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Helminthosporium solani: Biology, Occurrence, and Control
By
Chakradhar Mattupalli
A dissertation submitted in partial fulfillment of
the requirements for the degree of
Doctor of Philosophy
(Plant Pathology)
at the
UNIVERSITY OF WISCONSIN-MADISON
2013
Date of final oral examination: 04/29/2013
The dissertation is approved by the following members of the Final Oral Committee:
Amy O. Charkowski, Professor, Plant Pathology
Glen R. Stanosz, Professor, Plant Pathology
Douglas I. Rouse, Professor, Plant Pathology
James P. Kerns, Assistant Professor, Plant Pathology
Shelley H. Jansky, Associate Professor, Horticulture
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ABSTRACT
Potatoes are one of the top five organic vegetables purchased for home consumption in the
United States. Many biotic and abiotic factors limit organic potato production and to meet the
increasing consumer demand, especially from the fresh market, there is a need to produce good
quality blemish-free potatoes. This research shows that silver scurf(caused by Helminthosporium
solani) and black dot (caused by Colletotrichum coccodes) are two prevalent diseases in organic
potato production in Wisconsin, with high incidences across locations and cultivars. Seventy-five
percent and 94% of asymptomatic tubers assessed were positive for H. solani and C. coccodes
respectively. Because no cultivar has yet been identified that is completely resistant to silver
scurf, we performed minituber inoculation assays and identified a diploid interspecific hybrid
C287 that consistently supported little sporulation of H. solani, suggesting it has partial
resistance to silver scurf. Although important, H. solani is an understudied pathogen with poor
genomic information, and herein we report the draft genome of the fungus. The estimated
genome size of H. solani is ~ 35 megabases. In silico genomic analysis suggests that the fungus
possesses a large suite of genes encoding putative cell wall degrading enzymes.
We also explored alternate methods to control fungal pathogens of potato. We performed in
vitro tests with tobacco phylloplanins (Nt-phylloplanins) using an inexpensive microfluidic
approach to test for its inhibitory activity against plant and human fungal pathogens. Spores of
Colletotrichum coccodes and Aspergillus fumigatus treated with Nt-phylloplanins did not
germinate at the concentrations tested at 48 and 30 hours post treatment respectively. Inhibition
of spore germination of the human pathogen A. fumigatus by Nt-phylloplanins also reveals a new
venue for medicinal antifungal drug discovery that has not been explored previously.
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ACKNOWLEDGEMENTS
I am especially thankful to Dr. Amy Charkowski, my advisor, for accepting me into her
laboratory and instilling in me the confidence that led to the completion of this work. Her
enthusiasm, willingness to give her time, and constructive suggestions during the planning and
development of this research work were extraordinary. I am fortunate to work in her lab, which
provided me with various opportunities for conducting research, mentoring undergrads, writing
grant proposals and I am confident that these experiences will help shape my future career. I
would also like to thank my committee, Drs. Glen Stanosz, Doug Rouse, Jim Kerns and Shelley
Jansky for having provided me with valuable insights at critical stages of my research.
I would like to express my great appreciation to Dr. Ryan Shepherd, for introducing me
to phylloplanins and teaching protein-related work. I would like to offer special thanks to Dr.
Nancy Keller for her advice with phylloplanins and genome projects, and her lab group members
Joe Spraker, Erwin Berthier and Jon Palmer for their assistance with Aspergillus assays,
microfluidics and fungal transformations. Jeremy Glasner provided me with very valuable
assistance in sequencing the genome of Helminthosporium solani. I thank my lab group and
many undergrads for creating a friendly environment in the lab. I wish to acknowledge the care
and concern shown by my friends Nancy Koval, Angie Peltier, Bob Rand, Lenice Covert, Ana
Cristina Fulladolsa Palma and Gregorio Sanchez Perez. Karen Lackermann has been the most
encouraging and helpful friend I ever had and I also thank her for proofreading various chapters
of this dissertation.
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I thank my wife Radhika for her steady encouragement and patience, my son Neerajaksh
for his countless and precious smiles. Finally, I am indebted to my dear friend Vambarelli
Laxmikanth Varma for his unconditional support and motivation that enabled me to come to the
United States and pursue a PhD degree in Plant Pathology.
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TABLE OF CONTENTS
Abstract ……………………………………………………………………………….. i
Acknowledgements …………………………………………………………………… ii
List of tables ………………………………………………………………………....... v
List of figures ………………………………………………………………………….. vii
INTRODUCTION ……………………………………………………………………. 1
CHAPTER 1 ………………………………………………………………………….. 33
Evaluating incidence of Helminthosporium solani and Colletotrichum coccodes
on asymptomatic organic potatoes and screening potato lines for resistance to
silver scurf
CHAPTER 2 …………………………………………………………………………… 61
A microfluidic assay for identifying differential responses of plant and human fungal
pathogens to tobacco phylloplanins
CHAPTER 3 ……………………………………………………………………………. 83
A draft genome sequence reveals Helminthosporium solani’s arsenal for cell wall
degradation
SYNTHESIS ……………………………………………………………………………. 102
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LIST OF TABLES
INTRODUCTION
Table 1: Bacterial, fungal, and oomycete pathogens affecting potato tubers ………………. 31
Table 2: Comparison of four potato tuber surface blemish diseases that are common in
Wisconsin …………………………………………………………………………………… 32
CHAPTER 1:
Table 1. Soil type, previous crop, and planting and harvest dates for each sampling
location ……………………………………………………………………………………… 53
Table 2. Percentage of potato tubers asymptomatic for both silver scurf and black dot
after harvest from three locations in Wisconsin …………………………………………….. 54
Table 3. Percentage of asymptomatic tubers positive for Helminthosporium solani
and Colletotrichum coccodes after performing PCR and tuber incubation assays …………. 55
Table 4. Spore production on fourteen potato minituber lines inoculated with
Helminthosporium solani …………………………………………………………………… 56
Table 5. Kappa coefficient values showing agreement between tuber incubation
and PCR assays for detecting Helminthosporium solani and Colletotrichum coccodes
from asymptomatic tubers ……………………………………………………………………57
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CHAPTER 2
Table 1: Comparison of the effects of Nt-phylloplanins on fungal spore germination ……… 77
CHAPTER 3
Table 1: Comparison of the number of cellulose degrading genes among
ascomycete fungi …………………………………………………………………………….. 100
Table 2: Comparison of the number of hemicelluloses degrading genes among
ascomycete fungi …………………………………………………………………………….. 101
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LIST OF FIGURES
CHAPTER 1:
Figure 1. Percentage of asymptomatic tubers positive or negative for Helminthosporium
solani and Colletotrichum coccodes by tuber incubation (T) and PCR (P) assays ………….. 58
Figure 2. Percentage of tubers positive or negative for Helminthosporium solani (Hs) and
Colletotrichum coccodes (Cc) by either tuber incubation or PCR assay for asymptomatic
tubers and by tuber incubation assay for symptomatic tubers ……………………………….. 59
Figure 3. Conidia and conidiophores of Helminthosporium solani (left side) and sclerotia
of Colletotrichum coccodes (right side) observed in close proximity on a potato tuber
periderm under a dissecting microscope …………………………………………………….. 60
CHAPTER 2:
Figure 1: Schematic representation of microfluidic device fabrication process ……………. 78
Figure 2. Inhibition of Colletotrichum coccodes spore germination after treatment
with Nt-phylloplanins in tobacco leaf wash ………………………………………………… 79
Figure 3: Inhibition curve of Colletotrichum coccodes spore germination by
Nt-phylloplanins in tobacco leaf wash ……………………………………………………… 80
Figure 4. Inhibition of Aspergillus fumigatus spore germination after treatment
with Nt-phylloplanins in tobacco leaf wash ………………………………………………… 81
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Figure 5. Transient inhibition of Fusarium sambucinum spore germination by
Nt-phylloplanins in tobacco leaf wash …………………………………………………….... 82
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INTRODUCTION
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Potato (Solanum tuberosum L.) is an annual herbaceous plant belonging to the family
Solanaceae. The crop is well known to plant pathologists in particular due to the Irish potato
famine caused by the late blight disease in the mid 1840’s. In terms of human consumption,
potatoes are the third most important food crop in the world after rice and wheat. Potatoes are an
excellent source of low fat carbohydrates, vitamins B and C, as well as iron, potassium and zinc.
(CIP, 2013). Potatoes are consumed fresh, frozen, dehydrated, canned, or processed into forms
such as potato chips and French fries. In developed countries, up to 60% of potatoes are
consumed in a processed form (Kirkman, 2007). Potatoes are a critical crop in terms of food
security; as an example China, which is the world’s largest consumer of potatoes, intends to meet
50% of its increased food demand in the next 20 years through increased potato production (CIP,
2013).
Potatoes are propagated commercially using “seed” tubers. However, individual cells,
meristems, sprouts, true seeds, and stem cuttings can also be used as propagules (Struik and
Wiersema, 1999). A seed tuber produces sprouts in its eyes, which develop into shoots, and
produce roots from primordial on the sprouts. A potato plant is in fact a cluster of stems,
originating from one seed tuber or seed piece (Struik, 2007). Potatoes have modified under-
ground shoots called stolons, with elongated internodes, rudimentary leaves and hooked tips.
Tubers are the swollen parts of stolons bearing axillary buds and leaves, evident as eyes on the
tuber skin. The life cycle of a potato plant can be described through five growth stages: 1) sprout
development (sprouts develop from tubers and emerge from the soil) 2) vegetative growth
(leaves and stems develop from above-ground nodes, roots and stolons develop from below-
ground nodes) 3) tuber initiation (tubers start forming at stolon tips) 4) tuber bulking (tubers
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enlarge and develop as a strong sink organ), and 5) maturation (tuber growth ceases and aerial
parts start to senesce) (Miller and Hopkins, 2008).
Tubers are the economically significant portion of the potato plant and because they
develop in the soil, they are attacked by myriad pathogens that affect quality and yield. These
pathogens may cause symptoms on the tuber that are either external, internal or both (Table 1).
Fiers et al. (2012) recently reviewed various biotic, abiotic and management factors that
influence the occurrence and development of soil-borne potato pathogens. Understanding how
these pathogens attack the tuber requires some knowledge about the tuber periderm. In the first
part of this review, a general overview of how tubers naturally protect themselves from soil
micro-biota is provided. The second part of the review focuses on four pathogens that cause
potato tuber-surface blemish diseases that are commonly observed in Wisconsin.
Native and Wound Periderm:
The potato tuber surface is protected by a specialized suberin-rich layer called the native
periderm. This layer provides protection against water loss, mechanical damage, and pathogens
(Lulai, 2001). Epidermis exists on the developing tubers for a very short period which is
replaced by the native periderm and lenticels are formed below each stoma (Peterson and Barker,
1979). The native periderm is composed of three layers: phellem, phellogen, and phelloderm.
The phellem is the outermost layer consisting of suberized cells and is often referred to as the
skin of the potato tuber. The phellogen or the cork cambium is the middle layer and gives rise to
phellem and phelloderm. The phelloderm is the innermost layer of the periderm consisting of
parenchyma-like cells. Tubers with immature periderm are fragile and are susceptible to skinning
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and mechanical damage during harvest and handling operations. The reason for this fragile state
is because phellogen cells are meristematic and have thin cell walls, so they loosely hold the
phellem to the phelloderm, allowing easy scuffing from the tuber (Lulai and Freeman, 2001).
This wounding damage is overcome by killing the potato vines about 3 weeks before harvest
which promotes good skin-set (Lulai and Orr, 1993). Histological and immunolabeling studies
by Sabba and Lulai (2004) indicated that the phellogen walls are reinforced by pectin depositions
during native periderm maturation providing resistance to skinning injuries.
Tubers that are cut or wounded lack protection from native periderm and as the wounded
tuber begins to heal, it forms a wound periderm which is very similar in organization to the
native periderm (Sabba and Lulai, 2002). Before the formation of wound periderm, a suberized
closing layer may form. Suberin is a biopolymer composed of polyphenolic and polyaliphatic
domains. The two domains are spatially separate. The polyphenolic domain which is embedded
in the primary cell wall is cross-linked by glycerol to the polyaliphatic domain which is located
between the primary cell wall and the plasma membrane (Bernards, 2002). During wound
healing, differential deposition of these two domains play separate roles in conferring resistance
to bacterial and fungal pathogens. Lulai and Corsini (1998) observed that the polyphenolic
domain deposited within 2 to 3 days of wound healing provided resistance to the bacterial
pathogen Erwinia carotovora subsp. carotovora (causal organism of bacterial soft rot) but not
Fusarium sambucinum (causal organism of fungal dry rot). Resistance to F. sambucinum
developed after 5 to 7 days of wound healing process which corresponded with the completion of
deposition of the polyaliphatic domain. These results suggest that only complete suberin matrix
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(suberin polyphenolic and aliphatic domains) can provide resistance to both fungal and bacterial
infections.
The relative importance of potato diseases that cause external defects has been increasing
as the fresh market becomes a more significant source of potato sales. Fresh market consumers
demand washed potatoes with a good quality and healthy looking external appearance. Here we
will focus on the following four pathogens that cause external tuber defects and are common in
the state of Wisconsin: Rhizoctonia solani, Streptomyces scabies, Helminthosporium solani, and
Colletotrichum coccodes.
Rhizoctonia solani:
The Rhizoctonia disease complex of potatoes consists of two phases: Rhizoctonia canker
(infection of growing plants) and black scurf (infestation of tubers by sclerotia). This disease
complex is present wherever potatoes are grown (Banville and Carling 2001) and is caused by
the fungus, Rhizoctonia solani Kuhn (teleomorph: Thanatephorus cucumeris (Frank) Donk).
Isolates of R. solani are categorized into Anastomosis Groups (AGs) based on hyphal
incompatibility; AG-3 is the most prevalent AG found on potatoes (Tsror, 2010). Symptoms of
Rhizoctonia canker appear as reddish brown to black lesions leading to girdling of sprouts,
stolons and roots. Poor crop stands and delayed emergence occur as the fungus kills the growing
tip of the sprouts. Above ground symptoms may manifest in the form of aerial tubers in the axils
of branches and petioles, leaf rolling, stunting, rosetting and purple pigmentation of leaves
(Banville and Carling, 2001). Hymenia of the sexual stage of the fungus may be observed as a
white to gray powdery mass on the lower stems near the soil surface (Carling et al. 1989). The
other phase of the disease complex, black scurf, occurs on the tubers. The most obvious sign of
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this disease is the presence of flat or raised, irregularly shaped brown to black sclerotia. These
sclerotia are often referred as “the dirt that won’t wash off” as they adhere tightly to the tuber
skin (Banville et al. 1996). On tubers that are harvested before skin set, the pathogen can also
cause dark spots on tubers during storage (Buskila et al. 2011). Yield losses due to R. solani can
be as high as 20 to 30 % ( Banville 1989; Carling et al. 1989).
Disease cycle:
The primary sources of inoculum for this disease complex are mycelia and sclerotia
infested seed tubers and soil. Roots and stolons are infected at any time during the growing
season (Banville and Carling, 2001). After vine-killing and during tuber maturation, volatile
tuber exudates increase the formation of sclerotia on the tubers (Dijst, 1990). Chand and Logan
(1984) found an increase in the number of sclerotia per tuber on tubers harvested 4 weeks after
vine-killing as compared to those that were harvested one week after vine-killing. Incidence and
severity of black scurf have been shown to increase during storage (Chand and Logan, 1984;
Spencer and Fox, 1979).
Host-pathogen interactions:
Histopathological studies revealed that R. solani mycelia invade the tuber periderm
tissues inter-cellularly causing tissues surrounding the hyphae to collapse, arresting further
growth of the pathogen (Schaal, 1939; Chand and Logan, 1984). Hyphae did not penetrate
beyond the phellogen but in some cases extended to the cortical tissues (Chand and Logan, 1984;
Schaal, 1939; Spencer and Fox, 1979). However, when native periderm on tubers was removed
and the exposed areas of the tuber were artificially inoculated with R. solani, a thicker and less
organized closing layer with over suberized cell walls was formed. An increase in the expression
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of suberin biosynthesis potato genes such as StKCS6, CYP86A33, and POP_A was also observed
as a response to R. solani inoculation (Buskila et al. 2011).
Aoki et al. (1963) and Nishimura and Sabaki (1963) identified phytotoxic compounds such as
phenylacetic acid and its hydroxyl derivatives (meta-hydroxyphenylacetic acid and para-
hydroxyphenylacetic acid) produced from the culture filtrates of R. solani. Rioux et al. (2011)
compared putative pathogenesis related genes expressed by R. solani AG1 and AG3 in infected
rice leaves and potato sprouts respectively. They identified 12 genes similar between the two
pathosystems which play key roles in the early processes of infection such as appresorium
formation, cell wall degradation and toxin secretion. However, at present similar information
related to the genes involved in the tuber infection process is lacking.
Streptomyces spp.:
Common scab of potatoes is caused by Gram-positive, filamentous bacteria belonging to
the genus Streptomyces. Most of the 900 known species of Streptomyces are soil-inhabiting
saprophytes, except a few species which are pathogens on plants (Bignell et al. 2010a). Loria et
al. (2006) described various pathogenic species of Streptomyces that cause typical scab
symptoms. However, the most common pathogenic species of Streptomyces that cause common
scab of potato are: Streptomyces scabies (synonym S. scabiei), Streptomyces turgidiscabies and
Streptomyces acidiscabies. S. scabies is the oldest characterized scab pathogen with a worldwide
distribution while the other two Streptomyces species are newly emergent pathogens.
S. acidiscabies causes acid scab whose symptoms are indistinguishable from common scab, but
the pathogen occurs in soils with pH below 5.2 (Lambert and Loria, 1989). S. turgidiscabies
causes distinctly erumpent lesions on potato tubers in Japan (Miyajima et al. 1998). All three
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species are neither tissue nor host specific, causing scab lesions on various root and tuber crops.
They also cause root and shoot stunting and root tissue necrosis of model plants such as tobacco
and Arabidopsis (Bignell et al. 2010a). However, DNA homology studies suggest that these
species are not closely related (Miyajima et al. 1998; Healy and Lambert, 1991). Common scab
symptoms are mostly limited to the tubers, but the pathogen can also infect stolons (Loria,
2001). Initially, the lesions are circular or irregular and tan or brown-colored and enlarge as the
tuber swells. Commercially, infection of the first four to five formed internodes is critical for
scab infection as these early internodes expand more rapidly than the later-formed internodes,
causing more scabby lesions (Lapwood et al. 1970). As the lesions mature, they become rough in
texture, coalesce, grow up to 5 to 10 mm in diameter, and in severe cases cover most of the tuber
surface. When scab lesions are superficial, the disease is called russet scab, and when they are
sunken, it is called pitted scab. Likewise, if the lesions are raised, the disease is called erumpent
scab. Lesion types are influenced by several factors such as the pathogen strain, cultivar, age of
the tuber at the time of infection, and environmental factors (Loria, 2001). Common scab does
not affect tuber yield but unsightly corky lesions impact the marketability of the tubers (van der
Wolf and Boer, 2007).
Disease cycle:
Infected tubers and infested soil are the primary sources of inoculum for common scab
(Loria, 2001). The pathogen gains entry into the tuber through recently formed unsuberized
lenticels (Fellows, 1926). The early tuber development period (2 or 3 weeks after tuberization) is
the critical stage for common scab infection and symptoms develop within seven days after
exposure of the tuber to the pathogen (Lapwood and Hering, 1970; Khatri et al. 2011). Scab
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symptoms are more severe on the first formed internodes of the tuber as compared to the later-
formed internodes, because enlarging tubers are more susceptible to infection (Fellows, 1926;
Lapwood and Hering, 1970). Initially the pathogen grows vegetatively, producing a branched,
tangled network of filamentous cells, but as the tissues affected by scab mature the pathogen
produces reproductive hyphae that transform into spore chains (Jones, 1931).
Host-pathogen interactions:
After S. scabies gains entry into the tuber through immature lenticels, the meristem is
stimulated to form closely packed elongated cells which later collapse and turn brown. Cell
proliferation, expansion, and subsequent browning are attributable to thaxtomin, the toxin
produced by the pathogen (Lawrence et al. 1990). Gradually the meristematic activity ceases and
the last-formed cells suberize to form a wound cork which separates infected cells from the
healthy tissue below. The pathogen can further infect the healthy cells by growing through
incompletely suberized cells, leading to the formation of a second barrier of wound cork. These
barriers are formed as long as the tuber grows actively resulting in severe scab lesions (Jones,
1931). Khatri et al. (2011) studied pathogen-induced changes in the tuber periderm at 10 days
after tuber initiation in a hydroponic or a non-destructive pot culture system. They observed a
significant increase in phellem thickness, number of phellem cell layers and phellem suberin
content, and these changes were greater in the scab-resistant cultivar Russet Burbank than in the
susceptible cultivar Desiree. These results suggest that potato cultivars respond differentially to
scab infection by reinforcing their periderm structure.
Lawrence et al. (1990) were the first to establish the role of a toxin in S. scabies infection.
By artificially inoculating aspetically grown minitubers with either S. scabies or purified extracts
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obtained from scab lesions, they were able to reproduce common scab symptoms. They later
identified the phytotoxic agents as thaxtomin (in honor of Roland Thaxter who first described the
causal agent of common scab) A and B which are produced only by pathogenic Streptomyces
species. Thaxtomins are cyclic dipeptides (2, 5-diketopiperazines) derived from 4-nitro-
tryptophan and phenylalanine. The presence of a 4-nitroindole moiety plays a key role in
thaxtomin phytotoxicity (King and Calhoun, 2009). Inhibition of cellulose biosynthesis has been
identified as the primary mode of action of thaxtomins (Bignell et al. 2010a). So far, 11
members of the thaxtomin family have been characterized, 10 of which are produced by S.
scabies (King and Calhoun, 2009). Similarly, another class of phytotoxins called concanamycins
A and B were shown to be produced by S. scabies, but not by S. acidiscabies. Their role as a
virulence factor is yet to be determined (Natsume et al. 1998).
Another virulence factor that was accidently discovered while screening a cosmid library
is nec1 (Bukhalid and Loria 1997). When this gene was expressed in the non-pathogen
Streptomyces lividans, it caused necrosis of potato tuber disks and produced scab-like symptoms
on immature potato tubers. The nec1 gene is structurally conserved among S. scabies, S.
acidiscabies, and S. turgidiscabies, indicating that this gene may have been horizontally
transferred among these species (Bukhalid et al. 1998; Bukhalid et al. 2002). Although nec1 is
not directly involved in the biosynthesis of thaxtomin A, it is physically linked to thaxtomin A
biosynthesis genes (Bukhalid et al. 1998). Kers et al. (2005) described a large pathogenicity
island (discrete cluster of pathogenicity and virulence genes acquired by horizontal gene transfer)
of 325-660 kb in size conserved among the three pathogenic species of Streptomyces described
earlier. One of the genes identified in this pathogenicity island tomA, encodes a homologue of
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tomatinase enzyme, which can detoxify anti-microbial saponins secreted by plants. Although
Seipke and Loria (2008) showed that tomA is not important for pathogenicity in potatoes,
conservation of this gene among S. scabies, S. acidiscabies, S. turgidiscabies suggests a role in
plant-microbe interactions.
Coronofacic acid is a component of the phytotoxin coronatine, produced by plant
pathogens such as Pseudomonas syringae and Pectobacterium atrosepticum. Bignell et al.
(2010b) performed mutational studies and identified a secondary metabolite cluster that is
predicted to synthesize a compound similar to coronafacic acid. It is interesting to note that this
biosynthetic cluster is present only in S. scabies but not S. acidiscabies or S. turgidiscabies.
Genomic analysis of S. scabies 87-22 revealed the presence of other virulence factors such as
expansin-like proteins, cutinases and auxin biosynthetic genes. These results warrant further
investigations for their role and mechanism of action during host-pathogen interactions (Bignell
et al. 2010a).
Helminthosporium solani:
Silver scurf, caused by the imperfect fungus, Helminthosporium solani Durieu & Mont. is
a potato disease that affects tuber appearance. The disease is known to occur in various parts of
the world (Errampalli et al. 2001; Loria and Secor 2001). Symptoms are observed only on the
tubers and are localized to the periderm. At harvest, diseased tubers have grey lesions on the
periderm that appear silvery when moistened. The silvery appearance of the tubers is attributed
to air pockets that develop due to cellulolytic activity of the pathogen (Heiny and McIntyre,
1983). The outer cell layers of severely diseased tubers may slough off and H. solani infection
increases the permeability of the periderm to water vapor, resulting in tuber shrinkage and
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weight loss (Burke, 1938; Jellis and Taylor, 1974). In severely diseased tubers loss of
pigmentation occurs, especially on red-skinned cultivars reducing their fresh market value (Jellis
and Taylor, 1977). The disease affects the processing market as chips made from severely
diseased tubers have unacceptable black burnt edges (Holley and Kawchuck, 1996). Field studies
indicate that H. solani does not affect emergence and growth of the potato plants or tuber yields
(Mooi, 1968; Read and Hide, 1984; Cunha and Rizzo, 2004). Silver scurf has been gaining
economic importance in the past three decades due to the quality standards demanded by both
processing and fresh market consumers as well as the emergence of H. solani isolates resistant to
the broad-spectrum thiabendazole fungicides that are applied at postharvest by conventional
potato growers (Errampalli et al. 2001).
Disease cycle:
H. solani has a very narrow host range comprising only tuber-bearing Solanum species
and the weed species Solanum elaeagnifolium (Burke, 1938; Kamara and Huguelet, 1972;
Sethuraman et al. 1997; Rodriguez et al. 1995). The disease spreads both in the field and during
storage. Diseased seed tubers (Jellis and Taylor, 1977) and infested soil (Merida and Loria,
1994) act as the primary source of inoculum in the field while diseased seed tubers act as the
initial source of inoculum in storage (Rodriguez et al. 1996). After planting diseased seed tubers,
expansion of lesions occurs rapidly within 4 to 5 weeks so that the entire tuber surface is affected
(Jellis and Taylor, 1977). The mechanism of pathogen transmission from a diseased seed tuber to
its progeny tubers is unknown as the pathogen is not known to infect either stems or stolons
(Burke, 1938). Jellis and Taylor (1977) recovered viable conidia of H. solani from soil
surrounding progeny tubers at 14 weeks after planting and observed that the first infection sites
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on progeny tubers occur at the stolon end of the tubers. They attributed the increase in disease in
the field and during storage to the expansion of lesions on the diseased tubers as well as to the
spread of the pathogen from diseased tubers to healthy tubers. In studying H. solani conidial
production in storage, Rodriguez et al. (1996) found that conidia were produced throughout the
storage season, with production levels ranging from 0 to 12,000 conidia per day at 4oC and from
0 to 24,000 conidia per day at 10oC. H. solani colonized and sporulated in-vitro on senescent leaf
tissue of alfalfa, sorghum, rye, oats, corn and wheat suggesting the pathogen may be able to
survive saprophytically in soil (Merida and Loria, 1994). The pathogen can also survive in dry
soil of potato stores for a period of five months (Carnegie et al. 1996) and conidia may serve as
over-wintering structures for the pathogen (Kamara and Huguelet, 1972).
Host-pathogen interactions:
Histological studies by Hunger and McIntyre (1979) showed that the hyphae of H. solani
are localized to the periderm and that conidiophores arise from beneath the periderm. Fewer cell
layers were observed in infected (4 to 6 layers) compared to healthy (6 to 8 layers) periderm
(Hunger and McIntyre, 1979). Conidial germination occurred at 6 and 16 hours after inoculation
on tubers of Dark Red Norland (Martinez et al. 2004) and Norchip (Heiny and McIntyre, 1983),
respectively. Likewise, penetration by germ tubes into the periderm also differed between these
cultivars. For Dark Red Norland, germ tube penetration occurred 6 to 9 hours after inoculation,
whereas in Norchip germ tube penetration did not occur until five days after inoculation.
Appresoria-like structures and suberin deposits in cortical cell layers were noted by Heiny and
McIntyre (1983), but not by Martinez et al. (2004). Hyphae grow intra- or inter-cellularly and
move from one cell to another by direct cell wall penetration (Martinez et al. 2004). Martinez et
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al. (2004) observed necrosis in both colonized and neighboring cells and suggested that H. solani
may be a necrotroph.
Colletotrichum coccodes:
Colletotrichum coccodes (Wallr.) causes a tuber blemish disease called black dot of
potato, which has a world-wide distribution (Lees and Hilton, 2003). The presence of small black
sclerotia on tubers, roots, and above- and below-ground stems gives the disease its common
name. Diseased tubers may have gray-brown blemishes on the periderm, or less clearly delimited
discolored areas on which sclerotia may or may not be present (Jellis and Taylor, 1974). The
fungus has been shown to cause dark brown, deep sunken lesions following artificial inoculation
of mini-tubers stored at 5-15oC for 5 months (Glais and Andrivon, 2004). Diseased roots have a
stringy appearance when pulled out of the soil due to cortical tissue desiccation. The pathogen
infects the cortical tissues of the stem; after drying, amethyst discoloration on the inside of the
vascular cylinder is observed. Stolons may be attacked at any stage of plant growth and the
lesions cause severing, thus leaving a part of the stolon adhering to the stem end of the tuber
(Dickson, 1926). Diseased foliage bears dark brown to black lesions similar to those caused by
Alternaria solani, but the lesions of C. coccodes lack concentric rings (Johnson and Miliczky,
1993). The disease also causes a reduction in yield, in some cases even with very little symptom
expression (Johnson, 1994; Mohan et al. 1992; Barkdoll and Davis, 1992). Also, C. coccodes is
known to interact with other pathogens like Verticillum dahliae, causing increased disease
symptoms and more extensive fungal colonization in some potato cultivars (Tsror and
Hazanovsky, 2001). Fresh weight losses of 5 to 10 % were observed by Hunger and McIntyre
(1979) when tubers naturally infected with C. coccodes were stored for 18 weeks.
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Disease cycle:
Hosts of C. coccodes include vegetables such as tomato, pepper and squash in the
Solanaceae and Cucurbitaceae families (Dillard, 1992). Isolates of the pathogen collected from
potato were capable of producing typical anthracnose lesions on ripe tomato fruit and vice-versa
(Illman et al. 1959, Chesters and Hornby, 1965). The fungus can also colonize roots of winter
cherry, marrow and symptomless hosts such as chrysanthemum, white mustard, cress, cabbage
and lettuce (Chesters and Hornby, 1965). Also, C. coccodes colonizes the stems of rotation crops
such as yellow mustard, soybean, spring canola, alfalfa, oats, as well as the stems of a number of
weeds, including eastern black night shade, velvetleaf, giant foxtail and Timothy grass (Nitzan et
al. 2006a).
Black dot is a soil-borne and tuber-borne disease. Sclerotia are the over-wintering
structures. Greenhouse studies by Blakeman and Hornby (1966) found 53% of C. coccodes
sclerotia remained viable and no conidia remained viable after burying them in soil for 83 and 3
weeks respectively. Tuber-borne inoculum may be present as sclerotia on the tuber surface or as
hyphae in the vascular bundles (Tsror et al. 1999). Contaminated seed act as an important source
in introducing the disease to non-infested soils (Barkdoll and Davis, 1992). However, once
infested, soil-borne inoculum has a higher disease causing potential than tuber-borne inoculum
(Nitzan et al. 2008). Nitzan et al. (2008) conducted greenhouse studies and observed a nonlinear
trend in disease severity (sclerotial density on the roots and the length of the stem with visible
sclerotia) with increasing soilborne inoculum concentration. Aerial infection of potato foliage
can occur in certain regions where sandstorms and sprinkler irrigation are common. Johnson and
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Miliczky (1993) obtained infections by sandblasting potato foliage followed by inoculation with
a conidial suspension of C. coccodes. In this type of dispersal, blowing sand creates wounds on
the foliage and sprinkler irrigation aids in the dissemination of conidia and sclerotia.
Infection of below and above ground parts occurs early in the growing season (Davis and
Johnson, 2001). But, the pathogen has a latent period evident by restricted colonization in the
plant stems. As the plant enters the tuber bulking phase and starts to senesce, C. coccodes
colonization is activated and sclerotia develop on above and below-ground plant parts (Nitzan et
al. 2006b). However, Ingram and Johnson (2010) did not observe such a colonization pattern on
potato roots. Glais-Varlet et al. (2004) observed an increase in the development of black dot
symptoms upon artificial inoculation of minitubers stored at 5 to 15oC. These results suggest that
the disease may be able to spread on tubers during the storage period.
Host-pathogen interactions:
Not much literature is available for C. coccodes interactions with potato tubers. However,
considerable work was done in tomato on which the fungus causes anthracnose disease. The
extent to which these results can be extrapolated to potato is debatable owing to the differences
in the environment and the plant part affected. Reasons for slow progress on this subject may be
due to the under-estimation of the disease as the symptoms appear late in the cropping season,
typically coinciding with natural senescence. In addition, symptoms on stems and stolons
resemble those of Rhizoctonia canker, symptoms on leaves are similar to Verticillum wilt or
early blight and symptoms on tubers are similar to silver scurf (Davis and Johnson, 2001).
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Conclusion:
Potato growers must manage tuber surface defect diseases, but have few non-pesticide
based methods for this problem. All four diseases discussed in this review are soil- or tuber-
borne and are monocyclic, so control involves reducing the amount of initial inoculum. One way
to achieve this goal is by the use of certified seed potatoes. One challenge is to determine the
permissible level of surface defects in a certified seed tuber, which is especially difficult when
considering latent pathogens like C. coccodes (Glais-Varlet et al. 2004). The percentage of
surface defects that are permissible on certified seed tubers varies among states, with 5 % surface
defects permitted in some states like Wisconsin. However, in chapter 1, we show visual
assessment of tubers at harvest for silver scurf and black dot may be misleading because a very
high percentage of asymptomatic tubers are already infected with H. solani and C. coccodes.
As an alternative, growers could adopt new methods of producing seed tubers such as
seed potato sprout technology. This technology facilitates production of seed tubers by planting
detached sprouts, thus minimizing the risk of soil-borne pathogen movement (Souza-Dias et al.
2008). Reducing soil inoculum using fumigants is an option, but one that is limited due to
environmental concerns. Future efforts should focus on breeding for new cultivars resistant to
multiple diseases, a solution that is eco-friendly and has long-term viability. Although cultivated
potatoes are tetraploid, which complicates breeding efforts, the use of wild Solanum species in
potato breeding programs may be valuable as the progeny tubers following crosses with wild
relatives have long dormancy with good tuber size and set (Jansky and Peloquin 2006). For
instance, by crossing [H551 × S. chacoense] with [US-730 × (S. berthaultii × S. tarijense)], a
diploid interspecific clone C545 was obtained, which exhibited improved resistance to a number
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of important diseases including soft rot, common scab, pitted scab, early dying disease and early
blight (Jansky and Rouse 2003). Further in chapter 1, we show that these diploid inter-specific
clones exhibit partial resistance to other tuber diseases such as silver scurf.
Several cultural and chemical control methods are available for the management of the
tuber blemish diseases discussed here, although their effectiveness is limited (Errampalli et al.
2001; Lees and Hilton, 2003). As an addition to the existing disease management options, we
explored the possibility of utilizing leaf surface proteins that are naturally produced by tobacco
by testing for their inhibitory effects on fungal pathogens such as C. coccodes. The results of this
project are discussed in chapter 2.
This review demonstrates the dearth of information about the molecular basis of host-
pathogen interactions for the fungal pathogens discussed here (Table 2). With the advent of high
throughput sequencing techniques, the cost of obtaining genome data has dropped considerably.
This has led to the description of several fungal genomes and provided insights into the pathogen
biology, survival and infection strategies (Ellwood et al. 2010; de Wit et al. 2012; Islam et al.
2012; Manning et al. 2013). A next step to advance our understanding of fungal diseases of
potato is to sequence the genomes of fungi that infect tuber surfaces and to perform comparative
genomics to study factors involved in host-pathogen interactions. Based on what we know about
fungal pathogenicity, plant cell wall degrading enzymes and fungal toxins are likely candidates
for virulence factors important for infection of potato tubers. Because numerous fungi are present
in soil, but only these three species (H. solani, C. coccodes, and R. solani) commonly infect
potato tuber surfaces in Wisconsin, we anticipate that evidence of convergent evolution will be
found once the molecular determinants of pathogenicity have been identified for these fungi. In
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chapter 3, we address questions related to H. solani’s biology by generating the draft genome of
the fungus.
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87. Struik, P.C. 2007. In: Vreugdenhil, D., Bradshaw, J.E., Gebhardt, C., Govers, F., MacKerron,
D.K.L., Taylor, M.A., and Ross, H.A. Potato biology and biotechnology: Advances and
perspectives. p.219-233. Elsevier publications, Oxford, GB, UK.
88. Struik, P. C., and Wiersema, S. G. 1999. Seed Potato Technology. p. 383. Wageningen Pers,
Wageningen, the Netherlands.
89. Tsror, L., Aharon, M., and Erlich. O. 1999. Survey of bacterial and fungal seedborne diseases
in imported and domestic potato seed tubers. Phytoparasitica 27: 215-226.
90. Tsror, L., and Hazanovsky. M. 2001. Effect of coinoculation by Verticillium dahliae and
Colletotrichum coccodes on disease symptoms and fungal colonization in four potato
cultivars. Plant Pathol. 50: 483-488.
91. Tsror, L. 2010. Biology, epidemiology and management of Rhizoctonia solani on potato. J.
Phytopathol. 158: 649-658.
92. Van der Wolf, J. M., and De Boer, S. H. 2007. In: Vreugdenhil, D., Bradshaw, J.E., Gebhardt,
C., Govers, F., MacKerron, D.K.L., Taylor, M.A., and Ross, H.A. Potato biology and
biotechnology: Advances and perspectives. p.605. Elsevier publications, Oxford, GB, UK.
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Table 1: Bacterial, fungal and oomycete pathogens affecting potato tubers
Disease Causal agent Diagnosis
Bacterial ring rot Clavibacter michiganensis subsp. sepedonicus External and internal
Brown rot Ralstonia solanacearum External and internal
Soft rot Pectobacterium spp., Clostridium spp. External and internal
Common scab Streptomyces spp. External
Black dot Colletotrichum coccodes External
Black pit Alternaria alternata External and internal
Charcoal rot Macrophomina phaseolina External and internal
Early blight Alternaria solani External and internal
Fusarium dry rot Fusarium spp. External and internal
Gangrene Phoma foveata External and internal
Gray mold Botrytis cinerea External and internal
Late blight Phytophthora infestans External and internal
Leak Pythium spp. External and internal
Pink rot Phytophthora erythroseptica External and internal
Powdery scab Spongospora subterranea f. sp. subterranea External
Black scurf Rhizoctonia solani External
Rosellinia black rot Rosellinia spp. External and internal
Silver scurf Helminthosporium solani External
Skin spot Polyscytalum pustulans External
Stem rot Sclerotium rolfsii External and internal
Thecaphora smut Angiosorus solani External and internal
Verticillium wilt Verticillium spp. Internal
Wart Synchytrium endobioticum External
White mold Sclerotinia sclerotiorum External and internal
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Table 2: Comparison of four potato tuber surface blemish diseases that are common in
Wisconsin
Common Scab Black Scurf Black Dot Silver Scurf
Causal agent Streptomyces spp. Rhizoctonia
solani
Colletotrichum
coccodes
Helminthosporium
solani
Source of inoculum Infected tubers,
Infested soil
Infected tubers,
Infested soil
Infected tubers,
Infested soil
Infected tubers,
Infested soil
Primary inoculum Spores Mycelia and
sclerotia
Conidia Conidia
Invasion Filamentous cell
growth restricted
by formation of
wound cork
Hyphae grow
intercellularly
and localized to
the periderm, but
may sometimes
extend to cortex
Systemic and
hyphae colonize
cortical and
vascular bundle
cells
Hyphae grow
intra- and inter-
cellularly and
localized to the
periderm
Symptom/sign
development
Tuber Tuber Tuber, stolons,
roots, above-
ground stems
Tuber
Tuber
symptoms/signs
Corky lesions that
may be raised or
pitted or
superficial
Irregularly
shaped brown to
black sclerotia
adhering to the
periderm
Grey-brown
blemishes that
are less clearly
delimited and
with/without
sclerotia
Grey lesions on
the periderm that
appear silvery
when moistened
Overwintering
structures
Spores Mycelia and
sclerotia
Sclerotia Conidia
Active disease
spread in storage
No No No Yes
Pathogenicity and
virulence factors
during tuber-
pathogen
interactions
Thaxtomin, nec1,
concanamycins
Not known Not known Not known
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CHAPTER 1
Evaluating incidence of Helminthosporium solani and Colletotrichum coccodes on
asymptomatic organic potatoes and screening potato lines for resistance to silver scurf
This chapter has been published by the American Journal of Potato Research:
Mattupalli, C., Genger, R. K., and Charkowski, A. O. 2013. Evaluating incidence of
Helminthosporium solani and Colletotrichum coccodes on asymptomatic organic potatoes and
screening potato lines for resistance to silver scurf. American Journal of Potato Research doi:
10.1007/s12230-013-9314-3.
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Abstract
Silver scurf, caused by Helminthosporium solani, and black dot, caused by Colletotrichum
coccodes, cause tuber blemishes on potato (Solanum tuberosum L.) which affect processing and
fresh market trade. Tubers from ten cultivars were collected at harvest from three organic farms
in Wisconsin and categorized as symptomatic or asymptomatic based on visual symptoms of
silver scurf and black dot and/or signs of H. solani and C. coccodes. Tuber incubation and PCR
assays were performed on asymptomatic tubers to detect H. solani and C. coccodes. Tuber
incubation and PCR assays were in slight to fair agreement (kappa coefficient < 0.4) for
detecting both pathogens. Most asymptomatic tubers tested were positive by one or both assays
for H. solani (75%) or C. coccodes (94%). Minituber inoculation assays were also performed to
screen potato lines for resistance to silver scurf. Of the 14 lines tested, a diploid interspecific
hybrid, C287, had consistently low sporulation, suggesting it has partial resistance to silver scurf.
Since the majority of tubers harvested are already infected with one or both pathogens further
research should focus on organically acceptable management practices that may inhibit disease
development in field and in storage.
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Introduction
Potato (Solanum tuberosum L.) is the third most important food crop in the world for
human consumption (CIP 2012) and one of the top five organic vegetables purchased for home
consumption in the U.S. (Stevens-Garmon et al. 2007). Although organic potatoes comprise less
than one percent of total potato sales, Greenway et al. (2011) forecast a 19% annual growth rate
in U.S. per capita organic potato consumption from 2008 to 2013. Many biotic and abiotic
factors limit organic potato production. In order to meet the increasing consumer demand,
especially from the fresh market, there is a need to produce good quality blemish-free organic
potatoes.
Silver scurf, caused by the fungus Helminthosporium solani Durieu & Mont. is a little-
studied disease, particularly in organic potato production. At harvest, diseased tubers have grey
lesions on the periderm that appear silvery when moist. Under favorable storage conditions,
fungal sporulation makes tubers appear black and sooty. At later stages, the outer cell layers of
tubers slough off leading to tuber shrinkage and weight loss (Burke 1938; Jellis and Taylor
1974). Silver scurf has been gaining economic importance in the past three decades due to the
quality standards demanded by processing and fresh market consumers as well as the emergence
of H. solani isolates resistant to the broad-spectrum thiabendazole and thiophanate-methyl
fungicides (Errampalli et al. 2001; Geary et al. 2007). Initial infection of tubers by H. solani
occurs in the field either from the seed tuber (Jellis and Taylor 1977) or soil (Merida and Loria
1994). Disease spread occurs during storage, as air flow in storages aids in the dissemination of
fungal spores (Rodriguez et al. 1996).
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Colletotrichum coccodes (Wallr.) causes black dot, and the presence of small black
sclerotia on tubers, roots, and above- and below-ground stems gives the disease its common
name. Infected tubers may have gray-brown blemishes on the periderm, or less clearly delimited
discolored areas on which sclerotia may or may not be present (Jellis and Taylor 1974). The
pathogen can cause dark brown, deep sunken lesions on minitubers stored at 5 to 15 C for five
months (Glais and Andrivon 2004). Fresh weight losses of 5 to 10% occurred when tubers
naturally infected with C. coccodes were stored for 18 weeks (Hunger and McIntyre 1979). The
pathogen also infects roots, making them appear stringy when pulled out of the soil. Stolons
may also be attacked at any stage of tuber growth and the lesions can sever the stolon, leaving a
part of the stolon adhering to the tuber at harvest (Davis and Johnson, 2001). Foliar infections
manifest as dark brown to black lesions similar to those caused by Alternaria solani, but lacking
concentric rings (Johnson and Miliczky, 1993). Soil, infected seed tubers and air-borne inoculum
serve as sources of inoculum for this disease (Lees and Hilton 2003). The disease may be
overlooked as the symptoms appear late in the cropping season, coinciding with natural
senescence. In addition, symptoms on stems and stolons are similar to those of Rhizoctonia
canker, and on leaves similar to those of Verticillum wilt or early blight (Davis and Johnson,
2001).
Tuber symptoms of silver scurf and black dot are frequently confused owing to the
similarity of blemishes caused on the periderm, and because they may be present on the same
tuber (Jellis and Taylor, 1974). Depending on environmental conditions, both diseases can
increase during storage. Rodriguez et al. (1996) collected air-borne conidia of H. solani ranging
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from 0 to 12,000 conidia per day in storages at 4 C, and from 0 to 24,000 conidia per day in
storages at 10 C during one year of the study. They detected H. solani conidial production
throughout the storage season and also showed that greenhouse grown minitubers placed in the
storages could be infected by H. solani conidia. Although such extensive work has not been done
with black dot spread during storage, Mawson (1999) showed that preventing water
condensation during storage led to increased levels of black dot and decreased levels of silver
scurf on tubers. Latent infections by C. coccodes and H. solani have been documented (Glais-
Varlet et al. 2004; Tsror et al. 1999; Jellis and Taylor 1974) and may contribute to disease
increase during storage. To avoid continued underestimation of these diseases based on visual
symptoms alone, the frequency of latent infections on asymptomatic tubers, and the potential for
disease spread from such tubers, should be better understood. This information will help to
determine whether future efforts to manage these diseases in organic farming should focus on
reduction of inoculum spread or disease development in storage.
Organic potato production is unique from conventional production both in terms of
agronomic and plant protection practices. Fludioxonil, fludioxonil plus quintozene, and
azoxystrobin can reduce severity of silver scurf (Geary et al. 2007). However, these chemicals
cannot be used in organic potato production. Seed tuber treatments with salts acceptable in
organic production such as sodium carbonate and sorbic acid were not effective in controlling
silver scurf (Geary et al. 2007). In addition, to date, no potato cultivars with complete resistance
to silver scurf have been identified (Merida et al. 1994; Secor 1994; Thomas et al. 2005). Since
organic potato growers have few control strategies for silver scurf, we hypothesize that a high
percentage of organically grown potatoes, even if asymptomatic, are infected with H. solani
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and/or C. coccodes. With the availability of interspecific Solanum hybrids that have resistance to
multiple potato diseases (Jansky and Rouse 2003), resistance to silver scurf may be present in
potato germplasm suitable for breeding. Our objectives were to investigate the abundance of
asymptomatic tubers that harbor H. solani or C. coccodes in tubers harvested from organic potato
farms, and to identify potato germplasm that has resistance to silver scurf.
Materials and Methods
Tuber samples
Field trials were conducted during the 2011 growing season in conjunction with the on-farm
organic variety trials at three locations in Wisconsin: Antigo (A), Rosholt (B), and Cottage
Grove (C). At locations A and B, the experimental design was a randomized complete block
design with four replications at each location. The plots were hand planted and consisted of two
rows with 20 plants per row. Plots measured 1.8 m wide by 6.1 m long with 0.9 m row spacing.
At location C, a single replicate was planted, and cultivar location within the block was
randomized. The plots were hand planted, and consisted of single rows with 20 plants per row.
Plots measured 1.8 m wide by 6.1 m long with 0.9 m row spacing. Soil type, previous crop
history, and planting and harvest dates for each location are presented in Table 1. There were 31
cultivars planted at each location, including yellow, red, russet, and specialty cultivars. Of these
cultivars, 10 were selected that included multiple market classes and maturity groups, and are
commonly grown by conventional and/or organic farmers in Wisconsin (Table 2). Potatoes
were grown at these locations by farmers following organic management practices. The cultivars
planted had four different seed sources (Table 2) and all seed was grown under conventional
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management. Yield samples were harvested from measured lengths of each plot: 6.1 m lengths
for each plot row at location A, 3.05 m lengths of each plot row at location B, and 6.1 m lengths
of the single plot row at location C (data not shown). After harvest, the tubers from each plot
were bulked and thirty tubers of each cultivar were arbitrarily selected from one replicate plot at
each location. Sampled tubers were processed within a period of one week from the sampling
date, during which time the tubers were stored at 4 C. Each tuber was handled individually and
washed thoroughly under running tap water to remove soil deposits and then grouped as either
symptomatic or asymptomatic based on visual scoring. Tubers were considered to be
asymptomatic if they showed no signs of either H. solani or C. coccodes or symptoms of black
dot or silver scurf. Ten asymptomatic tubers were selected per cultivar per location for PCR and
tuber incubation assays, although in some cases there were fewer than ten asymptomatic tubers
available (Table 3). At location C there were fewer than ten asymptomatic tubers for some
cultivars. Hence, tubers that showed only signs of C. coccodes (sclerotia) but that lacked
symptoms of either disease and signs of H. solani were grouped as asymptomatic for silver scurf
but symptomatic for black dot (Table 3).
Tuber incubation assay
A humid chamber was made by placing a sterile paper towel in a 450 g deli container and
dispensing 10 ml of sterile distilled water onto the towel. A small rectangular portion on the side
of the container was cut out and the hole was sealed with Micropore 3M tape to facilitate
aeration. Each symptomatic (up to 15 tubers per cultivar per location) or asymptomatic tuber was
then placed in an individual deli container, and incubated in the dark at 24 C for a period of three
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40
weeks. Asymptomatic tubers were number coded so that the PCR and tuber incubation assays
were always conducted on the same tuber. At the end of the incubation period, the entire tuber
surface was examined for the presence of conidiophores and conidia of H. solani and sclerotia of
C. coccodes.
PCR assay
A one centimeter diameter circle of periderm was arbitrarily collected from each asymptomatic
tuber using a cork borer, excised and chopped into small pieces with a sterile blade and used for
DNA extraction. DNA was extracted from this tissue using the FastDNA SPIN Kit for Soil (MP
Biomedicals, Solon, OH) following the manufacturer’s protocol except that the homogenization
of the sample was carried out in a mini beadbeater (Biospec Products, Bartlesville, OK) for two
minutes. To detect H. solani, two sets of primer pairs (First-round: Hs1F1/Hs2R1; Nested:
Hs1NF1/Hs2NR1) developed by Cullen et al. (2001) were used to amplify ITS regions.
Sequences designed by Cullen et al. (2002) were used to amplify C. coccodes (First-round:
Cc1F1/Cc2R1; Nested: Cc1NF1/Cc2NR1). Each PCR reaction consisted of one µl of undiluted
potato DNA extract, 0.3 µM of each primer, and 1x PCR Master Mix (Promega Corporation,
Madison, WI) in 25 µl of total volume. One µl of the first-round PCR product was used for
nested PCR. Amplification of H. solani was carried out with cycling parameters slightly
modified from Cullen et al. (2001): 95 C for 150 s, followed by 30 cycles of 95
C for 45 s,
annealing at 66 C (first-round) and 60
C (nested) for 60 s, extension at 72
C for 90 s, and a final
elongation at 72 C for 5 min. All reactions were run in a Techne TC-512 thermocycler. H. solani
or C. coccodes DNA was used as the positive control and the negative control was nuclease free
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water (Promega Corporation, Madison, WI). PCR products were analyzed by electrophoresis on
2% agarose gel in 1X TAE buffer, stained with SYBR Safe DNA gel stain (Life Technologies,
Carlsbad, CA) and visualized under a UV transilluminator (Universal Hood II: Bio-Rad,
Hercules, CA). An asymptomatic tuber was considered positive for H. solani or C. coccodes if
the pathogen was detected by either PCR or the tuber incubation assay.
Minituber inoculation assay
Fourteen lines of minitubers were screened in two separate experiments (Table 4). Minitubers
grown in a nutrient film technique system were washed, surface disinfested with calcium
hypochlorite (0.5% active chlorine) for five minutes and rinsed twice in sterile water. B-AC-
16A isolate of H. solani recovered from an infected potato was grown on V8 medium containing
177 ml V8 juice, 20 g agar and 3 g CaCO3 per liter (Goth and Webb 1983). Cultures were
incubated in the dark at 24 C and a conidial suspension was prepared by scraping a four-week old
colony in a small volume of sterile water. The conidial suspension concentration was determined
with a hemacytometer and adjusted to the required concentration (105 spores per milliliter) by
dilution with 0.1% water agar. Lines C287, Dark Red Norland, Early Epicure and Epicure
Banana were screened in both experiments. A minimum of ten minitubers per line were
arbitrarily selected and inoculated with H. solani conidial suspension until wet using an
aspirator. Up to three inoculated minitubers of the same line were placed together in individual
humid chambers, which were designed as described earlier for the tuber incubation assay. Three
minitubers per line included as negative controls were sprayed with 0.1% water agar. After one
month, each minituber was placed in a tube containing 15ml of deionized water plus 10 µl of
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10% Tween20 and washed for five minutes on a laboratory shaker to extract conidia. The
minitubers were then removed from the suspension and weighed. To standardize across the
samples, the volume in each tube was adjusted to 20 ml with deionized water. A 10 µl portion of
this suspension was filtered through a membrane filter (Munck and Stanosz 2009) and conidia of
H. solani were counted with the aid of a dissecting microscope. The mean number of conidia
obtained from three such readings was adjusted for the total volume of water in which the
minituber was washed and divided by tuber weight to estimate the number of conidia extracted
per gram of minituber.
Statistical analyses
Tubers asymptomatic for silver scurf and black dot at harvest (Table 2), and the asymptomatic
tubers positive for H. solani and C. coccodes after performing tuber incubation and PCR assays
(Table 3) were binary responses. Hence, Fisher’s exact tests were performed to compare the
proportion of asymptomatic tubers among cultivars within a location and also to compare among
locations within a cultivar. PCR and tuber incubation assays were considered as two ratings
assessing an asymptomatic tuber, hence the inter-rating agreement between the assays was
measured using the kappa coefficient. The kappa coefficient lies between -1 and 1, where values
less than zero indicate less agreement than expected by chance and values greater than zero
indicate more agreement than expected by chance (Cohen 1960). Interpretation of the kappa
coefficient was made using an arbitrary benchmark scale (Landis and Koch 1977). For minituber
inoculation assays, significant differences were noted between the two experiments. Hence, a
linear model was fit treating the lines that were common in both experiments as separate lines.
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Comparisons among lines were made using ANOVA, with means separated when necessary
using Fisher’s LSD. All statistical analyses were performed with R software version 2.14.1.
Statistical significance was set at P < 0.05.
Results
Disease incidence at harvest
Ten potato cultivars were assayed for the incidence of black dot and/or silver scurf after harvest.
At each location, significant differences were observed among cultivars for the number of
asymptomatic tubers for silver scurf and/or black dot at harvest (P < 0.05). However, none of the
cultivars assayed were resistant to these diseases (Table 2). Barring three cultivars (Purple
Majesty, Freedom Russet and Dark Red Norland) from location A, and three cultivars (Freedom
Russet, Keuka Gold and Satina) from location B, more than 50% of the sampled tubers for each
cultivar were symptomatic for silver scurf and/or black dot. Significant differences (P < 0.05) for
the number of asymptomatic tubers at harvest were found among locations except for
Adirondack Blue and Adirondack Red (Table 2). The cultivar Freedom Russet was an extreme
example of this variability. All assayed Freedom Russet tubers from location B were
asymptomatic, while only 30% from location C appeared healthy.
Tuber incubation and PCR assays
Asymptomatic tubers were assayed for H. solani and C. coccodes by tuber incubation and PCR
assays to determine if these tubers carried either pathogen (Table 3). Tubers were considered
infected if they were positive for the pathogen by either assay. The majority of asymptomatic
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44
tubers assayed from these fields were infected with one or both of the pathogens: 75% of the
asymptomatic tubers assayed were positive for H. solani and 94% were positive for C. coccodes
(Fig. 1). The proportion of asymptomatic tubers that were positive for H. solani following tuber
incubation and PCR assays (Table 3), differed significantly among cultivars at each of the three
locations (P < 0.05). Only Adirondack Blue, Superior and Yukon Gold showed significant
differences among locations for incidence of H. solani on asymptomatic tubers. It is interesting
to note that tubers of Freedom Russet from location B showed no lesions at harvest (Table 1), yet
all tubers were infected with H. solani (Table 3). No significant differences were found among
locations or cultivars for C. coccodes (Table 3).
For both pathogens, among most of the cultivars and locations tested, there was slight to
fair agreement between the tuber incubation and PCR assays (Table 5). A moderate agreement
for H. solani was observed with Dark Red Norland and Yukon Gold. Location B had a
substantial agreement between the two H. solani assays. Tuber incubation and PCR assays did
not agree in 37% and 26% of the asymptomatic tubers assessed for H. solani and C. coccodes
respectively (Fig. 1). In samples where the two assays did not match, the tuber incubation assay
resulted in more positives for H. solani (29%) than C. coccodes (8%), while the PCR assay
detected more positives for C. coccodes (18%) than H. solani (8%).
The majority of the tubers that were assayed were infected with both H. solani and C.
coccodes. Both pathogens occurred on the same tuber in 69% of symptomatic and 65% of
asymptomatic tubers assessed (Fig. 2). In many cases, conidiophores of H. solani and sclerotia of
C. coccodes were observed in close proximity when the tuber periderm was viewed with the aid
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45
of a dissecting microscope (Fig. 3). Co-occurrence of these two fungi was reported earlier
(Nitzan et al. 2005), but there is essentially no information available on how these fungi may
interact on tuber surfaces.
Minituber inoculation assays
In the minituber inoculation assays, significant differences were observed among lines for H.
solani sporulation (Table 4). C287 consistently had the lowest conidial count per gram of
minituber, and Peruvian Blue had the highest level of sporulation. Epicure Banana from one
experiment had low sporulation that was not significantly different from C287, but it produced
higher number of conidia (24806 ± 3764) per gram of minituber in the other experiment,
suggesting variability either within the line or the experiments. However, since C287 supported a
low level of sporulation in both experiments, it appeared to have partial resistance to silver scurf.
Discussion
A high incidence of silver scurf and black dot was found across locations and cultivars
with 75% and 94% of asymptomatic tubers assessed positive for H. solani and C. coccodes
respectively. These results, coupled with the difficulty of distinguishing between the symptoms
of silver scurf and black dot, especially on blue-skinned cultivars, indicate that visual assessment
of the tubers at harvest for silver scurf and black dot symptoms is not predictive of pathogen
incidence. Since high incidences of both fungi were observed on asymptomatic tubers, the tuber
infections might have occurred shortly before harvest, or, alternatively infections may remain
latent in developing tubers. Consistent with these results, others have reported tuber latent
infections by C. coccodes (Glais-Varlet et al. 2004 and Tsror et al. 1999). Likewise, Jellis and
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Taylor (1974) did not observe symptoms of silver scurf until three to five weeks after infection.
Our results have direct implications for seed potato production. Black dot (Johnson et al. 1997)
and silver scurf (Geary and Johnson 2006) incidence increases with each year that potatoes are
replanted. Thus, organic farmers would likely face a very high incidence of both silver scurf and
black dot if they replant the tubers they harvested.
Since initial information about the presence of either pathogen on asymptomatic tubers
prior to our assays is unobtainable, this limited our ability to judge whether the tuber incubation
or PCR assay is more accurate. The fast, but relatively expensive PCR assay was in slight to fair
agreement with the slower, but inexpensive tuber incubation assay. The periderm samples for the
PCR assay were taken arbitrarily from each tuber and it is unlikely that these pathogens are
distributed evenly over the tuber surface at the time of harvest. Indeed, previous work showed
that C. coccodes is isolated at greater frequency from tuber stem ends than from either lateral
sections or bud ends (Johnson et al. 1997). These pathogens may also be present on the tuber but
lack sclerotia or fruiting bodies after incubation, which would result in low agreement between
the assays. Both scenarios likely occurred in this study since H. solani was detected more
frequently by the tuber incubation assay, suggesting that infected regions of some tubers were
not assayed, and C. coccodes was detected more frequently by PCR, suggesting that sclerotia did
not form consistently in our tuber incubation assay.
No cultivar has yet been identified that is completely resistant to silver scurf (Merida et
al. 1994; Secor 1994; Thomas et al. 2005). However, partial resistance to silver scurf was
reported in wild Solanum species such as S. demissum, S. chacoense, S. acaule, S. stoloniferum,
47
47
S. oxycarpum and S. hondelmannii (Rodriguez et al. 1995). Previous studies indicate that tuber
assays are likely to give a more accurate assessment of cultivar resistance to silver scurf than
field tests, since disease severity increases with the length of time that tubers remain exposed to
the pathogen in the field and this time period varies between early- and late-maturing cultivars
(Merida et al. 1994). In this study, minitubers were used instead of field-grown potatoes, because
when surface disinfested field-grown tubers were inoculated and incubated, the majority of the
samples were lost due to tuber decay pathogens. Minitubers grown by the nutrient film technique
remain nearly pathogen free, which minimized tuber decay.
Screening for resistance was performed based on the ability of H. solani to sporulate on
the tuber surface, a measure of resistance for silver scurf used previously by Merida et al. (1994)
and Rodriguez et al. (1995). Of the 14 lines examined, only C287 had consistently low spore
counts in both minituber inoculation experiments. C287, a diploid (2n = 2x = 24) interspecific
hybrid line has wild Solanum species in its parentage [Female parent: (US-W730 × (S.
berthaultii × S. tarijense) and male parent: (US-W730 × S. tarijense)]. C287 is resistant to
Verticillium wilt and other surface blemish diseases such as common scab and black scurf
(Jansky and Rouse, 2003). The results suggest that C287 may also have partial resistance to
silver scurf, which may be usefully transferred to commercial cultivars as has been done for
common scab (Murphy et al. 1995) and Verticillium wilt (Frost et al. 2006) resistance.
The high prevalence of H. solani and C. coccodes on organic potatoes and the lack of
tuber resistance in currently available cultivars suggest that future research should also focus on
preventing disease development during the growing season and in storage. Field data indicate a
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complex relationship between pathogen incidence, disease expression, cultivar, and location.
Although the percentage of asymptomatic tubers (Table 2) and the frequency of H. solani
infections (Table 3) in asymptomatic tubers differed significantly between some cultivars and
locations, no clear trends were seen when comparing cultivars and locations. Further research
into the influence of soil characteristics and management choices on disease development during
the growing season is needed. For organic growers with operational flexibility and a ready
market, harvesting and marketing immediately at plant maturity may be an adequate control for
silver scurf, since disease severity increases if tubers remain in the field (Merida et al 1994).
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Table 1. Soil type, previous crop, and planting and harvest dates for each sampling locationz
Aspect Location A Location B Location C
Soil type Langlade silt loam Sandy loam Dodge silt loam
Planting date 30 May 2011 20 May 2011 16 May 2011
Harvest date 11 October 2011 20 September 2011 17 October 2011
Previous crop Oats and Clover Alfalfa and Rye Popcorn
z Sampling locations A, B and C were farms located in Antigo, Rosholt, and Cottage Grove, WI
respectively.
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Table 2. Percentage of potato tubers asymptomatic for both silver scurf and black dot after harvest from three locationsx in Wisconsin
% Asymptomatic tubers
Seed Seed __________________________________________
Cultivar sourcey certification Maturity group Location A Location B Location C P-value
z
Market class: Blue
Adirondack Blue D Certified Medium 33.3 (30) 50.0 (30) 26.7 (30) 0.155
Purple Majesty R Certified Medium 82.1 (28) 31.0 (29) 10.0 (30) <0.001
Market class: Red
Adirondack Red D Certified Early to Medium 26.7 (30) 50.0 (30) 26.7 (30) 0.089
Chieftain C Certified Medium 20.7 (29) 40.0 (30) 0.0 (30) <0.001
Dark Red Norland L Foundation Early 60.0 (30) 0.0 (30) 10.0 (30) <0.001
Market class: Russet
Freedom Russet L Foundation Medium to late 65.4 (26) 100.0 (30) 30.0 (30) <0.001
Market class: Yellow
Keuka Gold D Certified Medium to late 46.7 (30) 63.3 (30) 10.0 (30) <0.001
Satina C Certified Medium early 30.4 (24) 53.3 (30) 23.3 (30) 0.047
Superior L Foundation Early 21.4 (28) 46.7 (30) 0.0 (30) <0.001
Yukon Gold L Foundation Early 26.7 (30) 36.7 (30) 10.0 (30) 0.045
P-valuez
<0.001 <0.001 0.002
x Sampling locations A, B and C were organic farms located in Antigo, Rosholt, and Cottage Grove, WI respectively. Numbers in
parentheses indicate sample size.
y Seed source: D= David Gallenberg farm, WI; C = Childstock farm, NY; L= Lelah Starks farm, WI; R= Rockey Farms, CO.
z Statistical analysis was carried out in R (Version 2.14.1) using Fisher’s exact test at 5% significance level.
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Table 3. Percentage of asymptomatic tubers positive for Helminthosporium solani and Colletotrichum coccodes after performing PCR
and tuber incubation assays
H. solani C. coccodes
_____________________________________________ _______________________________________________
Cultivarx Location
y
_______________________________________________________________________________________________
A B C P-valuez A B C P-value
z
____________________________________________________________________________________________________________
ADB 80.0 (10) 10.0 (10) 50.0 (10) 0.009 90.0 (10) 90.0 (10) 90.0 (8) 1.000
PMJ 60.0 (10) 33.3(9) 85.7 (7) 0.115 100.0 (10) 100.0(9) 71.4 (3) 1.000
ADR 75.0 (8) 100.0 (10) 100.0 (8) 0.172 87.5 (8) 90.0 (10) 62.5 (8) 0.458
CFT 83.3 (6) 100.0 (10) 100.0 (9) 0.240 100.0 (6) 100.0 (10) NA 1.000
DRN 30.0 (10) NA 66.7 (9) 0.179 100.0 (10) NA 100.0 (3) 1.000
FDR 100.0 (10) 100.0 (10) 100.0 (9) 1.000 100.0 (10) 80.0 (10) 100.0 (9) 0.310
KKG 90.0 (10) 80.0 (10) 100.0 (10) 0.754 80.0 (10) 100.0 (10) 100.0 (2) 0.567
STN 100.0 (7) 60.0 (10) 90.0 (10) 0.143 100.0 (7) 100.0 (10) 90.0 (4) 0.190
SUP 83.3 (6) 0.0 (10) 90.0 (10) <0.001 100.0 (6) 100.0 (10) NA 1.000
YKG 100.0 (8) 30.0 (10) 100.0 (10) <0.001 100.0 (8) 100.0 (10) 100.0 (3) 1.000
P-valuez 0.004 <0.001 0.004 0.483 0.507 0.410
x Cultivar: ADB= Adirondack Blue, PMJ = Purple Majesty, ADR = Adirondack Red, CFT = Chieftain, DRN = Dark Red Norland,
FDR= Freedom Russet, KKG = Keuka Gold, STN = Satina, SUP = Superior and YKG = Yukon Gold.
y Sampling locations A, B and C were organic farms located in Antigo, Rosholt, and Cottage Grove, WI respectively. Numbers in
parentheses indicate sample size.
z Statistical analysis was carried out using Fisher’s exact test at 5% significance level.
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Table 4. Spore production on fourteen potato minituber lines inoculated with Helminthosporium
solani
Line Number of conidia extracted per gram of minituber
Peruvian Blue 49283x ± 6437.7 a
Epicure Banana-Ay 24806 ± 3763.9 b
Peanut 23846 ± 6363.6 bc
Early Epicure-A 24307 ± 5815.1 bcd
Dark Red Norland-A 17793 ± 3845.4 bcd
Nosebag 12744 ± 2950.9 bcde
Superior 10957 ± 1854.3 cdef
Early Epicure-Bz 11717 ± 1882.1 def
Dark Red Norland-B 11892 ± 2456.1 def
White Cobbler 9920 ± 2633.9 ef
Scotia Blue 9728 ± 2278.9 ef
Trina 8514 ± 1596.4 ef
Pike 7021 ± 1372.3 ef
Round Blue Andean 10387 ± 6066.8 f
Tundra 3775 ± 1235.0 g
Epicure Banana-B 2360 ± 410.9 gh
C287-B 2031 ± 403.7 hi
C287-A 1216 ± 292.5 i
x Means (±SE) followed by the same lowercase letters are not significantly different within a
column (α = 0.05).
y Lines followed by the letter A are sporulation data from experiment 1
z Lines followed by the letter B are sporulation data from experiment 2
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Table 5. Kappa coefficient values showing agreement between tuber incubation and PCR assays
for detecting Helminthosporium solani and Colletotrichum coccodes from asymptomatic tubers
Kappa coefficienty
_______________________________________________
Cultivar / Locationz H. solani C. coccodes
Cultivar:
Adirondack Blue 0.11 0.24
Purple Majesty 0.21 0.0
Adirondack Red 0.16 0.19
Chieftain 0.04 0.0
Dark Red Norland 0.55 0.0
Freedom Russet 0.0 0.23
Keuka Gold 0.09 0.30
Satina 0.17 0.01
Superior 0.27 0.0
Yukon Gold 0.41 -0.07
Location:
A 0.24 0.25
B 0.62 0.09
C 0.03 0.30
y Kappa coefficients (Landis and Koch 1977): 0.81 to 1.00 = almost perfect agreement, 0.61 to
0.80 = substantial agreement, 0.41 to 0.60 = moderate agreement, 0.21 to 0.40 = fair agreement,
0.00 to 0.20 = slight agreement, <0.00 = poor agreement.
z Sampling locations A, B and C were organic farms located in Antigo, Rosholt, and Cottage
Grove, WI respectively.
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Figure 1. Percentage of asymptomatic tubers positive or negative for Helminthosporium solani
and Colletotrichum coccodes by tuber incubation (T) and PCR (P) assays. (T0P0) Negative for
both tuber incubation and PCR assays. (T1P1) Positive for both tuber incubation and PCR
assays. (T1P0) Positive for tuber incubation and negative for PCR assay. (T0P1) Negative for
tuber incubation and positive for PCR assay.
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Figure 2. Percentage of tubers positive or negative for Helminthosporium solani (Hs) and
Colletotrichum coccodes (Cc) by either tuber incubation or PCR assay for asymptomatic tubers
and by tuber incubation assay for symptomatic tubers. (Hs1Cc0) Positive for H. solani and
negative for C. coccodes. (Hs0Cc1) Negative for H. solani and positive for C. coccodes.
(Hs0Cc0) Negative for both H. solani and C. coccodes. (Hs1Cc1) Positive for both H. solani and
C. coccodes.
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Figure 3. Conidia and conidiophores of Helminthosporium solani (left side) and sclerotia of
Colletotrichum coccodes (right side) observed in close proximity on a potato tuber periderm
under a dissecting microscope.
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CHAPTER 2
A microfluidic assay for identifying differential responses of plant and human fungal
pathogens to tobacco phylloplanins
The structure of this chapter has been formatted with submission to PeerJ journal in mind:
Mattupalli C, Spraker JE, Berthier E, Charkowski AO, Keller NP, Shepherd RW. A microfluidic
assay for identifying differential responses of plant and human fungal pathogens to tobacco
phylloplanins. PeerJ.
Some of the data published in this chapter was done in Dr. Keller and Dr. Shepherd lab.
Berthier E fabricated the microfluidic plates and Spraker JE was responsible for the Aspergillus
assays.
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Abstract
Phylloplanins are defensive glycoproteins secreted onto leaf surfaces by trichome bearing plants
such as tobacco. They are of interest because of their anti-microbial properties, but like other
natural product bioactives, the assessment and screening of phylloplanin biological activity is
impeded by limited availabilities of active compounds. Here we report an inexpensive
microfluidic approach that requires only a few microliters of Nicotiana tabacum (tobacco)
phylloplanins (Nt-phylloplanins) to assess spore germination inhibition of plant and human
fungal pathogens. Spores of Colletotrichum coccodes and Aspergillus fumigatus treated with Nt-
phylloplanins did not germinate at 48 and 30 hours post treatment respectively. Nt-phylloplanins
transiently inhibited spore germination of Fusarium sambucinum, but had no detectable activity
against Alternaria alternata or Verticillium albo-atrum, demonstrating differential sensitivity of
fungi to Nt-phylloplanins. Inhibition of spore germination of the human pathogen A. fumigatus
by Nt-phylloplanins also reveals a new venue for medicinal antifungal drug discovery that has
not been explored previously.
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Introduction
The phyllosphere or microorganismal habitat of leaf surfaces is a complex and inhospitable
habitat exposed to continuous fluctuations in physical environment (Hirano and Upper, 2000). It
is inhabited by diverse genera of microbes including potential pathogens, that must also endure
the array of preformed and induced plant defences of the leaf surface or phylloplane (Lindow
and Brandl, 2003; Shepherd and Wagner, 2012). While many well-studied plant defenses and
defensive processes are triggered by pathogens as they access the plant interior or penetrate the
epidermal cell wall (Chisholm et al. 2006; Bent and Mackey 2007; Underwood and Somerville
2008), it is evident that sizable fortifications are also present at the outermost layers of the air-
plant interface, and consist mainly of secreted plant-produced biochemicals that act directly
against pathogens to hinder or obstruct their progress (Kennedy et al. 1992; Karamanoli and
Lindow 2006; Shepherd and Wagner 2007).
The leaf surface is composed of a pavement of epidermal cells interspersed with stomata,
trichomes and other leaf surface structures (Hirano and Upper, 2000). Trichomes are epidermal
protuberances developing outwards on the surface of various plant organs (Werker, 2000) and
can be broadly classified as simple/non-glandular and glandular secreting types. Glandular
secreting trichomes are present in about 30% of vascular plants and secrete secondary
metabolites such as terpenoids and phenylpropanoids that can provide disease and pest resistance
(Kelsey et al. 1984; Wagner et al. 2004). Shepherd et al. (2005) discovered that plants produce
and secrete to their leaf surfaces antifungal plant proteins termed phylloplanins. Native
phylloplanins of Nicotiana tabacum (Nt-phylloplanin) can be collected by gently agitating
undamaged leaves in water, lyophilizing the resulting solution and resuspending the lyophilized
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product in sterile water. With SDS-PAGE, Nt-phylloplanin is visible as four bands with
molecular masses of 16 kDa, 19 kDa, 21 kDa, and 25 kDa. The four bands all have the same N-
terminal amino acid sequence and are believed to arise from glycosyl additions to a single 13
kDa core polypeptide encoded by the phylloplanin gene (Accession AY705384). Homologues of
this gene have been reported to exist and be expressed only in N. tabacum trichome glands
(Harada et al. 2010).
Analyses of the phylloplanin promoter using reporter genes indicate that it directs expression
solely in short glandular secreting trichomes, thus revealing a highly specialized mechanism for
localization and delivery of proteins to leaf surfaces (Shepherd et al., 2005). Short glandular
trichomes are believed to continually secrete phylloplanins onto the leaf surface, although factors
such as leaf age might influence secretion. After leaf surface proteins are removed, their levels
are renewed within one week (Kroumova et al. 2007). Native Nt-phylloplanins have inhibitory
activity against spore germination and hyphal growth of the basidiomycete Rhizoctonia solani,
and the ascomycete Pyricularia oryzae (King et al. 2011). A recombinant 13 kDa Nt-
phylloplanin polypeptide produced in Escherichia coli inhibited spore germination of
Peronospora tabacina, causal agent of blue mold disease in tobacco (Shepherd et al. 2005).
Mechanized methods for harvesting phylloplane proteins on a large scale are not yet available.
This impeded our ability to screen for their antimicrobial properties and led us to explore for
techniques that are compatible with low volume of reagents. Microfluidic platforms are reputed
for temporal and spatial control of microenvironments concurrently allowing assays to be
performed in micro-channels using smaller volumes of samples (Paguirigan and Beebe, 2008).
To date, these microsystems have been successfully used for applications in drug discovery, cell
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culturing and characterization of fungal secondary metabolites (Dittrich and Manz, 2006; Gao et
al. 2011; Berthier et al. 2013). Here, we developed a simple microfluidic channel array that
enabled the in vitro study of antifungal effects of Nt-phylloplanin on plant and, for the first time,
human pathogens. We verified the hypothesis that Nt-phylloplanins have inhibitory effects on a
subset of fungi that have not been studied earlier and suggest that these molecules be considered
for development in medicinal venues.
Materials and Methods:
Nt-phylloplanin collection: Tobacco (Nicotiana tabacum TI 1068) plants were grown in 15.2-
centimeter diameter plastic pots in a greenhouse. MetroMix (Sun Gro Horticulture, BC, Canada)
was used as potting soil and a slow releasing fertilizer (Osmocote) was applied at the rate of 7.5
g per each pot. Nt- phylloplanins were collected by washing fully expanded leaves (undetached
from the plant) in 200 ml of distilled water with gentle agitation for 15 seconds. Leaves with
visible damage were not washed to avoid contamination from leaf sap. The wash solution was
then passed through a Miracloth (EMDMillipore, MA, USA) to filter debris and lyophilized to
dryness. The lyophilized powder was resuspended in 1 ml of sterile Milli-Q water, and
centrifuged at 12,000 × g for 5 minutes at 24 °C. The supernatant was filtered using a PVDF
membrane syringe filter (13 mm / 0.22 µm; Fisher Scientific, PA, USA) and stored at 4oC until
further use. Proteins were separated from the lyophilized product by sodium dodecyl sulfate –
polyacrylamide gel electrophoresis (SDS-PAGE) using a Mini-PROTEAN Tetra cell
electrophoresis system (Bio-Rad, Hercules, CA) and visualized with Coomassie Brilliant Blue.
Nt-phylloplanin concentration of 1 mg mL-1
was used in this study and the protein concentration
was estimated using BCA protein assay kit (Thermo Scientific Pierce, Rockford, IL).
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Proteinase treatment: Proteinase K immobilized on Eupergit C beads (200mg, Sigma-Aldrich,
MO, USA) was placed into mini-spin filters and 500 µL of leaf water wash samples were added
to these filters. The filters were then placed in a 1.5 mL microcentrifuge tube and incubated at
37oC overnight. Flow-through was collected by centrifuging the tubes at 5000 g for 10 minutes
which was again incubated at 95oC for 30 minutes, followed by a second centrifugation at 5000g
for 5 minutes to pellet any precipitate.
Fungal cultures: Alternaria solani (Asol11), Verticillium albo-atrum (AC186), Fusarium
sambucinum (PD1), Colletotrichum coccodes (DRN-MK), Aspergillus fumigatus (Af293) were
the fungal pathogens used for in vitro testing of phylloplanin activity. A. fumigatus was grown on
glucose minimal media (GMM) agar at 37ºC for 3 days. A. fumigatus spore suspensions were
made by applying 10 ml of sterile 0.01% Tween-20 solution to the plate and subsequently
agitating conidiophores with a cell spreader. A. solani was grown on clarified V8 media, F.
sambucinum on quarter strength PDA, C. coccodes and V. albo-atrum on full strength PDA.
Spore suspensions were prepared by scraping 10 to 15 day old colonies into a small volume of
sterile water. Spore suspensions were quantified using a haemocytometer and appropriate
dilutions were made with sterile ddH2O.
Microfluidic channel fabrication: Microfluidic channel assays were fabricated using a soft-
lithography method (Fig. 1). The mold was fabricated from a 15 cm diameter silicon wafer upon
which was spun 2 layers of SU8 epoxy photoresist, the first was set to a thickness of 150 µm and
the second to 500 µm. Masks for patterning the designs in the photoresist were drawn on Adobe
Illustrator (Adobe, USA) and printed at Imagesetter Inc (Madison, WI, USA). The soft-
lithography process was performed by mixing polydimethylsiloxane (PDMS) in a ratio of 1:10,
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degassing the mix in a vacuum desiccator, and pouring onto the mold. Following the application
of PDMS, a transparent sheet, a rectangle of silicone rubber, a rectangle of rigid polyester, and 4
kg of weights were placed on the mold. The molds were baked at 80˚C for 4 hours after which
the PDMS was removed from the mold and bonded via to a glass microscope slide following an
oxygen plasma surface activation treatment (FEMTO, Diener, Germany).
Spore Germination Assays: The spore germination assay was conducted in microfluidic channels
prepared according to the described method (Fig. 1). Each micro-channel was loaded with 20 µl
or 15 µl (A. fumigatus) total fluid volume, containing 10 µl or 13.5 µl (A. fumigatus) of
phylloplanin solution and 10 µl or 1.5 µl (A. fumigatus) spore suspension, premixed in
microcentrifuge tubes. Final spore concentration in each micro-channel was 4 × 102 for A. solani,
1.5 × 103 for A. fumigatus, 2 × 10
3 for C. coccodes and F. sambucinum, and 2 × 10
4 for V. albo-
atrum. Controls of proteinase treated phylloplanin solution were used and loaded in equal
volume to the treatment channels. Channels were incubated at room temperature in a moist
chamber to prevent evaporation from the channels. Germination rates were quantified
microscopically at 6, 12, 24, 30 or 48 hours post inoculation by counting 200 spores per
treatment. A spore was considered germinated if the length of the germ tube exceeds half the
length of the spore or multiple germ tubes developed. Each treatment had three biological
replicates.
Results and Discussion:
Due to the large volumes of water used for collecting phylloplanins from leaf aerial surfaces, the
samples first needed to be lyophilized to yield a concentrated product highly enriched in protein
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but very limited in volume and quantities. This aspect of sample recoverability was a significant
bottleneck in our ability to perform a large number of assays. Here, we developed a microfluidic
channel array enabling the efficient study of phylloplanins requiring only microliters of reagents
(Fig. 1). The accessible microscale assay leveraged simple pipette-based pumping methods
(Berthier and Beebe, 2007) allowing simple fabrication and use of the channels. The channels
positioned on a microscope slide allowed straightforward use and imaging at known locations
and devoid of typical meniscus effects that occur in microwell plates. Using this microscale
array, we performed tens of assays simultaneously with low volumes of Nt-phylloplanins, and
demonstrated that this assay will be broadly useful in further large-scale screening of natural
product bioactives against microorganisms.
Earlier studies indicated the antimicrobial effects of Nt-phylloplanins (Shepherd et al. 2005;
Kroumova et al. 2007; King et al. 2011) against an oomycete and two fungal pathogens. Here,
we show that the antimicrobial activity of Nt-phylloplanins is specific to some fungal species,
suggesting that fungal resistance mechanisms exist. Differential responses to antimicrobial
proteins by pathogens, which are likely driven by host-pathogen co-evolution, is not uncommon
(Segura et al. 1998; Segura et al. 1999; Van Damme et al. 1999; Vigers et al. 1992). For instance,
a 30 kDa protein isolated from leaves of Engelmannia pinnatifida strongly inhibited the growth
of plant pathogens such as Fusarium oxysporum, Alternaria solani, Gaeumannomyces graminis,
but not the human pathogen, Candida albicans (Huynh et al. 1996).
The inhibition of spore germination by tobacco leaf wash samples was tested in vitro against four
fungal plant pathogens (Alternaria solani, Verticillium albo-atrum, Fusarium sambucinum,
Colletotrichum coccodes), and a human pathogen (Aspergillus fumigatus). Nt-phylloplanins
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strongly inhibited C. coccodes and A. fumigatus, transiently inhibited Fusarium sambucinum,
and were inactive against A. solani and V. albo-atrum, thereby indicating a spectrum of pathogen
sensitivity (Table 1). C. coccodes spores treated with tobacco leaf wash sample did not germinate
at 48 hours post inoculation when treated with 1 mg mL-1
Nt-phylloplanins (Fig. 2). The LD50 for
Nt-phylloplanins against C. coccodes spores was approximately 0.1 mg mL-1
(Fig. 3) C.
coccodes spores failed to germinate when they were treated with tobacco leaf wash sample
overnight and washed twice with sterile water, thus the effect of Nt-phylloplanins was
irreversible (Fig. 2).
Few of the A. fumigatus spores germinated when incubated with tobacco leaf wash samples at
the Nt-phylloplanin concentrations tested, and this effect persisted for at least 30 hours post
inoculation (Fig. 4). The inhibition of A. fumigatus (Fig. 4) and C. coccodes (Fig. 2) spore
germination was eliminated when proteins were digested with proteinase-K, suggesting that Nt-
phylloplanins were necessary for inhibition.
Unlike C. coccodes and A. fumigatus, F. sambucinum spore germination was only transiently
inhibited by Nt-phylloplanins. F. sambucinum spores germinated in negative controls at 12 hours
post inoculation (hpi), but not when treated with Nt-phylloplanins. However, this inhibitory
effect started to degrade at 16 hpi and by 24 hpi no inhibitory effect was evident and extensive
hyphal growth occurred (Fig. 5).
The mechanism of action of Nt-phylloplanins remains unknown. Since, Nt-phylloplanins inhibit
C. coccodes and A. fumigatus (this work), Rhizoctonia solani and Pyricularia oryzae (King et al.
2011), but only transiently inhibit F. sambucinum and are ineffective against Alternaria solani
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and, Verticillium albo-atrum, this suggests that host-pathogen co-evolution may select for Nt-
phylloplanin resistance in some fungi. Knowledge of fungal resistance mechanisms and the ease
of acquisition of resistance to phylloplanins would aid in decisions on whether to develop these
antimicrobial proteins as a natural product for use in plant / human protection.
Aspergillus fumigatus is the most common Aspergillus species to cause human infections
(Dagenais and Keller, 2009) and it causes a spectrum of diseases such as allergic
bronchopulmonary aspergillosis, aspergilloma and invasive aspergillosis, depending on the
host’s immune status (Latge, 1999; Dagenais and Keller, 2009; Zaas and Alexander, 2009).
Currently there are four classes of antifungal agents with activity against Aspergillus: the
polyenes such as amphotericin B and nystatin; the triazoles such as itraconzole, voriconazole and
posaconazole; the echinocandins such as caspofungin and micafungin; and the allylamines such
as terbinafine (Chamilos and Kontoyiannis, 2005). However, treatments involving amphotericin
B and triazoles may cause nephric and hepatic toxicity (Wingard et al., 1999; Tan et al., 2006).
Lipid-based formulations of amphotericin B have reduced the incidence of nephrotoxicity, but
they do so at a significantly increased drug acquisition cost (Pound et al., 2011). Also, in some
countries such as UK, up to 15% of A. fumigatus strains have been found to be resistant to
various azoles (Mayr and Lass-Florl, 2011). This highlights the ongoing requirement for new
anti-fungal agents for this pathogen (Denning and Hope, 2010) and antimicrobial proteins are
now being explored as potential therapeutics. Over 1000 antimicrobial peptides are now known
to be produced by different classes of organisms, of which about 80 are plant defensins or plant
defense peptides produced by 50 plant species (AMSDatabase). To date, a majority of the
antimicrobial peptides utilized in drug development are of animal origin (Gordon et al., 2005).
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Our results suggest that examination of plant antimicrobial proteins, such as Nt-phylloplanins,
may prove fruitful for development of novel treatments for A. fumigatus.
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Table 1: Comparison of the effects of Nt-phylloplanins on fungal spore germination
Fungus Fungal family Disease Host Inhibition of
spore
germination
by Nt-
phylloplanins
Alternaria solani Pleosporaceae Early blight Plants No
Verticillium albo-atrum Plectosphaerellaceae Wilt Plants No
Fusarium sambucinum Nectriaceae Dry rot Plants Transient
Colletotrichum coccodes Glomerellaceae Black dot Plants Yes
Aspergillus fumigatus Trichocomaceae Invasive
aspergillosis
Humans Yes
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Figure 1: Schematic representation of microfluidic device fabrication process.
(1) A multilayered mold is fabricated using photolithography methods from a light-sensitive
negative photoresist SU8. Each layer of SU8 is spun on top of the previous one and patterned by
exposure to UV light. (2) Using soft-lithography, PDMS polymer is cured on the mold to create a
replicate of the channel array. (3) The PDMS layer is bonded to glass using an oxygen-plasma
surface activation method. (4) The result is a platform containing an array of microscale channels
on a microscope slide.
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Figure 2. Inhibition of Colletotrichum coccodes spore germination after treatment with Nt-
phylloplanins in tobacco leaf wash
C. coccodes spores were subjected to various treatments: water control, proteinase-K treated Nt-
phylloplanins, Nt-phylloplanins (1 mg mL-1
), treatment with Nt-phylloplanins overnight
followed by washing twice in sterile distilled water, and incubated in microfluidic channels (2 ×
103
spores per channel). Each point represents an average of three biological replicates with 200
spores assessed per each treatment. The data was arcsine transformed prior to being analyzed
using ANOVA, with means separated using Fisher’s LSD in R 2.15.3. Treatments with the same
lowercase letters are not significantly different at P < 0.05. Error bars represent standard error of
the mean.
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Figure 3: Inhibition curve of Colletotrichum coccodes spore germination by Nt-
phylloplanins in tobacco leaf wash
C. coccodes spores were treated with tobacco leaf wash containing various concentrations of Nt-
phylloplanins and incubated for a period of 12 hours. Each data point represents an average value
from three biological replicates.
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Figure 4. Inhibition of Aspergillus fumigatus spore germination after treatment with Nt-
phylloplanins in tobacco leaf wash
Spores of A. fumigatus were subjected to various treatments: GMM control, proteinase-K treated
Nt-phylloplanins, Nt-phylloplanins (1 mg mL-1
), and incubated in microfluidic channels (1.5 ×
103
spores per channel). Each point represents an average of three biological replicates with 200
spores assessed per each treatment. The data was arcsine transformed prior to being analyzed
using ANOVA, with means separated using Fisher’s LSD in R 2.15.3. Treatments with the same
lowercase letters are not significantly different at P < 0.05. Error bars represent standard error of
the mean.
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Figure 5. Transient inhibition of Fusarium sambucinum spore germination by Nt-
phylloplanins in tobacco leaf wash
A) F. sambucinum spores germinated in water controls at 12 hours post inoculation (hpi). B)
Non-germination of Nt-phylloplanin treated F. sambucinum spores at 12 hpi. C) Extensive
hyphal growth observed after treating F. sambucinum spores with Nt-phylloplanins at 24 hpi.
Bar = 20 µm.
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CHAPTER 3
A draft genome sequence reveals Helminthosporium solani’s arsenal for cell wall
degradation
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ABSTRACT
Helminthosporium solani, is a fungal pathogen belonging to the family Massarinaceae. It causes
blemishes on potato tubers, affecting processing and fresh market trade. Despite its world-wide
distribution, little is known about the biology of H. solani. Here we report the generation of a
draft genome sequence of H. solani with an estimated genome size of ~35 megabases, which is
also the first reference genome within the family Massarinaceae. We identified a large suite of
genes in the H. solani genome that encode putative cell wall degrading enzymes. Based on
comparison with other known genomes, we speculate that H. solani is a hemi-biotroph or
necrotroph. The presence of a large number of genes in the glycoside hydrolase (GH) 10 and 43
families, which aid in the hydrolysis of glucoronoarabinoxylan, also suggests that H. solani may
be able to survive on grass hosts or debris and indicates the need to re-examine the life cycle and
host range of this pathogen. The H. solani draft genome will be a valuable resource for
phylogenetic and pathogenicity studies on this pathogen.
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INTRODUCTION
The fungus Helminthosporium solani causes silver scurf disease of potatoes and has a worldwide
distribution (Errampalli et al. 2001). Potato tubers with silver scurf have grey lesions which are
localized to the periderm and appear silvery when moistened. Under favorable storage
conditions, fungal sporulation makes the tubers appear black and sooty. Loss of pigmentation
occurs in severely diseased tubers, especially on red-skinned cultivars (Jellis and Taylor, 1977),
reducing their fresh market value. Silver scurf also increases the permeability of the tuber skin to
water vapor, resulting in tuber shrinkage and weight loss (Burke, 1938; Jellis and Taylor, 1974).
Silver scurf negatively affects the processing market as chips made from severely infected tubers
have unacceptable black burnt edges (Holley and Kawchuck, 1996). Although silver scurf does
not affect tuber yields (Read and Hide, 1984; Cunha and Rizzo, 2004), the disease has been
gaining economic importance in the past three decades due to the quality standards demanded by
processing and fresh market consumers as well as the emergence of H. solani isolates resistant to
the broad-spectrum thiabendazole fungicides and thiophanate-methyl fungicides used to control
this disease (Errampalli et al. 2001; Geary et al. 2007).
Silver scurf spreads both in the field and during storage. Diseased seed tubers (Jellis and Taylor,
1977) and infested soil (Merida and Loria, 1994) act as the primary source of inoculum. H.
solani has a very narrow host range comprised of tuber-bearing Solanum species and the weed
species Solanum elaeagnifolium (Burke, 1938; Kamara and Huguelet, 1972; Sethuraman et al.
1997; Rodriguez et al. 1995). H. solani is an anamorphic fungus that can colonize senescent leaf
tissues of alfalfa, sorghum, rye, oats, corn, and wheat in vitro, and can also survive in dry soil of
potato stores for a period of five months (Merida and Loria, 1994; Carnegie et al. 1996). There
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are no known specialized over-wintering structures, however conidia may serve this purpose
(Kamara and Huguelet, 1972). Other aspects of the disease cycle such as pathogen spread from
mother tuber to daughter tubers and its overwintering phases are poorly understood.
Furthermore, genomic information for H. solani is scarce, as evidenced by the presence of only
36 nucleotide sequences of beta-tubulin and ribosomal genes in the National Center for
Biotechnology Information (NCBI). There is no information available regarding H. solani genes
required for virulence or pathogenesis. In this sense, H. solani is an understudied pathogen.
Helminthosporium solani is an ascomycete belonging to the order Pleosporales, the largest order
in the class Dothideomycetes (Kirk et al. 2008). Analyses of ribosomal DNA and ITS sequences
reveal that the closest relatives of H. solani is the wood saprophyte Helminthosporium velutinum
and Saccharicola bicolor, causal agent of leaf scorch disease of sugarcane (Olivier et al. 2000).
Currently H. solani is placed in the family Massarinaceae (Eriksson and Hawksworth, 2003).
With the exception of H. solani and S. bicolor, the majority of the members of Massarinaceae are
saprophytes (Liew et al. 2002; Zhang et al. 2009). Wood saprophytes possess multiple cell wall
degrading enzymes capable of hydrolyzing cellulose and hemicelluloses (Bucher et al. 2004;
Simonis et al. 2008; Sin et al. 2002). In a recent study, a large scale screening for hydrolytic
activity of pathogenic and non-pathogenic fungi has also revealed that plant pathogens are
promising sources for cell wall degrading enzymes (King et al. 2011). With the advent of high
throughput sequencing techniques, the cost of sequencing has dropped considerably. This has led
to the description of several fungal genomes with genes predicted to encode a number of
carbohydrate-active enzymes (Hane et al. 2007; Ellwood et al. 2010; de Wit et al. 2012; Islam et
al. 2012; Manning et al. 2013). Based on H.solani’s phylogeny and its growth in association with
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plants, we hypothesized that H. solani encodes genes involved in cell wall degradation. The
purpose of this study was to generate a draft genome of H. solani to gain insights into its biology
and identify candidate virulence factors that are important for colonization of potato tubers.
Materials and Methods:
Helminthosporium solani genomic DNA isolation
The H. solani isolate B-AC-16A was recovered from a diseased potato tuber. Pure culture of H.
solani was black in color when grown on V8 medium, conidiophores were septate and
unbranched, and conidia were dark brown obclavate (7-8 × 18-64 µm) with up to 8 septa (Goth
and Webb, 1983; Loria and Secor, 2001, Rivera-Varas et al. 2007). The single spore isolate of H.
solani was grown in minimal medium (Olivier and Loria, 1998) for 2 weeks in the dark at 24oC
on a rotary shaker. The fungal mycelia were then separated from the medium and placed
between paper towels to squeeze out the remaining medium. The mycelia were then lyophilized
overnight and DNA was extracted using phenol-chloroform-isoamyl alcohol (Bok and Keller
2012).
Sequencing of the H. solani genome and sequence assembly
H. solani genomic DNA was used for paired-end pyrosequencing on a full plate Roche (454)
Genome Sequencer FLX+ with Titanium chemistry. H. solani genomic DNA was also sequenced
in one lane of a 1X 100 bp flow cell on an Illumina HiSeq2000. We obtained approximately 1
million reads from the 454 sequencer and approximately 175 million reads from the Hiseq2000.
The genome was assembled from a combination of 454 and Illumina data using the Roche GS
De Novo Assembler (Newbler) Version 2.6 (Marqulies et al. 2005) The following parameters
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were used in the assembly- seed step 12, seed length 16, seed count 1, min overlap length 40,
minimum overlap identity 90, alignment identity score 2, alignment difference score –3. We used
all of the 454 data in the assembly. The first 25 million reads from the Illumina data were used in
the assembly to represent 50 X coverage of the genome assuming a genome size of 50 Mb.
Retrieval and gene modeling for cell wall degrading enzymes
Following genome assembly we retained all contigs larger than 100 bp and predicted genes using
the self-training gene prediction program GenMark-ES Version 2.3c (Borodovsky and Lomsadze
2011). The search for cell wall degradation-associated genes in the H. solani genome was also
performed with complete protein sequences from other ascomycetes such as Leptosphaeria
maculans and Cochliobolus heterostrophus. The sequences were used as query in a stand-alone
tblastn search with H. solani genome as the database (Tao, 2010). Preliminary annotations of
proteins sequences obtained from translation of the genes predicted by GeneMark were obtained
using InterProScan Version 4.8 (Zdobnov and Apweiler 2001) including membership in protein
families and Gene Ontology (Ashburner et al. 2000) categories.
Phylogenetic analyses:
Phylogenetic analyses of 16 putative cell wall degrading genes were conducted using MEGA
version 5.05 (Tamura et al. 2011). The amino acid sequences were aligned using Clustal W with
default parameters (Thompson et al. 1994). A neighbor-joining tree was constructed for each
sequence group and a bootstrap support for internal branches was estimated from 1000 pseudo-
replicates.
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RESULTS AND DISCUSSION
Helminthosporium solani is an understudied pathogen with few genomic resources. Here, we
report a draft genome sequence of H. solani, which is also the first reference genome within the
family Massarinaceae. The H. solani genome assembly consisted of 35.0 Mb in 9,248 contigs,
with about 34 Mb in contigs bigger than 500 bp (2939 contigs) that are contained in 486
scaffolds. The N50 scaffold size is 168,899 bp with an N50 contig size of 34,146 bp. The
assembly contained 874,415 Roche 454 reads and 19,976,474 Illumina reads.
We searched the H. solani contig sequences with the complete complement of predicted protein
sequences from the genomes of two closely related Ascomycete species Leptoshaeria maculans
and Cochiobolus heterostrophus C5 using tblastn (Altschul et al. 1990) with a threshold evalue
of 0.00001. The 36.36 Mb C. heterostrophus genome encodes13,336 predicted proteins and has
10,159 putative homologs in the H. solani genome. The 44.89 Mb L. maculans genome encodes
12,469 predicted proteins and has 8,459 putative homologs in the H. solani genome. The average
percentage identity between H. solani proteins and their homologs in either genome is about
60%. De novo gene prediction using GeneMark-ES revealed 14,862 predicted protein-coding
genes from the H. solani contig sequences. Approximately two-thirds of these predicted proteins
have some matching entry from the InterProScan searches.
H. solani genome sequence provided insight into several putative virulence genes, including
genes homologous to plant cell wall degrading enzymes, non-ribosomal peptide synthetases, and
polyketide synthetases. However, we focused our analysis on the plant cell wall degrading
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enzymes because of the large base of knowledge about the enzyme function and enzyme content
of plant-associated microbes.
H. solani encodes numerous plant cell wall degrading enzymes:
The complex polysaccharides of plant cell walls are hydrolyzed by a large variety of
carbohydrate-active enzymes (CAZymes) which are classified into families such as glycoside
hydrolases (GH), polysaccharide lyases, carbohydrate esterases and carbohydrate-binding
modules (Cantarel et al. 2008). Because the lifestyle of a fungus is generally well correlated with
CAZyme content, we compared the number of genes predicted to encode celluloytic and
hemicellulolytic enzymes in the genome of H. solani to a subset of ascomycetes that included a
saprotroph (Trichoderma reesei), necrotrophs (Fusarium graminearum, Sclerotinia sclerotiorum,
Botrytis cinerea), a hemibiotroph (Magnaporthe grisea) and a biotroph (Blumeria graminis). We
observed that H. solani has at least 71 putative genes encoding GH enzymes, which was
comparable to genomes of hemibiotrophs and necrotrophs (Table 1 and 2). For instance,
biotrophs generally possess fewer cell wall degrading enzymes than necrotrophs and
hemibiotrophs to evade plant defenses (deWit et al. 2012; Amselem et al. 2011; Goodwin et al.
2011). Based on GH enzyme content, H. solani likely has a hemibiotrophic or necrotrophic
lifestyle, which is in accordance with earlier histological studies where necrosis was observed in
both H. solani colonized and neighboring potato tuber cells (Martinez et al. 2004).
No evidence of horizontal gene transfer among putative cell wall degrading genes:
H. solani belongs to a family that consists mainly of saprophytes, suggesting that acquisition of
novel genes or gene duplication resulted in H. solani gaining the ability to colonize potato tuber
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surfaces. Because it is a tuber- and soil-borne pathogen and because soil is a complex
environment hosting diverse organisms, there are likely to be ample sources for horizontal gene
transfer for H. solani (Wenzl et al. 2005; Gorfer et al. 2011). Hence, we constructed phylogenetic
trees, a primary method for identification of horizontal gene transfer event, with a subset of 16
H. solani plant cell wall degrading gene homologs (Richards et al. 2011). However, all of the GH
enzyme gene sequence trees were congruent with the H. solani phylogeny (data not shown).
Hemicellulase enzyme content suggests that H. solani can degrade monocot plant cell walls:
Hemicelluloses are the second most abundant group of polysaccharides in the plant cell wall (van
den Brink and de Vries 2011). Xyloglucans and arabinoxylans constitute two major families of
hemicelluloses whose composition varies among plants. For instance, xyloglucans are
predominant in dicots and nongraminaceous monocots whereas arabinoxylans are primarily
present in monocot grasses in the family Poaceae (Albersheim et al. 2011). H. solani is a
pathogen on potato, which is a dicot plant. For this reason, we were surprised by the presence of
a single gene likely involved in xyloglucan degradation (family GH16) which was in contrast to
other fungi compared in this study and another eighteen fungal species in the class
Dothideomycetes (Ohm et al. 2012). This suggests that H. solani does not extensively digest
dicot hemicelluloses, which may explain, in part, its narrow host range. But the fact that H.
solani is well adapted to potato suggests that the pathogen may possess a different strategy. A
caveat to this analysis is that the gene number is not predictive of enzyme expression or activity,
as evidenced with T. reesei which produces highly efficient enzymes with a small number of
genes (Martinez et al. 2008).
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Glucuronoarabinoxylan, composed of a xylose backbone with arabinose and glucuronic acid side
chains is the major hemicellulose component of grass cell walls (van den Brink and de Vries
2011; Vogel 2008). Degradation of glucuronoarabionxylan is aided by endoxylanases (GH10)
and arabinofuranosidases (GH43). Fusarium graminearum and Magnaporthe grisea, which are
predominantly pathogens on grass hosts possesses 15 and 16 genes respectively that encode
GH10 and GH43 enzymes. It is interesting to note that H. solani possesses 13 GH10 and GH43
homologs, which is comparable with grass host pathogens. Although earlier in vitro studies
showed H. solani’s ability to colonize senescent grass leaf tissues (Merida and Loria 1994), our
genomic data suggest that H. solani may have a greater ability to survive on monocots than
dicots, which indicates the need to re-examine the life cycle and host range of H. solani. Further
studies should also focus on developing a transformation method for H. solani which would
allow functional analyses of candidate genes associated with cell wall degradation and elucidate
their role during pathogenesis.
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Table 1: Comparison of the number of cellulose degrading genes among ascomycete fungi
Fungal species
Lifestyle
Glycoside Hydrolase (GH) family
GH 1 GH 3 GH 5 GH 6 GH 7 GH12 GH45 GH61 GH74 Total
Trichoderma reesei a Saprotroph 2 8 3 1 2 1 1 3 1 22
Botrytis cinerea a Necrotroph 5 14 5 1 3 1 2 10 0 41
Sclerotinia sclerotiorum a Necrotroph 1 12 5 1 3 2 2 9 0 35
Fusarium graminearum a Necrotroph 3 17 3 1 2 2 1 13 1 43
Magnaporthe grisea a Hemi-biotroph 2 15 6 3 7 3 1 22 1 60
Helminthosporium solani b ? 3 15 7 2 5 1 1 17 0 51
Blumeria graminis f.sp. hordei a Biotroph 0 0 0 0 0 0 0 2 0 2
a Gibson et al. (2011)
b Data from this study
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Table 2: Comparison of the number of hemicelluloses degrading genes among ascomycete fungi
Fungal species
Lifestyle
Glycoside Hydrolase (GH) family
GH 10 GH 11 GH 16 GH 43 GH 51 GH 54 GH 62 GH 81 Total
Trichoderma reesei a Saprotroph 1 3 12 2 0 2 1 2 23
Botrytis cinerea a Necrotroph 2 2 11 4 3 1 1 2 26
Sclerotinia sclerotiorum a Necrotroph 2 3 10 3 2 1 0 1 22
Fusarium graminearum a Necrotroph 4 3 14 11 2 1 1 1 37
Magnaporthe grisea a Hemi-biotroph 6 5 10 10 3 1 4 2 41
Helminthosporium solani b ? 4 4 1 9 0 0 1 1 20
Blumeria graminis f.sp. hordei a Biotroph 0 0 3 0 0 0 0 1 4
a Gibson et al. (2011)
b Data from this study
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SYNTHESIS
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Silver scurf (causal agent: Helminthosporium solani) and black dot (causal agent: Colletotrichum
coccodes) are two potato surface blemish diseases that are frequently misidentified by potato
growers and whose importance in organic farming is poorly understood. Organic potato
production is unique from conventional production in terms of both agronomic and plant
protection practices. Synthetic pesticides are not permitted in organic farming which leaves the
crop vulnerable to various diseases. Very little on-farm research has been done to learn about the
importance of these diseases in organic potato farming. This dissertation is the first to
demonstrate that both H. solani and C. coccodes are highly prevalent in organic potato farms in
Wisconsin. This research also suggests that the visual assessment of tubers at harvest for silver
scurf and black dot may be misleading. Possible future work should focus on organic control
measures to prevent disease development during the growing season and in storage. With the
expanding fresh market for organic potatoes demanding blemish free potatoes, we believe there
is an immediate need for research on tuber surface defects.
In our experiments, we also observed H. solani and C. coccodes together on about 65% of the
asymptomatic and asymptomatic tubers assessed. Despite having different lifestyles, co-
occurrence of both pathogens in such a high frequency raises interesting ecological questions. At
one extreme, H. solani has a known host range limited to tuber-bearing Solanum species, some
saprophytic ability, and no known special over-wintering structures. At the other extreme, C.
coccodes has a broad host range, the ability to survive in crop debris, produce sclerotia, and
colonize the same host tissue as H. solani. Generally speaking, two species sharing a single niche
cannot co-exist for a long period of time, which raises questions about the nutrients that the
pathogens sequester from the tuber, the time of infection and the growth patterns affecting their
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reproductive potential. We have conducted preliminary experiments looking at carbon and
nitrogen utilization patterns of these pathogens, but more experiments need to be performed to
dissect this association and explore plausible interactions between these two pathogens.
As all current potato cultivars are susceptible to silver scurf, we screened potato cultivars and
lines for resistance to silver scurf. Mature tuber incubation assays performed to screen for
resistance were unsuccessful because 90% of the tubers were lost before the end of the assay due
to tuber decay pathogens. As an alternative to mature tubers, we explored the utilization of
hydroponically grown minitubers, which minimized tuber losses to decay pathogens. This work
demonstrated that minitubers can be successfully employed to screen for resistance to silver
scurf. This dissertation has identified a diploid inter-specific breeding line (C287) with
consistently low H. solani sporulation. C287 is known to have resistance to other significant
potato diseases such as verticillium wilt, common scab, and black scurf and findings from this
study suggest that C287 possesses partial resistance to silver scurf as well. Further work should
be focused on field and storage trials to evaluate silver scurf resistance of C287 at multiple
locations.
H. solani is a little studied pathogen with many unanswered questions regarding its biology and
lifecycle. For instance, little information is available regarding the genes involved in the
infection process, overwintering phase of the pathogen, and virulence factors. In order to address
some of these questions, we generated a draft genome sequence of H. solani, which is the first
reference genome in the family Massarinaceae and will be made available for public research.
This research led to the identification of a large suite of genes in the H. solani genome that
encode putative cell wall degrading enzymes. Based on comparison with other known genomes,
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we speculate that H. solani is a hemi-biotroph or necrotroph. Also, the presence of a large
number of genes in Glycoside Hydrolase (GH) 10 and 43 families, which aid in the hydrolysis of
glucoronoarabinoxylan, a major component of grass hemicelluloses, suggests that H. solani may
be able to survive on grass hosts and indicates the need to re-examine the host range of this
pathogen.
We believe draft genomic data will shed light on previously unknown aspects of the biology of
H. solani. In silico genomic analysis indicates H. solani may be a promising target for
identifying novel cell wall degrading enzymes for use in the biofuel industry. To this end, future
research should focus on performing enzymatic assays to obtain more detailed characterization
of the secretome of H. solani. It would also be useful to develop a new primer set for detecting
H. solani because the current nested primer set is time and cost prohibitive for use in diagnostics.
Analysis of the genome of H. solani also revealed several genes involved in secondary
metabolite production. Although our attempts to transform H. solani by biolistic transfection and
Agrobacterium-mediated transformation were unsuccessful, we strongly believe further work on
the development of a transformation procedure that would aid in performing functional
genomics.
Finally, we explored the antifungal properties of tobacco phylloplane (leaf surface) proteins to
elucidate their potential effects on pathogen development, and determine if they can be utilized
in post-harvest disease control. Due to the large volumes of water used for collecting
phylloplanins from leaf aerial surfaces, the samples first needed to be lyophilized to yield a
concentrated product highly enriched in protein. This aspect of sample recovery limited the
volume of phylloplanins and was a significant bottleneck in our ability to perform large numbers
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of assays. For this reason, we decided to use a microfluidic approach that has been successfully
used earlier for applications in drug discovery, cell culturing, and characterization of fungal
secondary metabolites. Using microfluidic channels, we were able to perform tens of assays
simultaneously with low volumes of phylloplanins, and demonstrated that this assay will be
broadly useful in further large-scale screening of natural product bioactives against
microorganisms.
Tobacco phylloplanins strongly inhibited spore germination of Colletotrichum coccodes and
Aspergillus fumigatus, transiently inhibited Fusarium sambucinum spore germination, and were
inactive against Alternaria solani and Verticillium albo-atrum, indicating a spectrum of pathogen
sensitivity that was not previously known. Most importantly, inhibition of spore germination of
the human pathogen A. fumigatus by tobacco phylloplanins reveals a new venue for medicinal
antifungal drug discovery that has not been explored previously. Future work should focus on
mass scale production of tobacco phylloplanins by expressing them in a heterologus system and
testing for inhibitory effects on additional species of Aspergillus that are human pathogens.
Lastly because the mechanism of action of phylloplanins remains unknown, further research
efforts should be directed at exploring the targets of phylloplanins.