

PROSITE motif scan revealed that the XCZ-3 transcriptional regulator contains a MarR-type

helix-turn-helix domain (Fig. 5).

Search for Additional Genes: Two additional mutants, termed MutD and MutE, were

obtained by EMS mutagenesis of XCZ-3 expressing the multicopy vector pLAFR6

containing both the transcriptional regulator and the oxidoreductase in an effort to identify

mutations in genes other than the transcriptional regulator and the oxidoreductase. Both

MutD and MutE showed the same non-degrading phenotype as the other mutants when

grown on cercosporin-containing medium (Fig. 2). Although the mutations in MutD and

59

MutE were generated while carrying multiple copies of the wild type transcriptional regulator

and oxidoreductase genes, they were still complemented by the putative operon containing

the wild-type transcriptional regulator and oxidoreductase genes, indicating that the new

mutations occurred in the same region already identified.

Sequencing of Mutants: The region spanning the transcriptional regulator and the

oxidoreductase genes was sequenced in each of the five non-degrading XCZ-3 mutants.

Sequence analysis identified single point-mutations in the oxidoreductase gene in all five

mutants, but no mutations in the transcriptional regulator (Fig. 5). Each mutation in the

oxidoreductase gene resulted in an amino acid change (Table 1). The predicted wild-type

oxidoreductase is 400 amino acids long and homology suggests that the active site resides

between amino acids 150 and 340. The mutations in MutA, MutB, and MutD fall within this

region. In MutC, the start codon was changed to a threonine, eliminating the protein start

site. In MutA, a tryptophan residue was changed to a stop codon, resulting in a truncated

protein. The mutations found in MutB, MutD, and MutE change three different glycines to

aspartic acid. When these three mutations were analyzed by NNPREDICT, the secondary

structure is predicted to be affected only in MutB. However, this substitution replaces a

single hydrogen with CH2COO-. It is not unexpected that substitution with a bulky and

acidic side chain would affect the tertiary structure and function of the protein.

Identification of Sequences Necessary for Mutant Complementation: As all mutants were

defective in the oxidoreductase only, we tested the ability of the oxidoreductase alone to

complement the mutants. The gene was amplified and cloned into the pLAFR6 vector

60

(Nucleotides 391-1948, Fig. 5). Surprisingly, this construct was unable to complement any

of the non-degrading mutants. By contrast, a construct containing the oxidoreductase gene

plus some upstream sequence (nucleotides 174-1948, Fig. 5) was able to complement all five

mutants. Although this construct does not include the full length gene for the transcriptional

regulator, it does include the potential active site (Fig. 5).

Presence of Cercosporin Upregulates Gene Expression: Quantitative RT-PCR was used to

determine if gene expression of either the wild-type oxidoreductase or transcriptional

regulator were regulated by the presence of cercosporin. Results were compared to

expression in cells grown in LB and in LB with 0.5% acetone, used to solubilize cercosporin.

Expression of the two genes in the acetone control was increased by approximately 3-4 fold

over the untreated control (Fig. 8). When treated with cercosporin, expression of the

transcriptional regulator and oxidoreductase were strongly upregulated, 17-fold and 136-fold,

respectively, compared to the untreated control.

Southern Analysis of Oxidoreductase Homologues in Degrading and Non-Degrading

Species: A series of cercosporin-degrading bacteria (XCZ-3, XAP-50, XAP-75, XAP-1,

XAP-39, XAP-49, XAP-34, XAP-31, and XAP-79) and non-degrading bacteria (XAP-76,

XAV-79, XAV-126, PST DC3000, and PSP 209-21-3R) were assayed by Southern analysis

for the presence of oxidoreductase homologues using a Digoxigenin-labeled probe amplified

from the wild-type XCZ-3 oxidoreductase (Fig. 9). All of the cercosporin-degrading isolates

were found to contain a sequence homologous to the XCZ-3 oxidoreductase. With one

exception, all of the non-degrading isolates lacked a detectable oxidoreductase homologue.

61

The one exception was PST DC3000. Although unable to degrade cercosporin, Southern

analysis detected a band that hybridized to the XCZ-3 oxidoreductase probe. A subsequent

NCBI BLAST search identified a similar sequence (Accession #NP_790149), with an E

value of 1e –85 (5).

The Transcriptional Regulator and Oxidoreductase are Not Sufficient for Cercosporin

Degradation in Non-Degrading Species: In order to determine if presence of the

transcriptional regulator and oxidoreductase genes alone are sufficient for cercosporin

degradation, cercosporin non-degrading bacteria were conjugated with the 23 kb

complementing library clone (in pLAFR6) containing the wild-type transcriptional regulator

and oxidoreductase genes. Quantitative RT-PCR was used to assess oxidoreductase gene

expression in each of the transconjugants upon exposure to cercosporin. Results show that

the oxidoreductase was expressed in all of the transconjugants and appeared to be regulated

similarly to XCZ-3, with increased expression upon treatment with cercosporin (Fig. 10).

PST DC3000 transconjugants, in particular, showed high levels of expression, equal to or

greater than that in the XCZ-3 wild type.

Although the oxidoreductase was expressed in the transconjugants, cercosporin-

degradation assays in both solid and liquid medium failed to detect cercosporin degradation

by the oxidoreductase-expressing transconjugants. A time-course assay of cercosporin

degradation in liquid medium is shown in Figure 11. Levels of cercosporin incubated in LB

medium alone remained constant over the 48 hours of the experiment. Incubation of

cercosporin with the wild-type control, XCZ-3, and the complemented mutant, MutA + p6-

23kb, resulted in almost complete loss of cercosporin by 48 hours. By contrast, incubation of

62

cercosporin with MutA and the non-degrading isolates XAV-79, XAV-126, and PstDC3000

resulted in only a small decrease in apparent cercosporin concentration, likely due to the

uptake of cercosporin into the cells and our inability to completely extract the cercosporin

from the cells. The transconjugants that were conjugated with p6-23kb were unable to

degrade cercosporin, showing no more loss of cercosporin than their wild type counterparts.

To ensure that other genes on the 23 kb clone were not in some way interfering with

degradation, the putative operon alone in pLAFR6 was conjugated into the non-degrading

bacteria. Cercosporin degradation by these transconjugants was still not achieved (data not

shown).

63

Discussion:

Many species in the genus Cercospora produce the molecule cercosporin and rely on

its toxicity to yield disease on their host plants. Research efforts to avoid or limit exposure of

a host plant to the toxin may be of great benefit in disease prevention. We have chosen to

focus on toxin degradation as a potential method to achieve toxin and disease resistance.

Previous reports of toxin degradation by transgenic plants have shown that this

method can be successful in yielding disease resistance. For example, Xanthomonas

albilineans causes leaf scald on susceptible sugarcane due to production of the toxin albicidin

(48). A gene for albicidin detoxification derived from Pantoea dispersa was used to

transform sugar cane. The resulting transgenic plants showed high levels of resistance to

pathogen invasion and multiplication, as well as a significant decrease in disease symptom

development. Another example is the use of the Tri101 gene derived from the fungus

Fusarium graminearum. F. graminearum utilizes the Tri101 gene as a means of self-

protection to inactivate the trichothecene toxins that it produces. When Tri101 was cloned

into tobacco, the transgenic tobacco was protected against the trichothecene DAS (22).

Although transgenic lines of susceptible cereals, the natural hosts of F. graminearum, have

not yet been generated, expression of Tri101 may prove to be a successful method for

generating resistant plants.

In previous work in our lab, we identified bacteria that are capable of degrading

cercosporin. Of 244 isolates tested, the most efficient degraders were pathovars of X.

campestris pathogenic on zinnia. XCZ isolates were shown to rapidly degrade cercosporin to

produce the non-toxic breakdown product xanosporic acid (30). The degradation reaction

was hypothesized to be caused by an oxygen insertion into one of the quinoid rings, adjacent

64

to the ketone carbonyl (29). The execution of this step would require enzymatic activity, but

the subsequent rearrangement that results in xanosporic acid may be spontaneous. We

hypothesized that cercosporin degradation is carried out by a single enzyme, perhaps by a

cytochrome P450.

The goal of this work was to isolate the gene(s) required for cercosporin degradation

by XCZ. We used EMS chemical mutagenesis to create non-degrading mutants of the wild-

type, and then identified the mutant genes by complementation. All mutants were

complemented by the same clone encoding a putative operon containing a transcriptional

regulator and an oxidoreductase. Homology searches showed that the transcriptional

regulator homologue has similarity to a transcriptional regulator for cryptic hemolysin and a

MarR transcriptional regulator. The oxidoreductase homologue has similarity to an

oxidoreductase and two monooxygenases. Although the regulator and oxidoreductase are

most similar to a putative operon in XAC, the function of these homologous genes has not

yet been experimentally confirmed in XAC.

Sequence analysis of the region spanning the transcriptional regulator and the

oxidoreductase in the mutants identified a single point-mutation present in the

oxidoreductase of each mutant, and no mutations in the transcriptional regulator. Our studies

suggest, however, that the transcriptional regulator may play an important role in regulation

of the oxidoreductase. A construct containing the wild-type oxidoreductase gene alone did

not complement the cercosporin non-degrading mutants, whereas a construct containing both

the transcriptional regulator and the oxidoreductase genes did complement the mutants. It is

not clear if the requirement is for the regulator or for upstream regulatory sequences.

Transformation of the mutants with a construct containing the oxidoreductase gene and

65

upstream sequences spanning 330 bases into the transcriptional regulator gene proved as

efficient in complementing the mutants as did the construct containing the intact regulator.

This construct did contain the putative active site of the transcriptional regulator as well as

sequences encoding five possible start sites that may allow the terminal portion of the

regulator to be translated in frame with the original sequence, resulting in expression of the

potentially necessary active site. It is interesting to note that we were unable to clone the

oxidoreductase gene alone into high expression vectors; recovered clones were slow to grow

and died after only five days (data not shown). When the gene for the transcriptional

regulator was included upstream of the oxidoreductase in constructs in the high expression

vectors, however, there were no problems in recovering clones that grew normally. These

results suggest that the transcriptional regulator plays a critical role in regulating expression

of the oxidoreductase. Finally, our qRT-PCR studies showed that both the regulator and the

oxidoreductase genes were upregulated after exposure to cercosporin, supporting our

hypothesis that expression of both genes is necessary to yield cercosporin degradation.

Although we attempted to disrupt the regulator gene in an effort to determine its role, we

were unsuccessful.

Since both the transcriptional regulator and oxidoreductase appear to have a role in

cercosporin degradation, the putative operon was transformed into other cercosporin non-

degrading bacteria in an attempt to determine if these two genes are sufficient for

degradation. Quantitative RT-PCR analysis confirmed that the two genes were expressed

and were upregulated in the presence of cercosporin. However, the transconjugants were

unable to degrade cercosporin, indicating that the two genes are not sufficient for cercosporin

degradation in bacteria.

66

We have hypothesized that cercosporin degradation involves the action of a

cytochrome P450 enzyme (29). Bacterial P450s require the participation of two additional

molecules in order to become activated: a flavodoxin and an iron sulfur ferredoxin, both of

which have a general oxidoreductase function. It is possible that the oxidoreductase gene

that we have isolated codes for one of the activating components. This would imply that the

P450 that actually catalyzes the degradation of cercosporin is absent in the transconjugants.

In an attempt to identify a P450 or other gene necessary for cercosporin degradation,

we mutagenized XCZ-3 that was transformed with a vector expressing multiple copies of the

region spanning the transcriptional regulator and the oxidoreductase, and screened for

cercosporin degradation. We hypothesized that expression of multiple copies of the wild-

type oxidoreductase would allow us to recover mutations in other genes. Surprisingly, the

two additional mutants that were recovered, MutD and MutE, also contained a mutation in

the oxidoreductase. We propose that additional genes are involved in the degradation

process, but that mutations in these genes are not recoverable, perhaps due to lethality.

Interestingly, Southern analysis showed that all bacterial isolates tested that are able

to degrade cercosporin contain a sequence homologous to the XCZ-3 oxidoreductase gene.

With one exception, bacterial isolates tested that are unable to degrade cercosporin, do not

contain a detectable oxidoreductase homologue. The exception is PST DC3000. It does not

have the ability to degrade cercosporin, but does contain a sequence homologous to the

oxidoreductase. These results are consistent with our hypothesis that additional genes may

be necessary for cercosporin degradation.

Because expression of the oxidoreductase gene has proven to be insufficient for

cercosporin degradation in non-degrading bacteria, further study is needed to identify

67

additional genes involved in this process. Once found, expression of these genes in plants

may provide for a novel strategy for control of Cercospora spp. and the diseases that they

cause.

68

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75

Table 1. Location of each mutation in the oxidoreductase gene of the five cercosporin non-

degrading mutants.

*Based on NNPREDICT analysis.

Mutant

Site of Mutation

(Amino Acid Residue) Nucleotide Change Amino Acid Change Change in Secondary Structure?*

MutA 251 G --> A tryptophan --> stop codon N/AMutB 158 G --> A glycine --> aspartic acid yesMutC 1 T --> C start codon --> threonine N/AMutD 196 G --> A glycine --> aspartic acid not predictedMutE 40 G --> A glycine --> aspartic acid not predicted

76

Figure 1. Chemical Structure of cercosporin and the breakdown product (xanosporic acid)

produced by X. campestris pv. zinniae, isolate XCZ-3.

Cercosporin

Xanosporic Acid

77

Figure 2. Cercosporin non-degrading XCZ-3 mutants. Mutants show a lack of a cleared halo

when grown on cercosporin-containing medium. The positive control (XCZ-3 wild-type)

appears at the top of the plate and the negative control (XAP-76) appears at the bottom of the

plate.

78

Figure 3. Agarose gel of Box PCR products. Lanes 1 and 10 = 1 kb Marker, Lanes 2 and 3 =

MutA, Lanes 4 and 5 = MutB, Lanes 6 and 7 = XCZ-3, Lanes 8 and 9 = MutC. Banding

patterns of mutants match those of the wild type strain.

79

Figure 4. Complementing fragment derived from sequencing the complementing 7.5 kb

subclone. Contig 9 alone complemented all five non-degrading mutants.

80

81

Figure 5. Sequence containing the putative transcriptional regulator and oxidoreductase

genes of XCZ-3 (GenBank Accession #DQ087176). The dashed line represents the

transcriptional regulator and the solid line represents the oxidoreductase. The darkened

triangles pointing to specific nucleotides in bold show the location of each of the mutations.

The shaded area denotes the sequence which is homologous to MarR genes and which

potentially contains a helix-turn-helix motif. The underlined, bold-faced codons are possible

alternative start sites that are in frame with the transcriptional

regulator.

82

Figure 6. Alignment of amino acid sequences of oxidoreductase genes from Xanthomonas

campestris pv. zinniae (XCZ-3) and Xanthomonas axonopodis pv. citri (XAC-306).

* = identical nucleotides;

: = conserved substitutions;

. = semi-conserved substitutions

XCZ3oxido ---MSRRILITGASVAGTTAAWWLDAYGVEVEVVERAPAFRDGGQNVDVRGSARDVLRRM 57 XACoxido VTAMPRRILITGASIAGNTAAWTLAHRGFDVTVAERATRFRDGGQNIDVRGVGREVLQRM 60 *.*********:**.**** * *.:* *.***. *******:**** .*:**:** XCZ3oxido GLEARAFERSTRELGTDWVDADDRVIARFKADASDTDSGPTADLEIRRGDLARMLYEASR 117 XACoxido GLERAALAQGTGEEGTAWIDAHGQAVATFKTEDVDGD-GPTAELEILRGDLARLLYDAAR 119 *** *: :.* * ** *:**..:.:* **:: * * ****:*** ******:**:*:* XCZ3oxido ERVAYRFGDSVCGVAQDQAG--VEITFHSGRCARYDAVIVAEGVGSHTREQVFPGENVPR 175 XACoxido SHVAYRFGDRIASIEDAAGSDAATVTFESGCSERFDAVIVAEGVGSSTREQVFPGENDPR 179 .:******* :..: : .. . :**.** . *:*********** ********** ** XCZ3oxido WMDLTLAYFSIPRQSHDSAYARQYNTVGGRGATLKPALDGKLGAYLGIQKKPGGENAWTP 235 XACoxido WMDLTIAYFTIPRSADDDRLWRWYHTTGGRSISLRPDRHGTTRAMLSLQKPPEGEQDWSV 239 *****:***:***.:.*. * *:*.***. :*:* .*. * *.:** * **: *: XCZ3oxido ERQRRFIETQFANDGWEFPRILAAMRDVDDFYFDVLRQVHMPRWSAGRVVLTGDAAWCPT 295 XACoxido DAQKAYLHERFADAGWQAARVLDGMHGTDDFYFDALRQVRMQRWHSGRTVLTGDAAWCAT 299 : *: ::. :**: **: .*:* .*:..******.****:* ** :**.*********.* XCZ3oxido SLSGIGTTLALVGSYVLAGELAQAVSPMHACMRYERIMRPFVKEGQNIPKLVPRLLWPHS 355 XACoxido PLAGIGATLAVTGAYVLANEIAQADTLDAAFAAYATAMRPMVEQAQGVPKIGPRLMNPHS 359 .*:***:***:.*:****.*:*** : * * ***:*::.*.:**: ***: *** XCZ3oxido AAGLTLLRGAMRVAGMPIVRRLVTDGFARDSNRIALPDYRPIATL 400 XACoxido RLGIHLLHGALKLASQPSVQNLAAKWMTPAIKAPDLSRYP----- 399 *: **:**:::*. * *:.*.:. :: : *. *

83

Figure 7. Alignment of amino acid sequences of transcriptional regulator genes from

Xanthomonas campestris pv. zinniae (XCZ-3) and Xanthomonas axonopodis pv. citri (XAC-

306).

* = identical nucleotides;

: = conserved substitutions;

. = semi-conserved substitutions

XCZ3reg VGYASRMTSDIHPEAAIAPGYLANHAARVFNRLVDARLREHGLTLALIGPLLLLSWKGPM 60 XACreg ---MLDQNELIHPQAAVAPGYLANHAARVFNRLVDAQLRPHGVSLALIGPIMLLSWKGPM 57 .. ***:**:*******************:** **::******::******** XCZ3reg LQRDLVRESAVKQPAMAALLDKLEAMALIAREASSTDKRAATVRLTAAGQDAAATGRTVL 120 XACreg LQRDLVIASAVKQPAMVALLDKLEGLKLIKRTPTPQDRRAALVALTARGRKIAELGGRAL 117 ****** ********.*******.: ** * .:. *:*** * *** *:. * * .* XCZ3reg LDVNALGVSGFSGKDAGLLTALLGRLIANLESAAGE- 156 XACreg LGMNAVALEGFSTDEVQQTVALMHRLIANMERHGQDV 154 *.:**:.:.*** .:. .**: *****:* . :

84

Figure 8. Quantitative RT-PCR analysis of expression of the transcriptional regulator and

oxidoreductase genes in XCZ-3 following exposure of the bacterium to cercosporin. CE =

cercosporin. Numbers above bars are fold-increase in expression as compared to the

untreated control, which was normalized to 1. The acetone control contained 0.5% acetone

used to solubilize cercosporin.

Trans criptional Regulator Regulation

1.0

3.3

17.0

0

2

4

6

8

10

12

14

16

18

Fold

Incr

ease

Untreated 50 µmCE

Acetonectrl

O x i do r e du c ta s e R e g u l a ti o n

1 .0 3 .6

1 3 6 .1

0

2 0

4 0

6 0

8 0

1 0 0

1 2 0

1 4 0

1 6 0

Fold

Incr

ease

Untreated 50 µmCE

Acetonectrl

85

Figure 9. Southern hybridization of total DNA isolated from cercosporin-degrading (XCZ-3,

XAP-50, XAP-75, XAP-1, XAP-39, XAP-49, XAP-34, XAP-31, and XAP-79) and non-

degrading (XAP-76, XAV-79, XAV-126, PST DC3000, and PSP 209-21-3R) isolates using a

PCR-amplified digoxigenin-labeled oxidoreductase gene as a probe. For each isolate, a total

of 5 μg DNA was digested with EcoRI and the entire digest was loaded on the gel. The wild-

type XCZ-3 was used in Lane 1 as a positive control. Sequences homologous to the

oxidoreductase are present in all degrading isolates and in the non-degrading isolate PST

DC3000.

86

Figure 10. Quantitative RT-PCR analysis of expression of the oxidoreductase in

transconjugants of the non-degrading bacteria XAV-79, XAV-126, and PST DC3000 as

compared to the XCZ-3 degrading strain. Figure shows estimated total oxidoreductase

transcript in non-transformed cells (wild-type) as compared to transconjugants grown with

and without cercosporin. CE = cercosporin; untreated = acetone control (used to solubilize

cercosporin). All transconjugants expressed the oxidoreductase and showed increased

expression in response to cercosporin.

0

500

1000

1500

2000

2500

3000

3500

4000

Est

imat

ed p

g/ug

Tot

al R

NA

XAV-79 XAV-126 PST DC3000

XCZ-3

Wild-type

Transconjugant / untreated

Transconjugant / 50 µM CE

XCZ-3 WT / 50 µM CE

87

Figure 11. Degradation of cercosporin in liquid culture by cercosporin-degrading and non-

degrading isolates and oxidoreductase-expressing transconjugants. Isolates tested were the

XCZ-3 wild-type, XCZ-3 non-degrading mutant MutA, wild-type non-degrading isolates

XAV-79, XAV-126, and PST DC3000, and the oxidoreductase-expressing transconjugants of

XAV-79, XAV-126, PST DC3000 and MutA. Cercosporin degradation was assayed by

measuring absorbance at 472 nm after extraction of cercosporin from culture filtrates at 0, 24,

and 48 hours as described in Materials and Methods. Cercosporin in LB alone (Untreated) is

included as a control. Small decreases in apparent cercosporin concentration with the non-

degrading bacteria are due to uptake of cercosporin by bacteria in the medium. Only XCZ-3

and the complemented MutA mutant were able to degrade cercosporin. Results shown are

from one of two experiments performed.

0

0.5

1

1.5

2

2.5

3

0 hrs 24 hrs 48 hrs

Time

Abs

orba

nce

Untreated

XCZ-3

MutA

MutA + p6-23kb

XAV-79

XAV-79 + p6-23kb

XAV-126

XAV-126 + p6-23kb

PST DC3000

PST DC3000 + p6-23kb

88

CHAPTER 3

The Xanthomonas campestris pv. zinniae oxidoreductase gene is regulated by plant

extracts, but is not necessary for pathogenesis.

89

Abstract:

Xanthomonas campestris pv. zinniae (XCZ) has been found to rapidly degrade

cercosporin, a photoactivated polyketide toxin produced by fungi in the genus Cercospora.

Studies have shown that XCZ contains a gene for a putative oxidoreductase that is necessary

for this process. As 32 of 32 isolates of XCZ tested had the ability to degrade cercosporin,

we hypothesized that zinnia may contain the natural substrate of this enzyme and that

degradation of the substrate was important in pathogenesis. Quantitative RT-PCR was used

to assay for oxidoreductase regulation in XCZ grown in the presence of zinnia and tobacco

extracts. The oxidoreductase gene was upregulated in the presence of both plant extracts, but

the response was greater with the tobacco extract. Five XCZ mutants with point mutations in

the oxidoreductase gene were tested for pathogenicity on zinnia. The mutants were not less

pathogenic than the wild-type. In addition, we previously identified other bacteria (XAP-50,

XAP-75, and PST DC3000) with oxidoreductase homologues using Southern analysis.

These bacteria were not able to infect zinnia. We conclude that the presence of a functional

oxidoreductase is not correlated with pathogenesis of zinnia.

90

Introduction:

The bacterium Xanthomonas campestris pv. zinniae (XCZ) is a pathogen of zinnia,

causing bacterial leafspot disease. This disease was first reported in Italy in 1929 and in

North America in 1972 (1, 15). The pathogen was originally identified as Xanthomonas

nigromaculans, but was later classified as a pathovar of Xanthomonas campestris (6).

Xanthomonads are Gram negative, aerobic, rod-shaped bacteria that do not form spores (1,

16). They may produce and secrete several hydrolytic enzymes, such as cellulases and

proteinases.

Xanthomonads also produce xanthan gum, a type of extracellular polysaccharide,

which surrounds the cells. This gel-like, slimy substance is thought to promote intercellular

bacterial colonization by protecting the bacteria against plant defense responses and

compounds and by providing a layer of moisture around the bacteria (16). Additionally,

xanthan gum may aid the bacteria during epiphytic growth, protecting them from desiccation

and UV radiation.

X. campestris can only enter the host through wounds or natural opening, such as

stomata and hydathodes (16). The bacteria colonize intercellular spaces and vascular tissue,

which allows them to spread throughout the plant. As the bacteria multiply, they may

eventually block the vascular tissue, causing wilts. Other symptoms resulting from

Xanthomonas infection include leaf, stem, and fruit necrosis and spots, cankers, and die-back

diseases (16).

XCZ primarily produces leaf spots on zinnia. The leaf spots are small, angular brown

lesions (1-4 mm), often surrounded by chlorotic halos (1, 16). As the leaf spots increase in

size, they may eventually coalesce. Water-soaking symptoms may also occur.

91

XCZ has been shown to be an excellent detoxifier of the fungal toxin cercosporin,

which is produced by Cercospora spp. (13). Out of 32 XCZ isolates collected from various

geographical areas, all 32 were able to degrade cercosporin. One isolate in particular, XCZ-

3, demonstrated a rapid degradation ability, with degradation beginning as early as 12 hours

after cercosporin exposure and leading to complete loss of the compound by 60 hours (13).

The ability of XCZ-3 to degrade cercosporin requires a gene that codes for a

putative oxidoreductase enzyme (GenBank Accession #DQ087176). Five out of five non-

degrading mutants recovered from mutagenesis experiments had point mutations in this

oxidoreductase gene (Taylor, Mitchell, and Daub, unpublished). Complementation of

mutants with an intact gene restored cercosporin-degrading activity.

It is unknown why an oxidoreductase in XCZ would be involved in cercosporin

degradation. It has previously been shown that XCZ cannot utilize cercosporin as a carbon

source (13). In addition, it is unlikely that cercosporin is the natural substrate for this enzyme

since zinnia does not produce cercosporin. Although Cercospora zinniae Ellis & G. Martin

is a reported pathogen of zinnia (7, 8), it does not appear to be a widespread or severe

problem and there is no indication that the two pathogens routinely co-exist together in the

same plant. Also, the ability to degrade cercosporin provides only a minimal benefit to the

bacterium when exposed to light (13).

We hypothesized that zinnia contains a chemical that has structural similarities to

cercosporin and that must be altered by the XCZ oxidoreductase in order for the bacterium to

use zinnia as a host. We chose to address this hypothesis by assaying for oxidoreductase

regulation in the presence of zinnia extract. We also tested for zinnia pathogenicity of XCZ-

92

3 oxidoreductase mutants, as well as by other bacteria that were found to have

oxidoreductase homologues.

93

Materials and Methods: Bacterial Cultures: The XCZ-3 wild-type strain (13) and XCZ-3 mutants were maintained on

Luria-Bertani (LB) medium (10 g of tryptone, 10 g of NaCl, and 5 g of yeast extract per liter;

pH 7.2) containing rifampicin at 100 μg/mL. The XCZ-3 mutants were obtained by EMS

mutagenesis, and all contain point mutations in the same oxidoreductase gene required for

cercosporin degradation (Taylor, Mitchell, and Daub, unpublished). Isolates of Xanthomonas

arboricola (pseudonym campestris) pv. pruni (14, 17) strains XAP-76, XAP-50 and XAP-75

were obtained from Dr. David Ritchie (North Carolina State University) and were maintained

on LB medium without antibiotics. Isolates of Xanthomonas axonopodis (pseudonym

campestris) pv. vesicatoria (14, 17) (XAV-79 and XAV-126), also obtained from David

Ritchie, were maintained on LB containing 200 μg/mL copper-sulfate. Isolate Pseudomonas

syringae pv. tomato (PST) DC3000 was obtained from Dr. Peter Lindgren (North Carolina

State University) and was maintained on LB containing rifampicin (as above). All of the

bacterial isolates were maintained at 28 ºC in the dark.

Plant Extract: Zinnia elegans and Nicotiana tabacum cv ‘Burley 21’ were grown in the

greenhouse. Leaves were extracted by adding 10 mL sterile dH2O for every 1 g of leaf tissue

and thoroughly homogenizing in a blender. The mixtures were filtered through sterile

cheesecloth and then centrifuged four times at 10000 g for 20 minutes at 4 °C, transferring

the supernatant to fresh centrifuge tubes after each spin. The extracts were filtered using

0.45 µm filter systems from Nalge Nunc International (Rochester, NY) and stored at –20 °C

until ready for use.

94

Quantitative RT-PCR of Oxidoreductase Expression in Wild-Type XCZ-3: XCZ-3 was

grown in LB liquid medium, LB medium containing 50% zinnia extract, or LB medium

containing 50% tobacco extract. RNA was isolated at 24 hours using the SV Total RNA

Isolation kit from Promega Corporation (Madison, WI). A total of 1 μg RNA for each

sample was used to make cDNA using Applied Biosystems (Foster City, CA) TaqMan®

Reverse Transcription Reagents kit. Primers for the oxidoreductase gene were designed

using Primer Express (Applied Biosystems, Foster City, CA) software and acquired from

Sigma Genosys (The Woodlands, TX). The primers F1 (5’-TGCGCTACGAACGGATCA-

3’) and R1 (5’-GGCACGAGCTTGGGAATGT-3’) were used. As an internal control, the

internal transcribed sequence (ITS) that resides between the 16S and 23S rRNAs was

amplified using primers F-ITS (5’-GTTCCCGGGCCTTGTACACAC-3’) and R-ITS (5’-

GGTTCTTTTCACCTTTCCCTC-3’) (9). Quantitative RT-PCR was carried out in the MJ

Research DNA Engine Opticon® 2 thermal cycler using 2X SYBR Green Master Mix

(Applied Biosystems, Foster City, CA) with the following conditions: 95 °C for 10 min.,

followed by 40 cycles of 95 °C for 15 sec., 60 °C for 1 min., 72 °C for 20 sec. The sample

plate was read after every cycle. Analysis of the quantitative RT-PCR results was carried out

according to Livak, et al. (11).

Zinnia Inoculation: The bacterial isolates XCZ-3, XAP-76, MutA, MutB, MutC, MutD,

MutE, XAP-50, XAP-75, XAV-79, XAV-126 and PST DC3000 were used for zinnia

inoculation (Fig. 1). The ability to degrade cercosporin was assayed on medium containing

the red-colored cercosporin. Degrading isolates are light-colored and produce a cleared halo

surrounding the colony; non-degrading isolates lack a halo and become dark in coloration

95

due to uptake of cercosporin. XCZ-3 is the cercosporin-degrading wild-type. Isolates MutA,

MutB, MutC, MutD, and MutE each contain a point mutation in the XCZ-3 oxidoreductase

gene and are not able to degrade cercosporin. Isolates XAP-76, XAV-79 and XAV-126 lack

a homologue to the oxidoreductase and are unable to degrade cercosporin. Isolates XAP-50

and XAP-75 are cercosporin-degrading isolates that contain a homologue to the

oxidoreductase gene. PST DC3000 also contains an oxidoreductase homologue, but is

unable to degrade cercosporin.

Overnight bacterial cultures were inoculated into antibiotic-free LB medium and

allowed to grow approximately four additional hours. The concentrations of the cultures

were determined by measuring absorbance at 600 nm using a Beckman DU 650

spectrophotometer. Cultures were then diluted using LB medium to obtain 106 and 108

cells/mL. Zinnia elegans that were approximately five weeks old were used for inoculations,

with a total of three zinnia plants inoculated for each concentration of all bacterial isolates.

Three to four leaves were inoculated per plant by pipetting 100 μL of each dilution onto the

leaf and rubbing the liquid over the adaxial and abaxial surfaces with a clean cotton swab.

The liquid on the leaves was allowed to dry and the plants were then covered with clear

plastic bags in order to maintain a humid environment. After three days, the bags were

removed and the plants were monitored for disease symptoms.

96

Results: Presence of Plant Extract Upregulates Oxidoreductase Gene Expression: Quantitative RT-

PCR was used to determine if gene expression of the wild-type oxidoreductase was regulated

by the presence of plant extracts. Results showed that transcript accumulation increases after

24 hours of exposure to both extracts from zinnia, the host plant, and tobacco, a non-host

(Fig. 2). Accumulation of the oxidoreductase transcript was increased 3.7-fold after

exposure to zinnia extract and 5.6-fold after exposure to tobacco extract when compared to

an untreated LB control set at one. Therefore, the oxidoreductase gene is responsive to

compounds in plants, but this response is not specific to the host plant zinnia.

Presence of the Oxidoreductase Not Correlated with Pathogenicity: Zinnia plants were

inoculated with the wild-type XCZ-3, the oxidoreductase mutants MutA, MutB, MutC,

MutD, and MutE, the cercosporin non-degrading isolates XAP-76, XAV-79, XAV-126 and

PST DC3000, and the cercosporin-degrading isolates XAP-50 and XAP-75 (Fig. 1).

Inoculation with all of the XCZ-3 oxidoreductase mutants yielded the bacterial leafspot

disease, with symptom severity that was equal to or greater than that of the wild-type XCZ-3

at both inoculum concentrations tested (Fig. 3). Inoculation with LB alone was used as the

negative control and did not produce disease on zinnia (image not shown).

Plants were also inoculated with other bacteria that are not pathogens of zinnia to

determine if the presence of the oxidoreductase may allow for some limited infection. These

included the cercosporin non-degrading isolates XAP-76, XAV-79 and XAV-126 that lack

an oxidoreductase homologue, isolates XAP-50 and XAP-75, which are cercosporin-

degrading isolates that contain an oxidoreductase homologue, as well as PST DC3000, a non-

97

degrader that does contain an oxidoreductase homologue. None of these isolates produced

any symptoms when inoculated onto zinnia (data not shown).

98

Discussion:

Bacteria are able to degrade a wide array of toxic compounds (2-5, 10, 12). In

previous efforts to identify enzymes that can degrade the fungal toxin cercosporin, Mitchell,

et al. tested 244 different bacterial isolates and identified 43 with cercosporin-degrading

activity (13). This activity was scattered among many different species, genera, and

pathovars. However, Xanthomonas campestris pv. zinniae (XCZ) was surprising in that all

32 isolates tested were able to degrade cercosporin. Since there is no known reason why

cercosporin-degrading activity is so ubiquitous in this pathovar, we hypothesized that zinnia

contains a defense compound similar to cercosporin that must be degraded for XCZ to be

pathogenic.

In previous work, we demonstrated that cercosporin degradation by XCZ-3 required a

gene encoding a putative oxidoreductase. Five XCZ-3 mutants that were unable to degrade

cercosporin had point mutations in the oxidoreductase open reading frame (Taylor, Mitchell,

Daub, unpublished). Complementation of these mutants with the wild-type gene restored

cercosporin-degrading activity. Further, with only one exception, degrading activity of other

bacteria could be correlated with the presence of an oxidoreductase homologue, based on

Southern analysis (Taylor, Mitchell, Daub, unpublished). Although the gene is necessary for

degradation, we have shown that it is not sufficient because expression in non-degrading

bacterial isolates did not impart cercosporin degrading activity (Taylor, Mitchell, Daub,

unpublished).

In order to test our hypothesis that the cercosporin-degrading oxidoreductase is

important in zinnia pathogenesis, we first examined regulation of the oxidoreductase in the

presence of plant extract using quantitative RT-PCR (Fig. 2). When XCZ-3, the cercosporin-

99

degrading wild-type isolate, was exposed to zinnia extracts, oxidoreductase transcript

increased 3.73-fold when compared to an LB control. However, transcript also increased

5.59-fold in response to tobacco extracts. Therefore, regulation was not specific to zinnia.

Our results suggest that plants contain a compound that induces expression of the

oxidoreductase and thus this gene could be important in pathogenesis. However, there is no

evidence for a specific inducing compound in zinnia.

To further test our hypothesis, we investigated zinnia pathogenicity of the XCZ-3

oxidoreductase mutants as well as other bacteria that degrade cercosporin and/or have a

homologue of the oxidoreductase gene. Contrary to our hypothesis, all of the XCZ-3 mutants

caused disease equivalent to that of the XCZ-3 wild-type (Fig. 3). Further, bacterial

pathogens of other plants were unable to infect zinnia even if they had an oxidoreductase

homologue or had cercosporin-degrading activity. Therefore, neither the presence of the

oxidoreductase or its homologues, nor cercosporin-degrading activity correlate with zinnia

pathogenicity.

To date, the function of the XCZ-3 oxidoreductase in pathogenicity remains

unknown. Upregulation by plant extracts suggests a possible role in the interaction of the

bacteria with plants. In order to fully understand the purpose of the oxidoreductase, future

work will need to focus on identifying the plant compound(s) that induce its upregulation.

100

References Cited:

1. Alippi, A. M. 1985. Bacterial leaf spot of Zinnia elegans, a new disease in Argentina.

Rev. Argen. Microbiol. 17:11-13.

2. Arnold, M., A. Reittu, A. von Wright, P. J. Martikainen, and M.-L. Suihko.

1997. Bacterial degradation of styrene in waste gases using a peat filter. Appl.

Microbiol. Biotech. 48:738-744.

3. Beller, H. R., A. M. Spormann, P. K. Sharma, J. R. Cole, and M. Reinhard.

1996. Isolation and characterization of a novel toluene-degrading, sulfate-reducing

bacterium. Appl. Environ. Microbiol. 62:1188-1196.

4. Cheng, T., S. P. Harvey, and G. L. Chen. 1996. Cloning and expression of a gene

encoding a bacterial enzyme for decontamination of organophosphorus nerve agents

and nucleotide sequence of the enzyme. Appl. Environ. Microbiol. 62:1636-1641.

5. Dagley, S. 1971. Catabolism of aromatic compounds by microorganisms. Adv.

Microb. Physiol. 6:1-46.

6. Dye, D. W., J. F. Bradbury, M. Goto, A. C. Hayward, R. A. Lelliot, and M. N.

Schroth. 1980. International standards for naming pathovars of phytopathogenic

bacteria and a list of pathovar names and pathotype strains. Rev. Plant Pathol.

59:153-168.

7. Ellis, J. B., and B. M. Everhart. 1885. Enumeration of the North American

Cercosporae. Journal of Mycology 1:17-24.

8. Farr, D. F., G. F. Bills, G. P. Chamuris, and A. Y. Rossman. 1989. Fungi on plants

and plant products in the United States. APS Press. The American Phytopathological

Society. St. Paul, Minnesota USA.

101

9. Gonçalves, E. R., and Y. B. Rosato. 2002. Phylogenetic analysis of Xanthomonas

species based upon 16S-23S rDNA intergenic spacer sequences. Int. J. Sys. Evol.

Microbiol. 52:355-361.

10. Hamann, C., J. Hegemann, and A. Hildebrandt. 1999. Detection of polycyclic

aromatic hydrocarbon degradation genes in different soil bacteria by polymerase

chain reaction and DNA hybridization. FEMS Microbiol. Letters 173:255-263.

11. Livak, K. J., and T. D. Schmittgen. 2001. Analysis of relative gene expression data

using real-time quantitative PCR and the 2 -Delta Delta C(T) Method. Methods 25:402-

408.

12. McGhee, I., and R. G. Burns. 1995. Biodegradation of 2,4-dichlorophenoxyacetic

acid (2,4-D) and 2-methyl-4-chlorophenoxyacetic acid (MPCA) in contaminated soil.

Appl. Soil Ecol. 2:143-154.

13. Mitchell, T. D., W. S. Chilton, and M. E. Daub. 2002. Biodegradation of the

polyketide toxin cercosporin. Appl. Environ. Microbiol. 68:4173-4181.

14. Rademaker, J. L. W., F. J. Louws, M. H. Schultz, U. Rossbach, L. Vauterin, J.

Swings, and F. J. de Bruijn. 2005. A comprehensive species to strain taxonomic

framework for Xanthomonas. Phytopathology 95:1098-1111.

15. Sleesman, J., D. G. White, and C. W. Ellett. 1973. Bacterial leaf spot of zinnia: a

new disease in North America. Plant Dis. Reptr. 57:555-557.

16. Swings, J. G., and E. L. Civerolo. 1993. Xanthomonas. London: Chapman and Hall.

17. Vauterin, L., B. Hoste, K. Kersters, and J. Swings. 1995. Reclassification of

Xanthomonas. Int. J. Sys. Bacteriol. 45:472-489.

102

Figure 1. Bacterial isolates used to inoculate zinnia. Cercosporin-degrading isolates XCZ-3,

XAP-50 and XAP-75 show a light coloration and the presence of a cleared halo when grown

on medium containing the red-colored cercosporin. Cercosporin non-degrading isolates

XAP-76, XAV-79, XAV-126, and PST DC3000, and the non-degrading XCZ-3 mutants

MutA, MutB, MutC, MutD and MutE show a lack of a cleared halo as well as a dark

coloration due to uptake of cercosporin.

103

Figure 2. Quantitative RT-PCR analysis of expression of the oxidoreductase gene in XCZ-3

following exposure of the bacterium to plant extracts. Numbers above bars are fold-increase

in expression as compared to the untreated control, which was normalized to 1.

1.00

3.73

5.59

0

1

2

3

4

5

6

24 hrs

Fold

Cha

nge

untreatedLB + zinnia extractLB + tobacco extract

104

A B

Figure 3. Zinnia leaves inoculated with the XCZ-3 wild-type and the cercosporin non-

degrading mutants MutA, MutB, MutC, MutD, and MutE. Plants were inoculated with (A)

106 cells/mL and (B) 108 cells/mL. Pictures were taken 20 days (A) and 16 days (B) after

inoculation. All of the mutants produced normal disease symptoms on zinnia.

105

Appendix – Search for proteins differentially expressed in XCZ-3 upon exposure to cercosporin.

Goal:

Cercosporin is a non-specific toxin produced by most Cercospora spp. Xanthomonas

campestris pv. zinniae, strain 3 (XCZ-3) has been shown to be an excellent degrader of

cercosporin (2). Using mutagenesis, we carried out a search for genes that are involved in

cercosporin degradation and found that a putative oxidoreductase is involved in the process.

Although we have shown that the XCZ-3 oxidoreductase is involved in cercosporin degradation,

it is not sufficient. This finding suggests that additional proteins play a role in degradation. In

this study, we attempted to identify additional proteins involved in XCZ-3 cercosporin

degradation by looking for proteins that were differentially expressed in the presence of

cercosporin.

Methods:

XCZ-3 was grown in LB medium with 50 μM cercosporin, or in LB with 0.5 % acetone

(used to solubilize cercosporin) as a control, for 24 hours. Bacterial cells were harvested by

centrifugation at 3400 rpm for 20 minutes. Three different methods were used to isolate protein

from the isolated cells.

For the first method (Set 1), cells were resuspended in 1 mL distilled water and sonicated

at 30 % power four times for 20 second intervals while on ice. The mixture was centrifuged and

the supernatant was separated from the pellet. The pellet was resuspended in 200 μL distilled

water. Both the resuspended pellet and the harvested supernatant were stored at –80 °C until

ready to use.

106

The second method (Set 2) followed the whole cell protein extraction protocol of Tahara,

et al. (3). Again, the pellet was separated from the supernatant and both were stored at –80 °C

until ready to use.

The third method (Set 3) was based on work by both Tahara, et al. (3) and Mot, et al. (1).

Bacterial cells were washed with phosphate buffer (30 mM Na2PO4, 16 mM KH2PO4, 137 mM

NaCl) (3) and harvested by centrifugation. Cells were then resuspended in extraction buffer (0.7

M sucrose, 0.5 M Tris, 30 mM HCl, 50 mM EDTA, 0.1 M KCl, 40 mM Dithiothreitol) and

incubated for 15 minutes at room temperature (1). Next, 750 μL buffer-saturated phenol (Gibco

BRL, Gaithersburg, MD) was added and the mixture was incubated for 15 minutes at room

temperature with shaking. The mixture was centrifuged in order to recover the phenol. This step

was followed by two rounds of the addition of an equal volume of extraction buffer, incubation

for 15 minutes at room temperature, and centrifugation. The phenol layer was recovered and

five volumes of cold ammonium acetate solution (100 mM ammonium acetate in methanol) was

added in order to precipitate the proteins (1). This mixture was incubated overnight at –20 °C,

centrifuged, and the supernatant was removed. The pellet was washed with cold ammonium

acetate solution twice, and again centrifuged with removal of the supernatant. The pellet was

washed with cold 80 % acetone, harvested by centrifugation and allowed to air dry. The pellet

was resuspended in 75 μL distilled water and stored at –80 °C until ready to use.

For each sample, 500 μg protein was boiled for three minutes and loaded onto a precast

polyacrylamide Ready Gel® from Bio-Rad (Hercules, CA). After running the gel, it was placed

in Coomassie gel stain (1.25 g Coomassie Blue, 50 mL acetic acid, 225 mL methanol, 225 mL

distilled water; filtered) overnight at room temperature with shaking. The gel was de-stained (50

mL acetic acid, 225 mL methanol, 225 mL distilled water; filtered) and set up to dry.

107

Results:

We hypothesized that other proteins, in addition to the XCZ-3 oxidoreductase, are

involved in the XCZ-3 degradation of cercosporin. We decided to test this hypothesis by looking

for proteins that may be increased after exposure to cercosporin. To do so, we investigated

banding patterns of total protein isolated from untreated XCZ-3 and XCZ-3 that has been

exposed to cercosporin (Fig. 1). Even though three different methods of protein extraction were

utilized, we were unable to detect any differences in banding patterns when the proteins were

analyzed by a one-dimensional polyacrylamide gel.

Conclusion:

We were unable to identify differences in protein banding patterns that might represent a

specific response to the presence of cercosporin (Fig. 1). However, this result may be due to the

low sensitivity of the assay. Although we have previously shown that presence of the XCZ-3

oxidoreductase gene transcript is upregulated when exposed to cercosporin, we were unable to

identify a protein band representing the oxidoreductase. Therefore, it was unlikely that we

would be able to identify additional proteins using this method. We know that the XCZ-3

oxidoreductase is not sufficient for cercosporin degradation, which suggests the involvement of

additional proteins. Using a more sensitive method, such as a 2-D gel, may reveal additional

proteins involved in this process.

108

References Cited: 1. de Mot, R., and J. Vanderleyden. 1989. Application of two-dimensional protein

analysis for strain fingerprinting and mutant analysis of Azospirillum species. Can. J.

Microbiol. 35:960-967.

2. Mitchell, T. D., W. S. Chilton, and M. E. Daub. 2002. Biodegradation of the polyketide

toxin cercosporin. Appl. Environ. Microbiol. 68:4173-4181.

3. Tahara, S. T., A. Mehta, and Y. B. Rosato. 2003. Proteins induced by Xanthomonas

axonopodis pv. passiflorae with leaf extract of the host plant (Passiflorae edulis).

Proteomics 3:95-102.

109

Figure 1. One-dimensional polyacrylamide gels displaying the banding patterns of XCZ-3 exposed to cercosporin or to 0.5 % acetone

as a control. Protein was extracted by three different methods, but the banding patterns of XCZ-3 exposed to cercosporin (CE) do not

visibly differ from those of the acetone controls (ctrl) within each set. Set 1, Set 2, and Set 3 refer to the three methods, respectively,

described in the Methods.

110

Appendix - Xanthomonas axonopodis pv. citri does not degrade cercosporin

Goal:

Xanthomonas campestris pv. zinniae (XCZ) has been shown to rapidly degrade the

fungal toxin cercosporin (2). We have previously shown that XCZ, strain 3 contains a gene for a

putative oxidoreductase that is involved in cercosporin degradation (GenBank Accession

#DQ087176). Sequencing of the region surrounding this gene has revealed a nearby gene for a

transcriptional regulator that is only 61 bp upstream from and transcribed in the same direction

as the oxidoreductase (GenBank Accession #DQ087176). Translated gene homology searches

using the Basic Local Alignment Search Tool (BLAST), version 2.2.11 have shown that the

closest matches to the XCZ-3 transcriptional regulator and oxidoreductase genes are found in

Xanthomonas axonopodis pv. citri (XAC), strain 306 (GenBank Accession #AAM36534, E = 2e

–114 and Accession #AAM36535, E = 1e-37, respectively). The program Clustal W 1.82 (EMBL-

EBI, Heidelberg, Germany) was used to establish amino acid alignments between the

transcriptional regulator and oxidoreductase translated genes from both XCZ-3 and XAC-306

(Fig. 1 and 2). As with XCZ-3, in XAC-306, the transcriptional regulator and oxidoreductase are

transcribed in the same direction and are only 30 bp apart. This observation suggests that in both

XCZ-3 and XAC-306 these two genes exist as an operon. This finding led us to question

whether or not the high similarity between the XAC-306 and XCZ-3 putative operons may result

in XAC also having the ability to degrade cercosporin. In this work, we show that XAC is

unable to degrade cercosporin.

111

Methods:

XAC can only be studied in a quarantined facility, so studies using this organism were

carried out at the approved facility of the Beltsville Agricultural Research Center, USDA, ARS

in Beltsville, MD by Tina Gouin, a researcher at this facility. The cercosporin-degrading isolate

XCZ-3 and the cercosporin non-degrading isolate XAP-76, as well as nutrient agar medium

plates containing 50 μM cercosporin (NACE), were sent to Tina Gouin. Isolate xc62 was the

only XAC isolate available at the facility and was used for the assay. It was incubated on the

NACE medium plates for 12 days in the dark and then analyzed for a cleared halo in the

surrounding medium.

Results:

The published genome of XAC-306 reveals a sequence that is highly homologous to the

region of XCZ-3 that spans the putative operon containing the transcriptional regulator and

oxidoreductase (Fig. 1 and 2) (1). Previous work in our lab has shown that the XCZ-3

oxidoreductase is involved in cercosporin degradation. Due to the high homology, we

hypothesized that XAC may also be able to degrade cercosporin. XAC isolate xc62 was tested

for cercosporin degradation by growing it on cercosporin-containing medium and assaying for

colony color and the presence of a cleared halo surrounding the colonies as compared to the

positive (XCZ-3) and negative (XAP-76) controls. In this assay, all colonies tested had small

halos, however, XAC-xc62 most closely resembled the negative control XAP-76 (Fig. 3). In

both cases, the colonies turned purple due to absorption of cercosporin. The small halo formed is

likely due to absorption of cercosporin from the medium and not to degradation

112

Conclusion:

Despite the high similarity that exists between the XCZ-3 and XAC-306 putative

operons, the XAC isolate that was tested (xc62) was unable to degrade cercosporin (Fig. 3). This

is perhaps due to the needed presence of an additional gene(s) not available in XAC.

Alternatively, the XAC-306 oxidoreductase, although strongly homologous to that of XCZ-3,

may differ in substrate specificity or regulation of expression in response to cercosporin. It is

also possible that the inability of XAC-xc62 to degrade cercosporin may be due to an absence of

the putative operon. The XAC isolate that was used in the cercosporin degradation test (xc62) is

not the same isolate that was sequenced (strain 306). It is possible that the putative operon does

not exist in all isolates of XAC. Therefore, we cannot make any strong conclusions based on the

data.

113

References Cited: 1. da Silva, A. C. R., J. A. Ferro, F. C. Relnach, C. S. Farah, L. R. Furlan, R. B.

Quaggio, C. B. Monteiro-Vitorello, M. A. Van Sluys, N. F. Almeida, L. M. Alves, A.

M. do Amaral, M. C. Bertolini, L. E. Camargo, G. Camarotte, F. Cannavan, J.

Cardozo, F. Chambergo, L. P. Ciapina, R. M. Cicarelli, L. L. Coutinho, J. R.

Cursino-Santos, H. El-Dorry, J. B. Faria, A. J. Ferreira, R. C. Ferreira, M. I. Ferro,

E. F. Formighieri, M. C. Franco, C. C. Greggio, A. Gruber, A. M. Katsuyama, L. T.

Kishi, R. P. Leite, E. G. Lemos, M. V. Lemos, E. C. Locali, M. A. Machado, A. M.

Madeira, N. M. Martinez-Rossi, E. C. Martins, J. Meidanis, C. F. Menck, C. Y.

Miyaki, D. H. Moon, L. M. Moreira, M. T. Novo, V. K. Okura, M. C. Oliveira, V. R.

Oliveira, H. A. Pereira, A. Rossi, J. A. Sena, C. Silva, R. F. de Souza, L. A. Spinola,

M. A. Takita, R. E. Tamura, E. C. Teixeira, R. I. Tezza, M. Trindade dos Santos, D.

Truffi, S. M. Tsai, F. F. White, J. C. Setubal, and J. P. Kitajima. 2002. Comparison of

the genomes of two Xanthomonas pathogens with differing host specificities. Nature

417:459-463.

2. Mitchell, T. D., W. S. Chilton, and M. E. Daub. 2002. Biodegradation of the polyketide

toxin cercosporin. Appl. Environ. Microbiol. 68:4173-4181.

114

Figure 1. Alignment of amino acid sequences of oxidoreductase genes from Xanthomonas

campestris pv. zinniae (XCZ-3) and Xanthomonas axonopodis pv. citri (XAC-306).

* = identical nucleotides;

: = conserved substitutions;

. = semi-conserved substitutions

XCZ3oxido ---MSRRILITGASVAGTTAAWWLDAYGVEVEVVERAPAFRDGGQNVDVRGSARDVLRRM 57 XACoxido VTAMPRRILITGASIAGNTAAWTLAHRGFDVTVAERATRFRDGGQNIDVRGVGREVLQRM 60 *.*********:**.**** * *.:* *.***. *******:**** .*:**:** XCZ3oxido GLEARAFERSTRELGTDWVDADDRVIARFKADASDTDSGPTADLEIRRGDLARMLYEASR 117 XACoxido GLERAALAQGTGEEGTAWIDAHGQAVATFKTEDVDGD-GPTAELEILRGDLARLLYDAAR 119 *** *: :.* * ** *:**..:.:* **:: * * ****:*** ******:**:*:* XCZ3oxido ERVAYRFGDSVCGVAQDQAG--VEITFHSGRCARYDAVIVAEGVGSHTREQVFPGENVPR 175 XACoxido SHVAYRFGDRIASIEDAAGSDAATVTFESGCSERFDAVIVAEGVGSSTREQVFPGENDPR 179 .:******* :..: : .. . :**.** . *:*********** ********** ** XCZ3oxido WMDLTLAYFSIPRQSHDSAYARQYNTVGGRGATLKPALDGKLGAYLGIQKKPGGENAWTP 235 XACoxido WMDLTIAYFTIPRSADDDRLWRWYHTTGGRSISLRPDRHGTTRAMLSLQKPPEGEQDWSV 239 *****:***:***.:.*. * *:*.***. :*:* .*. * *.:** * **: *: XCZ3oxido ERQRRFIETQFANDGWEFPRILAAMRDVDDFYFDVLRQVHMPRWSAGRVVLTGDAAWCPT 295 XACoxido DAQKAYLHERFADAGWQAARVLDGMHGTDDFYFDALRQVRMQRWHSGRTVLTGDAAWCAT 299 : *: ::. :**: **: .*:* .*:..******.****:* ** :**.*********.* XCZ3oxido SLSGIGTTLALVGSYVLAGELAQAVSPMHACMRYERIMRPFVKEGQNIPKLVPRLLWPHS 355 XACoxido PLAGIGATLAVTGAYVLANEIAQADTLDAAFAAYATAMRPMVEQAQGVPKIGPRLMNPHS 359 .*:***:***:.*:****.*:*** : * * ***:*::.*.:**: ***: *** XCZ3oxido AAGLTLLRGAMRVAGMPIVRRLVTDGFARDSNRIALPDYRPIATL 400 XACoxido RLGIHLLHGALKLASQPSVQNLAAKWMTPAIKAPDLSRYP----- 399 *: **:**:::*. * *:.*.:. :: : *. *

115

Figure 2. Alignment of amino acid sequences of transcriptional regulator genes from

Xanthomonas campestris pv. zinniae (XCZ-3) and Xanthomonas axonopodis pv. citri (XAC-

306).

* = identical nucleotides;

: = conserved substitutions;

. = semi-conserved substitutions

XCZ3reg VGYASRMTSDIHPEAAIAPGYLANHAARVFNRLVDARLREHGLTLALIGPLLLLSWKGPM 60 XACreg ---MLDQNELIHPQAAVAPGYLANHAARVFNRLVDAQLRPHGVSLALIGPIMLLSWKGPM 57 .. ***:**:*******************:** **::******::******** XCZ3reg LQRDLVRESAVKQPAMAALLDKLEAMALIAREASSTDKRAATVRLTAAGQDAAATGRTVL 120 XACreg LQRDLVIASAVKQPAMVALLDKLEGLKLIKRTPTPQDRRAALVALTARGRKIAELGGRAL 117 ****** ********.*******.: ** * .:. *:*** * *** *:. * * .* XCZ3reg LDVNALGVSGFSGKDAGLLTALLGRLIANLESAAGE- 156 XACreg LGMNAVALEGFSTDEVQQTVALMHRLIANMERHGQDV 154 *.:**:.:.*** .:. .**: *****:* . :

116

Figure 3. Xanthomonas axonopodis pv. citri isolate xc62 does not degrade cercosporin when

grown on cercosporin-containing medium. The positive control (XCZ-3) appears at the top of

the plate and the negative control (XAP-76) appears at the bottom of the plate. Although a small

halo is seen, the colonies of the non-degrading isolates turn purple due to absorption of

cercosporin from the medium.

117

Appendix 3 - Transformation of tobacco with the Xanthomonas campestris pv. zinniae

oxidoreductase gene involved in cercosporin degradation.

Goal:

Most Cercospora spp. produce the fungal toxin cercosporin, which plays a key role in

plant pathogenesis. The bacterium Xanthomonas campestris pv. zinniae (XCZ) has been shown

to rapidly degrade this toxin, possibly through the action of a cytochrome P450 (5). Previous

work in our lab has revealed that degradation of cercosporin by XCZ-3, a wild-type isolate,

requires the presence of a gene encoding an oxidoreductase. However, expression of the

oxidoreductase gene in cercosporin non-degrading bacteria has proven to be insufficient for

cercosporin degradation, suggesting a role for additional molecules. In general, cytochrome

P450s require the action of a reductase molecule in order to activate the P450. We hypothesize

that the XCZ-3 oxidoreductase may function as either the cytochrome P450 or as its reductase.

It is likely that the partner molecule is absent in the bacteria tested due to an absence of P450

systems in many bacteria.

Plants, however, are known to have numerous P450 systems (2, 7). Therefore, if

additional molecules are necessary to yield cercosporin degradation they may already be present

in plants due to the existing complexity of these enzyme systems. In order to test this

hypothesis, we decided to transform tobacco with the XCZ-3 oxidoreductase and assay for

cercosporin degradation. The XCZ-3 oxidoreductase was cloned into an Agrobacterium binary

vector and tobacco leaf disks were transformed. Calli were regenerated from the leaf disks and

screened for the ability to degrade cercosporin. Plants were also regenerated from the

transformed leaf disks in order to collect seed.

118

Methods: Cloning of Genes: The oxidoreductase gene was amplified from the wild-type bacterial isolate

XCZ-3 using Gateway®-compatible primers GF8kb1 (5’-

GGGGACAAGTTTGTACAAAAAAGCAGGCTTCCTGCTCGATGTCAATGC-3’) and

GR8kb1 (5’-

GGGGACCACTTTGTACAAGAAAGCTGGGTGAAGTTCTAGGGGCGACCAG-3’).

Gateway® Technology from Invitrogen (Carlsbad, CA) was used to move the oxidoreductase

gene into the kanamycin-resistance Gateway®-compatible binary vector pK2GW7 (4), yielding

pKoxido. The Agrobacterium tumefaciens strain EHA105 was made competent and then

transformed with pKoxido by the freeze-thaw method established by An, et al. (1). The

transformed oxidoreductase gene was sequenced to ensure accuracy. pK2GW7 alone could not

be used as a vector control because it contains the CmR-ccdB gene, which will result in cell

death when expressed. However, when a gene is cloned into the pK2GW7 multiple cloning site,

the CmR-ccdB gene is disrupted and no longer harmful. Therefore, the kanamycin-resistance

vector pBI121 (Clontech, Mountain View, CA) was transformed into the EHA105 cell line for

use as a vector control.

Plant Tissue Transformation: Tobacco leaf disks were transformed using Agrobacterium as

previously described by Horsch, et al. (3). Burley 21 tobacco leaves were sterilized in 70%

EtOH for 30 seconds, followed by five minutes in 10 % bleach, and three washes of sterile,

distilled water. Leaves were dried on sterile paper towels. Leaf disks were excised with a #5 (10

mm) cork borer from the sterilized leaves. Overnight bacterial cultures of EHA105 alone,

EHA105 transformed with pKoxido, and EHA105 transformed with the pBI121 vector alone

119

were diluted to 5 x 107 cells/mL in liquid MSBN medium (Murashige and Skoog [MS] medium

containing benzyladenine [BA] at 1 mg/L and naphthalene acetic acid [NAA] at 0.1 mg/L) (6).

Tobacco leaf disks were exposed to the bacterial cultures for 15 seconds with swirling, then

removed and slightly dried on sterile paper towels. For an untransformed control, leaf disks

were exposed to liquid MSBN medium without bacteria. The treated leaf disks were placed on

solid MSBN medium, adaxial side down, and incubated at 25 °C with 12 hours of light exposure

daily. After three days, untransformed leaf disks were transferred to fresh MSBN medium

without antibiotics and transformed leaf disks were transferred to MSBN medium containing 500

mg/L carbenicillin and 100 mg/L kanamycin. Disks were transferred to fresh medium every two

weeks until shoots formed.

Recovery of Plants and Callus: Once shoots formed, they were excised from the leaf tissue and

moved to MS medium without hormones (MSRT) containing 500 mg/L carbenicillin and 100

mg/L kanamycin for rooting. Untransformed shoots were treated in the same manner but placed

on MSRT without antibiotics. Shoots were transferred to fresh medium every two weeks until

roots formed. Rooted shoots were then transferred to soil and moved to the greenhouse. A layer

of cheesecloth was placed over the plants for the first three days while plants adjusted to the

change of environment. Plants were maintained in the greenhouse until seeds were produced and

collected.

In addition, a leaf was removed from each of the untransformed shoots upon transfer to

rooting medium and placed on 3/0.3 medium (MS medium lacking pyridoxine and nicotinic acid,

but containing 3 mg/L indoleacetic acid [IAA] and 0.3 mg/L kinetin) for callus formation.

Leaves from transformed shoots were treated in the same manner but placed on 3/0.3 medium

120

containing 500 mg/L carbenicillin and 100 mg/L kanamycin. Calli were transferred to fresh

medium every two weeks.

Screening of Callus: Calli were tested for cercosporin degradation by growing them on 3/0.3

medium with 50 μM cercosporin. They were incubated in the dark at 25 °C for 13 days and

assayed for a cleared halo representing cercosporin degradation.

Results:

Tobacco leaf disks were transformed with the XCZ-3 oxidoreductase gene in order to

determine if expression of this gene could yield resistance to the fungal toxin cercosporin. A

total of 18 transformed plants expressing this gene were recovered. Tobacco was also

transformed with a vector alone control, from which a total of 29 plants were recovered. As an

additional control, 49 tobacco plants that were submitted to the same treatment conditions, but

not transformed, were also recovered.

Callus tissue was generated from each of the recovered plants and used to screen for

cercosporin degradation. Calli were placed on callus growth medium containing cercosporin and

incubated in the dark. The screening plates were assayed for cleared halos surrounding the calli

after 13 days (Fig 1) and 21 days (Table 1). No halos were visible around any of the calli. The

calli transformed with the XCZ-3 oxidoreductase showed no differences when compared to the

calli transformed with the vector alone or the untransformed calli.

121

Conclusion:

Plants were regenerated from tobacco leaf disks transformed with the XCZ-3

oxidoreductase and with vector alone, as well as from untransformed tobacco leaf disks. Seed is

being harvested from these plants to screen for resistance to cercosporin and Cercospora

infection.

Callus tissue was also induced from each of the recovered plants and used to assay for

cercosporin degradation in culture. However, we were unable to detect a cleared halo, indicating

that tobacco callus expressing the XCZ-3 oxidoreductase could not degrade cercosporin (Table 1

and Fig. 1). In addition, the oxidoreductase-expressing calli showed no difference in phenotype

when compared with the untransformed and vector controls.

We know from previous work in our lab that expression of the XCZ-3 oxidoreductase

gene in cercosporin non-degrading bacteria is not sufficient to yield cercosporin degradation,

which suggests the involvement of additional proteins. Despite the many complicated P450

systems and their component molecules that exist in plants, the oxidoreductase was still unable

to yield cercosporin degradation in tobacco callus tissue. It is likely that the XCZ-3

oxidoreductase requires an additional molecule that is specific to XCZ. Further research should

focus on identifying this molecule. Then, perhaps, transgenic plants that are resistant to

Cercospora infection can be generated.

122

References Cited 1. An, G., P. R. Ebert, A. Mitra, and S. B. Ha. 1988. Binary vectors. Plant Molecular

Biology Manual, © Kluwer Academic Publishers, Dordrecht - Printed in Belgium: A3: 1-

19.

2. Chapple, C. 1998. Molecular-genetic analysis of plant cytochrome P450-dependent

monooxygenases. Annu. Rev. Plant Physiol. Plant Mol. Biol. 49:311-343.

3. Horsch, R. B., J. E. Fry, N. L. Hofmann, D. Eichholtz, S. G. Rogers, and R. T.

Fraley. 1985. A simple and general method for transferring genes into plants. Science

227:1229-1231.

4. Karimi, M., D. Inzé, and A. Depicker. 2002. GATEWAYTM vectors for

Agrobacterium-mediated plant transformation. Trends Plant Sci. 7:193-195.

5. Mitchell, T. D., W. S. Chilton, and M. E. Daub. 2002. Biodegradation of the polyketide

toxin cercosporin. Appl. Environ. Microbiol. 68:4173-4181.

6. Murashige, T., and F. Skoog. 1962. A revised medium for rapid growth and bioassays

with tobacco tissue cultures. Physiol. Plant. 15:473-494.

7. Schuler, M. A. 1996. Plant cytochrome P450 monooxygenases. Crit. Rev. Plant Sci.

15:235-284.

123

Table 1. Callus tissue transformed with the oxidoreductase gene was unable to degrade

cercosporin after 21 days incubation on cercosporin-containing medium.

Plant

#

# Callus pieces

Cercosporin degradation?

Untransformed 2-2 5 no 2-3 10 no 2-4 5 no 2-7 5 no 2-8 5 no 2-9 5 no 3-4 5 no 3-7 5 no 3-8 5 no 3-10 10 no 4-6 5 no 4-8 5 no

pBI121 2-2 10 no 2-3 5 no 2-6 5 no 2-10 5 no 3-6 10 no 4-1 5 no 4-2 5 no 4-4 5 no 4-5 5 no 4-9 5 no

pKoxido 3-1 3 no 3-2 3 no 3-5 5 no 3-6 7 no 3-8 7 no 3-9 7 no 3-10 10 no 3-11 10 no 3-12 8 no 3-13 10 no 3-14 10 no 3-16 5 no 3-b 10 no 3-d 5 no 4-1 5 no 4-2 5 no 4-a 5 no 4-b 5 no

124

Figure 1. Tobacco callus on cercosporin-containing medium. pKoxido calli have been

transformed with the pK2GW7 vector containing the XCZ-3 oxidoreductase gene. After 13 days

on callus medium containing cercosporin, pKoxido shows no cleared halo and no differences

when compared to the negative control pBI121 (transformed with vector alone) and the

untransformed calli.

pKoxido

untransformed pBI121