EVALUATION OF STRATEGIES FOR MANAGING SOILBORNE PATHOGENS OF
ASPARAGUS IN MICHIGAN
By
Chelsea Woods

A THESIS
Submitted to
Michigan State University
in partial fulfillment of the requirements
for the degree of
Plant Pathology – Master of Science
2014

ABSTRACT
EVALUATION OF STRATEGIES FOR MANAGING SOILBORNE PATHOGENS OF
ASPARAGUS IN MICHIGAN
By
Chelsea Woods
In 2012, Michigan produced 8.8 million kg of asparagus spears for fresh market and
processing worth $17.3 million. Asparagus is a perennial crop, with fields remaining profitable
for 12 to 15 years. Fusarium crown and root rot (caused by Fusarium oxysporum f. sp. asparagi
and F. proliferatum) and Phytophthora spear and root rot (caused by Phytophthora asparagi)
contribute to early decline, and decrease the vigor of young stands in fields previously cropped
with asparagus. Nine isolates of P. asparagi were tested for virulence using asparagus spears of
five commercial cultivars and asparagus seedlings of three cultivars. Significant differences in
isolate virulence and host susceptibility were observed on spears, but not on seedlings. Seedlings
of 18 and 11 cultigens were screened for resistance to P. asparagi and Fusarium spp.,
respectively. While all cultigens were susceptible to the pathogens, significant differences in the
degree of susceptibility were detected. Pathogen virulence varied, and the interaction between
pathogen and cultigen significantly affected root rot severity. Fungicides and biocontrol agents
were screened for efficacy as seed treatments against Fusarium spp. and P. asparagi in vitro.
Two fungicides and one biocontrol agent increased germination of seeds inoculated with P.
asparagi. Disease incidence was reduced when one biocontrol agent limited P. asparagi and
Fusarium spp. growth. This is the first evaluation of modern hybrids and seed treatments for
management of both Fusarium spp. and P. asparagi. Identifying resistant cultivars and
effective seed treatments is important for the development of an integrated pest management
system to control soilborne pathogens for Michigan asparagus grower

ACKNOWLEDGMENTS

I would like to thank my advisor, Dr. Mary Hausbeck, for her guidance, patience, and
support. I would also like to thank my committee members, Dr. Raymond Hammerschmidt and
Dr. Mathieu Ngouajio for their encouragement and advise. All of the Hausbeck members,
including former lab member Dr. Leah Granke, have generously volunteered their time and
expertise to help me finish my research and writing. Dr. Andrew Jarosz and Moslem Ladoni
have been tremendously helpful with statistical interpretations. Thanks to John Bakker and The
Oomen Brothers for providing fresh asparagus spears, and to Aspara Pacific, Bejo, Eden
Brothers, Rutgers University, University of Guelph, Vilmorin, and Walker Brothers for
providing asparagus seeds.
Thanks to all my family members and friends who have provided love and support and
occasional dinners

iii

TABLE OF CONTENTS

LIST OF TABLES ........................................................................................................................ vi
LIST OF FIGURES ...................................................................................................................... vii
LITERATURE REVIEW ............................................................................................................. 1
ASPARAGUS .................................................................................................................... 2
FUSARIUM CROWN AND ROOT ROT ......................................................................... 8
PHYTOPHTHORA SPEAR AND ROOT ROT ............................................................... 11
INTEGRATED PEST MANAGEMENT .......................................................................... 14
LITERATURE CITED ...................................................................................................... 19
CHAPTER I: VIRULENCE OF PHYTOPHTHORA ASPARAGI, SUSCEPTIBILITY OF
ASPARAGUS CULTIGENS TO SPEAR AND ROOT ROT, AND THE EFFICACY OF
SEED TREATMENTS ................................................................................................................. 25
ABSTRACT ....................................................................................................................... 26
INTRODUCTION ............................................................................................................. 27
MATERIALS AND METHODS ....................................................................................... 29
Isolates and inoculum preparation ................................................................................... 29
Susceptibility of asparagus cultivars to spear rot ..............................................................30
Seedling inoculation and incubation .................................................................................31
Virulence of isolates on seedling roots............................................................................ 33
Susceptibility of cultivars and breeding lines to seedling root rot ....................................33
Efficacy of fungicides and biocontrol agents as seed treatments ................................... 33
Pathogen isolation ............................................................................................................36
Statistical analysis .............................................................................................................36
RESULTS .......................................................................................................................... 38
Susceptibility of asparagus cultivars to spear rot ..............................................................38
Virulence of isolates on seedling roots............................................................................ 39
Susceptibility of cultivars and breeding lines to seedling root rot .................................. 39
Efficacy of fungicides and biocontrol agents as seed treatments .......................................
40
DISCUSSION .................................................................................................................... 49
LITERATURE CITED ...................................................................................................... 52
CHAPTER II: THE EFFECT OF CULTIGENS AND FUNGICIDES AND BIOLOGICAL
AGENTS ON FUSARIUM ROOT ROT OF ASPARAGUS....................................................... 56
ABSTRACT ....................................................................................................................... 57
INTRODUCTION ............................................................................................................. 58
MATERIALS AND METHODS ....................................................................................... 61
Fusarium spp. and inoculum preparation ........................................................................ 61
Seedling inoculation and incubation ............................................................................... 61
Susceptibility of cultivars and breeding lines to seedling root rot .................................. 62
Efficacy of fungicides and biocontrol agents as seed treatments .................................... 63

iv

Pathogen isolation .............................................................................................................64
Statistical analysis ........................................................................................................... 65
RESULTS .......................................................................................................................... 67
Susceptibility of cultivars and breeding lines to seedling root rot .................................. 67
Efficacy of fungicides and biocontrol agents as seed treatments .................................... 68
DISCUSSION .................................................................................................................... 72
LITERATURE CITED ...................................................................................................... 74

v

LIST OF TABLES

Table 1.1. Disease classes of inoculated asparagus seedlings used to calculate disease
severity index (DSI) ...................................................................................................................... 32
Table 1.2. Asparagus cultivars and breeding lines tested for susceptibility to Phytophthora
asparagi ........................................................................................................................................ 34
Table 1.3. Fungicides and biocontrol agents tested for efficacy against Phytophthora
asparagi ........................................................................................................................................ 34
Table 1.4. Scale of disease ratings on successfully germinated asparagus seedlings
inoculated with Phytophthora asparagi........................................................................................ 37
Table 1.5. Average disease severity index (DSI) and area under the disease progress curve
(AUDPC) of asparagus seedlings infected with Phytophthora asparagi isolates at 28 days
post inoculation ............................................................................................................................. 45
Table 1.6. The effect of Phytophthora asparagi isolate and seed treatment on germination
and disease incidence. ................................................................................................................... 46
Table 2.1. Asparagus cultivars and breeding lines tested for susceptibility to Fusarium
oxysporum f. sp. asparagi and F. proliferatum ............................................................................ 62
Table 2.2. Fungicides and biocontrol agents tested for efficacy against Fusarium
oxysporum f. sp. asparagi and F. proliferatum ............................................................................ 64
Table 2.3. Scale of disease ratings on successfully germinated asparagus seedlings
inoculated with Fusarium oxysporum f. sp. asparagi and F. proliferatum .................................. 64
Table 2.4. Average disease severity (%) of asparagus seedlings infected with Fusarium
oxysporum f. sp. asparagi and F. proliferatum at 28 days post inoculation................................ 69
Table 2.5. Area under the disease progress curve (AUDPC) of asparagus seedlings infected
with Fusarium oxysporum f. sp. asparagi and F. proliferatum .................................................... 69
Table 2.6. The effect of Fusarium spp. and seed treatment on germination and disease
incidence. ...................................................................................................................................... 70

vi

LIST OF FIGURES

Figure 1.1. The Phytophthora symptoms observed on inoculated asparagus seedling root
systems receiving the rating of a 0 to 4......................................................................................... 32
Figure 1.2. Asparagus seeds treated with fungicides and biocontrol agents on V8 agar
plates inoculated with Phytophthora asparagi isolates SP316 (A) and SP326 (B), 3 days
after planting ................................................................................................................................ 35
Figure 1.3. Symptoms of Phytophthora asparagi observed on germinated asparagus
seedlings. (A) 0 = no disease symptoms, (B) 1 < 50% radicle tissue water-soaked, (C) 2 >
50% radicle tissue water-soaked, and (D) 3 > 50% radicle tissue water-soaked with mycelia
growth ........................................................................................................................................... 37
Figure 1.4. Effect of cultivar (A) and Phytophthora asparagi isolate (B) on mean lesion
area on detached asparagus spears 7 days after inoculation. Bars with common letters are
not significantly different, LSD P=0.05 (A) and Tukey’s P=0.05 (B) ......................................... 41
Figure 1.5. Effect of Phytophthora asparagi isolates on disease severity index (DSI) where
0 = no disease and 100 = all seedlings roots with > 50% of their roots water-soaked (A) and
area under the disease progress curve (AUDPC) values (B). Bars with common letters are
not significantly different, LSD P=0.05 ....................................................................................... 42
Figure 1.6. Effect of cultivars on disease severity index (DSI) where 0 = no disease and
100 = all seedlings with > 50% of their roots water-soaked (A) and area under the disease
progress curve (AUDPC) values (B). Bars with common letters are not significantly
different, LSD P=0.05 .................................................................................................................. 43
Figure 1.7. The distribution of disease rating scores of germinated seedlings inoculated
with Phytophthora asparagi SP316 and treated with fungicides and biocontrol agents. ............. 47
Figure 1.8. The distribution of disease rating scores of germinated seedlings inoculated
with Phytophthora asparagi SP326 and treated with fungicides and biocontrol agents. ............. 48
Figure 2.1. Asparagus seeds treated with fungicides and biocontrol agents on V8 agar
plates inoculated with Fusarium oxysporum f. sp. asparagi (A) and F. proliferatum (B), 3
days after planting. ........................................................................................................................ 65
Figure 2.2. Fusarium symptoms observed on germinated asparagus seedlings. (A) 0 = no
disease symptoms, (B) 1 = lesions present on <50% radicle tissue, (C) 2 = lesions present
on >50% radicle tissue, and (D) 3 = all seed tissues covered in lesions ....................................... 66
Figure 2.3. The distribution of disease rating scores of germinated seedlings inoculated
with Fusarium oxysporum f. sp. asparagi and treated with fungicides and biocontrol agents .... 71

vii

LITERATURE REVIEW

1

ASPARAGUS
Asparagus is a dioecious, herbaceous perennial crop. Asparagus, formerly included in
the family Liliaceae (Drost, 1997), is the only genus of family Asparagaceae (Batchelor and
Scott, 2006; Kubitzki and Randall, 1990; The Angiosperm Phylogeny Group, 2009). The
number of species is estimated to be 170-300 (Kubitzki and Randall, 1990), but Asparagus
officinalis Linnaeus is the only species grown as an edible crop. Asparagus cultivation began in
Europe by the Romans and Greeks, and was produced for culinary and medicinal purposes
(Drost, 1997; Kubitzki and Randall, 1990; Schofield, 1946).
In 2009, the production of asparagus worldwide encompassed 195,819 hectares across 62
countries, a decrease of 30,176 ha since 2005. The major asparagus producing countries of
China, the United States, Spain, the Philippines, Australia, and New Zealand, have experienced
losses of production areas, while Peru and Mexico have increased production areas (Benson,
2012). Countries in the Southern Hemisphere are able to produce spears 12 months each year.
Peru and Mexico export nearly all of their asparagus in order to meet consumption demands in
regions such as the U.S. where the harvest season is relatively short (Benson, 2012; Elmer,
2001b).
Michigan ranks third in the U.S. for asparagus production, following California and
Washington. In 2012, Michigan produced 8.8 million kg of spears, with a total value of $17.3
million, for fresh market and processing (Anonymous, 2013). Over the last decade, the
production area in Michigan has decreased by 2,104 ha (Anonymous, 2013; Kleweno and
Matthews, 2002). All production in Michigan is used for domestic consumption, and
approximately 72% is utilized as canned or frozen (Benson, 2012).

2

The asparagus crown consists of a rhizome, adventitious or storage roots, and lateral or
feeder roots. Buds grow on the sympoidal rhizome in a bud cluster; individual shoots arise from
separate buds and new storage roots originate at the base of actively growing buds (Drost, 1997;
Kubitzki and Randall, 1990). Feeder roots grow from young tissue on the storage roots, and are
the primary nutrient and water absorbing structures (Drost, 1997). Fleshy storage roots contain
reserves of soluble carbohydrates, which are depleted during the harvest season (Drost, 1997;
Shelton and Lacy, 1980). Shoots emerge from the soil during the spring as succulent spears,
which mature into extensively branched ferns up to 2 m tall (Drost, 1997). A fern stand is
comprised of a primary stalk, secondary branches and needle-like cladophylls. Although the
entire fern tissue is able to undergo photosynthesis, the cladophylls are the primary
photosynthetic structures of asparagus plants.
Open-pollinated cultivars are dioecious and have substantial genetic variability, which
can limit the average vigor and yield of a field compared to fields planted with homogenous,
high performance cultivars (Stephens and Elmer, 1988; Zandstra et al., 1992). Male plants have
higher marketable yield and number of spears, earlier season production, reduced time between
emergence of individual spears, longer production duration, and lower mortality rate compared
to female plants (Drost, 1997). All-male hybrids from New Jersey (i.e. ‘Jersey Giant’) have been
developed to homogenize and increase average vigor and yield within each cultivar (Ellison and
Kinelski, 1985). ‘Jersey Giant’ and ‘Millennium,’ a cultivar developed in Canada, are
commonly grown in Michigan (Rodriguez Salamanca, 2010).
Asparagus requires well-drained soils; fields that retain standing water for prolonged
periods after rain events should be avoided (Sanders, 2001; Zandstra et al., 1992). Although
asparagus can tolerate soil with a pH between 5.0 and 7.5 (Zandstra et al., 1992), the optimum

3

range for healthy growth is 6.2 to 6.8 (Sanders, 2001). Soil pH below 6.0 favors Fusarium
infection (Zandstra et al., 1992).
Asparagus crowns are produced from seed that is planted in outdoor crown nurseries. To
limit soilborne pathogens, the soil in the seedbed should be fumigated prior to planting (Saude et
al., 2008). During preparation of the seedbed, fertilizer is applied such that approximately 224
kg each of P2O5 and K2O is available per hectare. Once the seedlings are 15 to 20 cm high,
approximately 56 kg N per hectare is broadcast along the rows. In Michigan, asparagus seeds
are planted between mid April and mid May, and placed approximately 2.5 to 3.8 cm deep and 5
cm apart in rows that are spaced 61 to 76 cm apart. Approximately 5.6 kg of seed are used per
hectare (Zandstra et al., 1992). After seedling emergence, weeds must be controlled by hand
(Sandsted et al., 1985). Approximately one year after planting, crowns are dug using a potato
digger in the early spring before bud break. The crowns are stored in loose piles or in bulk boxes
at 1.7 to 7.2°C for up to 2 months until they can be planted in production fields (Zandstra et al.,
1992).
Production fields are prepared the year prior to planting crowns by applying lime to
achieve a pH of 6.8 (Sanders, 2001; Zandstra et al., 1992), using herbicides to kill weeds, and
planting cover crops (i.e. sudangrass or clover). The cover crops are plowed into the soil in the
fall, and winter wheat or rye is planted. In the spring, prior to planting the crowns, fertilizer may
be applied so that approximately 280 kg per hectare each of P2O5 and K2O is available in the soil.
The fertilizer is typically spread over the cover crop and plowed down 30 cm, to ensure that the
nutrients will be in the soil below the crowns. Up to 78 kg P2O5 per hectare is applied in the
furrows at the time of planting (Zandstra et al., 1992). Furrows are 20.3 to 25.4 cm deep with
middle-buster plows, with 1.2 to 1.5 m between rows. Crowns are spaced 30.5 cm apart and

4

covered with 2.5 to 5.0 cm of soil. During the first season of growth, soil is gradually cultivated
into the furrow until the soil is level (Zandstra et al., 1992).
Harvesting of spears typically begins the third year after planting the crowns (Drost,
1997; Elmer, 2001b; Zandstra et al., 1992). In Michigan, harvesting typically occurs from early
to mid May through mid to late June, depending on the weather and geographical location
(Benson, 2012; Zandstra et al., 1992). Shoots grow rapidly when temperatures reach 27°C and
can be harvested daily. Under cooler temperatures (< 21°C), the spear growth rate may be slow
enough to harvest every 2 to 3 days (Zandstra et al., 1992). In Michigan, spears are hand
harvested by snapping at the base once they have grown 18 to 22 cm in length (Elmer, 2001b;
Zandstra et al., 1992). Commercial recommendations suggest harvesting for 2 weeks during the
third year and then for 6 weeks during the fourth and subsequent years. Spear diameter
decreases as crown carbohydrate supplies are depleted over the harvest period and it is
recommended that harvesting conclude when the spears are < 0.95 cm in diameter (Zandstra et
al., 1992). Overharvesting newly established fields can decrease marketable yield up to 15% in
the following season (Shelton and Lacy, 1980).
When the harvest season ends, the shoots are allowed to grow and mature into fern. As
the fern photosynthetic surface area increases, carbohydrate levels in the storage roots increase
rapidly, peaking in mid to late August (Shelton and Lacy, 1980). The fern growth period is
critical for replenishment of carbohydrate reserves and fern vigor and impacts the size and
number of spears the following spring (Drost, 1997). As Michigan temperatures cool in mid to
late September, the fern senesces and eventually becomes defoliated. The dead fern is left
standing until the following spring, when it is mowed down with a rotary chopper (Zandstra et al.,
1992).

5

Asparagus plantings are a long-term investment and can remain profitable for up to 20
years if properly maintained (Elmer et al., 1996; Zandstra et al., 1992; Zandstra et al., 2013).
Therefore, it is important for growers to promote plant vigor and minimize factors that
negatively affect stand health. Asparagus is considered to be a drought-tolerant crop, since overirrigation causes reduced yield and provides favorable conditions for pathogens (Drost, 1997).
However, adequate irrigation during dry years is needed to maintain yield and stand vigor (Drost,
1997). Up to 84.1 kg N per hectare should be reapplied every year and potassium levels should
be replenished at least every 3 years according to soil test recommendations (Zandstra et al.,
1992).
Weeds and volunteer asparagus seedlings compete for sunlight, nutrients, and water, and
can reduce plant vigor and yield. Common perennial weeds in asparagus fields include
quackgrass (Elytrigia repens), milkweed (Asclepias syriaca), bindweed (Convolvulus arvensis),
horsenettle (Conyza canadensis), and Canada thistle (Cirsium arvense) (Zandstra et al., 1992;
Zandstra et al., 2013). Michigan growers have adopted a no-tillage system (Zandstra et al.,
1992) to prevent crown damage, which can reduce stand vigor and marketable yield (Wilcox-Lee
and Drost, 1991) and lead to infection by soilborne pathogens. The no-till practice is effective
against volunteer asparagus, since most seeds do not germinate on the soil surface. Herbicides
are applied to the rows in the spring before spear emergence and in the summer after harvest
(Zandstra et al., 1992). Caution must be used in choosing preemergence herbicides, since
herbicide damage to asparagus stands can cause significant yield losses (Zandstra et al., 2013).
Insects can cause spear and fern damage, leading to yield loss and opening the asparagus
stands to pathogen infection. Pests include cutworms (Peridroma saucia and Euxoa messoria),
common and spotted asparagus beetles (Crioceris asparagi and Crioceris duodecimpunctata),

6

asparagus aphids (Brachycorynella asparagi), and asparagus miners (Zandstra et al., 1992). The
asparagus miner, Ophiomyia simplex, causes severe damage to fern stems and is believed to
transmit Fusarium spp. between plants (Morrison et al., 2011).
With proper maintenance, a healthy asparagus field will remain profitable for 12 to 15
years or more (Elmer et al., 1996; Sanders, 2001; Zandstra et al., 1992). However, asparagus
early decline can shorten field lifespan by 5 to 10 years (Elmer et al., 1996). As plant vigor
decreases gradually over time, the size and number of marketable spears and fern shoots are
reduced (Keulder, 1999). The associated “replant problem” occurs when new stands fail to
become successfully established in a field previously cropped with asparagus (Elmer et al., 1996;
Falloon et al., 1991; Keulder, 1999; Morrison et al., 2011).
Asparagus decline can be exacerbated by many factors, including environmental stress,
allelopathic residues, and damage from tillage, herbicides, and insects (Keulder, 1999; Morrison
et al., 2011; Zandstra et al., 2013). However, several diseases are considered to be the primary
contributors of premature decline and replant failure (Elmer et al., 1996). Soilborne pathogens,
Fusarium spp. and Phytophthora spp., infect the roots and crowns, leading to direct loss of yield
during the season of infection as well as long term decline in stand vigor. Michigan fields
recently planted have productive lifespans approximately one-half as long as the lifespan of
fields planted in the 1950s (Morrison et al., 2011). Although crown damage from soilborne
diseases often appears to progress slowly, it is irreversible. In plots infested with Phytophthora
spp. that had not been treated for 2 years, adding metalaxyl to the soil in third season did not
increase yield (Falloon et al., 1985).
The foliar diseases rust (causal agent Puccinia asparagi De Candolle) and purple spot
(causal agent Stemphylium vesicarium Wallr.) affect the main stem, secondary branches, and

7

cladophylls. Severe infections can result in premature defoliation, reducing the ability of the fern
to replenish carbohydrate supplies in the crown. Smaller and fewer spears are produced the
spring following severe foliar disease, and crown health is compromised. If severe disease
pressure continues for more than one season, the losses increase each year (Hausbeck et al.,
1999; Johnson and Lunden, 1992).
Three viruses: asparagus virus I (AVI), asparagus virus II (AVII), and tobacco mosaic
virus (TMV) are known to infect asparagus in the U.S. AVII causes the most damage and is
seedborne. AVI and TMV are transmitted by aphids and thrips, but are only harmful to stands
when AVII is also present, and the effect is greater than AVII infection alone. Although the
viruses themselves do not cause symptoms, viral infection reduces stand vigor and increases
susceptibility to Fusarium crown and root rot (Elmer, 2001b; Elmer et al., 1996).

FUSARIUM CROWN AND ROOT ROT
The genus Fusarium belongs to the order Hypocreales and includes saprophytic and
parasitic members that have adapted to a wide range of hosts and environmental conditions.
Most pathogenic species are strictly soilborne, although others have modes of air dispersal and
can colonize aerial tissues of hosts. Crescent-shaped macroconidia are produced on sporodochia
and microconidia are produced in chains or clusters on phialides. Some species also produce
chlamydospores (Agrios, 2005; Burgess, 1981). Fusarium species are distinguished through
morphological, biological, and phylogenetic characteristics (Leslie et al., 2001). Morphological
species concepts consider shape and size of the macroconidia, microconidia, and
chlamydospores. Physiological characters such as growth rates, mycotoxins, and secondary
metabolites are useful for morphological taxonomy. The biological species concept is based on

8

reproductive isolation, although problems can arise with mating type allele frequencies,
homothallic species, asexual reproduction, or outcrossing. Amplified fragment length
polymorphism (AFLP) markers aid in identifying isolates belonging to the same biological
species. Phylogenetic species concepts utilize DNA sequences such as ribosomal internally
transcribed spacer (ITS) sequences. The “coalescent approach” combines the phylogenetic and
biological species concepts by using molecular markers and reproductive barriers to confirm
conspecific isolates (Leslie et al., 2001).
Fusarium crown and root rot symptoms on asparagus begin as reddish brown
discoloration in the rhizome, shoot, and storage root vascular tissues (Zandstra et al., 1992). As
the disease progresses, the feeder roots completely rot away, the storage roots become hollow,
and the fern turns yellow and senesces (Elmer et al., 1996; Zandstra et al., 1992).
Several Fusarium spp., including F. redolens Wollenw., F. solani (Mart.) Appel &
Wollenw. emend. Snyd. & Hans., and F. culmorum (W. G. Smith) Sacc. (Schreuder et al., 1995;
Wong and Jeffries, 2006) have been reported to infect asparagus in various growing regions in
the world. However, F. oxysporum Schlectend.:Fr. f. sp. asparagi Cohen & Heald and F.
proliferatum (Matushima) Niremburg are the primary causal agents of Fusarium crown and root
rot, which is considered to be the most economically limiting disease to asparagus crops
worldwide (Elmer, 2001a). Since it was first documented in 1908 (Stone and Chapman, 1908),
Fusarium crown and root rot has been reported in North, Central, and South America, Australia,
Japan, New Zealand, Taiwan (Elmer, 2001b), South Africa (Schreuder et al., 1995), the
Netherlands (Van Bakel and Kerstens, 1970), Spain (Wong and Jeffries, 2006), and the United
Kingdom (Wong and Jeffries, 2006).

9

Fusarium oxysporum produces abundant chlamydospores but can also survive as hyphae
on plant residues in and on the soil. There is no known teleomorph of F. oxysporum (Burgess,
1981). Forty-three vegetative compatibility groups (VCGs) have been reported in F. oxysporum
f. sp. asparagi (Elmer et al., 1996) and it is believed that each VCG is a clonal lineage (Correll,
1991). Populations of the F. oxysporum species complex are also separated into formae
speciales based on host pathogenicity. Historically, VCGs were not expected to cross over
formae speciales (Correll, 1991; Puhalla, 1985). The pathogenicity of F. oxysporum on
asparagus does not appear to correspond with vegetative compatibility, which indicates that
pathogenicity on asparagus may be a common trait in F. oxysporum (Elmer, 2001a; Elmer et al.,
1996). Although pathogenic forme in the F. oxysporum complex are typically limited to one host
genotype, isolates of F. oxysporum pathogenic to A. officinalis have also been reported to infect
the roots of other Asparagus spp. as well as corn, pea, celery, gladiolus, lupine, and onion (Dan
and Stephens, 1995; Damicone et al., 1988; Elmer, 2001a; Stephens and Elmer, 1988). F.
oxysporum f. sp. asparagi usually infects young plants in nursery fields or newly established
production fields (Elmer et al., 1996; Keulder, 1999).
F. proliferatum (Teleomorph G. fujikuroi var. intermedia Kuhlman) is mating population
“D” of the Gibberella fujikuroi complex (Elmer, 1995; Samuels et al., 2001), although perithecia
have not been found in nature (Elmer, 2001a). The anamorph has not been found to produce
chlamydospores and it overwinters as mycelia on plant debris. F. proliferatum can colonize
asparagus stems, flowers, and berries as well as crowns (Burgess, 1981; Elmer, 2001a). It is
commonly found on mature and senescing plants (Keulder, 1999).
Fusarium spp., both nonpathogenic and pathogenic to asparagus, are ubiquitous in fields
in the major asparagus producing regions (Elmer, 2001a; Hartung et al., 1990). In a survey of

10

fields in major asparagus producing counties in Michigan, Hartung et al. (1990) found that 38%
of F. oxysporum isolates and all isolates of F. proliferatum sampled from both asparagus fields
and fields without any asparagus history were pathogenic. Both species are soilborne, seedborne,
transported on infested crowns, and vectored by insects (Elmer, 2001a; Elmer et al., 1996;
Morrison et al., 2011). F. proliferatum is also windborne (Elmer, 2001a). Plant stress (Keulder,
1999; Zandstra et al., 1992), allelopathic plant residues (Hartung and Stephens, 1983; Keulder,
1999), viral infection (Elmer, 2001b), and damage from tillage (Shelton and Lacy, 1980), insects
(Morrison et al., 2011), and herbicides (Zandstra et al., 2013) increase crop susceptibility to
Fusarium infection (Elmer, 2001a).

PHYTOPHTHORA SPEAR AND ROOT ROT
Members of the family Pythiaceae, including Phytophthora spp., are characterized by
coencytic mycelia with few to no septae (Agrios, 2005; Erwin and Ribeiro, 1996). When
conditions are favorable, sporangia (also called zoosporangia) form on sporangiophores or on
chains of internally proliferating sporangia. Obpyriform sporangia may or may not have an
apical bud called a papilla. Biflagellate, haploid zoospores are produced inside sporangia, and
then released. Asexual chlamydospores are spherical to oval, and form a protective cell wall that
enables survival in harsh conditions. Diploid oospores are formed when antheridia fuse with and
fertilize oogonia. Oospores have a thick cell wall, and can survive in soil for many years.
Heterothallic species are able to reproduce only when two mating types are present, resulting in
genetic recombination. Homothallic species can self-fertilize and have the potential to produce
large clonal populations (Erwin and Ribeiro, 1996). Morphological characteristics such as
colony growth and spore production on selective medium and at maximum temperatures are used

11

to identify Phytophthora species. DNA fingerprinting techniques including single-strand
conformation polymorphism (SSCP), restriction fragment length polymorphism (RFLP), and
AFLP analysis of ITS and mitochondrial DNA can distinguish among species which are
morphologically similar (Erwin and Ribeiro, 1996; Gallegly and Hong, 2008).
The P. megasperma complex was first described by Dreschler (1931) on Althaea rosea.
The sporangia are internally proliferating and nonpapillate. Paragynous antheridia and oogonia
produce oospores in abundance in homothallic cultures. Tompkins et al. (1936) and Hildebrand
(1959) expanded the criteria for oospore size (≥ 30 µm diameter) and host range to include
similar pathogens in what has been called the broadened species concept. In some cases, species
within the “Phytophthora megasperma complex” have been identified as a separate species
based on morphology or host specificity (Kuan and Erwin, 1980; Waterhouse, 1963).
Mitochondrial and chromosomal DNA RFLPs have defined nine distinct groups in the P.
megasperma complex, five of which are host-specific and the other four are broad host range.
Group E belongs to Phytophthora spp. that infects A. officinalis. The subgroups of E are each
from separate geographical locations, and suggest European origin for this genetic lineage
(Förster and Coffey, 1993).
Several species including P. megasperma Drechsler, P. megasperma var. sojae
Hildebrand, P. cryptogea Penthybridge & Lafferty, P. cactorum (Leb. & Cohn) Schröeter, and P.
richardiae Buisman infect asparagus crops worldwide (Falloon, 1990). P. megasperma var.
sojae is the most common cause of Phytophthora rot in New Zealand (Falloon, 1982; Falloon
and Grogan, 1991), but P. cryptogea is also present in California (Falloon and Grogan, 1991)
and the Netherlands (Falloon, 1990). Asparagus root rot has also been confirmed as a result of
infection from P. megasperma var. sojae in France (Baudry, 1995), P. nicotianae in Peru

12

(Aragon-Caballero et al., 2008), and P. megasperma in Canada (Vujanovic et al., 2003).
Phytophthora asparagi Saude & Hausbeck is the only known species to infect Michigan
asparagus crops (Saude et al., 2008). Results from the 99 isolates from Michigan tested with
AFLP fingerprinting indicated that all were from the same clonal lineage and could be due to the
homothallic sexual phase that is self-fertilizing (Saude et al., 2008).
On asparagus spears, Phytophthora infection often appears as soft, water-soaked, slightly
sunken lesions slightly above or below the soil line. Sometimes the lesions may also appear to
be brown or necrotic (Aragon-Caballero et al., 2008; Falloon et al., 1983). Infected spears may
“crook” or curl over as they wilt. Infected roots first show white or slightly transparent lesions,
then as the disease progresses lesions turn brown to reddish brown with hollow tissue (Falloon et
al., 1983). Water-soaked lesions and shriveling can occur on crowns in cold storage, but the
tissue can remain firm. Internal crown tissues sometimes turn yellow-brown, similar to
Fusarium wilt symptoms. Saude et al. (2008) observed that in drier conditions, lesions can
become necrotic on storage roots, and spears may show crooking and wilting but the tissue may
not appear to be water-soaked.
It has commonly been noted that Phytophthora rot on asparagus is favored during
conditions including cool temperatures and saturated soils (Baudry, 1995; Falloon and Grogan,
1991; Saude et al., 2008). Greenhouse trials confirmed that increasing the duration and
frequency of flooding along with cooling temperatures increased disease severity (Falloon and
Grogan, 1991). In newly established fields, increased flooding frequency (1X, 3X, and 4X)
increased spear emergence time and decreased survival by 20% to 43% during the first year
(Falloon, 1991). In established fields, wet and cool weather early in the harvest season increased

13

yield losses (Falloon et al., 1985). Overhead irrigation can also encourage disease (Falloon and
Fraser, 1991).
On AR - (150 ppm ampicillin and 30 ppm rifampicin) amended V8 agar (840 ml distilled
water, approximately 163 ml unclarified V8 juice, 16 agar, 3 g CaCO3) agar plates, P. asparagi
grows outward in a stellate rosaceous pattern. High oospore counts and low numbers of
sporangia are produced. On dilute V8 (1000 ml distilled water, approximately 40 ml unclarified
V8 juice, 16 agar, 3 g CaCO3), many sporangia are produced, and oospores are still present.
Sporangia are ovoid, obpyriform, nonpapillate, noncaducous, 42 to 47µm long x 23 to 40 µm
wide, produced on simply- or sparingly-branched sporangiophores. Internal proliferation may
occur after releasing zoospores. Oospores are amphigynous and 25 to 30 µm in diameter.
Chlamydospores have not been observed in this species. The minimum, optimum, and maximum
temperatures for growth on AR-V8 agar and spore production were found to be 10, 25, and >
30°C, respectively (Saude et al., 2008). When fresh ‘Millennium’ spears were inoculated with P.
asparagi isolates, larger lesions developed at 20°C than at 15°C and 25°C (Rodriguez Salamanca,
2010).

INTEGRATED PEST MANAGEMENT
Effective management of soilborne diseases requires an integrated approach. Because
many factors affect plant susceptibility and pathogen infection, direct as well as indirect
strategies are needed to minimize crown damage. Cultivar selection, cultural practices, soil
fumigation, fungicides, and biocontrol agents may be used to limit Fusarium and Phytophthora
rots. Managing foliar pathogens, viruses, insects, and weeds and selecting safe herbicides are
also important to maintain crop health and reduce susceptibility to Fusarium crown and root rot.

14

Currently, all A. officinalis cultivars are susceptible to F. oxysporum f. sp. asparagi, F.
proliferatum, and P. asparagi. All-male hybrids have higher plant vigor than open-pollinated
cultivars. Because good vigor reduces plant susceptibility to Fusarium pathogens, all-male
hybrids offer some protection against disease (Ellison and Kinelski, 1985). In vitro screens have
identified tolerance to Fusarium spp. in the ornamentals A. densiflorus ‘Sprengeri’ and ‘Myersii’
(Dan and Stephens, 1995; Stephens and Elmer, 1988; Stephens et al., 1989). Sexual
incompatibility between A. officinalis and A. densiflorus has prevented hybrids from being
developed (Stephens et al., 1989). Using recurrent selection, Falloon et al. (2002) have
identified experimental hybrids with significantly higher survival than susceptible ‘UC 157’
against P. megasperma var. sojae and P. cryptogea isolates from New Zealand, France,
Switzerland, and California. Seven hybrids also resulted in higher yield and quality equal to or
better than industry standards ‘UC 157’ and ‘JWC1’ (Falloon et al., 2002). As resistant cultivars
are identified and developed, cultivar selection will become a useful management tool for
growers.
Using cultural practices to avoid and prevent infection is recommended for Fusarium and
Phytophthora management. Fields with a history of Fusarium crown and root rot or
Phytophthora spear and root rot should be avoided if possible. Saturated soils create conditions
that are unfavorable for asparagus crown growth and increase plant susceptibility while creating
a favorable environment for pathogen development (Saude et al., 2008). Sand and sandy loam
soils can provide good drainage and irrigation should be applied as needed to prevent waterlogging and water-stress (Drost, 1997; Elmer, 2001a). Lime and fertilizer applications to
maintain soil pH above 6.0 and nutrient levels are important to maintain stand health and prevent
Fusarium infection (Zandstra et al., 1992). Falloon and Fraser (1991) suggested transplanting

15

crowns in late spring to avoid cool, wet conditions that are favorable to Phytophthora spear and
root rot. Using greenhouse-grown transplants has been recommended to avoid crown damage at
planting, but this practice has not been adopted by the Michigan industry because it represents a
significant change from current practices (M. Ngouajio, personal communication, 2014).
Overharvesting depletes the crown of nutrients and reduces stand vigor, and should therefore be
avoided (Shelton and Lacy, 1980). Michigan growers have adopted the no-till system, which
prevents crown damage and openings to infection (Wilcox-Lee and Drost, 1991; Zandstra et al.,
1992).
Crop rotation is a recommended method of reducing pathogen populations, but growers
should be careful in choosing a rotation crop. Although P. asparagi is the only known species to
cause asparagus spear and root rot in Michigan, Rodriguez Salamanca (2010) found that at 20
and 25°C P. nicotianae and P. capsici caused lesions on small ‘Jersey Knight’ spears
comparable with P. asparagi when incubated at 15 and 20°C. Although the asparagus harvest
season would normally end before soil temperatures reach 20 to 25°C, unusually warm springs
with heavy rainfall could create favorable conditions for infection by P. nicotianae and P.
capsici. Saude et al. (2008) tested the ability of P. asparagi to infect crops that are economically
important to Michigan. P. asparagi did not cause infection on soybean, alfalfa, or clover
seedlings, but was reisolated from roots of all seedlings. P. asparagi caused infection of cucurbit
fruits when the flesh tissue was wounded. Symptoms appeared as water-soaked lesions on
zucchini, water-soaked and necrotic lesions on yellow squash, and water-soaked lesions with
mycelial growth on slicing and pickling cucumbers (Saude et al., 2008).
Historically, sodium chloride was used on small parcels within the U.S. to manage weeds
and increase yields. When growers began to use herbicides, the number of reports of Fusarium

16

crown and root rot increased, so it was believed that sodium chloride could be used to control the
disease (Elmer, 2001a). Sodium chloride can improve yields in fields with severe Fusarium
crown and root rot symptoms, but studies in greenhouses and fields with less severe symptoms
have not resulted in consistent disease control (Elmer, 2004; Reid et al., 2001). Alternative
forms of chloride salt (calcium, manganese, and ammonium) were found to either have no effect
or to increase root rot symptoms (Reid et al., 2001). The mechanism of control and the
environmental impact of sodium chloride are unknown and the effect on environment as well as
future crops is not well understood (Elmer, 2001b; Elmer et al., 1996; Keulder, 1999).
During cool, wet periods, fungicides may be helpful to control Phytophthora spear and
root rot. When the fungicide metalaxyl was applied to production fields, marketable spear yield
increased by 22% to 43% (Falloon et al., 1983). In a newly established field, when the same soil
treatments were applied immediately after transplant they decreased the emergence time and
increased the stand survival in flooded fields, although the fungicide efficacy was decreased with
increased flooding frequency (Falloon, 1991). A 4-year-old field treated with 1.12 kg/ha
applications showed decreased rot and increased yield (38% and 118%) as well as increased
large spear counts (Falloon et al., 1985). Falloon and Fraser (1991) noted that mefenoxam
applied as 300 mg/liter crown soak was more effective than soil applications. The fungicide
soak allowed direct uptake, but soil applications can only reach the crowns after the water moves
the fungicides through the soil to the roots and crown. This can leave crowns vulnerable to
infection. However, crown soaks applied at rates higher than 150 ppm or 300 mg active
ingredient/liter have been phytotoxic to asparagus plants (Falloon and Fraser, 1991; Green et al.,
2008). Mefenoxam is registered for use on asparagus and can be applied before the harvest

17

begins. Saude et al. (2008) confirmed that 131 isolates of P. asparagi isolates from Michigan
were sensitive to 100 ppm of mefenoxam.

18

LITERATURE CITED

19

LITERATURE CITED

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Elmer, W. H. 1995. A single mating population of Gibberella fujikuroi (Fusarium proliferatum)
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Falloon, P. G., and Fraser, H. A. 1991. Control of establishment failures in asparagus
(Asparagus officinalis L.) caused by Phytophthora rot. New Zealand Journal of Crop and
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Falloon, P. G., and Grogan, R.G. 1991. Effect of root temperature, plant age, frequency, and
duration of flooding and inoculum placement and concentration on susceptibility of
asparagus to Phytophthora rot. New Zealand Journal of Crop and Horticultural Science
19:305-312.
Falloon, P. G., Mullen, R. J., Benson, B. L., and Grogan, R. G. 1985. Control of Phytophthora
rot with metalaxyl in established asparagus. Plant Disease 69:921-923.

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Förster, H., and Coffey, M. D. 1993. Molecular taxonomy of Phytophthora megasperma based
on mitochondrial and nuclear DNA polymorphisms. Mycological Research 97:11011112.
Gallegly, M. E., and Hong, C. 2008. Phytophthora: Identifying Species by Morphology and
DNA Fingerprints. The American Phytopathological Society, St. Paul, MN.
Green, K. R., Dyer, W., Falloon, P. G., Cooke, D. E. L., and Chimento, A. 2008. Management
of Phytophthora rot in UK asparagus crops. Acta Horticulturae 776:175-181.
Hartung, A. C., Stephens, C. T., and Elmer, W. H. 1990. Survey of Fusarium populations in
Michigan’s asparagus fields. Acta Horticulturae 271:395-401.
Hartung, A. C., and Stephens, C. T. 1983. Effects of allelopathic substances produced by
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Hausbeck, M. K., Hartwell, J., and Byrne, J. M. 1999. Epidemiology of Stemphylium leaf spot
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Hildebrand, A. A. 1959. A root and stalk rot of soybean caused by Phytophthora megasperma
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Johnson, D. A., and Lunden, J. D. 1992. Effect of rust on yield of susceptible and resistant
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Keulder, P. C. 1999. Asparagus decline and replant problem: A review of the current situation
and approaches for future research. Acta Horticulturae 479: 253-262.
Kleweno, D. D., and Matthews, V. 2002. Vegetable inventory 2001-2002 of the Michigan
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Kuan, T. L., and Erwin, D. C. 1980. Formae speciales differentiation of Phytophthora
megasperma isolates from soybean and alfalfa. Phytopathology 70:333-338.
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Samuels, G. J., Nirenberg, H. I., and Seifert, K. A. 2001. Perithecial species of Fusarium.
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24

CHAPTER I

VIRULENCE OF PHYTOPHTHORA ASPARAGI, SUSCEPTIBILITY OF ASPARAGUS
CULTIGENS TO SPEAR AND ROOT ROT, AND THE EFFICACY OF SEED
TREATMENTS

25

ABSTRACT
Phytophthora asparagi, causal agent of Phytophthora spear and root rot of asparagus, is
an important contributor to crop decline and replant problem in Michigan. Management
strategies are limited, and identifying resistant cultigens and seed treatments will provide
valuable strategies for the asparagus industry. Nine isolates of P. asparagi were tested for
virulence using asparagus spears of five commercial cultivars and asparagus seedlings of three
cultivars. On asparagus spears, differences in isolate virulence and host susceptibility were
observed with ‘Jersey Knight’ being the least susceptible and ‘Jersey Supreme’ the most
susceptible. No significant differences were detected among cultivars or isolates in the seedling
root rot experiment. Two virulent isolates were selected to inoculate asparagus seedlings of 18
cultigens. Isolate SP317 was significantly more virulent than SP316, and ‘Jersey Giant’ and
‘UG009’ were the least susceptible cultigens. The interaction between isolate virulence and
cultigen susceptibility significantly affected root rot severity. ‘Pacific 2000’ and ‘Jersey Giant’
were the least susceptible cultigens for isolates SP316 and SP317, respectively. Seven seed
treatments, including experimental fungicides and biocontrol agents, were screened for efficacy
against two isolates of P. asparagi on ‘Millennium’ in vitro. Seeds treated with mefenoxam and
Bacillus subtilis had the highest germination overall and on plates inoculated with SP326. When
seeds were inoculated with SP316, mefenoxam, DPX-QGU42, and B. subtilis treatments
germinated at the highest rates. Antagonism of mycelial growth was observed in the B. subtilis
treatment, which resulted in significantly decreased disease incidence compared to all other
treatments. This is the first reported evaluation of cultigen susceptibility to P. asparagi-induced
rot in asparagus seedling roots. These results are consistent with previous research suggesting

26

that virulence of isolates of Phytophthora spp. is a combination of host and pathogen genetics
and that some asparagus cultigens are less susceptible than others to P. asparagi.

INTRODUCTION
Michigan ranks third in the United States for asparagus production, following California
and Washington. In 2012, Michigan produced 8.8 million kg of spears for the fresh market and
processing, with a total value of $17.3 million, (Anonymous, 2013). With proper maintenance, a
healthy asparagus field will remain profitable for 12 to15 years or more (Elmer et al., 1996;
Sanders, 2001; Zandstra et al., 1992). However, asparagus early decline can shorten a field’s
lifespan by 5 to 10 years (Elmer et al, 1996). As plant vigor decreases gradually over time, the
size and number of marketable spears and fern shoots are reduced (Keulder, 1999). The replant
problem occurs when new stands fail to become successfully established in a field previously
cropped with asparagus (Elmer et al., 1996; Falloon and Fraser, 1991; Morrison et al., 2011;
Keulder, 1999). Historically, Fusarium crown and root rot was considered to be the primary soilborne disease affecting crop decline in Michigan, but recently Phytophthora spear and root rot
(caused by P. asparagi Saude & Hausbeck) has been reported as an important contributor as well
(Saude et al., 2008).
Phytophthora root rot infections begin underground, causing water-soaked lesions,
shriveling, and red-brown discoloration in crown and root tissue. As the disease progresses,
lesions spread longitudinally down the roots, which may become hollow. On crowns, affected
buds will fail to produce spears, and spears that grow from weakened crowns are usually smaller
than those from healthy crowns. Affected fields appear to have dead patches, especially in low
areas where water accumulates. Spear rot occurs when soil infested with zoospores splashes

27

onto shoots near the soil level. Symptoms appear as soft, water-soaked, slightly sunken lesions
slightly above or below soil line. Sometimes the lesions may also show necrosis (Falloon et al.,
1983; Aragon-Caballero et al., 2008).
P. asparagi is a homothallic, or self-fertilizing, species. DNA fingerprinting tests
indicated that isolates collected from asparagus fields in three Michigan counties were from one
clonal lineage, which suggests low genetic variability in Michigan populations (Saude et al.,
2008). Other Phytophthora spp., such as P. capsici, have significant genetic diversity and
varying levels of virulence within populations. Levels of disease severity depend on the
interaction between isolate and host genotypes (Glosier et al., 2008; Granke et al., 2012; Kim
and Hwang, 1992). Saude et al. (2008) found no differences in virulence among 115 isolates on
spears of 'Jersey Giant’, an all-male hybrid with little genetic variability. Testing multiple
isolates across several cultigens will increase differences between host genotypes, and may
increase differences in disease severity between the isolates.
Cultural practices to manage the disease are limited. Falloon and Fraser (1991) suggested
transplanting crowns in late spring to avoid cool/wet conditions. Crop rotation may be useful,
but growers are advised to be careful in choosing a rotation crop. Cultivar selection is a cultural
practice often used in vegetable crops, but host resistance has not been identified against P.
asparagi. Using recurrent selection, Falloon et al. (2002) identified experimental hybrids with
significantly higher survival than the susceptible ‘UC 157’ against Phytophthora spp. from New
Zealand, France, Switzerland, and California. Identifying cultivars with resistance or tolerance
to P. asparagi would be useful for the Michigan asparagus industry.
Traditionally, Michigan growers have not fumigated the soil of their crown nurseries,
though pathogens may be present. Saude et al. (2008) sampled crowns stored in bulk boxes from

28

a commercial producer and found that 40% of the samples were infected with P. asparagi. To
prevent P. asparagi inoculum from reaching production fields, growers should use only certified
clean material raised in nursery beds that are fumigated. However, fumigation can create a
biological vacuum, allowing new pathogens to colonize the soil (Keulder, 1999). There are
currently no fungicides labeled to protect young seedlings in nursery beds. Young plants are
especially susceptible to Phytophthora root rot (Falloon and Grogan, 1991) and effective
seedling treatments would be a valuable management tool.
The objectives of this study were to: i) Test the virulence of nine P. asparagi isolates on
asparagus spears and seedling roots of commercial cultivars, ii) Compare the relative
susceptibility of asparagus cultivars and breeding lines to Phytophthora spear and root rot, and
iii) Screen fungicides and biocontrol agents for efficacy against P. asparagi.

MATERIALS AND METHODS
Isolates and inoculum preparation. Michigan P. asparagi isolates C013, SP316,
SP317, SP318, SP319, SP324, SP325, SP326, and SP3236 from the collection of M. K.
Hausbeck were maintained on AR - (150 ppm ampicillin and 30 ppm rifampicin) amended V8
agar (840 ml distilled water, approximately 163 ml unclarified V8 juice, 16 agar, 3 g CaCO3) at
room temperature (23 ± 2°C) under continuous fluorescent lighting and transferred every two
weeks. Broth cultures were initiated by adding agar plugs from the mycelial margin of actively
growing cultures (4 to 9 days) to AR-amended V8 broth (840 ml distilled water, approximately
163 ml clarified V8 juice, 3 g CaCO3). For the spear rot experiments, a 7-mm mycelial plug was
added to 4 ml AR-V8 broth and for seedling root rot experiments a 4-mm mycelial plug was

29

added to 2 ml AR-V8 broth. Broth cultures were incubated under temperature and light
conditions as described above for 4 to 7 days before inoculating plant material.
Susceptibility of asparagus cultivars to spear rot. Commercial grade spears (1.5 to 2.0
cm in diameter and ≥ 14 cm tall) of ‘Millennium,’ ‘Tiessen,’ ‘Jersey Giant,’ ‘Jersey Knight,’ and
‘Jersey Supreme’ were harvested from fields in Oceana County, MI. In the first and third trials,
the spears were kept overnight in a cold room maintained at approximately 4.4°C. In the second
trial, the spears were inoculated within 12 hours of harvest.
Autoclaved Benona sandy loam from Oceana County was infested with one of the
following P. asparagi isolates; C013, SP316, SP317, SP318, SP319, SP324, SP325, SP326, or
SP3236 and used to inoculate spears. An AR-V8 broth culture of each P. asparagi isolate was
prepared as described above. After incubation, 4 ml of the broth culture of each isolate was
added to 10 g of autoclaved sandy loam that was placed in test tubes (25 x 150 mm). Sterile ARV8 broth was used for the uninoculated control treatments.
Asparagus spears were surface disinfested using the method of Saude et al. (2008) that
included washing in 5% sodium hypochlorite for 5 seconds, rinsing 3 times in distilled water for
5 seconds each. Spears were air dried on clean paper towels in a laminar flow hood, then
trimmed to 12 cm, with the apical tips and below ground tissue removed. Spears were
immediately placed into the test tubes with the base in direct contact with the infested soil. Five
spears of each cultivar were used for each isolate and for the uninoculated controls. Tubes were
arranged in a complete randomized design in test tube racks and incubated for 7 days at room
temperature under continuous fluorescent lighting. After incubation, the length and width of
each lesion was measured. Because lesions were not uniform in height but were similar to a
rectangle shape, area was calculated as A = W*H, where W was width at the spear base edge and

30

H was the average of 3 separate length measurements. Experiments were conducted three times
with ‘Millennium,’ ‘Jersey Giant,’ ‘Jersey Supeme,’ and ‘Jersey Knight’ spears, and twice with
‘Tiessen’ spears.
Seedling inoculation and incubation. Asparagus seeds were surface disinfested using
the method of Damicone et al. (1981) by agitating in a solution of 25,000 ppm benomyl in 99.9%
acetone on a rotary shaker (1000 rpm) for 24 hours, then rinsing 3 times in acetone and 3 times
in distilled water. Seeds were then agitated in 1.23% sodium hypochlorite at 1000 rpm for 1
hour and rinsed 3 times in distilled water. After being allowed to air dry for 1 to 2 hours in a
laminar flow hood, the seeds were placed on water agar (16 g agar and 1000 ml distilled water)
and germinated in the dark at room temperature for 7 to 14 days. Once the hypocotyls emerged,
the seeds were planted in 12 ml Hoagland’s agar (Stephens and Elmer, 1988) in test tubes (25 x
150 mm) and grown in growth chambers (16-hour photoperiod at 25/20°C day/night).
An AR-V8 broth culture (2 ml) of each P. asparagi isolate (see specific isolates in
experiments below) was prepared as described previously. When seedlings were approximately
11 cm in height and secondary branches with cladophylls had formed (approximately 2 weeks),
the broth culture was placed directly on the agar of each test tube. Following inoculation,
seedlings were allowed to grow and the infection to develop for 4 weeks under growth chamber
conditions described above. Severity of Phytophthora root rot was visually assessed weekly,
starting at 7 days post inoculation (dpi). A disease severity scale developed by Falloon (2002) of
0 to 4 was used (0 = no disease; 4 = water-soaked tissue, rot of > 50% of the roots) (Table 1.1,
Figure 1.1). A disease severity index (DSI) where 0 = no disease and 100 = all seedlings with >
50% of their roots water-soaked was calculated for each isolate/cultigen combination (Falloon et
al, 2002). For the fourth and final rating, seedling roots were removed from the agar prior to the

31

Table 1.1. Disease classes of inoculated asparagus seedlings used to calculate disease severity
index (DSI*).
Disease class
Symptoms
0
No disease
1
<10%**
2
10-20%
3
20-50%
4
>50%
*DSI=Σ (class x number plants/class) x 100/total plants x 4.
**Indicates area of root system water-soaked.

0

1

3

2

4

Figure 1.1. The Phytophthora symptoms observed on inoculated asparagus seedling root
systems receiving the rating of a 0 to a 4.

32

visual assessment. The area under the disease progress curve (AUDPC) was calculated (Madden
et al., 2007) to obtain the cumulative DSI throughout the experiments.
Virulence of isolates on seedlings roots. The virulence of nine P. asparagi isolates
(C013, SP316, SP317, SP318, SP319, SP324, SP325, SP326, and SP3236) were compared on
the seedling root systems of two-week-old ‘Millennium,’ ‘Mary Washington.’ and ‘Jersey Giant’
seedlings. Sterile AR-V8 broth (2 ml) was used for the negative controls. Five seedlings were
used for each cultivar/isolate combination and the controls in the first trial and 6 seedlings were
used in the second trial. Tubes were arranged in a complete randomized design in test tube racks
and incubated in growth chamber conditions previously described. Experiments were conducted
twice.
Susceptibility of cultivars and breeding lines to seedling root rot. Asparagus
seedlings of 18 cultivars and breeding lines (Table 1.2) were selected to evaluate differences in
susceptibility to P. asparagi isolates SP316 and SP317. Sterile AR-V8 (2 ml) was used as the
negative control. Six seedlings were used for each cultigen/isolate combination and the controls.
Seedlings were arranged in a complete randomized design in test tube racks and incubated in the
growth chamber conditions as described above.
Efficacy of fungicides and biocontrol agents as seed treatments. The methods used by
Broders et al. (2007) were adapted to screen five fungicides and two biocontrol agents (Table
1.3) for efficacy as seed treatments in vitro. A mycelial plug (7-mm in diameter) of SP316 or
SP326 was placed in the center of 15-cm-diameter petri plate containing 50 ml of V8 agar. The
isolates were allowed to colonize the plates for 3 days. Seeds of ‘Millennium’ were surface
disinfested as previously described and allowed to air dry in a laminar flow hood for
approximately 1 hour. Treatments were applied to seed lots (0.33 g) at the rates listed in

33

Table 1.2. Asparagus cultivars and breeding lines tested for susceptibility to Phytophthora
asparagi.
Cultigen
Source
Millennium
University of Guelph
UG5
University of Guelph
UG9
University of Guelph
UG10
University of Guelph
UG20
University of Guelph
Jersey Giant
Walker Brothers
NJ 938
Rutgers University
NJ 941
Rutgers University
NJ 1191
Rutgers University
Jersey Supreme*
Vilmorin
Mondeo*
Vilmorin
Pacific Challenger 1
Aspara Pacific
Pacific Challenger 2
Aspara Pacific
Pacific 2000
Aspara Pacific
74 x 22
Aspara Pacific
3 x Phy 99
Aspara Pacific
Cumulus (2735)
Bejo
Bacchus (2827)*
Bejo
*Packaged seeds were pretreated with thiram.

Table 1.3. Fungicides and biocontrol agents tested for efficacy against Phytophthora asparagi.
Treatment
Active ingredient
Rate/100 lb seed
Rate/20 seeds (0.48 g)
600.0 µl
Untreated
sterile deionized water
-0.15 µl
Apron
mefenoxam
14.2 ml
0.31 µl
Presidio
fluopicolide
29.6 ml
2.51 µl
Micora
mandipropamid
237.0 ml
0.19 µl
V-10208
experimental
17.7 ml
0.63 µl
DPX-QGU42
experimental
59.1 ml
*
3.60 mg
Streptomyces lydicus
Actinovate
340.2 g
Serenade Soil

Bacillus subtilis

*

813.2 ml

*

Biological control agent

34

8.61 µl

A

B

Figure 1.2. Asparagus seeds treated with fungicides and biocontrol agents on V8
agar plates inoculated with Phytophthora asparagi isolates SP316 (A) and SP326
(B), 3 days after planting.

Table 1.3. Seeds were soaked in each treatment for approximately 30 minutes, then air-dried in a
laminar flow hood for approximately 1 hour. Seeds from each treatment were equally spaced in
a randomized order around the plate, approximately 3 cm from the edge (Figure 1.2). Seeds
from each treatment were also plated on sterile V8 agar plates as the uninoculated controls.
At 10 days after plating the seeds, inhibition of mycelial growth was measured as the
distance between the seed and margin of mycelial growth along the radius of the agar plate.
Seeds were removed from culture and germinated seeds were rated for disease severity. The
number of seeds in each treatment that germinated on infested plates was compared with the
number that germinated on the uninoculated plates. Seeds were considered successfully
germinated if the hypocotyl had emerged or the radicle was > 5 mm long. Treatments were
35

scored according to a 0 to 3 rating scale (0 = no disease symptoms; 3 > 50% tissue water-soaked,
mycelial growth present) (Table 1.4, Figure 1.3). Each treatment was replicated five times for
each isolate and the uninoculated control, and the experiment was conducted three times.
Pathogen isolation. At the end of each study, plant tissue was surface disinfested in
0.525% sodium hypochlorite for approximately 60 seconds, rinsed in sterile distilled water, and
air dried in a laminar flow hood for 1 to 3 minutes. Tissue sections were placed into AR-V8 agar.
Plates were incubated for 3 to 7 days at room temperature (23-25°C) under continuous
fluorescent lighting to confirm the presence of the pathogen. For the spear rot experiment,
approximately 70% of the inoculated spears were sampled. Control spears with water-soaked
areas were also sampled to confirm the absence of pathogens. The epidermal layer was peeled
away and four small sections (approximately 5 x 5 mm) from the lesion margins were excised
with a flame-sterilized scalpel and disinfested and plated in the method described above. For the
seedling root rot experiments, approximately 44% and 56% of the inoculated plants were
sampled in the isolate virulence and variety screening trials. Control plants with discolored and
wilted roots were also sampled. Four small sections from lesion margins of root and crown
tissue were removed, disinfested, and plated. For the seed treatment experiment, whole seeds
were disinfested in the manner described above. After air-drying, radicle tissue was wounded
using a flame-sterilized scalpel and plated. All inoculated seeds were sampled, and control seeds
were also sampled if radicles had water-soaked or discolored tissue.
Statistical analysis. For all experiments, data was analyzed with a two-way analysis of
variance (ANOVA) using the PROC MIXED procedure with the REPEATED statement of SAS
version 9.3 (SAS Institute Inc., Cary, NC). Residuals were tested for normality and homogeneity
of variance using PROC UNIVARIATE. Trial repetition was treated as a random affect

36

Table 1.4. Scale of disease ratings on successfully germinated asparagus seedlings inoculated
with Phytophthora asparagi.
Rating
Symptoms
0
No disease
1
<50% radicle tissue water-soaked
2
>50% radicle tissue water-soaked
3
>50% radicle tissue water-soaked + mycelial growth present

A

B

C

D

Figure 1.3. Symptoms of Phytophthora asparagi observed on germinated asparagus
seedlings. (A) 0 = no disease symptoms, (B) 1 < 50% radicle tissue water-soaked, (C) 2 >
50% radicle tissue water-soaked, and (D) 3 > 50% radicle tissue water-soaked with mycelia
growth.

37

in all experiments, but was excluded from the model analyzing the incidence data in the seed
treatment trial. For the spear rot experiment, isolate was used as the group effect in the
REPEATED statement. Multiple comparisons among the cultivar and isolate means were
conducted using ANOVA and Tukey’s adjustments were used for separation of means when
effects were statistically significant at P = 0.05. For the seedling root rot experiments, the
REPEATED statement was used with cultigen as the group effect for the AUDPC data in the
virulence trial and the AUDPC and DSI data in the variety trial. Isolate was used as the group
effect for the DSI data in the virulence trial. Data from the control plants were removed prior to
analyses to avoid violating the assumption of equal variances. Multiple comparisons among the
cultigen and isolate means were conducted using ANOVA and Tukey’s adjustments were used in
the virulence screen and CONTRAST statements in the cultigen trial for separation of means
when effects were statistically significant at P = 0.05. For the seed treatment experiment, the
REPEATED statement was used with treatment as the group effect. Data for both variables was
square root transformed before analysis. Multiple comparisons among the isolate and treatment
means were conducted using ANOVA and Tukey’s adjustments were used when effects were
statistically significant at P = 0.05.

RESULTS
Susceptibility of commercial cultivars to spear rot. When spears were inoculated with
P. asparagi, dark green, water-soaked lesions developed regardless of isolate. Occasionally,
white mycelial growth was evident on the inoculated spears. All isolates were readily isolated
from the spear tissue (64 to 95%). By the end of the 7-day incubation period, some spears
developed a secondary infection on the scales and tip. Control spears either showed no water-

38

soaked lesions or only very small water-soaked areas along the base. P. asparagi was not
recovered from water-soaked tissue on the control spears.
The interaction between cultivar and isolate (P = 0.05) did not have a significant effect on
lesion area, but lesion size differed significantly among cultivars (P = 0.05) and isolates (P =
0.05). ‘Jersey Supreme’ spears had significantly larger lesions compared to ‘Jersey Giant’ and
‘Jersey Knight.’ ‘Tiessen’ spears had a larger average lesion area compared to ‘Jersey Knight.’
Lesion size was similar among ‘Jersey Knight,’ ‘Jersey Giant,’ and ‘Millennium’ and among
‘Millennium,’ Tiessen,’ and ‘Jersey Supreme’ (Figure 1.4A). When spears were inoculated with
isolates SP324, SP325, and SP3236, the lesions were larger than those resulting from isolates
SP318 and C013. Spears inoculated with isolates SP316, SP317, SP319, and SP326 produced
lesions that were statistically similar to all other isolates (Figure 1.4B).
Virulence of isolates on seedlings roots. Phytophthora rot symptoms appeared as
translucent water-soaked tissue that was sometimes accompanied by a red brown discoloration.
Symptoms were first observed on seedling roots at 7 days post inoculation (dpi). All isolates
were readily isolated from infected seedling roots (69 to 100%). There were no significant
differences among P. asparagi isolates (Figure 1.5) or among cultivars (Figure 1.6), but slight
trends were observed. Isolates SP316 and SP317 caused the highest DSI values and C013, SP316,
and SP317 had the highest AUDPC (Figure 1.5). Of the three cultivars tested, Millennium had
the smallest DSI and AUDPC (Figure 1.6).
Susceptibility of cultivars and breeding lines to seedling root rot. Root rot symptoms
were first observed at 7 dpi, and all cultigens were symptomatic at 14 dpi. The symptoms
appeared as described above. All isolates were readily recovered from root tissue (78 to 100%).

39

The interaction between cultigen and P. asparagi isolate (P = 0.05) had a significant
effect on DSI, but not on AUDPC (Table 1.5). SP317 inoculation resulted in significantly higher
DSI in cultigens Cumulus, Jersey Supreme, Mondeo, NJ1191, and Pacific Challenger 1. SP316
caused higher DSI in ‘Jersey Giant.’ ‘Jersey Supreme,’ ‘Pacific 2000,’ and UG009 had the
lowest DSI values, while ‘Cumulus,’ NJ1191, and ‘UG005’ had the highest DSI values when
inoculated with SP316. ‘Pacific Challenger 2’ and ‘3 x Phy99’ also had low DSI means when
inoculated with SP316, although unequal variances affected the DSI significance. ‘Jersey Giant,’
‘Pacific 2000,’ and ‘UG009’ were significantly less susceptible than ‘NJ1191,’ ‘Mondeo,’ and
‘Cumulus’ when inoculated with SP317. ‘Pacific Challenger 2,’ ‘UG010,’ and ‘74 x 22’ also
had low DSI means when inoculated with SP317, although variance size affected the DSI
significance.
Main effects isolate (P = 0.05) and cultigen (P = 0.05) significantly affected both DSI
and AUDPC. SP317 was significantly more virulent than SP316. ‘Jersey Giant’ and ‘UG009’
were significantly less susceptible than ‘Cumulus,’ ‘NJ1191,’ and ‘UG005.’ ‘Pacific 2000’ and
‘UG009’ had the smallest AUDPC values.
Efficacy of fungicides and biocontrol agents as seed treatments. At 10 dpi, both
pathogens had colonized most of the agar disc area, including the area immediately surrounding
seeds. Seeds from all treatments, except for Bacillus subtilis, were colonized with mycelium
growth. B. subtilis growth was observed on the treated seeds and the agar surrounded the seeds.
Average inhibition of mycelial growth caused by B. subtilis was observed surrounded the
immediate area outside of the bacterial growth for both P. asparagi isolates. SP316 were readily
isolated from seed tissues from most seed treatments (73 to 100%), and less readily from the B.
subtilis treatment (40%). SP326 was readily isolated from seed tissues from most seed

40

Figure 1.4. Effect of cultivar (A) and Phytophthora asparagi isolate (B) on mean lesion area on detached asparagus spears 7 days
after inoculation. Bars with common letters are not significantly different, LSD P=0.05 (A) and Tukey’s P=0.05 (B).

41

Figure 1.5. Effect of Phytophthora asparagi isolates on disease severity index (DSI) where 0 = no disease and 100 = all seedlings
roots with > 50% of their roots water-soaked (A) and area under the disease progress curve (AUDPC) values (B). Bars with
common letters are not significantly different, LSD P=0.05.

42

Figure 1.6. Effect of cultivars on disease severity index (DSI) where 0 = no disease and 100 = all seedlings with > 50% of their
roots water-soaked (A) and area under the disease progress curve (AUDPC) values (B). Bars with common letters are not
significantly different, LSD P=0.05.

43

treatments (73 to 93%), less readily from the mefenoxam and DPX-QGU42 treatments
(57 and 60%), and infrequently from the B. subtilis treatment (7%).
The interaction between isolate and treatment (P = 0.05) significantly affected seed
germination. P. asparagi isolates were statistically similar within treatments, although
germination of seeds inoculated with SP316 was usually lower than those inoculated with SP326
(Table 1.6). Mefenoxam, DPX-QGU42, and B. subtilis treatments had the highest germination
when seeds were inoculated with SP316 (Table 1.6). The remaining treatments were statistically
similar to the untreated seeds. Mefenoxam and B. subtilis were the only treatments with higher
germination compared to the untreated control when seeds were inoculated with SP326 (Table
1.6). Both main effects isolate and treatments (P = 0.05) were significant for seed germination.
Seeds inoculated with SP316 had significantly lower germination (approximately 18%)
compared to those inoculated with SP326 (42%). Mefenoxam (approximately 73%) and B.
subtilis (approximately 97%) were most effective in increasing germination percentage
compared to the untreated control (approximately 1%).
The interaction between isolate and treatment (P = 0.05) did not significantly affect
disease incidence, but the main effects isolate (P = 0.05) and treatment (P = 0.05) were
significant. Seeds inoculated with SP316 (approximately 79%) had higher root rot incidence
than those inoculated with SP326 (approximately 63%). B. subtilis-treated seeds had the lowest
disease incidence (Table 1.6), although seedling radicals in the treatment group appeared stunted
and discolored. All remaining treatments were statistically similar to the untreated control.
Of the seedlings inoculated with SP316 and that germinated successfully, mandipropamid
(N = 2) and B. subtilis (N = 13) had the highest percentage of 0 and 1 disease severity ratings
(100% and 85%, respectively) (Figure 1.7). All seedlings treated with mefenoxam (N = 5) and

44

Table 1.5. Average disease severity index (DSI) and area under the disease progress curve
(AUDPC) of asparagus seedlings infected with Phytophthora asparagi isolates at 28 days post
inoculation.
z
Disease severity index
Cultigen
AUDPC
P. asparagi SP316
P. asparagi SP317
y
y
x
Millennium
50.0 ABC
65.3 BCDE
1130.2 BCD
UG005
72.2 C
68.1 CDE
1227.4 D
UG009
36.1 A
40.3 ABC
585.8 AB
UG010
40.3 AB
44.4 ABCDE
663.5 ABC
UG020
47.2 ABC
48.6 BCDE
824.0 BCD
Jersey Giant
40.3 *AB
31.9 A
544.5 ABC
NJ938
44.4 AB
55.6 ABCDE
743.8 ABCD
NJ941
44.5 AB
54.2 ABCDE
853.2 BCD
NJ1191
55.6 BC
68.0 *DE
896.9 CD
Jersey Supreme
37.5 A
61.1 *CDE
717.0 C
Mondeo
44.4 AB
70.8 *DE
999.0 CD
Pacific Challenger 1
43.1 AB
65.3 *CDE
947.9 CD
Pacific Challenger 2
44.0 ABC
42.0 ABCDE
805.8 ABCD
44.4 ABC
449.7 A
Pacific 2000
31.9 A
3 x Phy99
37.8 AB
64.9 CDE
827.6 BCD
74 x 22
54.2 ABC
43.1 ABCD
753.5 BC
Bacchus
55.6 ABC
66.6 CDE
904.2 CD
Cumulus
57.0 BC
70.8 *E
909.0 CD
z
Seedlings were given disease ratings (0 to 4) that were used to calculate a disease severity index
(DSI) between 0 and 100, where 0 = no disease and 100 = all seedlings with >50% of their roots
water-soaked. Each DSI represents the average of three repeated tests with six asparagus
seedlings per cultigen/isolate combination per test.
y
DSI values with the same letter are not significantly different among cultigens for each isolate
(within columns). P = 0.05.
*Isolates within a cultigen are significantly different (within rows). P = 0.05
x
AUDPC values with the same letter are not significantly different between cultigens. P = 0.05.

Streptomyces lydicus (N = 1) received a severity rating of 3 when inoculated with SP316 (Figure
1.7). All seedlings in the fluopicolide (N = 2) and V-10208 (N = 1) treatments received a rating
of 2 when inoculated with SP316 (Figure 1.7). More than half of the seedlings treated with
DPX-QGU42 (N = 10) and inoculated with SP316 were given a rating of 3, and the remaining
seedlings were given either a rating of 1 or 2 (Figure 1.7). No seedlings germinated successfully
in the untreated control when inoculated with SP316, and therefore not given a disease severity

45

Table 1.6. The effect of Phytophthora asparagi isolate and seed treatment on germination and
disease incidence.
z
Germination
(%)
Treatment
Disease incidence (%)
P. asparagi SP316
P. asparagi SP326
Untreated
Mefenoxam
Fluopicolide
Mandipropamid
V-10208
DPX-QGU42
Streptomyces lydicus
Bacillus subtilis
z

0.0
50.0
20.0
13.3
11.1
76.7
11.1
100.0

y

A
BCD
ABCD
ABCD
AB
CD
ABC
D

8.3
100.0
23.3
40.0
55.6
53.3
52.8
93.3

A
B
AB
AB
AB
AB
AB
B

71.6
75.7
93.1
82.3
93.1
78.1
88.1
13.0

B
B
B
B
B
B
B
A

x

Germination percentages were calculated for each treatment as (GI/G0)*100, where GI =

number of inoculated seeds that germinated and G0 = number of uninoculated seeds that
germinated
y
Germination percentages with the same letter are not significantly different among P. asparagi
isolates for each treatment (lower case) or between treatments for each isolate (upper case).
Tukey’s, P=0.05.
x
Disease incidence values with the same letter are not significantly different between treatments.
Tukey’s, P=0.05.

rating. All seedlings treated with B. subtilis (N = 12) and inoculated with SP326 received a
rating of 0, except for one seedling that was given a rating of 1 (Figure 1.8). Approximately
27% and 20% of seedlings treated with mefenoxam (N = 11) and S. lydicus (N = 5) were given a
rating of 0 when inoculated with SP326 (Figure 1.8). Most of the seedlings (approximately
83%) inoculated with SP326 and treated with mandipropamid (N = 6) were given a rating of 1
(Figure 1.8). Varying numbers of seedlings in the remaining seed treatment groups were given
ratings between 1 and 3. Only one seedling in the untreated control successfully germinated and
was given a rating of 1 (Figure 1.8).

46

Disease rating distribution (%)

100

N=5

N=2

N=2

N=1

N = 10

N=1

N = 13

80

60

40

20

0
id
ide
am
am
ox
col
p
i
n
o
e
p
f
o
ipr
Me
Flu
nd
a
M

s
08
42
us
tili
dic
b
102
GU
y
u
l
Q
s
V
Xces
us
DP
my
cill
o
a
t
B
ep
Severity=3
Str
Severity=2
Seed treatment
Severity=1
Severity=0

Figure 1.7. The distribution of disease rating scores of germinated seedlings inoculated
with Phytophthora asparagi SP316 and treated with fungicides and biocontrol agents.

47

Disease rating distribution (%)

100

N=1

N = 11

N=3

N=6

N=7

N=7

N=5

N = 12

80

60

40

20

0
2
id
us
ed
de
08
lis
U4
xam
02
oli
am
dic
bti
eat
o
c
G
1
y
p
u
r
i
n
l
t
s
o
Q
V
fe
op
XUn
ces
ipr
lus
Me
Flu
cil
nd
my
DP
a
a
o
t
B
M
ep
Str

Severity=3
Severity=2
Severity=1
Severity=0

Figure 1.8. The distribution of disease rating scores of germinated seedlings inoculated
with Phytophthora asparagi SP326 and treated with fungicides and biocontrol agents.

48

DISCUSSION
Phytophthora asparagi is a homothallic species, and molecular tests suggest that
Michigan populations have risen from one clonal lineage (Saude et al., 2008). Other
Phytophthora spp., such as P. capsici, have substantial genetic variation within populations and
virulence varies between isolates. Pathogen virulence, however, is not solely based on pathogen
genotype, but is also influenced by host genetic variability. A previous study concluded that P.
asparagi virulence levels were similar across Michigan isolates. The asparagus spears used were
from one all-male hybrid, and did not represent genetic differences between cultigens or within
open-pollinated cultivars. In the current study, the results suggest that cultigens affected the
virulence levels among P. asparagi isolates.
The differences in isolate virulence on five cultivars of asparagus spears were significant,
although the interaction between cultivar and isolate was not. When all nine isolates were
screened for virulence on seedling roots of three cultivars, there were no statistical variations
between cultivars or isolates. However, in the cultigen screen, there were significant differences
in virulence between SP316 and SP317, depending on the cultigen. The increased diversity of
asparagus genotypes likely made the interaction between isolate and host detectable.
Host resistance against Phytophthora spp. is a valuable management tool in crops such as
soybean (Lin et al., 2013) and pepper (Mallard et al., 2013). True host resistance against P.
asparagi has not been developed in asparagus, but a breeding program in New Zealand has
identified several breeding lines with tolerance to P. megasperma var. sojae and P. cryptogea.
Although all cultigens used in this study were susceptible to P. asparagi, several varieties were
relatively less susceptible, including the New Zealand cultivar Pacific 2000. ‘Jersey Giant’ and
UG009 were least susceptible. Other cultigens from the Pacific, Jersey, and Guelph lines were

49

highly susceptible to P. asparagi, indicating that while each line contains better hybrids, an
entire line of hybrids that is consistently superior to the other breeding programs has not yet been
developed.
When asparagus seeds were inoculated with P. asparagi isolates in vitro, damping-off
incidence was high for all seed treatments except for Bacillus subtilis. B. subtilis seeds also
germinated at higher percentages than most of the treatments. Seeds not affected by
Phytophthora symptoms were surrounded by B. subtilis growth, which antagonized mycelial
growth on the agar. This is consistent with prior studies, where B. subtilis effectively inhibited
Phytophthora spp. in vitro (Chung et al., 2008). B. subtilis produces several antibiotics with
antifungal and antibacterial properties, which has resulted in effective control of a wide range of
plant pathogens in laboratory and greenhouse conditions (Berger et al., 1996; Chung et al., 2008).
A number of seedling radicles within the B. subtilis treatment group were stunted and discolored,
with an absence of root hairs. Because the in vitro conditions were highly favorable for the
bacterium, it is likely that bacterial growth overwhelmed the young seedling tissues. B. subtilis
should be studied further in soil-based assays to determine if the seed treatment is safe for young
asparagus roots before it can be recommended for use on seeds or young seedlings growing in
nursery fields.
Mefenoxam increased germination in plates inoculated with either isolate. Both isolates
were previously reported to be mefanoxam-sensitive (Saude et al., 2008). Mefanoxam has been
used to successfully control Phytophthora root rot as a crown soak and as a soil drench (Falloon
and Fraser, 1991; Green et al., 2008) and is registered for use in production fields. Our results
suggest that mefanoxam has the potential to successfully protect young seedlings from
Phytophthora root rot as well.

50

In this study, the unequal number of germinated seedlings across treatments made
germination and disease incidence a more reliable measure of treatment effectiveness than
disease severity. The same active ingredients used in this study could also be applied to seedling
roots following germination to measure the effect of fungicides and biocontrol agents on
Phytophthora root rot severity. In vitro conditions, with an abundant nutrient supply, lack of
competition and antagonism from microbial communities, and favorable environmental
conditions for growth and infection may have increased the pathogens ability to rapidly colonize
seed tissues from what would normally occur in soil. Future research should examine the
efficacy of these and other seed treatments in soil-based assays to better represent field
conditions.
In order to effectively manage Phytophthora spear and root rot, growers are advised to
combine control strategies. Careful selection of asparagus cultivars and chemical and biological
treatments is useful in an integrated approach to disease management. Our results suggest that
certain hybrids from the Guelph, Jersey, and New Zealand breeding programs and mefanoxam
may be helpful in management programs.

51

LITERATURE CITED

52

LITERATURE CITED

Aragon-Caballero, L. M., Hurtado-Gonzales, O. P., Flores-Torres, J. G., Apaza-Tapia, W., and
Lamour, K. H. 2008. First report of Phytophthora nicotianae causing asparagus spear
and root rot in Peru. Plant Disease 92:982-982.
Anonymous. 2013. Michigan vegetable summary 2012. USDA National Agriculture Statistics
Service. NR-13-09.
Baudry, A. 1995. Phytophthora megasperma var. sojae on Asparagus officinalis in France.
Plant Disease 79:1188.
Berger, F., Li, H., White, D., Frazer, R., and Leifert, C. 1996. Effect of pathogen inoculum,
antagonist density, and plant species on biological control of Phytophthora and Pythium
damping-off by Bacillus subtilis Cot1 in high-humidity fogging glasshouses.
Phytopathology 86: 428-433.
Broders, K. D., Lipps, P. E., Paul, P. A., and Dorrance, A. E. 2007. Characterization of Pythium
spp. associated with corn and soybean seed and seedling disease in Ohio. Plant Disease
91:727-735.
Chung, S., Kong, H., Buyer, J. S., Lakshman, D. K., Lydon, J., Kim, S. D., and Roberts, D. P.
2008. Isolation and partial characterization of Bacillus subtilis ME488 for suppression of
soilborne pathogens of cucumber and pepper. Applied Microbiology and Biotechnology
80: 115-123.
Damicone, J. P., Cooley, D. R., and Manning, W. J. 1981. Benomyl in acetone eradicates
Fusarium monoliforme and F. oxysporum from asparagus. Plant Disease 65:892-893.
Elmer, W. H., Johnson, D. A., and Mink, G. I. 1996. Epidemiology and management of the
diseases causal to asparagus decline. Plant Disease 80:117-125.
Falloon, P. G. 1982. Baiting, pathogenicity, and distribution of Phytophthora megasperma var.
sojae in New Zealand asparagus soils. New Zealand Journal of Agricultural Research
25:425-429.
Falloon, P. G. 1990. Field screening of asparagus for tolerance to Phytophthora rot. Acta
Horticulturae 271:69-76.
Falloon, P. G., and Fraser, H. A. 1991. Control of establishment failures in asparagus
(Asparagus officinalis L.) caused by Phytophthora rot. New Zealand Journal of Crop and
Horticultural Science 19:47-52.

53

Falloon, P. G., and Grogan, R.G. 1991. Effect of root temperature, plant age, frequency, and
duration of flooding and inoculum placement and concentration on susceptibility of
asparagus to Phytophthora rot. New Zealand Journal of Crop and Horticultural Science
19:305-312.
Falloon, P. G., Falloon, L. M., and Anderson, A. M. 2002. Breeding asparagus varieties
resistant to Phytopthora. Acta Horticulturae 589:185-191.
Glosier, B. R., Ogundiwin, E. A., Sidhu, G. S., Sischo, D. R., and Prince, J. R. 2008. A
differential series of pepper (Capsicum annuum) lines delineates fourteen physiological
races of Phytophthora capsici. Euphytica 162:23-30.
.
Granke, L. L., Quesada-Ocampo, L. M., and Hausbeck, M. K. 2012. Differences in virulence of
Phytophthora capsici isolates from a worldwide collection on host fruits. European
Journal of Plant Pathology 132:281-296.
Keulder, P. C. 1999. Asparagus decline and replant problem: A review of the current situation
and approaches for future research. Acta Horticulturae 479: 253-262.
Kim, E. S., and Hwang, B. K. 1992. Virulence to Korean pepper cultivars of isolates of
Phytophthora capsici from different geographical areas. Plant Disease 76:486-489.
Lin, D., Zhao, M., Jieqing, P., Johnson, A., Zhang, B., Abney, T. S., Hughes, T. J., and Ma, J.
2013. Molecular mapping of two genes conferring resistance to Phytophthora sojae in a
soybean landrace PI 567139B. Theoretical and Applied Genetics 126:2177-2185.
Madden, L. V., Hughs, G., and van den Bosch, F. 2007. The Study of Plant Disease Epidemics.
The American Phytopathological Society, St. Paul, MN.
Mallard, S., Cantet, M., Massire, A., Bachellez, A., Ewert, S, and Lefebvre, V. 2013. A key
QTL cluster is conserved among accession and exhibits broad-spectrum resistance to
Phytophthora capsici: a valuable locus for pepper breeding. Molecular Breeding 32:349364.
Morrison, W. R. III, Tuell, J. K., Hausbeck, M. K., and Szendrei, Z. 2011. Constraints on
asparagus production: The association of Ophiomyia simplex (Diptera: Agromyzidae)
and Fusarium spp. Crop Science 51:1414-1423.
Rodriguez Salamanca, L. 2010. Management of asparagus rots. M. S. Thesis. Michigan State
University, East Lansing.
Sanders, D. C. 2001. Commercial asparagus production. North Carolina State University
Horticulture Information Leaflet HIL-2-A. Online. February 2014.

54

Saude, C., Hurtado-Gonzales, O. P., Lamour, K. H., and Hausbeck, M. K. 2008. Occurrence
and characterization of a Phytophthora sp. pathogenic to asparagus (Asparagus
officinalis) in Michigan. Phytopathology 98:1075-1083.
Stephens, C. T., and Elmer, W. H. 1988. An in vitro assay to evaluate sources of resistance in
Asparagus spp. to Fusarium crown and root rot. Plant Disease 72:334-337.
Vujanovic, V., Hamel, C., Jabaji-Hare, S., and St-Arnaud, M. 2003. First report of root rot on
asparagus caused by Phytophthora megasperma in Canada. Plant Disease 87:447-447.

55

CHAPTER II

THE EFFECT OF CULTIGENS AND FUNGICIDES AND BIOLOGICAL AGENTS ON
FUSARIUM ROOT ROT OF ASPARAGUS.

56

ABSTRACT
Fusarium crown and root rot (caused by Fusarium oxysporum f. sp. asparagi and F.
proliferatum) is considered to be the most economically limiting disease to asparagus crops
worldwide. Pathogenic isolates of F. oxysporum f. sp. asparagi and F. proliferatum are nearly
ubiquitous in soils, and methods to protect asparagus from disease are limited. F. oxysporum f.
sp. asparagi (isolate FOA-50) and F. proliferatum (isolate P-67) were used to inoculate
seedlings of 11 cultigens under controlled conditions. Significant differences were detected in
isolate virulence and cultigen susceptibility. The F. oxysporum f. sp. asparagi isolate was more
virulent than the F. proliferatum isolate. Reduced susceptibility to Fusarium crown and root rot
was identified in commercial cultivars Jersey Supreme, Mondeo, and Mary Washington, and
breeding line UG005. The interaction between pathogen and cultigen had a significant effect on
disease severity. Commercial cultivars Jersey Supreme and Mondeo had reduced susceptibility to
both pathogens. Five seed treatments, including fungicides and biocontrol agents, were
evaluated against FOA-50 and P-67 on ‘Jersey Giant’. No significant difference in germination
was detected between treatments, but the biocontrol, Bacillus subtilis, reduced disease incidence
compared to the remaining treatments. This is the first reported study on the efficacy of seed
treatments against Fusarium spp. on asparagus. Future studies should evaluate these and
additional seed treatments to determine their efficacy in managing disease in greenhouse and
field conditions.

57

INTRODUCTION
F. oxysporum Schlectend.:Fr. f. sp. asparagi Cohen & Heald and F. proliferatum
(Matushima) Niremburg are the primary causal agents of Fusarium crown and root rot, which is
considered to be the most economically limiting disease to asparagus crops worldwide (Elmer,
2001a; Elmer, 2001b; Schreuder et al., 1995; Van Bakel and Kerstens, 1970; Wong and Jeffries,
2006). F. proliferatum can colonize below- and above-ground tissues of asparagus plants and is
commonly found on mature crowns, while F. oxysporum f. sp. asparagi usually infects young
plants (Elmer et al., 1996; Burgess, 1981; Elmer, 2001a; Keulder, 1999). Fusarium crown and
root rot symptoms begin as reddish brown discoloration in the rhizome, shoot, or storage root
vascular tissues (Zandstra et al., 1992). As the disease progresses, the feeder roots completely
rot, storage roots become hollow, and the fern stand turns yellow and senesces (Elmer et al.,
1996; Zandstra et al., 1992).
F. oxysporum abundantly produces chlamydospores, but can also survive as hyphae on
plant residues in and on the soil. There is no known teleomorph of F. oxysporum (Burgess,
1981). Forty-three vegetative compatibility groups (VCGs) have been reported in F. oxysporum
f. sp. asparagi (Elmer et al., 1996) and it is believed that each VCG is a clonal lineage (Correll,
1991). The pathogenicity of F. oxysporum on asparagus does not appear to correspond with
vegetative compatibility, indicating that pathogenicity on asparagus may be a common trait in F.
oxysporum (Elmer, 2001a; Elmer et al., 1996).
F. proliferatum (Teleomorph G. fujikuroi var. intermedia Kuhlman) is mating population
“D” of the Gibberella fujikuroi complex (Elmer, 1995; Samuels et al., 2001). Perithecia have
not been found in nature (Elmer, 2001a). The anamorph has not been found to produce
chlamydospores and it overwinters as mycelia on plant debris.

58

Pathogenic isolates of F. oxysporum f. sp. asparagi and F. proliferatum are nearly
ubiquitous within the major asparagus producing regions, so choosing fields with pathogen-free
soils may not be an option to prevent Fusarium crown and root rot (Hartung et al., 1990; Elmer,
2001a). Planting disease-free crowns from fumigated nurseries, applying protective crown soaks,
and following good cultural practices to maintain stand vigor are the best management strategies
available (Drost, 1997; Elmer, 2001b; Morrison et al., 2011; Zandstra et al., 1992). Maintaining
crop health by managing foliar pathogens, viruses, insects, and weeds and selecting safe
herbicides is also important to reduce susceptibility to Fusarium crown and root rot (Elmer,
2001b; Hausbeck et al., 1999; Johnson and Lunden, 1992; Morrison et al., 2011; Zandstra et al.,
1992).
Host resistance against Fusarium spp. is a valuable management tool in crops such as
wheat (Burgess et al., 2001) and snap beans (Snapp et al., 2003). Currently, all A. officinalis
cultivars are susceptible to F. oxysporum f. sp. asparagi and F. proliferatum. All-male hybrids
from Rutgers University have higher plant vigor than open-pollinated cultivars, which might
offer some protection against disease (Ellison and Kinelski, 1985). In vitro screens have
identified tolerance to Fusarium spp. in the ornamentals A. densiflorus ‘Sprengeri,’ and
‘Myersii’, however sexual incompatibility between the two species has prevented hybrids from
being deveoped (Dan and Stephens, 1995; Stephens and Elmer, 1988; Stephens et al., 1989). As
resistant cultivars are identified and developed, cultivar selection could become a useful
management tool for growers.
Research of chemical and biological strategies has shown varying levels of disease
control. Historically, sodium chloride was believed to control the Fusarium crown and root rot
(Elmer, 2001a), but growth chamber, greenhouse, and field studies have not resulted in

59

consistent disease control. NaCl does not appear to benefit healthy stands, the mode of action
and environmental impact is unknown, and it exacerbates Phytophthora crown and root rot
(Elmer, 2004; Morrison et al., 2011; Reid et al., 2001). Therefore it is not currently
recommended for use. Seed disinfestation with benomyl in acetone is useful to remove seedborne Fusarium spp. (Damicone, 1981), but does not provide protection from other soil-borne
pathogens. Fungicides such as Cannonball and Topsin, applied as a crown soak, have resulted in
improved spear and fern production (Hausbeck and Cortright, 2009). Biological agents
including nonpathogenic Fusarium spp., arbuscular mycorrhizae, and Trichoderma harzianum
have reduced disease and increased root weight in some studies, but in other trials these
treatments did not affect crown health (Arriola et al., 2000; Counts and Hausbeck, 2008; Elmer,
2004; Elmer, 2008).
Young asparagus seedlings are especially susceptible to F. oxysporum f. sp. asparagi
(Elmer et al., 1996 and Keulder, 1999). Nursery beds are sometimes fumigated to remove
pathogen populations, and growers are advised to surface disinfest their seeds before planting
(Zandstra et al., 1992). Currently, Michigan growers do not use fungicides or biocontrols to
protect seedlings grown in nursery fields (Hausbeck, personal communication, 2014).
Identifying and developing effective seed treatments against Fusarium crown and root rot would
be useful for producing disease-free crowns.
The objectives of this study were to: i) screen seedling varieties for relative susceptibility
and ii) test chemical and biological products for efficacy as seed treatments against Fusarium spp.

60

MATERIALS AND METHODS
Fusarium spp. and inoculum preparation. Michigan isolates of F. oxysporum f. sp.
asparagi (FOA-50) and F. proliferatum (P-67) from the collection of M. K. Hausbeck were
taken from long-term storage by aseptically transferring infested silica crystals to PDA (39 g
potato dextrose agar and 1000 ml distilled water) plates. Cultures were maintained at room
temperature (23 ± 2°C) under continuous fluorescent lighting and transferred every 7 to 10 days.
Spore suspensions were prepared by flooding cultures after fungal growth covered the
entire disc with sterile distilled water and scraping with a flame-sterilized glass rod. The
resulting spore suspensions were collected and diluted to 106 conidia/ml for FOA-50 and 105
conidia/ml for P-67 using a hemocytometer (Stephens and Elmer, 1988). Suspensions were used
within 30 minutes of preparation.
Seedling inoculation and incubation. Asparagus seeds were surface disinfested using
the method of Damicone et al. (1981) by agitating in solutions of 25,000 ppm benomyl in
acetone on a rotary shaker (1000 rpm) for 24 hours, then rinsing 3 times in acetone and 3 times
in distilled water. Seeds were then agitated in 1.23% sodium hypochlorite at 1000 rpm for 1
hour and rinsed 3 times in distilled water. After being allowed to air dry for 1 to 2 hours in a
laminar flow hood, the seeds were placed on water agar (16 g agar and 1000 ml distilled water)
and germinated in the dark at room temperature for 7 to 14 days. Once hypocotyls emerged, the
seeds were planted in 13 ml of Hoagland’s agar (Stephens and Elmer, 1988) in test tubes (25 x
150 mm) and grown under continuous fluorescent light at room temperature.
When seedlings were approximately 11 cm in height and secondary branches with
cladophylls had formed (approximately 2 weeks), a 0.5 ml aliquot of FOA-50 (106 conidia/ml) or
P-67 (105 conidia/ml) was placed directly on the agar of each test tube. Following inoculation,

61

seedlings were allowed to grow and infection to develop for 4 weeks under laboratory conditions
described above.
Susceptibility of cultivars and breeding lines to seedling root rot. Eleven cultivars
and breeding lines (Table 2.1) were selected to compare relative susceptibility against F.
oxysporum f. sp. asparagi and F. proliferatum. Seeds were disinfested and germinated using the
methods described above. When the seedlings were approximately 11 cm in height and
secondary branches with cladophylls had formed (approximately 2 weeks), conidial suspensions
(0.5 ml) were added to test tubes. Sterile distilled water (0.5 ml) was used for uninoculated
controls. Four seedlings were used for each cultigen. Seedling test tubes were arranged in a
randomized complete block design in test tube racks.

Table 2.1. Asparagus cultivars and breeding lines tested for susceptibility to Fusarium
oxysporum f. sp. asparagi and F. proliferatum.
Cultigen
Source
Millennium
University of Guelph
UG005
University of Guelph
UG009
University of Guelph
UG010
University of Guelph
UG020
University of Guelph
Mary Washington
Eden Brothers
NJ 938
Rutgers University
NJ 941
Rutgers University
NJ 1191
Rutgers University
Jersey Supreme*
Vilmorin
Mondeo*
Vilmorin
*Packaged seeds were pretreated with thiram.

Seedlings were rated for disease incidence and severity of root rot every 7 days beginning
7 days post inoculation (dpi). Disease incidence was recorded, and severity was measured as a
visual estimate of the area (%) of each root system with Fusarium lesions. For the final rating
(week 4), seedlings were removed from the test tubes and the roots were mechanically separated

62

from the agar. The area under the disease progress curve (AUDPC) was calculated (Madden et
al., 2007) to obtain the cumulative severity (%) throughout the experiments. The experiment
was conducted 4 times.
Efficacy of fungicides and biocontrol agents as seed treatments. The methods used by
Broders et al. (2007) were adapted to screen three fungicides and two biocontrol agents for
efficacy as seed treatments in vitro. A mycelial plug (7-mm in diameter) of FOA-50 or P-67 was
placed in the center of 15-cm-diameter petri plate containing 50 ml of PDA. Seeds of
‘Millennium’ were surface disinfested according to the methods of Damicone et al. (1981) and
allowed to air dry in a laminar flow hood (approximately 1 hour). Treatments were applied to
seed lots (0.33 g) at the rates listed in Table 2.2. Seeds soaked in the treatments for
approximately 30 minutes, then air-dried in a laminar flow hood for approximately 1 hour.
Seeds from each treatment were equally spaced in a randomized order around the plate,
approximately 3 cm from the edge (Figure 2.1).
At 11 days after plating the seeds, inhibition of mycelial growth was measured as the
distance between the seed and margin of mycelial growth along the radius of the agar plate. The
seeds were removed from culture and successfully germinated seeds were rated for disease
severity. The number of seeds that successfully germinated on inoculated plates was compared
with the number that germinated on the uninoculated plates. Seeds were considered successfully
germinated if the hypocotyl had emerged or the radicle was > 5 mm long. Treatments were
scored according to a 0 to 3 rating scale, where 0 = 100% germination with no disease symptoms
and 3 = all seed tissues colonized (Table 2.3, Figure 2.2). Each treatment was replicated 5 times
for each isolate and the uninoculated control, and the experiment was conducted a total of 4
times: twice with ‘Jersey Giant.’

63

Table 2.2. Fungicides and biocontrol agents tested for efficacy against Fusarium oxysporum f.
sp. asparagi and F. proliferatum.
Treatment
Active ingredient
Rate/100 lb seed
Rate/20 seeds (0.48 g)
Sterile deionized water
Untreated
600.0 µl
azoxystrobin
44.4 ml
Dynasty
0.47 µl
penthiopyrad
29.6
ml
Fontelis
0.31 µl
fludionoxil
4.5 g
Maxim
0.05 mg
*
340.2 g
Streptomyces lydicus
Actinovate AG
3.60 mg
*
813.2
ml
Bacillus subtilis
Serenade Soil
8.61 µl
*

Biological control agent

Table 2.3. Scale of disease ratings on successfully germinated asparagus seedlings inoculated
with Fusarium oxysporum f. sp. asparagi and F. proliferatum.
Rating
Symptoms
0
no disease
1
<50% radicle tissue covered in lesions
2
>50% radicle covered in lesions
3
All viable seed tissues covered in lesions

Pathogen isolation. At the end of the observation period for the seedling root rot
experiment, small sections of root tissue from lesion margins were plated on Komada’s agar
(1000 ml distilled water, 20 g D-galactose, 15.0 g agar, 2.0 g L-asparagine, 1.0 g K2HPO4, 1.0 g
pentachloronitrobenzene, 0.5 g KCl, 0.5 g MgSO4 7H2O, 0.1 g Fe-Na-EDTA, 10 ml oxgall
solution, and 6 ml streptomycin), a media selective for Fusarium spp. growth. All seedlings
were sampled.
For the seed treatment experiment, whole seeds were disinfested in 0.525% sodium
hypochlorite for approximately 60 seconds, rinsed in sterile distilled water, and air dried in a
laminar flow hood for 1 to 3 minutes. Radicle tissue was wounded using a flame-sterilized
scalpel and plated in the method described above. All inoculated seeds were sampled, and
control seeds were also sampled if radicles had water-soaked or discolored tissue.

64

A

B

Figure 2.1. Asparagus seeds treated with fungicides and biocontrol agents on V8
agar plates inoculated with Fusarium oxysporum f. sp. asparagi (A) and F.
proliferatum (B), 3 days after planting.
Statistical analysis. In the cultigen experiment, disease severity data was tested by
analysis of variance (ANOVA) using the PROC GLIMMIX procedure of SAS version 9.3 (SAS
Institute Inc., Cary, NC). The PROC MIXED procedure was used to analyze AUDPC data. Trial
repetition was treated as a random affect for both data sets. Cultigen was treated as the group
effect in the REPEATED statement for AUDPC data. Residuals were tested for normality and
homogeneity of variance using PROC UNIVARIATE. Data from the control plants were
removed prior to analyses to avoid violating the assumption of equal variances. Multiple
comparisons among the cultigen means were conducted using ANOVA and Tukey’s adjustments
were used for separation of means when effects were statistically significant at P = 0.05.

65

A

B

C

D

Figure 2.2. Fusarium symptoms observed on germinated asparagus seedlings. (A) 0 = no
disease symptoms, (B) 1 = lesions present on <50% radicle tissue, (C) 2 = lesions present on
>50% radicle tissue, and (D) 3 = all seed tissues covered in lesions.

For the seed treatment experiment, germination data was tested by analysis of variance
(ANOVA) using the PROC MIXED procedure. Treatment was treated as the group effect in the
REPEATED statement. Residuals were tested for normality and homogeneity of variance using
PROC UNIVARIATE. Data was square root transformed prior to analysis to avoid violating the
assumption of equal variances. Disease incidence data were analyzed using analysis of variance
with the AOV command of R version 3.01 (The R Foundation for Statistical Computing, Vienna,
Austria). Multiple comparisons among the cultigen means were conducted using a Fischer’s
Least Significant Differences (LSD) test (Agricolae package) at P = 0.05.

66

RESULTS
Susceptibility of cultivars and breeding lines to seedling root rot. At four weeks post
inoculation, all inoculated plants were diseased. Seedlings infected with both Fusarium spp.
developed dark red-brown lesions with distinct margins. Root tissue of uninoculated controls
remained vigorous without discoloration. The respective pathogen was successfully isolated
from seedlings infected with F. oxysporum f. sp. asparagi and F. proliferatum. An unidentified
Fusarium spp. was also isolated from approximately 3% of the asymptomatic uninoculated
controls (NJ938 and NJ1191).
The interaction between pathogen and asparagus cultigen was significant (P >0.05) for
root rot severity. The main effects, cultigen (P = 0.05) and isolate (P = 0.05) were significant. F.
oxysporum f. sp. asparagi caused significantly higher severity in cultivars Mary Washington and
Mondeo and breeding lines NJ941, UG009, and UG020 compared to F. proliferatum. F.
proliferatum caused higher disease severity in UG010. NJ941 was most susceptible to both
pathogens. ‘Jersey Supreme’ was significantly healthier than ‘Millennium,’ UG009, NJ1191,
and NJ941 when inoculated with F. oxysporum f. sp. asparagi. UG005, UG009, UG020,
‘Mondeo,’ and NJ938 were less susceptible than NJ941 and NJ1191 when inoculated with F.
oxysporum f. sp. asparagi. Cultivars Mary Washington, Mondeo, Supreme, and breeding line
UG020 were significantly less susceptible than NJ938, NJ941, NJ1191, and UG010 when
inoculated with F. proliferatum (Table 2.4).
The interaction between pathogen and cultigen significantly affected AUDPC. Infection
by F. oxysporum f. sp. asparagi resulted in a higher AUDPC than F. proliferatum in most
cultigens, but was only significantly higher in NJ941. There were no significant differences
between cultigens when inoculated with F. proliferatum (Table 2.5).

67

Efficacy of fungicides and biocontrol agents as seed treatments. At 11 days post
inoculation, both pathogens had colonized most of the agar disc area, including the area
immediately surrounding seeds. Seeds from all treatments, except for Bacillus subtilis, were
colonized with mycelial growth. B. subtilis growth was observed on the treated seeds and the
agar surrounded the seeds. Average inhibition of mycelial growth caused by B. subtilis was
observed at an average length of 8.5 mm from the bacterial growth in plates inoculated with F.
oxysporum f. sp. asparagi and immediately surrounding the bacterial growth in F. proliferatum
cultures. F. oxysporum f. sp. asparagi and F. proliferatum were readily isolated from seed
tissues in all treatments (100%), except for the B. subtilis treatment (0 and 10%).
The interaction between pathogen and treatment (P = 0.05) was did not significantly
affect germination or incidence. The main effect pathogen (P = 0.05) significantly affected
germination, but main effect treatment (P = 0.05) was not significant. Both pathogens reduced
seed germination compared to the uninoculated seeds, and F. proliferatum infection resulted in
significantly lower germination (2.5%) than seeds inoculated with F. oxysporum f. sp. asparagi
(40%). Pathogen (P = 0.05) did not have a significant effect on disease incidence. Seeds treated
with B. subtilis had significantly lower disease incidence than the remaining treatments (Table
2.6). All germinated seedlings inoculated with F. proliferatum were given the highest severity
rating. Of the seedlings inoculated with F. oxysporum f. sp. asparagi, only one that was treated
with B. subtilis (N = 1) had a severity rating of 0 (Figure 2.2). However, the seedling was
stunted with discoloration at the radicle tip. All germinated seedlings treated with azoxystrobin
(N = 3), penthiopyrad (N = 3), and Streptomyces lydicus (N = 2) were given a rating of 3 (Figure
2.2).

68

Table 2.4. Average disease severity (%) of asparagus seedlings infected with Fusarium
oxysporum f. sp. asparagi and F. proliferatum at 28 days post inoculation.
x
Disease
severity
(%)
Cultigen
F. oxysporum f. sp. asparagi
F. proliferatum
y
Millennium
12.7 BCD
14.8 BC
UG005
11.5 AB
10.6 ABC
UG009
15.0 *BC
12.4 ABCD
UG010
11.5 AB
16.2 *D
UG020
12.7 *AB
10.0 AB
Mary Washington
14.2 *ABC
7.1 A
NJ938
13.3 AB
14.5 CD
NJ941
34.2 *D
28.0 E
NJ1191
18.6 C
15.9 D
Jersey Supreme
10.3 A
8.9 AB
Mondeo
12.1 *AB
8.0 A
x
Seedlings were rated by a visual estimation of the percentage of root system infected. Each
disease severity value represents the average of 4 repeated tests with 4 replicate asparagus
seedlings per pathogen/cultigen combination per test.
y
Disease severity values with the same letter are not significantly different among cultigens for
each pathogen (within columns). Tukey's, P=0.05.
*Pathogens within a cultigen are significantly different (within rows). LSD, P = 0.05

Table 2.5. Area under the disease progress curve (AUDPC) of asparagus seedlings infected with
Fusarium oxysporum f. sp. asparagi and F. proliferatum.
x
AUDPC
Cultigen
F. oxysporum f. sp. asparagi
F. proliferatum
y
Millennium
233.0 A
318.3 A
UG005
365.3 A
242.8 A
UG009
271.3 A
262.5 A
UG010
248.3 A
220.9 A
UG020
309.5 A
225.3 A
Mary Washington
340.2 A
249.4 A
NJ938
352.6 A
356.6 A
NJ941
1097.2 *B
448.0 A
NJ1191
378.9 A
296.6 A
Jersey Supreme
362.0 A
263.6 A
Mondeo
254.8 A
251.6 A
x
AUDPC values were calculated from weekly severity ratings over the period of 4 weeks.
y

AUDPC values with the same letter are not significantly different among pathogens for each
cultigen (lower case) or between cultigens for each pathogen (upper case). Tukey's, P=0.05.
*Pathogens within a cultigen are significantly different (within rows). LSD, P = 0.05

69

Table 2.6. The effect of Fusarium spp. and seed treatment on germination and disease incidence.
z
Germination (%)
Treatment
Disease incidence (%)
F. oxysporum f. sp.
F. proliferatum
asparagi
100.0 By
Untreated
33.3
10.0
Azoxystrobin
Penthiopyrad
Fludionoxil
Streptomyces lydicus
Bacillus subtilis
z

75.0
62.5
58.3
50.0
50.0

50.0
0.0
0.0
10.0
0.0

100.0
93.8
100.0
100.0
6.3

B
B
B
B
A

Germination percentages were calculated for each treatment as (GI/G0)*100, where GI =

number of inoculated seeds that germinated and G0 = number of uninoculated seeds that
germinated
y
Disease incidence values with the same letter are not significantly different among seed
treatments. LSD, P=0.05.

70

Disease rating distribution (%)

100

N=2

N=3

N=3

N=3

N=2

N=1

80

60

40

20

0
ed
eat
r
t
Un

d
in
yra
rob
p
t
s
o
i
y
ox
nth
Az
Pe

il
s
us
tili
xon
dic
b
o
y
l
i
u
s
d
ces
lus
Flu
my
cil
o
a
t
B
ep
Str

Seed treatment

Severity=3
Severity=2
Severity=1
Severity=0

Figure 2.3. The distribution of disease rating scores of germinated seedlings inoculated
with Fusarium oxysporum f. sp. asparagi and treated with fungicides and biocontrol agents.

71

DISCUSSION
Effective management strategies against Fusarium crown and root rot are limited, and
currently the best control methods are good cultural practices (Drost, 1997; Elmer, 2001b;
Zandstra et al., 1992). Resistant cultivars, a useful tool to manage disease, are not currently
available in Asparagus officinalis. In this study, all cultigens were susceptible to both Fusarium
species. However, cultigens Jersey Supreme, Mondeo, and UG020 were less susceptible to both
pathogens; NJ938, UG005, and UG010 were less susceptible to F oxysporum f. sp. asparagi; and
Mary Washington was less susceptible to F. proliferatum. NJ941 was consistently the most
susceptible to Fusarium crown and root rot. This information may be valuable to growers as
well as to breeding programs for cultivar selection.
When asparagus seeds were inoculated with Fusarium spp. in vitro, damping-off
incidence was high for all seed treatments except for Bacillus subtilis. Seeds not affected by
Fusarium symptoms were surrounded by B. subtilis growth, which antagonized mycelial growth
on the agar. This is consistent with prior studies, where B. subtilis effectively inhibited
Fusarium spp. in vitro (Chung et al., 2008). B. subtilis produces several antibiotics with
antifungal and antibacterial properties, which has resulted in effective control of a wide range of
plant pathogens in laboratory and greenhouse conditions (Berger et al., 1996; Chung et al., 2008).
A number of seedling radicles within the B. subtilis treatment group were stunted and discolored,
with an absence of root hairs. Because the in vitro conditions were highly favorable for the
bacterium, it is likely that bacterial growth overwhelmed the young seedling tissues. B. subtilis
should be studied further in soil-based assays to determine if the seed treatment is safe for young
asparagus roots before it can be recommended for use on seeds or young seedlings growing in
nursery fields.

72

In this study, the unequal number of germinated seedlings across treatments made disease
incidence a more reliable measure of treatment effectiveness than disease severity. The same
active ingredients used in this study could also be applied to seedling roots following
germination to measure the effect of fungicides and biocontrol agents on Fusarium root rot
severity. In vitro conditions, with an abundant nutrient supply, lack of competition and
antagonism from microbial communities, and favorable environmental conditions for growth and
infection may have increased the pathogens ability to rapidly colonize seed tissues from what
would normally occur in soil. Future research should examine the efficacy of these and other
seed treatments in soil-based assays to better represent field conditions.
In order to effectively manage Fusarium crown and root rot, growers are advised to
combine control strategies. Careful selection of asparagus cultivars and integrating chemical and
biological treatments is useful in a coordinated approach to disease management. The results of
this study suggest that specific hybrids from the Guelph, Jersey, and German asparagus lines
may be helpful in asparagus cultivation.

73

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74

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