Xanthomonas campestris - NCSU
Transcript of Xanthomonas campestris - NCSU
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
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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
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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
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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
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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
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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
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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).
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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.
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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
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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
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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
1. Alderman, S. C., and M. K. Beute. 1986. Influence of temperature and moisture on
germination and germ tube elongation of Cercospora arachidicola. Phytopathology
76:715-719.
2. Assante, G., R. Locci, L. Camarda, L. Merlini, and G. Nasini. 1977. Screening of
the genus Cercospora for secondary metabolites. Phytochem. 16:243-247.
3. Balis, C., and M. G. Payne. 1971. Triglycerides and cercosporin from Cercospora
beticola: fungal growth and cercosporin production. Phytopathology 61:1477-1484.
4. Batchvarova, R., V. Nikolaeva, S. Slavov, S. Bossolova, V. Valkov, S.
Atanassova, S. Guelemerov, A. Atanossov, and H. Anzai. 1998. Transgenic
tobacco cultivars resistant to Pseudomonas syringae pv. tabaci. Theory Appl. Genet.
97:986-989.
5. Batchvarova, R. B., V. S. Reddy, and J. Bennett. 1992. Cellular resistance in rice
to cercosporin, a toxin of Cercospora. Phytopathology 82:642-646.
6. Beckman, P. M., and G. A. Payne. 1982. External growth, penetration, and
development of Cercospora zeae-maydis in corn leaves. Phytopathology 72:810-815.
7. Black, S. D., and M. J. Coon. 1987. P-450 cytochromes: structure and function.
Adv. Enzymol. Relat. Areas Mol. Biol. 60:35-87.
8. Callahan, T., M. Rose, M. Meade, M. Ehrenshaft, and R. Upchurch. 1999. CFP,
the putative cercosporin transporter of Cercospora kikuchii, is required for wild type
cercosporin production, resistance, and virulence on soybean. MPMI 12:901-910.
9. Calpouzos, L. 1966. Action of oil in control of plant disease. Annu. Rev. Phytopath.
4:369-386.
30
10. Calpouzos, L., and G. F. Stalknecht. 1967. Symptoms of Cercospora leaf spot of
sugar beets influenced by light intensity. Phytopathology 57:799-800.
11. Cavallini, L., A. Bindoli, F. Macrì, and A. Vianello. 1979. Lipid peroxidation
induced by cercosporin as a possible determinant of its toxicity. Chemico-biol.
Interact. 28:139-146.
12. Choquer, M., K. L. Dekkers, H. Chen, L. Cao, P. P. Ueng, M. E. Daub, and K. R.
Chung. 2005. The CTB1 gene encoding a fungal polyketide synthase is required for
cercosporin biosynthesis and fungal virulence of Cercospora nicotianae. MPMI
18:468-476.
13. Chou, W., and T. M. Kutchan. 1998. Enzymatic oxidations in the biosynthesis of
complex alkaloids. Plant J. 15:289-300.
14. Chung, K. R., M. E. Daub, K. Kuchler, and C. Schüller. 2003. The CRG1 gene
required for resistance to the singlet oxygen-generating cercosporin toxin in
Cercospora nicotianae encodes a putative fungal transcription factor. Biochem.
Biophys. Res. Comm. 302:302-310.
15. Chung, K. R., M. Ehrenshaft, D. K. Wetzel, and M. E. Daub. 2003. Cercosporin-
deficient mutants by plasmid tagging in the asexual fungus Cercospora nicotianae.
Mol. Genet. Gen. 270:103-113.
16. Chung, K. R., A. E. Jenns, M. Ehrenshaft, and M. E. Daub. 1999. A novel gene
required for cercosporin toxin resistance in the fungus Cercospora nicotianae. Mol.
Gen. Genet. 262:382-389.
17. Chupp, C. 1953. A monograph of the fungus genus Cercospora. Ithaca, NY:9-17.
31
18. Coates, S. T., and D. G. White. 1998. Inheritance of resistance to gray leaf spot in
crosses involving selected resistant inbred lines of corn. Phytopathology 88:872-982.
19. Colrat, S., C. Deswarte, A. Latché, A. Klaébé, M. Bouzayen, J. Fallot, and J.
Roustan. 1999. Enzymatic detoxification of eutypine, a toxin from Eutypa lata, by
Vitis vinifera cells: partial purification of an NADPH-dependent aldehyde reductase.
Planta 207:544-550.
20. Daub, M. E. 1982. Cercosporin, a photosensitizing toxin from Cercospora species.
Phytopathology 72:370-374.
21. Daub, M. E. 1982. Peroxidation of tobacco membrane lipids by the photosensitizing
toxin, cercosporin. Plant Physiol. 69:1361-1364.
22. Daub, M. E., and S. P. Briggs. 1983. Changes in tobacco cell membrane
composition and structure caused by the fungal toxin, cercosporin. Plant Physiol.
71:763-766.
23. Daub, M. E., and M. Ehrenshaft. 2000. The photoactivated Cercospora toxin
cercosporin: contributions to plant disease and fundamental biology. Annu. Rev.
Phytopath. 38:437-466.
24. Daub, M. E., and R. P. Hangarter. 1983. Light-induced production of singlet
oxygen and superoxide by the fungal toxin, cercosporin. Plant Physiol. 73:855-857.
25. Daub, M. E., G. B. Leisman, R. A. Clark, and E. F. Bowden. 1992. Reductive
detoxification as a mechanism of fungal resistance to singlet oxygen-generating
photosensitizers. Proc. Natl. Acad. Sci. USA 89:9588-9592.
32
26. Daub, M. E., M. Li, P. Bilski, and C. F. Chignell. 2000. Dihydrocercosporin singlet
oxygen production and subcellular localization: a possible defense against
cercosporin phototoxicity in Cercospora. Photochem. Photobiol. 71:135-140.
27. Daub, M. E., and G. A. Payne. 1989. The role of carotenoids in resistance of fungi
to cercosporin. Phytopathology 79:180-185.
28. de la Fuente-Martínez, J. M., G. Mosqueda-Cano, A. Alvarez-Morales, and L.
Herrera-Estrella. 1992. Expression of a bacterial phaseolotoxin-resistant ornithyl
transcarbamylase in transgenic tobacco confers resistance to Pseudomonas syringae
pv. phaseolicola. Bio/tech. 10:905-909.
29. Ding, Y., S. Li, X. Li, F. Sun, J. Liu, N. Zhao, and Z. Rao. 2003. Site-directed
mutagenesis and preliminary x-ray crystallographic studies of the tabtoxin resistance
protein. Protein Pept. Lett. 10:255-263.
30. Duvick, J. 1987. Detection of cercosporin in grey leaf spot-infected maize leaf tissue.
Phytopathology (abstr.) 77:1754.
31. Ehrenshaft, M., P. Bilski, M. Li, C. F. Chignell, and M. E. Daub. 1999. A highly
conserved sequence is a novel gene involved in de novo vitamin B6 biosynthesis.
Proc. Natl. Acad. Sci. USA 96:9374-9378.
32. Ehrenshaft, M., K. R. Chung, A. E. Jenns, and M. E. Daub. 1999. Functional
characterization of SOR1, a gene required for resistance to photosensitizing toxins in
the fungus Cercospora nicotianae. Curr. Genet. 34:478-485.
33. Ehrenshaft, M., and M. E. Daub. 2001. Isolation of PDX2, a second novel gene in
the pyridoxine biosynthesis pathway of eukaryotes, archaebacteria, and a subset of
eubacteria. J. Bacteriol. 183:3383-3390.
33
34. Ehrenshaft, M., A. E. Jenns, K. R. Chung, and M. E. Daub. 1998. SOR1, a gene
required for photosensitizer and singlet oxygen resistance in Cercospora fungi is
highly conserved in divergent organisms. Molec. Cell 1:603-609.
35. Ehrenshaft, M., A. E. Jenns, and M. E. Daub. 1995. Targeted gene disruption of
carotenoid biosynthesis in Cercospora nicotianae reveals no role for carotenoids in
photosensitizer resistance. MPMI 8:569-575.
36. Ehrenshaft, M., and R. G. Upchurch. 1991. Isolation of light-enhanced cDNAs of
Cercospora kikuchii. Appl. Environ. Microbiol. 57:2671-2676.
37. Estabrook, R. W., D. Y. Cooper, and O. Rosenthal. 1963. The light reversible
carbon monoxide inhibition of the steroid C21-hydroxylase system of the adrenal
cortex. Biochem. Zeit. 338:741-755.
38. Estabrook, R. W., M. R. Franklin, B. Cohen, A. Shigamatzu, and A. G.
Hildebrandt. 1971. Biochemical and genetic factors influencing drug metabolism.
Influence of hepatic microsomal mixed function oxidation reactions on cellular
metabolic control. Metabolism 20:187-199.
39. Estabrook, R. W., M. S. Shet, C. W. Fisher, C. M. Jenkins, and M. R.
Waterman. 1996. The interaction of NADPH-P450 reductase with P450: an
electrochemical study of the role of the flavin mononucleotide-binding domain. Arch.
Biochem. Biophys. 333:308-315.
40. Eyer, C. S., and W. L. Backes. 1992. Relationship between the rate of reductase-
cytochrome P450 complex function and the rate of first electron transfer. Arch.
Biochem. Biophys. 293:231-240.
34
41. Fajola, A. O. 1978. Cercosporin, a phytotoxin from Cercospora spp. Phys. Plant
Path. 13:157-164.
42. Fajola, A. O., and S. O. Alasoadura. 1973. Chemical control of the frog-eye disease
(Cercospora nicotianae) of tobacco (Nicotiana tabacum) in Nigeria. Ann. Appl. Biol.
74:219-224.
43. Fore, S. A., M. E. Daub, and M. K. Beute. 1988. Phytotoxic substances produced
by some isolates of Cercospora arachidicola are not cercosporin. Phytopathology
78:1082-1086.
44. Gonzalez, F. J. 1989. The molecular biology of cytochrome P450s. Pharmacological
Rev. 40:243-288.
45. Goodwin, S. B., L. D. Dunkle, and V. L. Zismann. 2001. Phylogenetic analysis of
Cercospora and Mycosphaerella based on the internal transcribed spacer region of
ribosomal DNA. Phytopathology 91:648-658.
46. Gordon, S. G., M. Bartsch, I. Matthies, H. O. Gevers, P. E. Lipps, and R. C.
Pratt. 2004. Linkage of molecular markers to Cercospora zeae-maydis resistance in
maize. Crop Sci. 44:628-636.
47. Guillén, P., M. Guis, G. Martínez-Reina, S. Colrat, S. Dalmayrac, C. Deswarte,
M. Bouzayen, J. Roustan, J. Fallot, J. Pech, and A. Latché. 1998. A novel
NADPH-dependent aldehyde reductase gene from Vigna radiata confers resistance to
the grapevine fungal toxin eutypine. Plant. J. 16:335-343.
48. Gwinn, K. D., D. A. Stelzig, and J. L. Brooks. 1987. Effects of corn plant age and
cultivar on resistance to Cercospora zeae-maydis and sensitivity to cercosporin. Plant
Dis. 71:603-606.
35
49. Halkier, B. A. 1996. Catalytic reactivities and structure/function relationships of
cytochrome P450 enzymes. Phytochem. 43:1-21.
50. Hartman, P. E., W. J. Dixon, T. A. Dahl, and M. E. Daub. 1988. Multiple modes
of photodynamic action by cercosporin. Photochem. Photobiol. 47:699-703.
51. Higa, A., M. Kimura, K. Mimori, T. Ochiai-Fukuda, T. Tokai, N. Takahashi-
Ando, T. Nishiuchi, T. Igawa, M. Fujimura, H. Hamamoto, R. Usami, and I.
Yamaguchi. 2003. Expression in cereal plants of genes that inactivate Fusarium
mycotoxins. Biosci. Biotechnol. Biochem. 67:914-918.
52. Jenns, A. E., and M. E. Daub. 1995. Characterization of mutants of Cercospora
nicotianae sensitive to the toxin cercosporin. Phytopathology 85:906-912.
53. Jenns, A. E., M. E. Daub, and R. G. Upchurch. 1989. Regulation of cercosporin
accumulation in culture by medium and temperature manipulation. Phytopathology
79:213-219.
54. Jenns, A. E., D. L. Scott, E. F. Bowden, and M. E. Daub. 1995. Isolation of
mutants of the fungus Cercospora nicotianae altered in their response to singlet-
oxygen-generating photosensitiziers. Photochem. Photobiol. 61:488-493.
55. Kim, J. H., N. Mahoney, K. L. Chan, R. J. Molyneux, and B. C. Campbell. 2004.
Secondary metabolites of the grapevine pathogen Eutypa lata inhibit mitochondrial
respiration, based on a model bioassay using the yeast Saccharomyces cerevisiae.
Curr. Microbiol. 49:282-287.
56. Kimura, M., I. Kaneko, M. Komiyama, A. Takatsuki, H. Koshino, K. Yoneyama,
and I. Yamaguchi. 1998. Trichothecene 3-O-Acetyltransferase protects both the
36
producing organism and transformed yeast from related mycotoxins. J. Biol. Chem.
273:1654-1661.
57. Krinsky, N. I. 1979. Carotenoid protection against oxidation. Pure Appl. Chem.
51:649-660.
58. Kuyama, S., and T. Tamura. 1957. Cercosporin. A pigment of Cercospora kikuchii
Matsumoto et Tomoyasu. I. Cultivation of fungus, isolation and purification of
pigment. J. Am. Chem. Soc. 79:5725-5726.
59. Kuyama, S., and T. Tamura. 1957. Cercosporin. A pigment of Cercospora kikuchii
Matsumoto et Tomoyasu. II. Physical and chemical properties of cercosporin and its
derivatives. J. Am. Chem. Soc. 79:5726-5729.
60. Leisman, G. B., and M. E. Daub. 1992. Singlet oxygen yields, optical properties,
and phototoxicity of reduced derivatives of the photosensitizer cercosporin.
Photochem. Photobiol. 55:373-379.
61. Lipps, P. E. 1998. Gray leaf spot: a global threat to corn production. On-line
publication. APSNet Feature (May), Published by the American Phytopathological
Society, ST. Paul, MN.
62. Lipps, P. E., D. G. White, J. E. Ayers, and L. D. Dunkle. 1998. Gray leaf spot of
corn: update. Technical Report of the North Central Region (NCR-25) Committee on
Corn and Sorghum Diseases. On-line publication. May feature, full report.
63. Lousberg, R. J. J. C., U. Weiss, C. A. Salemink, A. Arnone, L. Merlini, and G.
Nasini. 1971. The structure of cercosporin, a naturally occurring quinone. Chem.
Commun. 22:1463 - 1464.
64. Lucas, G. B. 1991. The compendium of tobacco disease. St. Paul, MN, APS Press.
37
65. Lynch, F. J. 1977. Production of cercosporin by Cercospora species. Trans. Brit.
Mycol. Soc. 69:496-498.
66. Macrì, F., and A. Vianello. 1979. Photodynamic activity of cercosporin on plant
tissues. Plant Cell Environ. 2:267-271.
67. Maher, E. A., N. J. Bate, W. Ni, Y. Elkind, R. A. Dixon, and C. J. Lamb. 1994.
Increased disease susceptibility of transgenic tobacco plants with suppressed levels of
preformed phenylpropanoid products. Proc. Natl. Acad. Sci. USA 91:7802-7806.
68. Menkir, A., and M. A. Adepoju. 2005. Registration of 20 tropical midaltitude maize
line sources with resistance to gray leaf spot. Crop Sci. 45:803-804.
69. Menkir, A., and M. Ayodele. 2005. Genetic analysis of resistance to gray leaf spot
of midaltitude maize inbred lines. Crop Sci. 45:163-170.
70. Mitchell, T. D., F. Alejos-Gonzalez, H. S. Gracz, D. A. Danehower, M. E. Daub,
and W. S. Chilton. 2003. Xanosporic acid, an intermediate in bacterial degradation
of the fungal phototoxin cercosporin. Phytochem. 62:723-732.
71. Mitchell, T. D., W. S. Chilton, and M. E. Daub. 2002. Biodegradation of the
polyketide toxin cercosporin. Appl. Environ. Microbiol. 68:4173-4181.
72. Mulder, J. L., and P. Holliday. 1974. Cercospora nicotianae. CMI Descriptions of
Pathogenic Fungi and Bacteria 416:1-2.
73. Mumma, R. O., F. L. Lukezic, and M. G. Kelly. 1973. Cercosporin from
Cercospora hayii. Phytochem. 12:917-922.
74. Munro, A. W., and J. G. Lindsay. 1996. Bacterial cytochromes P-450. Mol.
Microbiol. 20:1115-1125.
38
75. Nakahara, K., and H. Shoun. 1994. Crystallization and preliminary X-ray
diffraction studies of nitric oxide reductase cytochrome P450nor from Fusarium
oxysporum. J. Mol. Biol. 239:158-159.
76. Nilsson, N.-O., M. Hansen, A. H. Panagopoulos, S. Tuvesson, M. Ehlde, M.
Christiansson, I. M. Rading, M. Rissler, and T. Kraft. 1999. QTL analysis of
Cercospora leaf spot resistance in sugar beet. Plant Breeding 118:327-334.
77. Orth, C. E., and W. Schuh. 1994. Resistance of 17 soybean cultivars to foliar,
latent, and seed infection by Cercospora kikuchii. Plant Dis. 78:661-664.
78. Payne, G. A., and J. K. Waldron. 1983. Overwintering and spore release of
Cercospora zeae-maydis in corn debris in North Carolina. Plant Dis. 67:87-89.
79. Peterson, J. A., R. E. Ebel, D. H. O'Keeffe, T. Matsubara, and R. W. Estabrook.
1976. Temperature dependence of cytochrome P-450 reduction. A model for
NADPH-cytochrome P-450 reductase:cytochrome P-450 interaction. J. Biol. Chem.
251:4010-4016.
80. 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.
81. Roberts, G. A., G. Grogan, A. Greter, S. L. Flitsch, and N. J. Turner. 2002.
Identification of a new class of cytochrome P450 from a Rhodococcus sp. J.
Bacteriol. 184:3898-3908.
82. Robeson, J. R., M. A. F. Jalal, and R. B. Simpson. 1993. Patent #5,262,306 USA.
39
83. Rollins, J. A., M. Ehrenshaft, and R. G. Upchurch. 1993. Effects of light- and
altered-cercosporin phenotypes on gene expression in Cercospora kikuchii. Can. J.
Microbiol. 39:118-124.
84. Rossi, V. 1995. Effect of host resistance in decreasing infection rate of Cercospora
leaf spot epidemics on sugarbeet. Phytopath. medit. 34:149-156.
85. Setiawan, A., G. Koch, S. R. Barnes, and C. Jung. 2000. Mapping quantitative trait
loci (QTLs) for resistance to Cercospora leaf spot disease (Cercospora beticola
Sacc.) in sugar beet (Beta vulgaris L.). Theory Appl. Genet. 100:1176-1182.
86. Sollod, C. C., A. E. Jenns, and M. E. Daub. 1992. Cell surface redox potential as a
mechanism of defense against photosensitizers in fungi. Appl. Environ. Microbiol.
58:444-449.
87. Spikes, J. D. 1989. Photosensitization. In The Science of Photobiology, ed. K.C.
Smith, New York: Plenum:79-110.
88. Steinkamp, M. P., S. S. Martin, L. L. Hoefert, and E. G. Ruppel. 1979.
Ultrastructure of lesions produced by Cercospora beticola in leaves of Beta vulgaris.
Phys. Plant Path. 15:13-26.
89. Steinkamp, M. P., S. S. Martin, L. L. Hoefert, and E. G. Ruppel. 1981.
Ultrastructure of lesions produced in leaves of Beta vulgaris by cercosporin, a toxin
from Cercospora beticola. Phytopathology 71:1272-1281.
90. Tertivanidis, K., C. Goudoula, C. Vasilikiotis, E. Hassiotou, R. Perl-Treves, and
A. Tsaftaris. 2004. Superoxide dismutase transgenes in sugarbeets confer resistance
to oxidative agents and the fungus C. beticola. Transgenic Res. PC1193:1-9.
40
91. Thorold, C. A. 1940. Cultivation of bananas under shade for the control of leaf spot
disease. Trop. Agric. Trin. 17:213-214.
92. Upchurch, R. G., M. Rose, and M. Eweida. 2001. Over-expression of the
cercosporin facilitator protein, CFP, in Cercospora kikuchii up-regulates production
and secretion of cercosporin. FEMS Microbiol. Letters 204:89-93.
93. Upchurch, R. G., M. Rose, M. Eweida, and T. Callahan. 2002. Transgenic
assessment of CFP-mediated cercosporin export and resistance in a cercosporin-
sensitive fungus. Curr. Genet. 41.
94. Upchurch, R. G., D. C. Walker, J. A. Rollins, M. Ehrenshaft, and M. E. Daub.
1991. Mutants of Cercospora kikuchii altered in cercosporin synthesis and
pathogenicity. Appl. Environ. Microbiol. 57:2940-2945.
95. Urlacher, V. B., S. Lutz-Wahl, and R. D. Schmid. 2004. Microbial P450 enzymes
in biotechnology. Appl. Microbiol. Biotech. 64:317-325.
96. Vauterin, L., B. Hoste, K. Kersters, and J. Swings. 1995. Reclassification of
Xanthomonas. Int. J. Sys. Bacteriol. 45:472-489.
97. Velicheti, R. K., and J. B. Sinclair. 1994. Production of cercosporin and
colonization of soybean seed coats by Cercospora kikuchii. Plant Dis. 78:342-346.
98. Venkataramani, K. 1967. Isolation of cercosporin from Cercospora personata.
Phytopath. Z. 58:379-382.
99. Ward, J. M. J., E. L. Stromberg, D. C. Nowell, and J. F. W. Nutter. 1999. Gray
leaf spot: a disease of global importance in maize production. Plant Dis. 83:884-895.
100. Werck-Reichhart, D., S. Bak, and S. Paquette. 2002. Cytochromes P450. The
Arabidopsis Book, eds. C.R. Somerville and E.M. Meyerowitz, American Society of
41
Plant Biologists, Rockville, MD, doi/10.1199/tab.0028,
http://www.aspb.org/publications/arabidopsis/.
101. White, R. E., and M. J. Coon. 1980. Oxygen activation by cytochrome P-450.
Annu. Rev. Biochem. 49:315-356.
102. Willets, A. 1997. Structural studies and synthetic applications of Baeyer-Villiger
monooxygenases. Tibtech 15:55-62.
103. Windels, C. E., H. A. Lamey, D. Hilde, J. Widner, and T. Knudsen. 1998. A
Cercospora leaf spot model for sugar beet. In practice by an industry. Plant Dis.
82:716-726.
104. Yamazaki, S., and T. Ogawa. 1972. The chemistry and stereochemistry of
cercosporin. Agric. Biol. Chem. 1707-1718.
105. Yamazaki, S., A. Okubo, Y. Akiyama, and K. Fuwa. 1975. Cercosporin, a novel
photodynamic pigment isolated from Cercospora kikuchii. Agric. Biol. Chem.
39:287-288.
106. Zhang, L., and R. G. Birch. 1997. The gene for albicidin detoxification from
Pantoea dispersa encodes an esterase and attenuates pathogenicity of Xanthomonas
albilineans to sugarcane. Proc. Natl. Acad. Sci. USA 94:9984-9989.
107. Zhang, L., J. Xu, and R. G. Birch. 1999. Engineered detoxification confers
resistance against a pathogenic bacterium. Nature Biotech. 17:1021-1024.
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
References Cited:
1. Assante, G., R. Locci, L. Camarda, L. Merlini, and G. Nasini. 1977. Screening of
the genus Cercospora for secondary metabolites. Phytochem. 16:243-247.
2. Ausubel, F. M., R. Brent, R. E. Kingston, D. D. Moore, J. G. Seidman, J. A.
Smith, and K. Struhl. 2002. Curr. Prot. Mol. Biol.
3. Balis, C., and M. G. Payne. 1971. Triglycerides and cercosporin from Cercospora
beticola: fungal growth and cercosporin production. Phytopathology 61:1477-1484.
4. Batchvarova, R. B., V. S. Reddy, and J. Bennett. 1992. Cellular resistance in rice
to cercosporin, a toxin of Cercospora. Phytopathology 82:642-646.
5. Buell, C. R., V. Joardar, M. Lindeberg, J. Selengut, I. T. Paulsen, M. L. Gwinn,
R. J. Dodson, R. T. Deboy, A. S. Durkin, J. F. Kolonay, R. Madupu, S.
Daugherty, L. Brinkac, M. J. Beanan, D. H. Haft, W. C. Nelson, T. Davidsen, N.
Zafar, L. Zhou, J. Liu, Q. Yuan, H. Khouri, N. Fedorova, B. Tran, D. Russell, K.
Berry, T. Utterback, S. E. Van Aken, T. V. Feldblyum, M. D'Ascenzo, W. L.
Deng, A. R. Ramos, J. R. Alfano, S. Cartinhour, A. K. Chatterjee, T. P. Delaney,
S. G. Lazarowitz, G. B. Martin, D. J. Schneider, X. Tang, C. L. Bender, O.
White, C. M. Fraser, and A. Collmer. 2003. The complete genome sequence of the
Arabidopsis and tomato pathogen Pseudomonas syringae pv. tomato DC3000. Proc.
Natl. Acad. Sci. USA 100:10181-10186.
6. Calpouzos, L., and G. F. Stalknecht. 1967. Symptoms of Cercospora leaf spot of
sugar beets influenced by light intensity. Phytopathology 57:799-800.
7. Choquer, M., K. L. Dekkers, H. Chen, L. Cao, P. P. Ueng, M. E. Daub, and K. R.
Chung. 2005. The CTB1 gene encoding a fungal polyketide synthase is required for
69
cercosporin biosynthesis and fungal virulence of Cercospora nicotianae. MPMI
18:468-476.
8. Chung, K. R., M. Ehrenshaft, D. K. Wetzel, and M. E. Daub. 2003. Cercosporin-
deficient mutants by plasmid tagging in the asexual fungus Cercospora nicotianae.
Mol. Genet. Gen. 270:103-113.
9. Coates, S. T., and D. G. White. 1994. Sources of resistance to Gray leaf spot of
corn. Plant Dis. 78:1153-1155.
10. 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.
11. Daub, M. E. 1982. Cercosporin, a photosensitizing toxin from Cercospora species.
Phytopathology 72:370-374.
70
12. Daub, M. E. 1982. Peroxidation of tobacco membrane lipids by the photosensitizing
toxin, cercosporin. Plant Physiol. 69:1361-1364.
13. Daub, M. E., and S. P. Briggs. 1983. Changes in tobacco cell membrane
composition and structure caused by the fungal toxin, cercosporin. Plant Physiol.
71:763-766.
14. Daub, M. E., and M. Ehrenshaft. 2000. The photoactivated Cercospora toxin
cercosporin: contributions to plant disease and fundamental biology. Annu. Rev.
Phytopath. 38:437-466.
15. Daub, M. E., and R. P. Hangarter. 1983. Light-induced production of singlet
oxygen and superoxide by the fungal toxin, cercosporin. Plant Physiol. 73:855-857.
16. Ditta, G., S. Stanfield, D. Corbin, and D. R. Helinski. 1980. Broad host range DNA
cloning system for Gram-negative bacteria: construction of a gene bank of Rhizobium
meliloti. Proc. Natl. Acad. Sci. USA 77:7347-7351.
17. Duvick, J. 1987. Detection of cercosporin in grey leaf spot-infected maize leaf tissue.
Phytopathology (abstr.) 77:1754.
18. Fajola, A. O. 1978. Cercosporin, a phytotoxin from Cercospora spp. Phys. Plant
Path. 13:157-164.
19. Feil, H., W. S. Feil, P. Chain, F. Larimer, G. Dibartolo, A. Copeland, A. Lykidis,
S. Trong, M. Nolan, E. Goltsman, J. Thiel, S. Malfatti, J. E. Loper, A. Lapidus,
J. C. Detter, M. Land, P. M. Richardson, N. C. Kyrpides, N. Ivanova, and S. E.
Lindow. 2005. Comparison of the complete genome sequences of Pseudomonas
syringae pv. syringae B728a and pv. tomato DC3000. Proc. Natl. Acad. Sci. USA
102:11064-11069.
71
20. 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.
21. Hartman, P. E., W. J. Dixon, T. A. Dahl, and M. E. Daub. 1988. Multiple modes
of photodynamic action by cercosporin. Photochem. Photobiol. 47:699-703.
22. Higa, A., M. Kimura, K. Mimori, T. Ochiai-Fukuda, T. Tokai, N. Takahashi-
Ando, T. Nishiuchi, T. Igawa, M. Fujimura, H. Hamamoto, R. Usami, and I.
Yamaguchi. 2003. Expression in cereal plants of genes that inactivate Fusarium
mycotoxins. Biosci. Biotechnol. Biochem. 67:914-918.
23. Kuyama, S., and T. Tamura. 1957. Cercosporin. A pigment of Cercospora kikuchii
Matsumoto et Tomoyasu. I. Cultivation of fungus, isolation and purification of
pigment. J. Am. Chem. Soc. 79:5725-5726.
24. Leisman, G. B., and M. E. Daub. 1992. Singlet oxygen yields, optical properties,
and phototoxicity of reduced derivatives of the photosensitizer cercosporin.
Photochem. Photobiol. 55:373-379.
25. 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.
26. Lousberg, R. J. J. C., U. Weiss, C. A. Salemink, A. Arnone, L. Merlini, and G.
Nasini. 1971. The structure of cercosporin, a naturally occurring quinone. Chem.
Commun. 22:1463 - 1464.
27. Louws, F. J., D. W. Fulbright, C. T. Stephens, and F. J. de Bruijn. 1994. Specific
genomic fingerprints of phytopathogenic Xanthomonas and Pseudomonas pathovars
72
and strains generated with repetitive sequences and PCR. Appl. Environ. Microbiol.
60:2286-2295.
28. Lynch, F. J. 1977. Production of cercosporin by Cercospora species. Trans. Brit.
Mycol. Soc. 69:496-498.
29. Mitchell, T. D., F. Alejos-Gonzalez, H. S. Gracz, D. A. Danehower, M. E. Daub,
and W. S. Chilton. 2003. Xanosporic acid, an intermediate in bacterial degradation
of the fungal phototoxin cercosporin. Phytochem. 62:723-732.
30. Mitchell, T. D., W. S. Chilton, and M. E. Daub. 2002. Biodegradation of the
polyketide toxin cercosporin. Appl. Environ. Microbiol. 68:4173-4181.
31. Mumma, R. O., F. L. Lukezic, and M. G. Kelly. 1973. Cercosporin from
Cercospora hayii. Phytochem. 12:917-922.
32. Nakamura, Y., T. Kaneko, S. Sato, M. Mimuro, H. Miyashita, T. Tsuchiya, S.
Sasamoto, A. Watanabe, K. Kawashima, Y. Kishida, C. Kiyokawa, M. Kohara,
M. Matsumoto, A. Matsuno, N. Nakazaki, S. Shimpo, C. Takeuchi, M. Yamada,
and S. Tabata. 2003. Complete genome structure of Gloeobacter violaceus PCC
7421, a cyanobacterium that lacks thylakoids. DNA Res. 10:181-201.
33. Orth, C. E., and W. Schuh. 1994. Resistance of 17 soybean cultivars to foliar,
latent, and seed infection by Cercospora kikuchii. Plant Dis. 78:661-664.
34. 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.
35. Robeson, J. R., M. A. F. Jalal, and R. B. Simpson. 1993. Patent #5,262,306 USA.
73
36. Rossi, V. 1995. Effect of host resistance in decreasing infection rate of Cercospora
leaf spot epidemics on sugarbeet. Phytopath. medit. 34:149-156.
37. Steinkamp, M. P., S. S. Martin, L. L. Hoefert, and E. G. Ruppel. 1979.
Ultrastructure of lesions produced by Cercospora beticola in leaves of Beta vulgaris.
Phys. Plant Path. 15:13-26.
38. Steinkamp, M. P., S. S. Martin, L. L. Hoefert, and E. G. Ruppel. 1981.
Ultrastructure of lesions produced in leaves of Beta vulgaris by cercosporin, a toxin
from Cercospora beticola. Phytopathology 71:1272-1281.
39. Thorne, L., L. Tansey, and T. Pollock. 1987. Clustering of mutations blocking
synthesis of xanthan gum by Xanthomonas campestris. J. Bacteriol. 169:3593-3600.
40. Thorold, C. A. 1940. Cultivation of bananas under shade for the control of leaf spot
disease. Trop. Agric. Trin. 17:213-214.
41. Upchurch, R. G., D. C. Walker, J. A. Rollins, M. Ehrenshaft, and M. E. Daub.
1991. Mutants of Cercospora kikuchii altered in cercosporin synthesis and
pathogenicity. Appl. Environ. Microbiol. 57:2940-2945.
42. Vauterin, L., B. Hoste, K. Kersters, and J. Swings. 1995. Reclassification of
Xanthomonas. Int. J. Sys. Bacteriol. 45:472-489.
43. Venkataramani, K. 1967. Isolation of cercosporin from Cercospora personata.
Phytopath. Z. 58:379-382.
44. Ward, J. M. J., E. L. Stromberg, D. C. Nowell, and J. F. W. Nutter. 1999. Gray
leaf spot: a disease of global importance in maize production. Plant Dis. 83:884-895.
74
45. Windels, C. E., H. A. Lamey, D. Hilde, J. Widner, and T. Knudsen. 1998. A
Cercospora leaf spot model for sugar beet. In practice by an industry. Plant Dis.
82:716-726.
46. Yamazaki, S., and T. Ogawa. 1972. The chemistry and stereochemistry of
cercosporin. Agric. Biol. Chem. 1707-1718.
47. Yamazaki, S., A. Okubo, Y. Akiyama, and K. Fuwa. 1975. Cercosporin, a novel
photodynamic pigment isolated from Cercospora kikuchii. Agric. Biol. Chem.
39:287-288.
48. Zhang, L., J. Xu, and R. G. Birch. 1999. Engineered detoxification confers
resistance against a pathogenic bacterium. Nature Biotech. 17:1021-1024.
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