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THESIS

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LIBRARY

Michigan State
University 5

 

 

 

This is to certify that the

thesis entitled

Evaluation of Strategies for the Control of Fusarium Crown
and Root Rot of Asparagus

presented by

Taylor C. Reid

has been accepted towards fulﬁllment
of the requirements for

Masters degree in Science

    

Major professor

Date December 8, 2000

O~7 639 MS U is an Afﬁrmative Action/Equal Opportunity Institution

 

PLACE IN RETURN BOX to remove this checkout from your record.
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DATE DUE

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EVALUATION OF STRATEGIES FOR THE CONTROL OF F USARIUM CROWN
AND ROOT ROT OF ASPARAGUS

By

Taylor C. Reid

A THESIS

Submitted to
Michigan State University
in partial fulﬁllment of the requirements
for the degree of

MASTER OF SCIENCE

Department of Botany and Plant Pathology

2000

ABSTRACT

EVALUATION OF THE STRATEGIES FOR THE CONTROL OF FUSARIUM
CROWN AND ROOT ROT OF ASPARAGUS

By
Taylor C. Reid

Potential strategies for the control of F usarium crown and root rot of asparagus, caused by
the fungal pathogens F usarium proliferatum, and F usarium oxysporum f. sp. asparagi
were explored. The effectiveness of sodium chloride (N aCl), and lime applications were
investigated in a research ﬁeld and two ﬁelds (one healthy, one declining) in commercial
asparagus production. NaCl applications (1120 kg/ha) were effective in signiﬁcantly
increasing yield and fern health in the severely declined research plot, but neither NaCl or
lime applications had a signiﬁcant effect in commercial ﬁelds. Alternative forms of
chloride salt were tested for their ability to control Fusarium disease on seedlings in
growth chamber and greenhouse studies. No salt treatment was more effective than NaCl
at reducing disease in these tests. The effectiveness of fungicides and biological controls
in controlling Fusarium disease was tested in growth chamber, greenhouse and ﬁeld
studies. Five non-pathogenic strains of F oxysporum signiﬁcantly reduced disease
caused by both F. proliferatum, and F. oxysporum f. sp. asparagi in growth chamber tests
on asparagus seedlings in culture tubes, but not in greenhouse studies with seedlings in
soil. The fungicide ﬂudioxonil was effective in controlling seedling death in greenhouse
tests with high pathogen inoculum, and ﬂudioxonil, benomyl, and Trichoderma
harzianum signiﬁcantly reduced disease compared to the control in greenhouse tests with

a low pathogen inoculum. Field experiments with fungicides and biological controls

were not successful due to low disease pressure.

DEDICATION

To my father,
for instilling in me
a passion for the natural world,
and for giving me the freedom
to explore it;
and to the memory of
my ﬁend

Jeph Stemheim

iii

ACKNOWLEDGMENTS

/

I would like to thank my major advisor, Dr. Mary Hausbeck for her invaluable
insights, enthusiasm, criticism, and support, and for believing in me even when I was
unsure. I would like to thank Dr. Wade Elmer for his time and his tutelage, and the other
members of my committee, Dr. Darryl Warncke, Dr. Gene Saﬁr, and Dr. Dennis
Fulbright for their assistance and suggestions. Kadir Kizilkaya has been extremely
helpful and patient with statistical analysis, and I have been helped in many ways by
people working in the Hausbeck lab and in The Department of Botany and Plant
Pathology. 1 would also like to extend my gratitude to Mary Jo and John Bakker, and
Norm Myers in Oceana County, and to growers Ken and Ralph Oomen, Tom and Rick
Oomen, and Bob Schaffer for their cooperation and generosity.

I would also like to thank Amy, Lola, Alana, Cathy, Jim, Sue, Robyn, Whitney,
Charlie, Kelly, Ngoc, Julie, Chris, and all the folks at PB, and give special thanks to Kurt,

Kierstyn, and Iris Lamour for their friendship and company which has been a great gift to

me during my time here.

iv

TABLE OF CONTENTS

LIST OF TABLES ...................................................... vi
LIST OF FIGURES .................................................... vii
LITERATURE REVIEW .................................................. 1
Asparagus 2
Fusarium Crown and Root Rot 13
Literature Cited 42
SECTION I.

EFFECT OF SODIUM CHLORIDE AND LIME APPLICATIONS ON COMMERCIAL
ASPARAGUS AND THE EFFECT OF ALTERNATE FORMS OF CHLORIDE SALT

ON F USARIUM CROWN AND ROOT ROT ................................ 62
Abstract 63
Introduction 64
Materials and Methods 65
Results 71
Discussion 84
Literature Cited 89

SECTION II.

THE USE OF FUNGICIDES AND BIOLOGICAL CONTROLS IN THE
SUPPRESSION OF FUSARIUM CROWN AND ROOT ROT OF ASPARAGUS . . . 92

Abstract 93
Introduction 94
Materials and Methods 95
Results 102
Discussion 1 11
Literature Cited 114
APPENDIX .......................................................... 120
Section I. 120
Section II 126

LIST OF TABLES

Table

1. Mean data for soil samples from 4/98 (before any treatments were

applied) and 5/00 (after two years of treatment) in Oceana County ﬁeld

plots, tables show combined data for both ﬁelds unless signiﬁcant ﬁeld

by treatment effects were found, letter notation indicates signiﬁcant differences
between treatment means from 2000 with 1998 data used as a covariate.

2. Mean data for soil samples from 4/98 (before any treatments were
applied) and 5/00 (after two years of treatment) at MSU Botany and
Plant Pathology Farm, letter notation indicates signiﬁcant differences
between treatment means from 2000 with 1998 data used as a covariate
for top and bottom data separately.

vi

114

115

LIST OF FIGURES

Figure Page

1. EFFECT OF SODIUM CHLORIDE AND LIME APPLICATIONS ON
COMMERCIAL ASPARAGUS AND THE EFFECT OF ALTERNATE FORMS OF
CHLORIDE SALT ON FUSARIUM CROWN AND ROOT ROT

1. Effect of different forms of chloride salt on root rot (%) in two experiments with
asparagus seedlings grown in Hoagland’s agar in test tubes and infested with F.
proliferatum. 73

2. Effect of different forms of chloride salt on root rot (%) in two experiments with
asparagus seedlings grown in Hoagland’s agar in test tubes and infested with F
oxysporum f. sp. asparagi. 74

3. Effect of different forms of chloride salt on root rot (%) in two greenhouse
experiments with asparagus plants grown in soil infested with F. oxysporum
f. sp. asparagi and F. proliferatum. 75

4. Effect of different forms of chloride salt on root weight (g) in two greenhouse
experiments with asparagus plants grown in soil infested with F. oxysporum
f. sp. asparagi and F. proliferatum. 76

5. Effect of different forms of chloride salt on shoot weight (g) in two greenhouse
experiments with asparagus plants grown in soil infested with F. proliferatum

and F. oxysporum f. sp. asparagi. 78

6. Mean harvest weight of asparagus spears from commercial ﬁeld plots in Oceana
County, M1 1998-2000. 79

7. Percentage of asparagus stalks > 0.79 cm in diameter in commercial ﬁeld plots
in Oceana County, M1 1998-2000. 80

8. Mean harvest weight of asparagus spears from research plot at MSU. Botany
and Plant Pathology Farm 1998-2000. . 81

9. Percentage of asparagus stalks >' 0.79 cm in diameter from research plot at
MSU. Botany and Plant Pathology Farm 1998-2000. 82

vii

LIST OF FIGURES (Continued)

Fi ure

II. THE USE OF FUNGICIDES AND BIOLOGICAL CONTROLS IN THE
SUPPRESSION OF FUSARIUM CROWN AND ROOT ROT OF ASPARAGUS

10. Average number of asparagus plants dead in two greenhouse experiments

11.

12.

13.

14.

15.

when grown in soil infested with a high level of F. proliferatum and F.
oxysporum f. sp. asparagi and treated with a chemical or biological control
product.

Shoot weight of asparagus seedlings from two greenhouse experiments
when grown in soil infested with a low level of F. proliferatum and F.
oxysporum f. sp. asparagi and treated with a chemical or biological control
product.

Root weight of asparagus seedlings from two greenhouse experiments
when grown in soil infested with a low level of F. proliferatum and F.
oxysporum f. sp. asparagi and treated with a chemical or biological control
product.

Roots of asparagus seedlings exhibiting lesions (%) in two greenhouse
experiments when grown in soil infested with a high level of F.
proliferatum and F. oxysporum f. sp. asparagi and treated with a chemical
or biological control product.

Root system of asparagus seedlings exhibiting lesions (%) in two growth
chamber experiments when grown in culture tubes infested with F usarium
oxysporum f. sp. asparagi and treated with non-pathogenic F usarium
oxysporum.

Root system of asparagus seedlings exhibiting lesions (%) in two growth
chamber experiments when grown in culture tubes infested with F usarium
Proliferatum and treated with non-pathogenic F usarium oxysporum.

APPENDIX 1.

16.

Mean sodium level in soil samples from MSU Botany and Plant Pathology
Farm ﬁeld 4/98-5/00 in parts per million.

viii

103

105

106

107

109

110

123

LIST OF FIGURES (Continued)

Figge

17. Mean sodium level in soil samples from healthy ﬁeld in Oceana County

18.

between 4/98 and 5/00 in parts per million.

Mean sodium level in soil samples from declined ﬁeld in Oceana County
between 4/98 and 5/00 in parts per million.

APPENDIX II.

19.

20.

21.

22.

23.

Mean fern height in two seedling nursery trials with chemical and
biological controls.

Number of shoots per plant in two seedling nursery trials with chemical
and biological controls.

Fresh weight of shoots in two seedling nursery trials with chemical and
Biological controls.

Fresh weight of roots in two seedling nursery trials with chemical and
Biological controls.

Percentage of root system exhibiting lesions in two seedling nursery trials
with chemical and biological controls.

ix

124

125

127

128

129

130

131

LITERATURE REVIEW

ASPARAGUS

Asparagus (Asparagus ofﬁcinalis L.) is an important vegetable crop throughout
the world. A member of the Liliaceae (lily) family, the genus has 150 species, many of
which are grown for ornamental purposes (Kotecha and Kadam, 1998). Although its
center of origin is not known with certainty, it is generally considered to be native to the
eastern Mediterranean region (Nonnecke, 1989). Reports of asparagus cultivation date
back as far as Cato (200 B.C.), and its name comes from the ancient Greeks (Kidner,
1959). Pliny, writing in the time of Christ, described the plant much as we see it today
(Kidner, 1959). In ancient Europe asparagus was used as a diuretic, sedative, and pain
killer (Drost, 1997). Cultivation was perfected by the Romans who grew their asparagus
in trenches, the favored method of culture until the 1800's (Kidner, 1959).

Asparagus is a dioecious, herbaceous perennial, the below ground parts of which
are collectively referred to as the crown, while the above ground portion is called either
spears or fern, depending on its stage of growth (Drost, 1997). The crown is composed of
the rhizome which produces the buds, adventitious roots, and storage roots (Drost, 1997).
Each crown produces shoots throughout the season from the buds borne on the rhizome.
These shoots, often called spears, are harvested when they are young and tender (Kotecha
and Kadam, 1998). When left to grow after the harvest season, the shoots grow into
bushy plants up to two meters in height which are referred to as ferns. The true leaves of
the plant are inconspicuous ﬂat scales pressed close to the stalk, while the major
photosynthetic organs are the myriad needle-like cladodes (cladophylls, phylloclades) that

proliferate along the stems (Anonymous, 1960). The ﬂowers of the plant are greenish-

white or yellow, and male and female ﬂowers are borne on separate individuals, though
some may be perfect, having both pistils and stamens (Green, 1896). Female and perfect
ﬂowers will produce red berries in the fall, bearing a single hard black seed.

North America and Europe are the largest asparagus producers, though world
production has increased greatly in the last few years. Taiwan, Japan, China, Peru, Chile,
Australia, New Zealand, and South Africa all have signiﬁcant production acreage (Drost,
1997). Europeans are partial to white asparagus, etiolated by mounding earth over the
beds, while most U.S. consumers prefer green asparagus, grown aboveground. In 1997,
29,800 ha of asparagus were grown in the U.S., with a production value of over 180
million dollars (National Agricultural Statistics Service, 1998). This represents just a
fraction of world production, which is estimated at 218,395 ha (Benson, 1997). The U.S.
is the world’s second largest producer after China whose estimated production is 55,000
ha and is increasing (Benson, 1997). Peru and Spain follow with about 20,000 ha each,
while Germany, France, and Mexico each produce over 10,000 ha (Benson, 1997). Most
U.S. asparagus is grown in California (12,180 ha), Washington (9,310 ha), and Michigan
(7,085 ha), with other states growing a signiﬁcantly smaller fraction for local fresh
markets (Michigan Department of Agriculture, 1998). In Michigan, 79% of the total
production value was from processing in 1997, though the fresh market has seen growth
in recent years (Michigan Department of Agriculture, 1998).

Asparagus grows best on a light, well drained soil like the loamy sands of Western
Michigan (Kotecha and Kadam, 1998). Some asparagus is direct seeded, but most is

transplanted from seedling nurseries after a year of growth. Seeds are planted 2.5-5 cm

deep, at a rate of 7-9 kg per ha, with rows spaced 60-76 cm apart (Kee et al., 1999;
Kotecha and Kadam, 1998; Ware and McCollum, 1975). In the spring of the second year,
before growth commences, crowns are dug from these seedling nurseries for planting in
the ﬁeld. One ha of seedlings will produce enough crowns for approximately 5 ha of
production ﬁeld (Zandstra etal., 1992). Furrows for crown planting are set 1.22-1.525 m
apart, and are plowed to a depth of 15-25 cm (Kee et al., 1999; Kotecha and Kadam,
1998; Ware and McCollum, 1975). Most publications recommend a crown spacing of
30.5-45.7 cm apart within the furrows, although recent research has shown a closer
spacing to be favorable, and Michigan farmers are beginning to adopt this practice (Kelly
et al., 1997). Crowns are covered by about 5 cm of soil, and trenches are ﬁlled in
gradually as plants emerge (Ware and McCollum, 1975). The older, open pollinated
cultivars such as ‘Viking KB3’ and ‘Mary Washington’, have been replaced with the
more vigorous all male hybrids, such as ‘Jersey Giant’ and ‘Jersey Knight’, or with other
hybrids such as ‘Synthetic 4-56’, which is about 85% male (Foster et al., 1999).

Harvesting of asparagus typically begins in the third season of growth, but is
limited to two weeks (Kotecha and Kadam, 1998; Zandstra et al., 1992). Beginning the
fourth year, harvesting occurs for approximately six weeks or until the majority of spears
become thin and unmarketable (Zandstra et al., 1992). In Michigan and Washington,
harvest usually begins in early May, while Califomia’s season begins in February (Ware
and McCollum, 1975; Zandstra et al., 1992). Spear harvest varies among regions, and is
treated differently for the fresh or processing markets. In some areas, spears are

mechanically harvested or hand cut below the soil surface (Kotecha and Kadam, 1998). In

Michigan, spears are harvested once they reach approximately 12.7 cm in height by hand
snapping them at the base (Zandstra et al., 1992). Workers ride on motorized carts that
span 3-7 rows, and harvest the asparagus in front of them as they pass. Fresh market
asparagus is picked longer to allow trimming for uniformity. Asparagus is capable of
rapid growth (up to 1.6 cm/day) dun'ng warm weather (Kotecha and Kadam, 1998).
Thus, the crop must be harvested every day when temperatures are high. Hot weather can
also cause premature opening of heads which will result in a decrease in quality (Zandstra
etaL,1992)

Two federal grades (U.S. No. 1, and U.S. No. 2) apply to fresh market asparagus,
and these are further divided by spear diameter into Colossal, Fancy and Choice sizes,
(N onnecke, 1989). The Michigan asparagus industry has established guidelines for
processing asparagus which include a minimum spear diameter of 0.79 cm (5/16 inches)
12.7 cm (5 inches) below the tip, no white butts, no long spears, and no beetle eggs
(Zandstra etal., 1992). The tolerance of the industry often depends on the time of year
and the demand for product.

It is recommended that fresh market asparagus be cooled as soon as possible after
harvest (Zandstra et al., 1992) (Kotecha and Kadam, 1998). Hydrocooling or vacuum-
cooling are both feasable, though the latter is less effective (Kotecha and Kadam, 1998;
Wills et al., 1989). Warm asparagus will very quickly develop increased ﬁber, open tips,
wilted stalks, a loss of ﬂavor, and a loss of vitamin C (Kotecha and Kadam, 1998;
Zandstra et al., 1992). Asparagus can be kept for no longer than three weeks at 200 C

(Kotecha and Kadam, 1998; Zandstra et al., 1992). Biochemical and physiological

changes in storage can include spear lengthening, increased ﬁber, and decreased sugar
(Kotecha and Kadam, 1998). Lower temperatures will decrease these changes, and
asparagus can be stored at 0° C for up to 10 days, but is subject to chilling injury if held
longer at this temperature (Kotecha and Kadam, 1998; Wills et al., 1989). A high relative
humidity must also be maintained in order to prevent dessication (Kotecha and Kadam,
1998; Zandstra et al., 1992).

Since most Michigan asparagus is grown for the processing market, several local
and national processing plants, are set up in the major production areas. Most processing
asparagus is dumped into bulk boxes in the ﬁeld, and taken to the processing plants at the
end of each day (Zandstra etal., 1992). Most processing asparagus is canned rather than
frozen (Kotecha and Kadam, 1998). From the conveyer belts asparagus is sorted for size
and color and placed in the cans, to which a dilute brine of 2% salt is added (Kotecha and
Kadam, 1998). The cans are exhausted and sealed in an atmosphere of live steam
(Kotecha and Kadam, 1998). Sealed cans are processed at 115-120° C for 25-35 minutes
depending on their size (Kotecha and Kadam, 1998).

The nutritional requirements of asparagus are complicated somewhat by the fact
that it is an extremely deep rooted, perennial crop. Nitrogen applications, for instance,
have been shown to be more effective when applied after harvest than before, and
phosphorus is commonly applied in furrows before planting (Drost, 1997; Kotecha and
Kadam, 1998; Nonnecke, 1989). An asparagus crop will annually remove 7, 2, and 5% of
applied nitrogen, phosphorus, and potassium, respectively (Drost, 1997). Most of the

plant’s nutrient utilization occurs post-harvest during fern and root growth (Drost, 1997).

Commercial recommendations for fertilizer applications call for the band application of
phosphorus at 33.9 kg/ha in the trench furrow during crown planting, an application of
not more than 90.4 kg/ha of nitrogen, split between pre- and postharvest, and potassium
applied every second year at 67.8 kg/ha K20 (Warncke et al., 1992). There has been no
yield response shown to higher rates of N, and an excess could potentially contribute to
groundwater contamination (Zandstra et al., 1992). For asparagus crown nurseries,
commercial recommendations call for 56.5 kg/ha of nitrogen to be disced in before
seeding, and 56.5 kg/ha to be sidedressed when plants are about 6 inches high (Warncke
etal., 1992). Phosphorus and potassium should be applied so that 226 kg/ha of P205 and
K20 are available to the crop (Zandstra et al., 1992). Most commercial growers in
Michigan apply their nitrogen as ammonium nitrate.

Another important component of asparagus production is maintaining proper pH.
Commercial recommendations call for growers to maintain a soil pH of 6.8, both in the
crown nursery and harvested ﬁelds (Warncke et al., 1992). Research has shown that
asparagus produces best at a soil pH of 7.5, and that low pH can increase the potential for
early decline (Hodupp, 1983). Most asparagus growers are aware of this factor, and many
test their soil and treat regularly with lime. However, a 1983 study indicated that
commercial ﬁelds in Michigan showed soil pH levels as low as 3.7 and averaging 5.6
(Hodupp, 1983).

Asparagus is one of the most salt tolerant crops known, and historically salt has
been considered an important element in healthy asparagus growth (Elmer, 1989).

References to, and recommendations of salt applications on asparagus can be found in

many old gardening books, and books on asparagus culture (Anonymous, 1960; Barnes
and Robinson, 1881; Bennett, 1910; Brill, 1884; Buist, 1855; Burr, 1865; Green, 1896;
Greiner, 1890; Harris, 1883; Kidner, 1959). Some of the references include using salt as
a weed control, but most suggest using it for its growth beneﬁts, or simply note that its
use is common practice. Interestingly, some authors indicate that, while salt will not
harm the plant, is not speciﬁcally beneﬁcial to growth (Bennett, 1910; Green, 1896;
Greiner, 1890). Barnes (Barnes and Robinson, 1881), however, suggests that salt may be
beneﬁcial only inland where saltwater is not present. Green (Green, 1896) speciﬁcally
cites experiments that indicate that salt is not of value as a manure.

Early references to salt application vary, and may include personal observations,
folk wisdom, or scientiﬁc studies. Barnes and Robinson (1881) suggest that salt is most
beneﬁcial on older plantings, and claim that it has insecticidal properties. They also
recommend that it not be applied while the roots are in a dormant state, while it is dry and
sunny, or to recently transplanted crowns. Brill (1884) comments that some people assert
that a layer of salt '/2 inch thick will be beneﬁcial, and notes that asparagus can tolerate
almost any amount of salt, though using more than 5 bushels/acre is not necessary.
According to Bennett (1910), salt is commonly applied at a rate of 600 lb/acre, and may
serve as a weed killer for a short time. According to Harris (1883), salt was usually
broadcast in early spring at a rate of 10 bushels/acre, though he also notes that it has been
recommended to apply it at much higher rates (120 bushels/acre). Rates of up to 2 lb. per
square yard (9680 lb/acre) are suggested in some texts, split into two applications (Barnes

and Robinson, 1881). Burr (1865) suggests mixing salt into manure to apply both in the

planting furrow and each autumn, but does not indicate an appropriate rate. Kidner
(1959) opines that sodium liberates more available potash and thus is useful in nutrition,
and goes on to note some of the negative effects of salt on soil structure, concluding that
salt use is excellent in moderation but dangerous in excess.

In 1905, Walker (1905) demonstrated a beneﬁt from salt applications (1000 lb.
per acre) on 5 varieties, showing a 13.5% increase in growth in the third year after
planting, resulting in an increase of 100 lb/acre of harvested asparagus. Rudolfs (1921)
also demonstrated the beneﬁt of salt to asparagus growth and determined an optimum rate
of application. It is interesting to note that Rudolfs made applications during rainy
weather, a practice that at least one of the early references recommends (Barnes and
Robinson, 1881). Better fern vigor and a higher yield resulted from increasing amounts
of salt (up to 500 leacre) following the second year of application on both 12 year old
and 3 year old ﬁelds (Rudolfs, 1921). No weed control beneﬁt was attributable to any of
the treatment rates (150, 300, and 500 lb./acre) (Rudolfs, 1921).

Weeds can be a serious problem on asparagus. Because research has shown
increased yields when asparagus fern is mowed, rather than disced in the fall or spring
(Putnam and Lacy, 1977), most Michigan asparagus is produced in a no-till system.
Generally, a cover crop of winter rye is planted in order to combat soil erosion. Weed
control is obtained solely by chemical means. A broad spectrum herbicide is applied
before harvest, and again after harvest is completed. Spears are mowed or picked close to
the ground before applications in order to minimize herbicide damage to the crowns.

Greenhouse studies have shown that the herbicides diuron, metribuzin, simazine, and

terbacil all cause damage to young asparagus crowns, and reduce yield (Hodupp, 1983).
Other research has shown that the herbicide terbacil, despite providing good, season long
control of weeds in asparagus plantings, causes injury to the fern when applied at a rate of
2.2 kg/ha or greater (Welker and Brogdon, 1972). Simazine has also been shown to
reduce yield in ﬁeld plots (Damicone etal., 1987).

Weeds can cause damage to asparagus plantings by competing for water and
nutrients and by producing alle10pathic substances (Putnam et al., 1983). Perennial
weeds, including horsenettle (Solanum carolinense), quackgrass (A gropyron repens),
ﬁeld bindweed (Convolvulus arvensis), common milkweed (Asclepias syriaca), swamp
smartweed (Polygonium lapathifolium), northern dewberry (Rubusfragellaris), ﬁeld
horsetail (Equisetum arvense), and yellow nutsedge (C yperus esculentus) are the most
problematic in asparagus because it is a perennial crop (Putnam et al., 1983) (Zandstra et
al., 1992). It is recommended that asparagus growers kill perennial weeds with a
registered herbicide the year before planting a ﬁeld (Zandstra et al.. 1992). Volunteer
asparagus, which grows from seeds dropped by the female plants in the fall. can be a
serious pest in ﬁelds that are not planted with the all-male varieties (Zandstra et al..
1992). Common annual weeds include crabgrass (Digitaria sanguinalis), yellow foxtail
(Setaria glauca), redroot pigweed (A maranthus retroﬂexus), common lambsquarters
(Chenopodium album), fall panicum (Panicum dichotomiﬂorum), barnyardgrass
(Echinochola crusgalli), stinkgrass (Eragrostis major), and ﬁeld sandbur (Cenchrus
tribuloides) (Putnam et al., 1983). Fall panicum, in particular, has become a problem in

no tillage ﬁelds, and is a vigorous competitor with asparagus fern (Putnam et al., 1983).

10

A number of insects are pests in asparagus. Cutworrns (Euxoa messoria and E.
scandens) can cause damage to growing spears by feeding on the tips or sides of new
shoots, causing losses of up to 90% of the harvestable crop (Graﬁus, 1983; Zandstra et
al., 1992). Most farmers control this pest with a pre-emergence insecticide application at
the time of spring mowing. Common asparagus beetles (Criocerus asparagi) can be a
major pest on spears, where they feed and lay eggs, and on fern, where larvae can cause
severe defoliation (Putnam et al., 1983; Zandstra et al., 1992). Farmers often spray
during harvest to control these insects because the presence of eggs on spears can lead to
rejection by processors. Spotted asparagus beetles (C. duodecimpunctata), which appear
later in the season, are usually a lesser problem. They feed on spears, but lay their eggs
on fern. Larvae feed only on ﬁ'uit, and do not cause defoliation (Putnam et al., 1983;
Zandstra et al., 1992). Plant bugs (Adelphocoris lineolatus and Lygus lineolaris) can
cause severe fern damage late in the season by injecting toxic saliva into the plant and
causing tip dieback and tissue collapse (Putnam et al., 1983; Zandstra et al., 1992).
Asparagus aphids (Brachycorynella asparagi) are a common inhabitant of asparagus
ﬁelds, though they are of minor economic importance in Michigan where their numbers
are generally kept low by predators (Putnam et al., 1983; Zandstra et al., 1992). Aphids
tend to be a problem in young ﬁelds, and can be very damaging in the asparagus growing
areas of Washington (Putnam etal., 1983; Zandstra et al., 1992). Asparagus miners
(Ophiomyia simplex) are Dipteran pests that lay two generations of eggs each year
(Putnam et al., 1983). Although they are not particularly damaging alone, they are closely

associated with stem rot caused by F usarium spp. (Damicone et al., 1987; Gilbertson et

11

al., 1985). Insecticide treatments used to control the miner have led to decreased
incidence of stem and crown rot, and increased survival and yield of asparagus plantings
(Damicone et al., 1987).

Nematode problems have been sporadically reported in asparagus (Crittenden.
1952; Putnam et al., 1983; Rohde and Jenkins, 1958). Most research, however, indicates
that asparagus is not very susceptible to nematodes, presumably because of toxic
substances excreted by the roots (Crittenden, 1952; Rohde and Jenkins, 195 8).
Crittenden (1952) observed very low infection rates and no hypertrophy when asparagus
was planted in soils that were heavily infested with root-knot nematodes. Similarly,
Rohde and Jenkins (1958) showed that populations of T richodorus christiei died out in
the soil around asparagus plants, and isolated a root exudate that was antagonistic to
several species of plant parasitic nematodes.

There are several disease problems that cause losses on asparagus in Michigan
each year. Purple spot is a disease caused by the fungus Stemphylium vesicarium and was
ﬁrst described in Michigan in 1982 (Lacy, 1982). The disease causes small purple lesions
with a necrotic center on asparagus stems and cladophylls that can build up in numbers
great enough to cause premature defoliation and fern death. The disease also attacks
spears and can cause processors to reject asparagus because of asthetics. The fungus
enters the plant through wounds, or natural openings, and is favored by warm, wet
conditions (Meyer, 1997). The causal organism overwinters on plant debris, and
consequently is a much more severe problem in no tillage systems (Zandstra et al., 1992).

Control of the disease is attained by the use of fungicide sprays, and the development of a

12

disease forecasting system has allowed for better application timing in Michigan (Meyer,
1997). Asparagus rust is a fungal disease caused by Puccinia asparagi (Zandstra et al.,
1992). Several spore stages are produced during the growing season. that cause damage
to all aboveground plant parts (Putnam et al., 1983). The disease is most prevalent in wet
years, and severe outbreaks can cause premature fern death and yield loss (Putnam et al.,
1983; Zandstra et al., 1992). Asparagus virus I (AV-I), a potyvirus, asparagus virus 11
(AV-II), an ilarvirus, and tobacco streak virus, an ilavirus, have been reported to occur in
asparagus throughout the world (Evans et al., 1990; Falloon et al., 1986). These viruses,
common in commercial asparagus plantings, have been associated with a decline in ﬁeld
productivity of up to 50%, and a diminished rooting capacity (Evans et al., 1990; Jaspers
and F alloon, 1997). These viruses have also been linked with an increased susceptibility
to Fusarium diseases (Evans and Stephens, 1989). All are mechanically transmitted, and
asparagus virus 11 is seedbome (Evans and Stephens, 1989; Falloon et al., 1986). No

measures to control virus are currently practiced by growers in Michigan.

FUSARIUM CROWN AND ROOT ROT‘

One of the major limitations to asparagus production, both in Michigan and
throughout the world, is asparagus decline syndrome. This condition was ﬁrst described
in California as “a slow decline in the productivity of old asparagus plantings, and the
inability to re-establish productive plantings” (Grogan and Kimble, 1959). Fields
afﬂicted with this disorder exhibit a premature reduction in the size and number of spears,

death of crowns, and eventually become unproﬁtable to harvest (Elmer et al., 1996;

13

Grogan and Kimble, 1959). In addition, ﬁelds replanted in asparagus after an asparagus
crop is removed often exhibit reduced emergence, yellowing, wilting, and death (Grogan
and Kimble, 1959). This phenomenon is commonly referred to as “replant bound early
decline” (Elmer et al., 1996). These problems often have serious economic impacts on
farmers, and are responsible for the abandonment of asparagus production in Western
Massachusetts, Connecticut, and New Jersey (Hartmann, 1996; Damicone et al., 1988;
Taylor and Rathier, 1985; Johnston et al., 1979). There are a number of cultural and
biological factors that contribute to asparagus decline syndrome. Acidic soils (Hodupp,
1983), alle10pathic root exudates (Blok and Bollen, 1993), over-harvesting (Shelton,
1978), and other abiotic factors that weaken the plant can contribute signiﬁcantly to the
problem (Elmer et al., 1996). Although insects, foliar diseases, and weeds all play a role
(Elmer et al., 1996), the single most important factor contributing to decline is F usarium
crown and root rot (Grogan and Kimble, 1959; Fantino, 1990; Schoﬁeld et al., 1996;
Taylor and Rathier, 1985).

The fungal genus F usarium was ﬁrst described by Link in 1809 as containing
species with fusiform spores borne on a stroma (Booth, 1971). The genus was validated
in the lntemational Botanical Code by Fries in 1821, who placed it in the order
Tuberculariae (Booth, 1971). F usarium spp. may produce both macroconidia and
microconidia, so named for their respective sizes (Alexopoulos et al., 1996).
Macroconidia are crescent shaped spores that are often borne on sporodochia, and have a
foot-shaped basal cell (Windels, 1992; Alexopoulos et al., 1996). Microconidia are small,

aseptate spores that are generally round or oval in shape (Alexopoulos et al., 1996). In

14

many species, they are produced in chains or false heads on aerial mycelia (Nelson et al..
1983). Some species may also produce thick-walled chlamydospores, and some produce
sclerotia (Alexopoulos et al., 1996; Windels, 1992). The perfect stage of the fungus,
though only known in some species, produces perithecia and belongs to the Hypocreales
(Booth, 1971). The taxonomy of the genus F usarium is the subject of some debate, and
different systems separate the genus into as few as nine or as many as 90 species
(Toussoun, 1981; Windels, 1992; Wollenweber, 1913; O'Donnell et al., 1998). More
recently, molecular techniques have been employed in an attempt to elucidate
evolutionary relationships, and to organize species along these lines (O'Donnell etal.,
1998)

F usarium spp. are among the most common fungi in the world, and are widely
distributed in soil and on organic substrates (Booth, 1971). They have been isolated from
permafrost in the arctic, and from the sand of the Sahara, and can be pathogens, parasites.
or saprophytes (Booth, 1971; Windels, 1992). F usarium can be found on almost every
plant species in most parts of the world, and can cause disease in humans and animals as
well as plants (Booth, 1971; Windels, 1992). Some species of the fungus produce
mycotoxins that can be contaminants of human and animal food, and were reportedly
used as biological warfare agents in both Vietnam and Afganistan (Alexopoulos et al.,
1996; Booth, 1971). Well adapted for survival in a broad range of environments,
tenacious isolates have been found in many preserved foods, stored chemicals, and in
airplane fuel tanks (Booth, 1971). Though generally considered a soil-home fungus,

many species are also efﬁcient at airborne dispersal (Matos et al., 1997; Burgess, 1981).

15

The main interest in the genus F usarium, however, is not strictly mycological, but
rather in its role as a plant pathogen. F usarium causes two distinct types of disease on
plant roots: cortical rots, and vascular wilts (Toussoun, 1981). Several economically
devastating diseases on important agricultural crops have made F usarium infamous in the
last 100 years. Cotton wilt, pea wilt, tomato wilt, and banana wilt, to name just a few,
have been so catastrophic that they threatened to eliminate entire agricultural industries
(Toussoun, 1981). Today, Fusarium diseases affect an astoundingly broad range of crops,
and F usarium is among the most economically important genuses of plant pathogens
worldwide.

Fusarium crown and root rot of asparagus is caused by the pathogens F usarium
oxysporum (Schlect.) f. sp. asparagi (Cohen & Heald) and F usarium proliferatum (T.
Matsushima) Nirenberg (formerly F usarium moniliforme Sheld.). The disease was ﬁrst
described in Massachusetts in 1908 as a wilt of young shoots during the cutting season,
followed by a yellowing and rot of the mature stalks, and was associated with an
unidentiﬁed F usarium sp. (Cohen and Heald, 1941 ). Cook (1923) described two cases in
New Jersey where circular areas in asparagus ﬁelds were dwarfed and dying. The disease
was most prominent in the center of the circular area, becoming less severe toward the
margins, and the plants were characterized by “a pronounced brown or rusty discoloration
on the stems below the surface of the ground”. Cook went on to satisfy Koch’s postulates
with an organism he identiﬁed as F usarium sp. Between the time of Cook’s observations
and 1940, root rot, crown rot and wilt of asparagus were reported from New York, New

Jersey, North Carolina, South Carolina, Pennsylvania. Illinois, Massachusetts, Missouri,

16

California, and Washington (Cohen and Heald, 1941). Cohen and Heald (1941)
described the symptoms of the disease in depth and noted both vascular discoloration,
wilt, and death of the stalks, dried out roots, red-brown lesions on the cortical areas of the
root, and discoloration extending through the vascular system of the roots from the site of
the lesion. They also noted that it was often possible to trace vascular discolorations
down through an injured or harvested shoot, into the crown, and through the root system,
suggesting that the point of infection may actually be the injured above-ground parts.
These researchers isolated F. oxysporum from diseased plant material, and satisﬁed
Koch’s postulates with single spored isolates of the fungus. Investigations of asparagus
material throughout the country led to the conclusion that the disease-causing organism
was present in all asparagus growing areas. Cohen and Heald (1941) also observed
disease affecting asparagus seedlings, a problem that was later described by Graham
(1955) as poor seedling emergence, stunting, yellowing, and wilting of the older
seedlings, collapse of sections of primary root, and elliptical root lesions of varying size.
Graham also described an inability to establish crown plantings in a ﬁeld that had been
afflicted with the disease, a situation that Grogan and Kimble (1959) would later refer to
as the asparagus replant problem. Graham (1955) conﬁrmed Cohen and Heald’s (1941)
observation that the pathogen is widespread, and noted that it could survive for long
periods of time without its preferred host, was efﬁcient at colonizing dead organic matter,
and could be found on wild asparagus plants. Grogan and Kimble (1959) were the ﬁrst to
call the causal agent of decline F. oxysporum f. sp. asparagi.

Both Grogan and Kimble (1959) and Graham (1955) reported ﬁnding pathogenic

17

isolates of F moniliforme. The former found 21 isolates in their studies, only two of
which killed fewer than 25% of their test plants, while most killed many more. Graham
(195 5) implicated F. moniliforme as a contributor to asparagus decline, and described it
as pathogenic on seedlings, but was unable to reproduce the disease symptoms observed
in the ﬁeld. The ﬁrst mention of F. moniliforme as a primary disease-causing organism
was in 1971 when Endo and Burkholder (1971) linked the fungus to a dry, brown crown
rot and stand decline in California. Johnston et a1. (1979) determined the role of F.
moniliforme in asparagus decline, and compared its pathogenicity with that of F
oxysporum f. sp. asparagi. This study showed that F moniliforme was more commonly
associated with crown and stem tissue than with root tissue, and that its virulence was
similar to that of F. oxysporum f. sp. asparagi. Further, F. moniliforme was dominant in
older plantings while F usarium oxysporum f. sp. asparagi tended to predominate in
younger ones. Damicone and Manning (1985), however, found that F. moniliforme was
more virulent than F. oxysporum f. sp. asparagi, and was dominant in ﬁrst year asparagus
from disease free crowns in Massachusetts. After F. proliferatum was differentiated
from F. moniliforme based on the presence of polyphialides (Nelson et al., 1983), Elmer
(1990b) found that F. proliferatum isolates were highly virulent on crown tissue and were
commonly isolated from plants in the ﬁeld. It is now generally accepted that F.
proliferatum, not F. moniliforme is the primary cause of crown and stem rot of asparagus,
though the latter is also a pathogen (Elmer and F errandino, 1992).

Other F usarium spp. have also been shown to be pathogenic on asparagus.

F usarium culmorum can be an important stem pathogen in EurOpe, while F. proliferatum

18

is generally absent (Blok and Bollen, 1995). Schreuder et a1. (1995) found F. solani
caused root lesions on seedlings in South Africa, though it was less virulent than either F.
proliferatum or F. oxysporum f. sp. asparagi. Damicone and Manning (1985) also
reported ﬁnding this species in 1.4% of sampled crowns and found it was a mild pathogen
on seedlings. Elmer and Ferrandino (1992) found that F. subglutinans could be slightly
pathogenic to asparagus. Armstrong and Armstrong (1969) reported that F. oxysporum f.
sp. apii, which causes celery wilt, as well as formae speciales’ cubense (banana, some
grasses), and medicaginis (alfalfa, pea, vetch) could cause symptoms on asparagus.
Elmer and Stephens (1989) observed root lesions on asparagus caused by F. oxysporum f.
sp. cepae (onion) and F oxysporum f. sp. gladioli (gladiolus). Blok and Bollen (1997). in
contrast reported that asparagus was not susceptible to F. oxysporum f. sp. pisi (pea),
Iupini (lupine), cepae (onion), Iilii (lily), or gladioli (gladiolus). In general, most other
species are considered to be of minor importance in comparison to F. proliferatum and F.
oxysporum f. sp. asparagi.

Early reports of disease caused by F usarium recognized that the pathogen is
widely distributed (Grogan and Kimble, 1959; Cohen and Heald, 1941; Graham, 1955).
Graham (195 5) suggested that the pathogen may be indigenous to soil never planted in
asparagus. Gilbertson et a1. (1981) isolated pathogenic F. oxysporum from a meadow, a
pasture, and a woodland. Hartung et a1. (1990) conﬁrmed that, although pathogen
populations were higher in the vicinity of asparagus plants, soils that had no history of or
proximity to asparagus plantings still contained pathogenic isolates of both F. oxysporum

f. sp. asparagi and F. proliferatum.

l9

Although F. oxysporum is not commonly thought of as being wind dispersed, this
can be an important factor in some situations (Burgess, 1981). One report has linked air-
borne microconidia of F. oxysporum f. sp. radicis-lycopersici, an important pathogen of
tomato, with re-colonization of steamed greenhouse soil (Burgess, 1981). Air dispersal
of the fungus on wind-blown dust and soil particles has been suggested as well (Burgess,
1981). The extent to which this is a factor with F oxysporum f. sp. asparagi is unknown.
F. proliferatum, in contrast, is known to be wind dispersed (Burgess, 1981), and airborne
spores of this fungus are common in ﬁelds, often infecting asparagus ﬂowers and fruit
(Gilbertson and Manning, 1983). Damicone and Manning (1985) have also reported
ﬁnding the pathogen sporulating proliﬁcally on asparagus stubble in the spring,
conﬁrming that air-home dispersal may be very common.

Both F. oxysporum f. sp. asparagi and F. proliferatum may be seed home.
Contrary to published reports (Damicone etal., 1981; Inglis, 1980), Graham (1955) did
not ﬁnd any contamination of seed with pathogenic F usarium despite repeated attempts.
Grogan and Kimble (1959), however, did ﬁnd that unsterilized seeds planted in steamed
soil could sometimes (though rarely) result in wilt and death of seedlings from F.
oxysporum f. sp. asparagi. Lewis and Shoemaker (1964a) sampled seeds from New
Jersey, Massachusetts, Ontario, and Michigan, and found that seeds from all the samples
were infected with F. oxysporum f. sp. asparagi. Inglis (1980) found that many seed lots
were contaminated with both F. oxysporum f. sp. asparagi and F. proliferatum, and
though seeds were not infected internally, deep crevices and beetle feeding sites provided

ideal places for spores to lodge and escape disinfestation. Gilbertson and Manning

20

(1983) reported that seed grown in Massachusetts had a F. proliferatum contamination
rate of up to 10%. Damicone et a1. (1981) found F usarium contamination on every seed
lot they tested, and determined that the only way to completely disinfect seeds was to
soak them in 25,000 ppm benomyl in acetone for 24 hours. Blok (1996b) reported that
Dutch seed lots were consistently infected with F. oxysporum f. sp. asparagi, and showed
that infestation usually occurs during the harvesting process when infested soil adheres to
fallen berries.

Another important source of inoculum is infected crowns from seedling nurseries.
According to Gilbertson et al. (1981), F usarium infections of one-year-old crowns used
to plant asparagus ﬁelds can exceed 50%. Blok (1996b) similarly reported that 75% of
one-year—old Dutch crowns were infected with F. oxysporum f. sp. asparagi. Fantino
(1990) reported that in Italy. F usarium contamination is common in seedling nurseries
throughout the country.

Asparagus miners can be an important disease vector. Gilbertson et al. (1985)
reported that larval feeding by asparagus miners resulted in increased stem rot caused by
F. proliferatum, and that asparagus miners can carry both F. oxysporum f. sp. asparagi
and F. proliferatum. Damicone et a1. (1987) found that insecticide treatment could
reduce crown rot incidence by 30%. Asparagus miner control is not currently practiced in
Michigan, and the insects may be an important factor in disease spread.

The ubiquity of F usarium isolates pathogenic to asparagus in nature has led
researchers to question the biological relationships between isolates of these fungi. To

this end, several researchers have investigated vegetative compatibility groups in an

21

attempt to understand the diversity and relatedness of F usarium spp. in the ﬁeld. Elmer
and Stephens (1989) investigated 97 strains of pathogenic F. oxysporum from Michigan
ﬁelds, and world collections. Their research identiﬁed 43 distinct vegetative compatibility
groups of F. oxysporum that could infect asparagus, some members of which were
isolated from other crops. LaMondia and Elmer (1989) tested isolates from three plants
in a single plot in Connecticut and found that eight of 18 F. oxysporum isolates belonged
to unique vegetative compatibility groups. Blok and Bollen (1997) similarly found that
24 Dutch isolates of F. oxysporum f. sp. asparagi belonged to 18 different vegetative
compatibility groups. Because vegetative compatibility groups are presumed to be
genetically isolated from one another (Correll et al., 1987), this study suggests that the
ability to cause disease on asparagus may have evolved independently in a number of
different strains. Although this kind of diversity has also been observed in F. oxysporum
f. sp. lycopersici (Elias and Schneider, 1991), most pathogenic forms of F. oxysporum are
comprised of one or only a few vegetative compatibility groups (Burgess, 1981; Jacobson
and Gordon, 1990). With F proliferatum, Elmer (1990c) found 20 vegetative
compatibility groups among 110 isolates from four ﬁelds in Connecticut, though most
isolates fell into six groups that were found in all the ﬁelds. LaMondia and Elmer (1989)
found 13 vegetative compatibility groups in 97 isolates from three plants in a single test
plot. Thus, the virulence trait is present among a number of diverse isolates, though some
may predominate in asparagus ﬁelds. The signiﬁcance of these results cannot be
overstated. The fact that many different groups of the pathogens may be able to infect

asparagus has important implications on disease control strategies, and may preclude the

22

development of robust gene for gene resistance through breeding efforts.

It is also interesting to note the range of virulence between different isolates of the
pathogens. Grogan and Kimble (1959) noted a wide range of pathogenicity in both F
oxysporum and F proliferatum, from those that were nonpathogenic, to those that killed
100% of inoculated seedlings. Elmer and F errandino (1992) described a wide range of
pathogenicity for isolates in section Liseola (F. proliferatum, and F. moniliforme).
Similarly, Damicone and Manning (1985) observed a wide range of pathogenicity with
both F oxysporum f. sp. asparagi and F proliferatum, and LaMondia and Elmer (1989)
noted differences in mean virulence ratings of isolates belonging to different vegetative
compatibility groups of F proliferatum. Different kinds of infections (cortical, vascular)
have been reported for F oxysporum f. sp. asparagi (Smith and Peterson, 1983; Grogan
and Kimble, 1959; Graham, 195 5; Cohen and Heald, 1941), and it is possible that
different strains of the fungus could exhibit different types of infection, though this idea is
largely speculative.

The host range of the F usarium pathogens involved in crown and root rot of
asparagus have been investigated by several researchers. Cohen and Heald (1941) tested
F oxysporum f. sp. asparagi against a range of crops. In these experiments, the pathogen
did not cause disease on potatoes, tomatoes, or camations, and did not rot potato tubers or
onion bulbs held in a moist chamber. It did cause disease on ornamental asparagus, and
caused a slight wilt of Alaska peas. Graham (1955) also tested F. oxysporum f. sp.
asparagi against onions, spinach, luceme, gladious, oats, barley, rye, and corn. Of these,

only gladiolus developed lesions from which the pathogen was re-isolated. Armstrong

23

and Armstrong (1969) found that F. oxysporum f. sp. asparagi did not cause disease on
celery, lupine, vegetable sponge, Maltese cross, cotton, Chrysanthemum, eggplant, alfalfa,
Chinese lantern plant, Mexican sunﬂower, garden pea, safﬂower, tobacco, tomato, or
carnation. Blok and Bollen (1997) tested a number of different plants with their Dutch
isolates, and found that only asparagus, and occasionally pea and lupine were susceptible.
While the host range of F proliferatum has been investigated less extensively, it is
important to note that cross-pathogenicity has been observed between asparagus and corn
(Damicone et al., 1988), a ﬁnding that has important implications for crop rotation.

A number of different factors contribute to the severity of F usarium infection, and
thus contribute to asparagus decline. Ni gh (1990) states that asparagus serves as a host to
F usarz'um spp. without disease symptoms until one or a combination of factors causes
plant stress. Once these conditions are met, the plant becomes susceptible and the fungus
is able to cause root rot and wilt. Nigh’s research indicates that any factor causing plant
stress was found to increase the incidence and severity of the disease. In his studies on
moisture level, infection was increased when water was limiting. It is likely that water is
less often limiting in Michigan and other production regions than it is in Yuma, AZ where
these studies were conducted, but farmers do correlate dry years with increased death mid
seedling blight (N. Myers, County Extension Agent, personal communication). Wilcox-
Lee (1987), similarly, found decreases in fern height, number of shoots per plant, fern dry
weight, and root dry weight with decreasing soil matric potentials. Although Wacker
(1988) found signiﬁcantly increased disease severity and higher populations of F.

oxysporum f. sp. asparagi at higher soil moisture levels (0 MPa vs. -1.5 MPa),

24

experiments by Elmer and LaMondia (1997) have shown the opposite effect with both F.
oxysporum f. sp. asparagi and F. proliferatum. Graham (1955) also found that low levels
of soil moisture were more conducive to seedling damping off than high levels.

Hodupp (1983), Welker and Brogdon (1972), and Damicone et al. (1987) have all
shown decreased yield or injury from herbicide applications (terbacil, simazine, diuron,
metribuzin) indicating that herbicide use may be an important stress factor. Weed
competition, however, can also cause additional plant stress. Shelton (1978) showed
increased crown survival and yield in both F usarium-infected, and non-infected plots
with herbicide use, as compared to hand-weeded plots which he was not able to keep
weed free throughout the season. Shelton (1978) also showed that over-harvesting can
weaken asparagus plants by reducing the level of carbohydrates stored in the roots. Evans
and Stephens (1989) demonstrated that asparagus infected with the viruses AV-I or AV-II
developed signiﬁcantly more disease from F oxysporum f. sp. asparagi than did virus-
free plants, and correlated the presence of the virus with increased root exudation and
susceptibility of root tissue to infection. Hodupp (1983) has shown that low soil pH can
inhibit asparagus growth. Though these experiments never correlated low pH with
F usariurn disease, acidic soil may contribute to plant stress and increase disease
susceptibility. Hodupp (1983) also correlated more soil compaction with decreased
asparagus vigor, suggesting that this may be an important stress factor as well.

The stress factor that has been studied most extensively is the effect of autotoxic
root exudates from asparagus plants. Beginning with the earliest disease reports,

researchers have observed that decline is most severe in ﬁelds that have previously grown

25

asparagus, and that the problem may persist for many years (Elmer et al., 1996). A
number of alle10pathic compounds have been identiﬁed from asparagus, including
caffeic, ferulic, malic, citric, methylenedioxycinnamic, asparagusic and chlorogenic acids,
saponins, and glycosides (Keulder, 1997). Yang (1982) found that extracts from both
ﬁeld-grown asparagus plants. and plants grown in sterile culture delayed the germination
of asparagus seed and inhibited root and shoot development. Because activated charcoal
and autoclaving did not reduce the toxicity of the extracts, Yang suggested that the toxic
substances were very stable and could persist in ﬁelds. Hartung and Stephens (1983)
looked at the effects of asparagus toxins on F usarium populations and on the
susceptibility of roots to infection. These investigations revealed that disease caused by
F usarium spp. was much more severe in the presence of asparagus tissue, but asparagus
tissue did not enhance the growth of the pathogens. Pierce and Miller (1990) conﬁrmed
this synergistic relationship, and showed that soluble solids from asparagus extract
enhanced the severity of disease caused by F usarium, though there was also an autotoxin
effect independent of this. Their research also indicated that the activity of asparagus
autotoxin dissipated over time. Hartung et al. (1989) showed that respiration and
peroxidase activity decreased, and electrolyte efﬂux increased in the presence of
asparagus extracts, suggesting that the chemicals may have direct physiological and
biochemical effects on asparagus plants that make them more susceptible to disease.
Blok and Bollen (1993) found inhibitory effects of root extracts up to 10 years after
asparagus ﬁelds had been taken out in laboratory experiments, but suggest that the

amount of root biomass present in the ﬁeld after termination of the crop in the previous

26

season is not high enough to have a direct growth inhibition effect. It is likely, they
conclude, that the effect of root residues is of secondary importance to the buildup of
pathogen populations in the soil which have been shown to correlate directly with the
severity of replant disease (Blok and Bollen, 1996a). However, as Blok and Bollen
suggest (1993), the toxic residues may contribute to the problem by affecting the
populations of soil microﬂora and predisposing roots to F usarium infection. Further
investigations by these researchers (Blok and Bollen, 1996c) conﬁrm, in fact, that root
extracts from asparagus residues retard the growth of a wide variety of fungi including
T richoderma and Gliocladium spp., but do not affect F usarium spp. In the same study.
however, the researchers did not ﬁnd any increase in disease severity in soils containing
plant residues in the Netherlands.

A signiﬁcant amount of research has been dedicated to developing strategies for
the control of F usarium crown and root rot. Cohen and Heald (1941) detennined that
nutrition and pH are ineffective as controls, and stated that the only real control for the
disease would be development of plant resistance. Lewis and Shoemaker (1964b) tested
a number of different species in the genus Asparagus for resistance to F usarium
infections. They found only one species that was resistant to the disease and speculated
that it was too distantly related to commercial asparagus to be useful in breeding.
Stephens et a1. (1989) tested a number of different varieties and species for disease
resistance and found slightly less susceptibility in some all-male hybrids as well as A.
densiﬂorus, but no true disease resistance. The original publications on these varieties

also report increased tolerance to disease, but not disease resistance (Ellison, 1985;

27

Ellison et al., 1990; Ellison and Kinelski, 1986). Despite the development of an in vitro
assay to evaluate resistance (Stephens and Elmer, 1988), and considerable interest on the
part of growers for resistant varieties, this type of disease control does not appear to be
forthcoming. Only one of the more recent cultivar trials speciﬁcally addresses Fusarium
crown and root rot resistance (Elmer et al., 1997). In one ﬁeld trial, moderate resistance
to disease was found with the varieties Emerald and Rutgers Syn #2 which had
consistently high yields and lost less than 40% of their crowns over seven years,
compared to an average loss of over 60%. In another trial, cultivars Lucullus and Jersey
General lost only 8% of crowns over seven years compared to a loss of almost half of the
crowns in susceptible cultivar Mary Washington (Elmer et al., 1997). Caution should be
exercised in extrapolating these results to Michigan conditions since cultivars that exhibit
a high productivity in one region do not always do so in another (Elmer et al., 1997).
Because of the ubiquitousness of the pathogens, and the perennial growth habit of
the crop, chemical control strategies have thus far demonstrated. at best. only marginal
success. Damicone et al. (1981) have shown that the difﬁculty of disinfesting seeds can
be overcome by soaking them in benomyl (25,000 ppm) in acetone. Knaﬂewski and
Sadowski (1990) showed that although seed treatment with benomyl in acetone did not
exclude F usarium infection, disease severity in one-year-old crowns was reduced slightly
with this treatment, and no other chemical treatment completely eliminated F usarium
contamination on seeds. Di Lenna and F otello (1990), in contrast, found no advantage to
seed disinfestation with benomyl in acetone for disease reduction, suggesting that seed

contamination may not be important in all cases. Manning and Vardaro (1977) reported

28

that soil fumigation (methyl isothiocyanate/chlorinated C3 hydrocarbons) in addition to
fungicide crown soaks (benomyl or captafol) of F usarium-free crowns (grown in a
greenhouse) resulted in increased vigor in plots infested with F usarium spp. Damicone et
al. (1987) also found increased yield when disease-free crowns were transplanted into
fumigated (methyl isothiocyanate/chlorinated C3 hydrocarbons) soil. Lacy (1979), in
contrast, found that fungicide soaks of crowns grown in naturally infested nursery soils
did not result in better survival or spear production in subsequent years. This study also
showed that crowns grown in infected nurseries did not show higher yields in fumigated
(DD-MENCS) plots than in non-fumigated plots. Lacy (1979) did show, however, that
crowns grown in fumigated soil and transplanted to fumigated soil had higher yields than
those grown in non-fumigated soil, and transplanted into fumigated soil. These studies
seem to indicate that chemical treatment is ineffective once seedlings have been infected
in the nursery. Molot et al. (1985) found that crowns grown in fumigated (methyl
bromide) nursery soils were healthier than those grown in non-fumigated soils, but that
subsequent grth was unexpectedly much better for infected crowns than for healthy
ones, and that the healthy crowns from the fumigated ﬁeld actually became more
susceptible to infection than unhealthy crowns. These researchers suggested that the lack
of mycorrhizal colonization in the fumigated soils limited the ability of these crowns to
thrive, and made them more susceptible to disease. This study also found little advantage
to pre-plant crown treatments (bleach, carbemdazim/captafol, “Ceretal”/bleach).

DiLenna and F oltello (1990) reported that the location from which crowns originated was

much better correlated to disease reduction than chemical soil disinfestation, noting that

29

nurseries in regions where decline was less problematic produced much healthier crowns
than those in regions where decline is severe. These researchers suggested biological soil
suppressiveness as the reason for this geographical difference in disease incidence.
Faloon and Fauser-Kevem (1996) demonstrated F usarium disease control in New Zealand
with thiabendazole crown soaks. Though it was not clear whether these experiments
were conducted with disease-free or infested crowns, the question may be moot for
Michigan growers because the manufacturer (N ovartis Crop Protection, Inc. Greensboro,
NC) has indicated that they have no plans to register this fungicide for use on asparagus
in the United States (A. Talley, Fungicide Development Director, personal
communication).

The role of nutrition in disease development has been studied in a number of
different host-pathogen systems. Many different macro- and micronutrients have been
implicated in both increased disease susceptibility and increased control (Huber, 1990).
Disease suppression with mineral nutrition can be due to effects on the nutrition of the
plant, effects on the nutrition of the pathogen, effects on other organisms in the system, or
a combination of factors (Woltz and Jones, 1981). Because of the complexity of
interactions between plants, pathogens, other microbes, and the soil, mechanisms for
disease control with nutrients are poorly understood. A signiﬁcant amount of research
has focused on the effects of nutrients on plant disease because nutrition can be managed
easily and cheaply. Though little research speciﬁcally addresses the asparagus-Fusarium
system, the role of nutrition in F usarium control has been studied in a number of crops.

Although not technically a plant nutrient, lime can be extremely important in the

30

nutritional regime of the plant because it affects nutrient availability and contains
calcium, which is often implicated in disease suppression. Contrary to published reports
(Stephens and Elmer, 1988), increased pH has never been speciﬁcally linked with
decreased Fusarium disease in asparagus. Increased pH has been shown to increase yields
(Hodupp, 1983), but the mechanism by which this increase occurs has never been studied.
Fusarium wilt of cotton caused by F oxysporum f. sp. vasinfectum has been associated
with acidic soils (Bell, 1989). Increased pH has been linked with the suppression of
Fusarium wilt of tomato caused by F oxysporum f. sp. lycopersici (Jones et al., 1989).
This may simply be a calcium effect, since it has been suggested that calcium inhibits the
activity of polygalacturonase produced by F usarium (Corden, 1965). Calcium is also
important in carbohydrate removal, neutralization of cell acids, cell wall deposition, and
the formation of pectates in the middle lamella, which suggests that it may have a role in
forming physical barriers to plant invasion (Huber, 1990; Huber, 1989). However, other
researchers have shown that high tissue calcium was not important in the reduction of the
Fusarium wilt of tomato, so lime may be affecting disease by some other mechanism
(Jones et al., 1989).

Both the form and the amount of nitrogen supplied to plants have been implicated
as important factors in disease development. High levels of nitrogen have been found to
encourage a number of plant diseases, including Fusarium corrn rot of gladiolus caused
by F oxysporum f. sp. gladioli (Jones et al., 1989). A number of researchers have also
studied how the form of nitrogen (ammonium vs. nitrate) affects the development of

Fusarium disease. Most studies have found that the nitrate form of nitrogen decreases

31

Fusarium disease as compared to ammonium, especially when combined with high soil
pH (Jones etal., 1989). This phenomenon has been reported for tomato, watermelon,
Chrysanthemum, muskmelon, cucumber, radish, lime, cotton, celery, and other crops
(Jones et al., 1989; Schneider, 1985). Elmer (1989) has tested nitrogen form on Fusarium
crown and root of asparagus and found, similarly, that weights of seedlings were
increased with nitrate-nitrogen in the presence of F oxysporum f. sp. asparagi or F
proliferatum.

The effects of other nutrients on the development of Fusarium disease have been
less widely studied. Several researchers have found decreased disease with increased
levels of potassium (Jones et al., 1989). Potassium chloride has been shown to decrease
Fusarium yellows of celery caused by F oxysporum f. sp. apii, though this researcher
attributed the effect to the ratio of potassium to chloride in the plant, rather than to either
ion alone (Schneider, 1985). Several studies have shown that high rates of phosphorus
can increase disease severity. Levels of phosphorus above what is needed for plant
growth favored F usarium disease in tomato, muskmelon, and cotton (Jones et al., 1989).
Micronutrient nutrition may also be important in F usarium disease development. Studies
with F oxysporum f. sp. lycopersici have demonstrated repeatedly the relatively high
requirement of this organism for copper, iron, manganese, and molybdenum, and that
increasing the amounts of the nutrients was beneﬁcial to the growth and sporulation of
the pathogen (Jones et al., 1989). This, in fact, may be the reason that lime is effective in
suppressing disease (Jones et al., 1989). Most micronutrients are much less available as

pH increases, and may therefore be limiting to F usarium growth.

32

Another interesting factor in F usarium disease development is the suppressive
effect of chloride salt. For many years, applying salt to asparagus plantings was a
common practice. Asparagus is one of the most salt tolerant crops grown commercially,
and researchers working in the early part of this century showed empirically that salt
applications can increase yields (Walker, 1905; Rudolfs, 1921), though the mechanism is
not clearly understood. More recently, research has focused on the role of salt in the
control of Fusarium crown and root rot. Elmer (1989) ﬁrst identiﬁed the potential
suppressiveness of salt on Fusarium disease while exploring the effect of different
nutrient formulations on disease development. In these experiments, NaCl reduced
disease development and root colonization on asparagus seedlings. KCl was also
effective in disease control, especially when the amount of potassium otherwise supplied
was limited, leading to the conclusion that disease suppression was due to a chloride
effect, and that the ratio of potassium to chloride may be the most important factor in
disease suppression. Another interesting observation in this study was that fresh weights
of seedlings increased with chloride applications, while dry weights did not always
increase, indicating that chloride applications may have affected osmotic regulation in the
plants.

Further research (Elmer, 1992) conﬁrmed that NaCl applications were suppressive
to Fusarium crown and root rot in both greenhouse and ﬁeld plots. NaCl was also more
effective in controlling disease in these plots than KCl, and plants were shown to actually
accumulate some Na+ (up to 5.5 mg/g of tissue). Fern concentrations of sodium were also

correlated with increased plant weights and decreased the fraction of diseased roots,

33

leading to the speculation that not just chloride, but also sodium was having an effect on
disease development. Since applications of NaCl did not reduce populations of F usarium
in the soil, and did not control weeds in this experiment, it was speculated that
applications affected the susceptibility of the host to the disease. Elmer (1992) proposed
that the increase in CI' concentration in the plant balanced the inﬂux of cations (N Hf, K‘,
Ca2+, etc.) that may otherwise be offset by the synthesis of organic acids such as malate,
and thereby reduce the exudation of organic substrates into the rhizosphere, which could
play a role in reducing F usarium colonization and infection. Other experiments have
shown that NaCl applications can effect carbohydrate and malate production in the roots
of asparagus plants (Elmer, 1990a).

Other investigations into the mechanism by which NaCl suppresses crown and
root rot have focused on plant water relations. A number of researchers have explored
the effect of water relations on F usarium disease (Cook, 1973; Cook, 1981; Cook and
Papendick, 1972; Papendick, 1974; Schneider, 1993; Schneider and Pendery, 1983). In
general, Fusarium diseases are most severe in dry years, with water stress predisposing
plants to disease (Cook, 1973; Cook, 1981; Schneider and Pendery, 1983). Cultural
practices, and the levels of different nutrients in the plant are known to affect water
potentials and can increase the rate of F usarium disease development (Papendick, 1974).
In a study by Elmer and LaMondia (1997), NaCl applications resulted in lower fern water
potential, and increased turgor pressure. It was hypothesized that the uptake of chloride
was responsible for the maintenance of lower osmotic potential in the plant, thereby

increasing turgidity and decreasing plant stress during periods of drought. Other

34

researchers have also suggested that lowered water potentials can limit pathogen invasion
(Cook and Papendick, 1972).

Salt may be eliciting not simply a plant effect, but one that involves other aspects
of the soil community. Elmer (1995) has investigated the effect of NaCl applications on
the density and composition of root colonizing bacteria. This study showed a signiﬁcant
increase in the fraction of manganese-reducing bacteria on the roots of NaCl treated
plants, and an increase in the level of manganese in the roots of these plants compared to
the control. In addition, several strains of these manganese-reducing bacterial were able
to decrease disease incidence in greenhouse trials.

In contrast to other studies on NaCl applications to asparagus (Elmer, 1989;
Elmer, 1992; Elmer, 1995; Elmer and LaMondia, 1997; Rudolfs, 1921; Walker, 1905),
Francois (1987) showed decreasing yields with increasing salinity levels. In these
experiments, salt was applied in irrigation water every 12 days, rather than in a single
yearly application to the soil. In light of the investigations by Elmer (1989, 1992, 1995,
1997) it is possible that the constant salinity in the solution supplied to the plants in this
experiment never allowed for an osmotic potential differential to be developed, and
thereby negated the mechanism by which salt aids the growth of plants. It is plausible
that this differential in osmoticum between soil and root allows for differences in either
nutrient uptake or root exudation which may result from applications of NaCl to
asparagus. Similarly, a study by Porter et al. (1917) showed increased yield with a single
application of NaCl (1120 kg/ha), and decreased yield with multiple applications (280

kg/ha four times). An alternate explanation for these differences could be simply that the

35

yield increases seen with NaCl are simply the result of decreases in F usarium disease,
which offset decreases in plant yield with salt applications. Elmer (1989, 1992, 1995,
1997) clearly showed that NaCl applications reduce Fusarium disease and increase yield,
and it is possible that the experiments of Walker (1905) and Rudolphs (1921), which
showed increased yield with salt applications, were done in a system infected with

F usarium pathogens. Nor is this explanation necessarily inconsistent with the results
obtained by Porter et al. (1917), who showed increased yields with a single salt
application and not with multiple salt applications, but applied their treatments to
different parts of the ﬁeld, and thus may have had different levels of F usarium infection
in each treatment.

The mechanism by which salt affects disease may involve a number of factors and
is not ﬁllly understood, yet it is clear that NaCl has the potential to decrease the severity
of Fusarium disease and increase yields in asparagus. Because studies with other crops
have actually shown increases in the severity of F usarium disease with the application of
NaCl (Standaert et al., 1978) it is likely that the beneﬁt of salt applications may be related
to the status of asparagus as a salt tolerant plant. Furthermore, it is possible that the
mechanism of salt tolerance will lend some insight to the question of why NaCl decreases
disease in asparagus. Salt tolerance in asparagus is an organismal trait, not present in
undifferentiated tissue to the extent that it is in the whole plant (Mills, 1988). Hence, it is
supposed that there are physiological and anatomical mechanisms at work that allow the
plant its halophytic characteristics. Asparagus, like other halophytes, has developed a

mechanism by which it is able to take up Cl' while excluding Na+ (Elmer, 1992; Mills,

36

1988). All halophytic plants seem to respond positively to NaCl applications (Flowers et
al., 1977). Salt applications commonly result in increased fresh weight in halophytes,
indicating a rise in water content (Flowers et al., 1977), and the osmotic effects of high
salt concentrations can be quite dramatic (Waisel, 1972). Other research has also shown
an increase in lignin and cellulose in halophytes when salt was applied (Flowers et al.,
1977), factors that have often been implicated in disease resistance (Agrios, 1997).
Another strategy that may have a place in the control of Fusarium crown and root
rot is the use of biological control organisms. The suppressiveness of some soils to
Fusarium diseases has led researchers to the conclusion that microbial activity may be an
important factor in disease inhibition (Louvet et al., 1981). Several different organisms
have been identiﬁed as having potential for the biological control of Fusarium diseases in
asparagus and other crops. The potential of non-pathogenic strains of F oxysporum to
protect plants against disease caused by pathogenic strains was ﬁrst recognized in the
1960's (Davis, 1967; Davis, 1968). Since this time cross-protection, or biological control
with non-pathogens, has been demonstrated for a number of crops including tomato
(Fuchs et al., 1997), chickpea (Hervas et al., 1995), watermelon (Larkin et al., 1996),
cucumber (Mandeel and Baker, 1991), pea (Oyarzun et al., 1994), carnation (Postrna and
Rattink, 1992), and asparagus (Blok et al., 1997; Damicone and Manning, 1981;
Damicone and Manning, 1982; Tu et al., 1990). Damicone and Manning (1981; 1982)
demonstrated signiﬁcant reduction in disease symptoms in greenhouse trials with
asparagus. Blok et al. (1997) reported more than a 50% decrease in root rot severity with

each of 13 nonpathogenic isolates from asparagus roots or ﬁeld soils. Both studies

37

reported differences in the ability of individual isolates to reduce disease, where some
were recognized as particularly useful while others were not. The mechanism of disease
suppression in asparagus is not clearly understood, though studies with both watermelon
and tomato have suggested that non-pathogenic isolates induce resistance to pathogenic
isolates in these plants (Fuchs et al., 1997; Larkin et al., 1996). Field experiments using
non-pathogenic F usarium isolates have shown the potential for use on young plants, but
haven’t been effective on older plantings already infected with the disease (Damicone and
Manning, 1981; Damicone and Manning, 1982; Tu et al., 1990; Blok et. a1, 1997).

Another organism with the potential for biological control is the fungus
Trichoderma. T richoderma spp. can be antagonistic to pathogenic ﬁmgi by competition,
antibiosis, or the production of lytic enzymes, and are able to grow on hyphal cell walls or
living mycelium (Benharnou and Chet, 1993; Elan et al., 1983; Sivan and Chet, 1986).
Trichoderma has been shown to attach its self to other fungi either by appressoria or
hyphal coils (Elad et al., 1981), and can even inhibit the germination of F oxysporum
chlarnydospores (Sivan and Chet, 1989). Gennari et al. (1990) and Nipoti et al. (1990)
have shown increased germination of asparagus seedlings with soil and seed applications
of T. harzianum. Arriola (1997) has also shown decreased disease and increased plant
weights when T. harzianum was applied to peat infested with F oxysporum f. sp.
asparagi.

Vesicular-arbuscular mycorrhizae are common inhabitants in soil, and commonly
form symbiotic associations with a number of plant species (Wacker, 1988). These

organisms are capable of increasing growth and yield in crop plants by increasing nutrient

38

uptake and resistance to salinity and drought (Menge, 1983; Nelsen and Saﬁr, 1982).
Mycorrhizal associations have also been shown to affect the incidence and development
of plant diseases (Dehne, 1982; Schenck, 1981). Several studies have emphasized the
importance of mycorrhizal colonization of asparagus roots (Burrows et al., 1990; Hussey
et al., 1984; Pederson et al., 1991). Especially interesting is a study by Pederson et al.
(1991) who showed increases in asparagus dry weight and bud formation when
mycorrhizae were applied to plants grown in fumigated ﬁeld soil. Other researchers have
observed decreases in the severity of F usan'um crown and root rot with mycorrhizal
inoculations (Arriola, 1997; Wacker et al., 1990a), underscoring the importance of these
relationships in disease development. Applications of mycorrhizal fungi show potential
for use in disease suppression, but because different mycorrhizal fungi predominate in
asparagus of different ages (Wacker et al., 1990b), the species introduced may be an
important consideration.

Elson (1993; Elson et al., 1994) isolated a species of Streptomyces from asparagus
soil that was found to be antagonistic to F oxysporum f. sp. asparagi and F
proliferatum. Chemical extraction yielded an inhibitory compound which reduced the
populations of pathogenic F usarium in the soil but also reduced root length. Cells of the
bacteria colonized asparagus roots, but not in sufﬁcient concentrations to control
F usarium disease. Elmer (1995) also identiﬁed bacterial species (Pseudomonas
corrugata, Serratia spp.) capable of reducing disease without negative effects on plant
growth. Further study is needed to identify the potential of these organisms for biological

control.

39

Since the causal organisms were ﬁrst discovered, investigators have struggled to
ﬁnd a solution to the problem of Fusarium crown and root rot. In Michigan, no speciﬁc
control strategies are practiced other than a modest amount of crop rotation, and the use
of high yielding varieties with which a certain amount of loss can be tolerated. It is clear
from decades of research that a single conventional strategy (chemical control, avoidance,
resistance breeding) for effective disease control is not forthcoming. There are, however,
a number of factors that, when employed together, may constitute an effective strategy for
reducing disease to an acceptable level. Several researchers have investigated integrated
strategies for F usarium disease control in different systems (Beale and Pitt, 1990;
Chakravarty et al., 1990; Garibaldi and Lodovica—Gullino, 1984; Minuto et al., 1995;
Sivan and Chet, 1993). Damicone and Manning (1985; Damicone et al., 1987) attempted
to develop an integrated strategy for F usarium disease control in asparagus that did not, it
seems, save asparagus production in Massachusetts. However, many advances in disease
control have been made since that time.

Clearly, more research is needed on the potential for integrating various disease
control strategies into a modern asparagus production system. However, the following
components warrant consideration for inclusion in an integrated disease management
system:

1) Selection of a vigorous, high yielding, and moderately resistant variety of asparagus
suited to the particular region in which it will be planted (Elmer et al., 1997).
2) Selection of a seedling nursery ﬁeld with low concentrations of F usarium pathogens

(Didelot et al., 1996) have developed a prediction test for root rot that could be easily

40

modiﬁed for the Michigan industry), or ﬁelds that have been out of asparagus production
for as long as possible, and have not been rotated with corn (Damicone et al., 1988), peas
(Cohen and Heald, 1941), or their relatives.

3) Treatment of asparagus seed prior to planting with benomyl in acetone (Damicone et
al., 1981; Knaﬂewski and Sadowski, 1990), or Trichoderma spp. (Gennari et al., 1990;
Nipoti et al., 1990).

4) Fumigation of asparagus seedling nurseries (Lacy, 1979; Molot et al., 1985), followed
by reintroduction of mycorrhizal fungi (Molot et al., 1985; Wacker, 1988).

5) Fumigation or chemical treatment of asparagus ﬁelds into which healthy asparagus
crowns are planted (Damicone et al., 1987; Lacy, 1979; Manning and Vardaro, 1977).

6) Application of biological control agents at the time of crown planting (Arriola, 1997;
Blok et al., 1997; Damicone and Manning. 1982; Tu et al., 1990; Wacker, 1988).

7) Maintenance of proper pH for healthy asparagus growth (Hodupp, 1983).

8) Use of the nitrate form of nitrogen on ﬁelds with a high potential for decline, or ﬁelds
that have begun to decline (Elmer, 1989).

9) Judicious use of NaCl on declining ﬁelds if it can be established that this does not pose
a risk to the nutrient status of the crop, soil structure, rotation crops, or groundwater
quality (Elmer, 1992).

10) Irrigation of asparagus plantings, especially crown nurseries, in times of severe water

stress (Nigh, 1990).

41

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SECTION 1

EFFECT OF SODIUM CHLORIDE AND LIME APPLICATIONS ON COMMERCIAL

ASPARAGUS AND THE EFFECT OF ALTERNATE FORMS OF CHLORIDE SALT

ON FUSARIUM CROWN AND ROOT ROT

62

ABSTRACT

Experiments with NaCl (high and low rates) and lime applications were
conducted in asparagus ﬁelds in commercial production (one healthy and one declining),
and an experiment with NaCl was conducted in a badly declined research ﬁeld at
Michigan State University from 1998 to 2000 to test the ability of these treatments to
control F usarium crown and root rot of asparagus. Growth chamber and greenhouse
studies were conducted to test the ability of alternative forms of chloride salt to reduce
disease caused by F usarium oxysporum f. sp. asparagi and F usarium proliferatum. NaCl
applications led to increased yield and fern health in the research plot, but no treatment
effects were observed in commercial ﬁelds. Soil tests in these ﬁelds indicated that NaCl
did not decrease levels of calcium, magnesium, or potassium, and did not affect pH. In
growth chamber studies with asparagus seedlings in Hoagland’s agar test tubes, and in
greenhouse studies with asparagus seedlings in soil, no forms of chloride salt tested were

more effective than NaCl in controling Fusarium crown and root rot.

63

INTRODUCTION

Fusarium crown and root rot, caused by F usarium proliferatum (T. Matsushima)
Nirenberg and F usarium oxysporum (Schlect.) f.sp. asparagi (Cohen & Heald) is the
major cause of asparagus early decline, an economically important disease worldwide.
Symptoms of the disease include yellowing and senescence of ferns, destruction of feeder
roots, collapse of storage roots, and death of crowns (Elmer et al., 1996). In a perennial
crop such as asparagus, the lack of effective chemical controls (Lacy, 1979) and the
limited resistance of available cultivars (Stephens etal., 1989), makes Fusarium crown
and root rot an especially difﬁcult disease to control.

A disease management strategy that has shown promise is the use of sodium
chloride (NaCl) salt as a soil amendment. Asparagus is an extremely salt tolerant crop,
and salt applications were recommended by gardening experts until the early part of this
century (Bennett, 1910; Burr, 1865; Harris, 1883). Elmer (1989;1992) demonstrated a
reduction in F usarium crown and root rot with NaCl applications in the greenhouse and in
small research ﬁeld plots, and some commercial asparagus growers have adopted the
practice.

Salt applications on ﬁeld soils can potentially cause problems such as loss of soil
structure, soil acidiﬁcation, nutrient displacement, groundwater contamination, and
damage to sensitive rotation crops (Brady, 1990). Although the mechanism by which
NaCl suppresses disease is not clearly understood, it is likely that the chloride molecule is
playing a role (Elmer, 1989; Elmer, 1992). Research has suggested that potassium

chloride (KCl) is not as effective as NaCl in disease suppression (Elmer, 1992), but the

64

effects of other forms of chloride salt on disease, have never been studied.

A high pH has been shown to decrease the severity of F usarium diseases in some
crops including cotton (Bell, 1989), and tomato (Jones et al., 1989). Hodupp (1983) has
demonstrated an increase in asparagus yields at high pH, and soil pH can affect the
availability of nutrients to the plant, which can often be a factor in F usarium disease
suppression (Huber, 1989; Huber, 1990).

The purpose of this study was to determine the effects of yearly lime and NaCl
applications on the yield and fern vigor of commercial asparagus plantings in Michigan,
both declining and healthy; to determine the effect of these treatments on pH, and the
quantity of sodium, magnesium, potassium, and calcium in the soil; and to determine
whether alternative forms of chloride salt could be as effective as NaCl in disease

suppression.

MATERIALS AND METHODS

Growth chamber studies with chloride salts. Growth chamber studies were
conducted to test the ability of different forms of chloride salt to suppress disease caused
by F oxysporum f. sp. asparagi and F proliferatum. In tests with F oxysporum f. sp.
asparagi, treatments consisted of the following: i) a control (no salt added), ii) NaCl
(17.1 mM), iii) NaCl (34.2 mM), iv) CaCl2 (17.1 mM), v) NH4C1 (34.2 mM). For tests
with F proliferatum, treatments consisted of i) a control, ii) NaCl (34.2 mM), iii) CaCl2
(17.1 mM), iv) MnCl2 (8.55 mM), v) NH4C1 (34.2 mM). Hoagland’s solution agar

(Damicone and Manning, 1982; Davis, 1967) was prepared and chloride salt added to

65

provide an equal amount of chloride to each treatment except the control, and the MnCl2
treatment, which was toxic to the plants when too much was applied. All treatments were
standardized to a pH of 6.0 using sodium hydroxide (NaOH). Agar (12-16 g/L) was
added to the control and each treatment to achieve a uniform consistency. Each solution
was autoclaved and added aseptically to sterile glass test tubes (16x150 mm).

Asparagus seeds (cv. ‘Mary Washington’, Rupp Seeds, Wauseon, OH ) were
sterilized according to the method of Damicone et al. (1981) by spinning on a magnetic
stir plate for 24 hours in a mixture of 100 ml acetone and 2.5 g benomyl (5 g Benlate 50
DF, E. I. du Pont de Nemours & Co. Inc., Wilmington, DE), rinsing them in acetone (3x)
and in distilled water (3x), then spinning for 1 hour in 1.05% sodium hypochlorite (20%
household bleach), and rinsing again in sterile distilled water (3x). Seeds were air dried,
and placed on water agar under sterile conditions. Once hypocotyls began to emerge,
healthy, uniform seedlings were transferred to the test tubes under sterile conditions.
Tubes were capped, and seedlings were grown for 10 days. Twenty and 40 seedlings
were used for each treatment in tests with F oxysporum f.sp. asparagi and F
proliferatum, respectively.

F oxysporum f. sp. asparagi and F proliferatum were isolated from diseased
asparagus crowns and stems in 3 commercial asparagus ﬁelds in Oceana County,
Michigan, and a research plot at the Michigan State University Botany and Plant
Pathology Farm. Four hundred and ﬁfty nine isolates were tested for pathogenicity by
inoculating seedlings growing in test tubes ﬁlled with Hoagland’s Solution agar in the

method described by Stephens and Elmer (1988). Isolates judged to be highly virulent on

66

asparagus seedlings were put into storage on silica gel according to the procedure
described by Windels et al. (1988). Highly virulent isolates of F oxysporum f. sp.
asparagi (isolate 2-7) and F proliferatum (isolate P-67), were grown on carnation leaf
agar for two weeks under ﬂuorescent lights. The surface of the plates were ﬂooded,
scraped with a scalpel to loosen spores, ﬁltered through three layers of sterile cheesecloth,
and diluted with sterile water to 1x106 conidia/ml using a hemacytometer. A spore
suspension (0.5 ml) was added to each seedling tube, and plants were grown for 4 weeks
in a 25°/20° C (day/night) growth chamber under ﬂuorescent plant lights, with a 16 hour
photoperiod.

Seedlings were assessed for disease based on the percentage of the root system
that exhibited lesions or had collapsed. Dead plants were considered to be 100%
infected. Each experiment was conducted twice.

Greenhouse studies with chloride salts. ‘Mary Washington’ asparagus seeds
were sterilized as previously described and germinated in the dark on sterile water agar
petri plates. Seedlings were then planted into 64 cc plug trays ﬁlled with High Porosity
Professional Planting Mix (Michigan Peat Co., Houston, TX) as soon as hypocotyls
emerged. Plants were grown for 14 days, then clean, uniform seedlings were transfered to
512 cc plastic pots.

Inoculum was prepared using a modiﬁcation of the method described by Stephens
and Elmer (1988). Distilled water (100ml) was added to breathable autoclave bags (Field
& Forest Products Inc., Peshtigo, WI) containg 200g of pearl millet seed (Rupp Seeds,

Wauseon, OH). Bags were autoclaved for 1 hour on consecutive days, and allowed to

67

cool. Several pieces of silica gel infested with the isolates were added to separate bags
and allowed to grow for 2 weeks. Bags were shaken occasionally to prevent clumping.
Millet seed was dried in autoclaved paper bags for 7 days, then ground in a Thomas Mill
(Arthur H. Thomas Co. Philadelphia, PA). Ground millet (150 g) infested with a mixture
of F osysporum f. sp. asparagi (isolates 2-7 and F OA-50) was combined with ground
millet (150g) infested with a mixture of F proliferatum (isolates P-67 and M-63 74), and
added to 40 L of a 4:1 mixture of steamed sandy loam to Bacto High Porosity
Professional Planting Mix (Michigan Peat Co., Houston, TX), to yield an inoculum
density of 320,000 CFUs per gram of soil. Isolate F OA-50 originating from a Michigan
asparagus ﬁeld and isolate M-63 74 from an asparagus ﬁeld in Connecticut were obtained
from W. Elmer, Connecticut Agricultural Experiment Station. Soil for the uninfested
control treatments consisted of a 4:1 mixture of steamed sandy loam to Bacto High
Porosity Professional Planting Mix which no pathogen was added.

Seedlings were washed free of soil, and inspected for lesions or discoloration, and
uniform, healthy crowns were planted into either infested or uninfested soil in 512 cc
plastic pots. The following treatments were applied to each of 16 plants in infested soil in
amounts adjusted to give equivalent CI' content in each pot: i) no salt (control), ii) NaCl
(0.32 g/pot), iii) CaC12*2HZO (0.40 g/pot), iv) NH4C1 (0.29 g/pot), v) KCl (0.40 g/pot),
and vi) Mn2Cl*4HzO (0.53 g/pot). An uninfested control was also included in this
experiment. Plants were allowed to grow for 3 weeks, and were fertilized twice weekly
with 15 ml of Hoagland’s solution. Plants were watered as needed with distilled water.

Treatments were arranged in a completely randomized design, and the experiment was

68

conducted twice. Fresh weights of roots and shoots were taken, and the percentage of the
root system exhibiting lesions was estimated.

NaCl and lime applications in established asparagus ﬁelds. A single research
plot (688 m2) was established at the Botany and Plant Pathology Research farm in East
Lansing, Michigan in a 17-year-old asparagus ﬁeld (cv. ‘Mary Washington’) with severe
symptoms of F usarium related decline including thin stands, missing crowns, and poor
fern growth. Rows were spaced 1.52 m apart, and the soil was a sandy loam Treatments
consisted of an untreated control, and rock salt (N aCl) (PureMelt Ice and Snow Melting
Salt, ICM Kalium Co., Bannockbum, IL) applications (1120 kg/ha). Salt was applied on
17 April (1998), 30 April (1999), and 25 April (2000), using a Gandy drop spreader
(Gandy Co., Owatonna, MN).

Treatments were arranged in a randomized complete block design with 5 blocks,
each separated by an untreated buffer row. Treatment plots consisted of a 6.1 In section
of a single row. Pelletized lime (Nutralime, Mineral Processing Co., Carey, OH) was
applied to all treatments in the spring of each year at a rate of 6719 kg/ha, and fertilizer
(12-12-12, Northern Star Minerals, E. Lansing, MI) was applied at 336 kg/ha. Pesticide
sprays were applied as needed for the control of insects, weeds, and foliar diseases.
Spears greater than 12.71 cm in height were harvested by hand snapping between 5 and
26 May in 1998 (10 harvests), between 10 May and 9 June 1999 (17 harvests), and
between 2 May and 6 June in 2000 (14 harvests).

Two research plots were established in Oceana County, Michigan in loamy-sand

asparagus ﬁelds (cv. ‘Syn-456') in commercial production, that had been previously

69

cropped in asparagus. One plot was established in a healthy-appearing 2.02 ha ﬁeld of
11-year-old asparagus. A second plot was established nearby in a declining (thin, and
unproductive) 3.44 ha ﬁeld of 10-year-old asparagus. In each plot, crowns were planted
approximately 25 cm apart in rows spaced 1.52 m apart. Treatments consisted of the
following: i) an untreated control, ii) a low rate of salt (560 kg/ha), iii) a high rate of salt
(1120 kg/ha), iv) a pelletized lime application (6719 kg/ha), v) a low rate of salt and
pelletized lime, vi) and a high rate of salt and pelletized lime. All treatments were applied
using a Gandy drop spreader on 10 April (1998), 27 April (1999), and 19 April (2000).

Treatments were arranged in a randomized complete block design with 6 blocks.
Treatment plots consisted of a 6.1 m section of 3 parallel rows. Two buffer rows
separated each treatment and 3.05 m separated each block within the rows. Fertilizer and
pesticide sprays, with the exception of lime were applied to these plots as needed by the
growers.

All spears greater than 12.70 cm in height were harvested by hand snapping
between 2 May and 15 June (26 harvests) in 1998, 3 May and 12 June in 1999 (25 and 24
harvests for the declining and healthy ﬁelds, respectively), and 2 May and 13 June and 30
April and 13 June for the declining and healthy ﬁelds respectively (22 harvests each) in
2000. All plots were evaluated for fem health by counting the number of stalks
(excluding volunteer seedlings) greater or less than 0.79 cm in diameter 12.70 cm above
the soil surface approximately 5 weeks after the termination of harvest.

Soil samples were taken from each treatment plot in both the experimental and

commercial ﬁelds in April (before treatments were applied), July, and October 1998, in

70

April and October 1999, and in April 2000. Samples consisted of eight 30.48 cm soil
cores split into the top 15.24 cm and bottom 15.24 cm and homogenized. All soil
samples were analyzed at the Michigan State University Soil Testing Laboratory for pH,
and concentrations of calcium, potassium, magnesium, and sodium.

Statistical analysis. The Proc Mixed procedure was used with SAS statistical
analysis soﬂware (SAS Institute Inc., Cary, NC) for all analyses. Data from all tests were
evaluated by comparing Bonferroni adjusted Least Squared means for all treatments. In
soil tests, data from treatments in spring 2000 (after two treatment applicatations) were
compared using spring 1998 data (before any treatments were applied) as a covariate.
Differences in Na levels between fall 1998 and fall 1999 were compared to determine
whether Na had built up in the soil between these years. Harvest data from the three
years were evaluated as repeated measures. Percentage data (root system covered with
lesions) for growth chamber trials with the pathogen F proliferatum were transformed
using the arcsine transformation to normalize data. Because of the heterogeneous
variance in trial by treatment combinations in percentage data (root system covered with
lesions) for greenhouse trials, variance component estimation was carried out by using the
Repeated statement with Group Option in SAS. L. S. means differences were considered
signiﬁcant at P=0.05, and data for the two replications of each experiment is presented

together unless trial by treatment interactions were signiﬁcant.

RESULTS

Growth chamber studies with chloride salts. A large fraction (84.4%) of the

71

root system of control plants exhibited lesions or collapse caused by F. proliferatum,
while plants treated with NaCl (57.0%) or CaCl2 (69.6%) exhibited signiﬁcantly less root
rot (Figure 1). MnCl2 did not signiﬁcantly limit root rot compared to the control, but was
not signiﬁcantly different than the NaCl and CaCl2 treatments. NH4C1 treatments were
not effective and resulted in signiﬁcantly more disease than the control.

Root rot caused by F oxysporum f. sp. asparagi was limited signiﬁcantly by a
high rate (32.4 mM) of NaCl (44.4%) compared to the control (77.5%) and all other
treatments with the exception of the low rate (17.1 mM) of NaCl (62.3%) (Figure 2).
Neither CaCl2 or the low rate of NaCl were signiﬁcantly more effective than the control
in reducing disease caused by this pathogen. Plants treated with NH4C1 were signiﬁcantly
more diseased than the control.

Greenhouse studies with chloride salts. In experiment 2, only NaCl-treated
plants exhibited signiﬁcantly less root rot (18.6%) than the infested control (33.9%), but
were not signiﬁcantly different from the other treatments (Figure 3). In both experiments,
the uninfested control remained virtually disease free (< 1%).

In experiment 1, NaCl treated plants had signiﬁcantly greater fresh root weight
(1.50 g) than the infested control (0.95 g) and all other treatments except KCl (1.13 g)
(Figure 4). Signiﬁcant differences in fresh root weight were not observed among the
treatments in experiment 2, although NaCl had a higher mean weight (1.30 g) than the
infested control (0.89 g). The uninfested control had signiﬁcantly greater fresh root
weights than all other treatments in both experiments.

There were no signiﬁcant differences in fresh shoot weight when treatments were

72

 

a.

O

O
1

be

O
O
l

h
O
l

N
O
l

 

 

Root system exhibiting lesions (%)
8
l

 

O
l

NH4C| Control MnClz CaCIz NaCl

Figure 1. Effect of different forms of chloride salt on root
rot (%) in two experiments with asparagus seedlings grown
in Hoagland's agar in test tubes and infested with F.
proliferatum. Different letter notation indicates signiﬁcant
differences in L 8 means comparisons.

73

 

 

 

Root system exhibiting lesions (%)

 

Figure 2. Effect of different forms of chloride salt on
root rot (%) in two experiments with asparagus seedlings
grown in Hoagland's agar in test tubes, and infested with
F. oxysporum f. sp. asparagi. Error bars represent
standard errors, and different letter notation indicates
significant differences in L S means comparisons.

74

Root system exhibiting lesions (%)

 

60
Experiment 1
50 ~
40 4
30 ~

204

10*

 

 

NI-I4CI MnClz CaCIZ KCI Infested NaClUnlnfested

 

 

60
Experiment 2
50 ~
40- ab
30 -

20a

10*

 

 

Infested NH4CI CaCIZ KCI MnCIZ NaClUnlnfested

Figure 3. Effect of different forms of chloride salt on
root rot (%) in two greenhouse experiments with
asparagus plants grown in soil infested with F.
oxysporum f. sp. asparagi and F. proliferatum.

Error bars represent standard error, and different
letter notation indicates signiﬁcant differences in

L 8 means comparisons.

75

 

 

Experiment 1 c

 

 

 

NH4CI CaC12 MnCIZ Infested KCI NaClUninfested

 

Experiment 2 9

Root weight (g)

 

 

 

NH4¢I Infested CaC12 KCI MnCIZ NaClUnlnfested

Figure 4. Effect of different forms of chloride salt on root
weight (g) in two greeenhouse experiments with asparagus
plants grown in soil infested with F. oxysporum f. sp.
asparagi and F. proliferatum. Error bars represent
standard error, and different letter notation indicates
signiﬁcant differences in L 8 means comparisons.

76

compared to the control (Figure 5). However, NaCl was superior to CaClz, MnClz, and
NH4Cl treatments. The uninfested control had signiﬁcantly greater shoot weight than all
treatments.

Field trials with NaCl and lime applications. No signiﬁcant treatment or year
by treatment differences were found in asparagus yield in the commercial ﬁelds (Figure
6). Signiﬁcant differences between years were observed, and were due to differences in
the number of harvests taken each year, as well as environmental conditions. Although
signiﬁcant treatment differences were found between the control and 1120 kg NaCl
treatments in the diseased ﬁeld in 1999 (P=0.020) no overall treatment or year by
treatment effect was found in the percentage of stalks larger than 0.79 cm in diameter
(Figure 7).

Signiﬁcant differences (P=0.0420) in year by treatment effect were found in the
comparison of yield between salt treated and untreated plots in the research ﬁeld at the
Michigan State University Botany and Plant Pathology Farm in 2000, but not in 1998 or
1999 (Figure 8). Signiﬁcant differences were found in the percentage of stalks larger than
0.79 cm in diameter in 2000 (P=0.0049), but not in 1998 or 1999 (Figure 9).

Soil tests. NaCl applications did not signiﬁcantly affect levels of pH, potassium,
magnesium, or calcium after two treatments in the research plot at the MSU Botany and
Plant Pathology Farm (see appendix). Sodium levels were signiﬁcantly higher in salt-
treated plots (90 ppm) than in untreated plots (21 ppm) in the top 15 cm of soil.
Similarly, in the 15-30 cm soil level sodium levels were higher in the treated plots (94.6

ppm) than in untreated plots (26.6 ppm).

77

 

N
L

Shoot weight (g)

 

 

 

Nl-I4CI MnCI2 CaClZ Infested KCI NaClUninfested

Figure 5. Effect of different forms of chloride salt on
shoot weight (g) in two greeenhouse experiments with
asparagus plants grown in soil infested with F. oxysporum
f. sp. asparagi and F. proliferatum. Error bars represent
standard error, and different letter notation indicates
signiﬁcant differences in L 8 means comparisons.

78

Mean harvest weight (g)

8000

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

7000 - I
x i
6000 - / a i /
/ / w
/ A f M
5000 - A / C A
A / x
A / g /
4000 4 x / / ,
x / /
x / i x
3000 - x / j v
/ / v
/ V j /
2000 - v / / z
r / r
/ / / V
1000 - / x A ,
x / ’ ,
/ / / /
0 - 'r‘ f 4 1'
Control 500 lb 1000 lb Lime 500 81 L 1000 81 L
8000
7000 - - '1998'
[Z] '1999'
6000 1 - '2000'
5000 -
4000 4
3000 -
2000 —
1000 -
o _
Control 500 lb 1000 lb Lim 500 a L 1000 a L

 

 

 

 

 

 

 

 

 

Healthy Field

Declining Field

Figure 6. Mean harvest weight of asparagus spears from

commericial ﬁeld plots in Oceana County, M1 1998-2000
during 1998 to 2000. Error bars represent standard error.

79

Percentage of fern > 0.79 cm in diameter

 

80

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

Healthy Field
I
60 - i x I E I I
/ / / / / V
/ / x / K
/ / / / /
V / V / j /
/ / / / / x
40 A ” f / x x r
M / M / r /
/ / A / x r
/ / / x / /
/ j j x / /
/
2° ‘ j / x S C /
r A V / / /
/ / i; / / /
’ ’ / v /
’ ’ / / x /
d ’ x
0 _ 4 1‘ "f 4 1"
Control 500lb 10001b Lime SOOSL 1000&L
80
-1998 . . .
ﬂ] 1999 Declining Field
- 2000
60 '1
g I
/
, s
40 a j /
/ i
/ x
/ z
/
20 - y j
/ z
/ /
/ z
’ r
/
0 " 14 K

 

 

 

 

 

 

 

 

 

Control 560kg 1120kg Lime 560&L 1120&L

Figure 7. Percentage of asparagus stalks > 0.79 cm
in diameter in commercial ﬁeld plots in Oceana County,
MI 1998 to 2000. Error bars represent standard error.

80

3000

 

 

- No Salt pgooo42
EICZSMI

2500 ns

 

 

 

2000 ~

1500 - "s

1000 -

Mean harvest weight (g)

500 4

 

\\\\\\\\\\\\\\14
d\\\\\\\\\\\\\\\\\\\‘l—+

 

 

 

 

 

 

o ' . / .
1998 1999 2000

 

Figure 8. Mean harvest weight of asparagus from research
plot at MSU Botany and Plant Pathology Farm 1998 to 2000.
Error bars represent standard error.

81

60

 

- No NaCl
m NaCl

 

 

50

404

309

20‘

1o—

 

 

P=0.0049

\K\\\\\\\\\\\\\\\\\\l-+

 

 

\\\\\\\\\\\\\\\V-4l

 

 

 

Percentage of fern > 0.79 cm in diameter (%)

1 998

 

 

 

1 999 2000

Figure 9. Percentage of asparagus stalks > 0.79 cm in
diameter in research plot at MSU Botany and Plant
Pathology Farm 1998-2000. Error bars represent
standard error.

82

In the commercial ﬁeld plots in Oceana County, no signiﬁcant differences were
found in sodium levels in the top 15 cm of soil (see appendix). Signiﬁcantly higher
sodium levels were found in the 1120 kg NaCl (51 ppm) and 1120 kg NaCl + lime
treatments (56 ppm) than in any other treatment in the 15-30 cm soil level. In addition,
the 560 kg NaCl treatment showed signiﬁcantly higher sodium levels (36 ppm) than
control (24 ppm), lime (24 ppm) or 560 kg NaCl + lime (29 ppm) treatments. In general,
increasing levels of salt application led to increased sodium levels in the soil both with
and without lime and at both soil levels. Analysis of sodium levels at the two fall
samplings (1998 and 1999) did not show signiﬁcant increases in sodium levels in either
of the commercial ﬁelds at either soil level in any treatment (data not shown). Signiﬁcant
increases in sodium levels (+35 ppm) were shown from these sampling dates from the 15-
30 cm level in the heavier soil at the MSU Botany and Plant Pathology Farm. pH,
magnesium and calcium levels were signiﬁcantly affected by lime applications in the
commercial ﬁelds. Signiﬁcantly higher pH levels were found in the 1 120 kg NaCl + lime
(6.50), lime (6.55), and 560 kg NaCl + lime (6.58) treatments than in the 560 ngaCl
treatment (6.19) and signiﬁcantly higher levels were found in the lime treatment, than in
the 1 120 kg NaCl treatment (6.27) in the top 15 cm of soil in combined results from both
ﬁelds. Combined results also show that in the treatment that received 1120 kg NaCl +
lime had signiﬁcantly higher pH levels (6.26) in the 15-30 cm soil level than either the
control treatment (5.87) or the 560 kg NaCl treatment (6.00). Differences among
treatments in potassium levels were not signiﬁcant in either the top 15 cm or 15-30 cm

level of soil. Signiﬁcantly greater levels of calcium were found in the top 15 cm of soil in

83

the 560 kg NaCl + lime (970 ppm) and the 1120 kg NaCl + lime (979 ppm) treatments
than in the 560 kg NaCl treatment (755 ppm). No signiﬁcant differences in calcium level
were found in the 15-30 cm soil level. Signiﬁcantly higher magnesium levels were found
in the top 15 cm of soil in the lime (168 ppm) and 560 kg NaCl + lime (162 ppm)
treatments than in the untreated control (80 ppm) in the declining ﬁeld, but no signiﬁcant
differences were found in the healthy ﬁeld at this soil level. Signiﬁcantly higher levels of
magnesium were also found in the lime treatment (154 ppm) than in the 560 kg NaCl

treatment (116 ppm) at the 15-30 cm soil level.

DISCUSSION

F usarium crown and root rot has become limiting to asparagus production in
Michigan as growers are forced to rely more on ﬁelds that had been previously cropped to
asparagus. In our study, NaCl treatments suppressed disease in a research plot at the
MSU Botany and Plant Pathology Farm verifying earlier reports (Elmer, 1989; Elmer,
1992). However, NaCl treatments did not signiﬁcantly affect disease in commercial
replant ﬁelds.

Yield increases with NaCl applications are closely related to its ability to suppress
Fusarium disease (Elmer, 1990b; Elmer, 1992; Elmer, 1995; Elmer and LaMondia, 1997).
The effectiveness of this treatment may be correlated to the amount of disease pressure in
the ﬁeld. Experiments conducted with NaCl and its effect on asparagus yield have been
conducted in Connecticut, where asparagus production was abandoned because Fusarium

decline was so severe (Elmer, 1992). Our own study showing the effectiveness of salt

84

was conducted in a severely declined plot at the Botany and Plant Pathology Farm at
MSU. Yield increases have yet to be demonstrated in commercial ﬁelds, which are
generally taken out of production once yield becomes limited by disease. It is possible
that salt treatments will show signiﬁcant yield effects with continued applications as
commercial ﬁeld plots decline further. Similarly, lime applications, which have been
shown to decrease Fusarium disease on other crops (Bell, 1989; Jones et al., 1989) did not
signiﬁcantly affect yield or stalk diameter.

In the commercial ﬁelds, treatment areas were separated by at least 3.05 m on all
sides. However, sodium levels and pH levels increased over time in plots not treated with
NaCl or lime. Increases in pH may be partially explained by the fact that each of the
ﬁelds was accidentally treated with a single lime application during the experiment by the
growers. There was some movement of sodium ﬁ'om treated into untreated plots over
time, especially in the lower strata of soil. Although data from soil tests seems to suggest
that much of the movement occured while the crowns were dormant, the fact that there
was some salt movement into untreated plots may have had an effect on the results of this
study. This may partially explain why yield differences in these plots have not been
shown, and suggests that in future tests, treated areas should be separated by a greater
distance.

The effect of NaCl on the nutrient content and pH of soils, on rotation crops and
groundwater, and the potential development of soil sodicity are all concerns that must be
carefully considered. Our results did not show decreases in the levels of magnesium,

potassium, or calcium with NaCl applications, and salt does not appear to be affecting

85

pH. Our analyses also suggest that salt moves through the soil proﬁle quickly in
Michigan asparagus ﬁelds, and we did not show signiﬁcant buildup of sodium in soils in
commercial ﬁelds between October 1998 and October 1999. Mechanical analysis have
shown that these ﬁelds can have a sand content of 85% or more, and NaCl may leach
easily through the proﬁle with rainwater. Because there is little structure in these soils to
begin with, sodicity buildup is less of a concern than with other soil types. Furthermore,
no changes in bulk density of soil have been detected in previous experiments with NaCl
at these levels (Elmer, 1992). Because NaCl does not appear to build up in soils, and
because asparagus ﬁelds are often left fallow for a season after being taken out of
production, the affect of salt on rotation crops may not be of great concern in this region.
Furthermore, no weed control has been observed with direct salt applications at these
levels (Elmer, 1992) suggesting that the affect of salt applications on other plants grown
in these ﬁelds a year or more later may be relatively benign. Caution must be exercised,
however, in drawing these conclusions based on less than three years of soil data, and
observations with particular weeds. We have also shown, with soil tests in the sandy
loam in the research plots at MSU, that salt buildup is more of a concern in heavier
soils. It may be wise for asparagus growers to test their soils for sodium levels after
repeated salt applications, and to avoid salt-sensitive crops such as carrots or fruit trees in
their rotations for several years. Salt buildup, if it does become a problem, can be treated
with applications of gypsum or by irrigating the affected soils (Brady, 1990).

Because of the potential problems with NaCl applications to commercial ﬁeld

soils, we also compared the potential of alternative forms of chloride salt, that could also

86

supply nutrients necessary for plant health, to that of NaCl in disease suppression. The
effectiveness of NaCl in suppressing disease in our studies using agar ﬁlled test tubes
conﬁrms earlier reports (Elmer, 1989, Elmer, 1992). NaCl was more effective than other
forms of chloride salt in controlling disease caused by F proliferatum, and plants treated
with NaCl had lower levels of disease caused by F oxysporum f. sp. asparagi than any
other treatment. These results suggest that disease control is not simply the result of
increased chloride nutrition.

Calcium has been shown to be important in disease resistance in some plant-
disease systems by protecting pectate from maceration, and ameliorating the effects of
toxins (Huber, 1990). The increased disease control with CaCl2 shown in tests against the
pathogen F proliferatum may be due to the effect of increased calcium or to an unknown
disease control mechanism involving chloride. Manganese has been implicated as an
important element in disease control in a number of different plant systems (Huber,
1990). Its lack of effectiveness in these tests may be due to its immobility at the relatively
high pH of the agar (6.0). The increased chloride nutrition provided did not appear to
affect disease in this treatment. It is interesting to note that NH4Cl-treated plants were
more diseased than control plants in tests with both pathogens. In other systems, it has
been suggested that increased nitrogen can make plants more susceptible to disease by
increasing succulence, and decreasing physical barriers to invasion (Huber, 1990). Elmer
(1989) has also shown that plants fertilized with ammonium nitrogen were more
susceptible to disease than those fertilized with nitrate nitrogen. It appears that increased

ammonium levels increased the plants susceptibility to the pathogens in these studies.

87

The results of greenhouse experiments with different forms of chloride salt
generally conﬁrm the results of the test tube studies. No other form of chloride salt tested
was more effective in controlling disease or showed greater fresh weight or roots or
shoots than NaCl when equivalent levels of chloride were applied. In these experiments,
NH4C1 again seemed to favor disease.

Although we did not ﬁnd signiﬁcant increases in plant health with NaCl
applications in commercial ﬁelds, our research has conﬁrmed the potential of NaCl and
other chloride salts for use in disease control. Future studies on the long term effects of
salt applications to commercial ﬁelds, both on plant health, and soil could add
signiﬁcantly to our present understanding of its usefulness. Despite considerable effort
(Elmer, 1990a; Elmer, 1995; Elmer and LaMondia, 1997), the mechanism by which NaCl
suppresses disease is not yet clearly understood, and continued research in this area could
be beneﬁcial to the effort to ﬁnd an effective control for Fusarium crown and root rot of
asparagus. It is also possible that higher levels of KCl or CaCl2 could have a greater
effect on disease suppression, and since the potentially negative effects of NaCl are not a
concern with these salts, further experimentation with these salts should not be

discounted, since some disease suppression was detected with each.

88

LITERATURE CITED

Bell, A. A. 1989. Role of nutrition in diseases of cotton. Pages 167-204 in:
Soilbome Plant Pathogens: Management of Diseases with Macro- and
Microelements. A. W. Engelhard ed. APS Press, St. Paul.
Bennett, 1. D. 1910. The Vegetable Garden. Doubleday, Page & Company, New
York.
Brady, N. C. 1990. The Nature and Properties of Soils. MacMillan Publishing
Company, New York.
Burr, F., 1865. Field and Garden Vegetables of America. J. E. Tilton and
Company, Boston.
Damicone, J. P., Cooley, D. R., and Manning, W. J. 1981. Benomyl in acetone
eradicates F usarium moniliforme and F usarium oxysporum from asparagus seed.
Plant Disease 65: 892-893.
Damicone, J. P., and Manning, W. J. 1982. Avirulent strains of F usarium
oxysporum protect asparagus seedlings from crown rot. Canadian Journal of Plant
Pathology 4:143-146.
Davis, D. 1967. Cross-protection in Fusarium wilt diseases. Phytopathology

57:311-314.
Elmer, W. H. 1989. Effects of chloride and nitrogen form on growth of
asparagus infected by F usarium spp. Plant Disease 73:736-740.
Elmer, W. H. 1990a. Effect of NaCl on carbohydrates and malate production in

asparagus roots and on infection by Fusarium. (Abstract). Phytopathology

89

80: 1025.

Elmer, W. H. 1990b. Suppression of F usarium crown and root rot of asparagus
with chloride and different forms of nitrogen fertilizers. Acta Horticulturae
271:323-329.

Elmer, W. H. 1992. Suppression of Fusarium crown and root rot of asparagus
with sodium chloride. Phytopathology 82:97-104.

Elmer, W. H. 1995. Association between Mn-reducing root bacteria and NaCl
applications in suppression of F usarium crown and root rot of asparagus.
Phytopathology 85: 1461 -1467.

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.

Elmer, W. H., and LaMondia, J. A. 1997. Studies on the supression of Fusarium
crown and root rot of asparagus with NaCl. Pages 54-67 in: The IX international
Asparagus Symposium, Pasco, WA, B. Benson ed.

Harris, J. 1883. Gardening for Young and Old. Orange Judd Company, New
York.

Hodupp, R. M. 1983. Investigation of factors which contribute to asparagus
decline in Michigan. M. S. Thesis-Michigan State University, E. Lansing, 54 pp.
Huber, D. M. 1989. The role of nutrition in the take-all disease of wheat and
other small grains. Pages 46-74 in: Soilbome Plant Pathogens: Management of

Diseases with Macro- and Microelements, A. W. Engelhard ed. APS Press, St.

90

Paul

Huber, D. M. 1990. Pages 357-394 in: The use of fertilizers and organic
amendments in the control of plant disease. Handbook of Pest Management in
Agriculture, D. Pinentel (ed.), CRC Press, Boca Raton, FL.

Jones, J. P., Engelhard, A. W., and Woltz, S. S. 1989. Management of Fusarium
wilt of vegetables and omamentals by macro- and microelement nutrition. Pages
18-32 in: Soilbome Plant Pathogens: Management of Diseases with Macro- and
Microelements, A. W. Engelhard ed. APS Press, St. Paul.

Lacy, M. L. 1979. Effects of chemicals on stand establishment and yields of
asparagus. Plant Disease Reporter 63:612-616.

Stephens, C. T., De Vries, R. M., and Sink, K. C. 1989. Evaluation of Asparagus
species for resistance to F usarium oxysporum f. sp. asparagi and F moniliforme.
HortScience 24: 365-368.

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.

Windels, C. E., Burnes, P. M., and Kommedahl, T. 1988. Five-year preservation

of F usarium species on silica gel and soil. Phytopathology 78:107-109.

91

SECTION II

THE USE OF FUNGICIDES AND BIOLOGICAL CONTROLS IN THE

SUPPRESSION OF FUSARIUM CROWN AND ROOT ROT OF ASPARAGUS

92

ABSTRACT

Growth chamber, greenhouse, and ﬁeld experiments were conducted with
fungicides and biological controls including experimental non-pathogenic isolates of
F usarium oxysporum to test their ability to control disease caused by F oxysporum f. sp.
asparagi and F proliferatum. In greenhouse studies with asparagus seedlings in soil,
Trichoderma harzianum, benomyl, and ﬂudioxonil treatments increased root weight and
decreased root disease compared to the infested control when a low level of F oxysporum
f. sp. asparagi and F proliferatum was used. The fungicide ﬂudioxonil limited plant
death caused by F usarium spp. at high inoculum levels, while T. harzianum was not
effective. Non-pathogenic F oxysporum isolates were effective in limiting F usarium
disease on asparagus seedlings in culture tubes, although isolates differed in their ability
to control disease caused by F oxysporum f. sp. asparagi and F proliferatum. In
greenhouse studies no signiﬁcant differences in plant death were found between
asparagus plants growing in media infested with F oxysporum f. sp. asparagi and F
proliferatum and left untreated, and those treated with non-pathogenic F oxysporum.
Our attempt to test fungicides and biological control products under commercial ﬁeld

conditions was not successful due to low disease pressure.

93

INTRODUCTION

The fungal pathogens F usarium proliferatum (T. Matsushima) Nirenberg and
F usarium oxysporum (Schlect.) f. sp. asparagi (Cohen & Heald) cause Fusarium crown
and root rot, the primary factor in asparagus early decline which is an economically
important disease in Michigan and throughout the world (Blok et al., 1997; Elmer et al.,
1996; Faloon and F auser-Kevem, 1996; Fantino, 1990; Grogan and Kimble; Tu et al.,
1990). Efforts to manage the disease through the development of resistant cultivars have
met with limited success (Elmer et al., 1997; Stephens et al., 1989). Although fungicides
have not provided adequate disease suppression (Di Linnea and Foletto, 1990; Lacy,
1979), the chemical 2-(4-Thiazolyl)benzimidazole (Novartis Crop Protection Inc.,
Greensboro, NC) was effective in New Zealand study (Faloon and Fauser-Kevem, 1996)
but is not registered for use on asparagus in the U.S. Currently, fungicides are not used to
manage the disease in Michigan.

Asparagus is a deep-rooted perennial, making application of chemical or
biological controls difﬁcult once the crop is established. Commercial asparagus is seeded
and grown in specialized seedling nurseries for one year before crowns are dug out and
planted into production ﬁelds. Infection of seedlings while in the nursery predisposes a
crop to severe decline (Di Lenna and F oletto, 1990; Lacy, 1979) making the seedling
stage a critical time to apply control measures for F usarium crown and root rot.

Researchers have investigated the potential of biological control agents to

suppress Fusarium disease on a number of crops (Beale and Pitt, 1990; Cassini et al.,

1985; Davis, 1967; Hervas et al., 1995; Larkin et al., 1996; Mandeel and Baker, 1991;

94

Marois, 1993; Oyarzun et al., 1994) The fungus T richoa’erma harzianum has been shown
to be effective against F oxysporum f. sp. asparagi on asparagus seedlings in greenhouse
trials (Arriola, 1997). Nonpathogenic strains of F oxysporum have also consistently
shown effectiveness against F usarium oxysporum f. sp. asparagi in growth chamber and
greenhouse studies, and were shown to be effective in one ﬁeld study, but not in others
(Blok et al., 1997; Damicone and Manning, 1982; Tu et al., 1990).

The objective of this study was to determine whether fungicides and biological
controls including experimental non-pathogenic isolates of F oxysporum could control
disease caused by F oxysporum f. sp. asparagi and F proliferatum in growth chamber,

greenhouse, and ﬁeld experiments.

MATERIALS AND METHODS

Greenhouse experiments with fungicides and a biological control product.
Asparagus seeds were sterilized by spinning on a magnetic stir plate for 24 hours in a
mixture of 100 ml acetone and 2.5 g benomyl (5 g Benlate 50 DF, E. I. du Pont de
Nemours & Co. Inc., Wilmington, DE), rinsing them 3 times each in acetone then
distilled water, then spinning for 1 hour in 1.05% sodium hypochlorite (20% household
bleach), rinsing 5 times with sterile distilled water and blotting on a sterile paper towel
(Damicone et al., 1981; Elmer and Stephens, 1989). Seeds were germinated in the dark
on sterile water agar petri plates, and planted into 96 cc plug trays ﬁlled with Bacto High
Porosity Professional Planting Mix (Michigan Peat Co., Houston, TX) as soon as

hypocotyls emerged.

95

Plants were grown for 14 days, then clean, uniform seedlings were transfered to
512 cc plastic pots ﬁlled with steam sterilized sandy loam soil. Soil for treatments
containing F oxysporum f. sp. asparagi and F proliferatum was infested by growing
cultures of the pathogens on sterile millet seed for two weeks, drying the seed in sterile
paper bags, grinding the millet, and mixing equal parts infested with each pathogen into
steam sterilized soil in a cement mixer for 4 hours (Wacker, 1988). Separate experiments
were performed with a high (200,000 cfu/g soil) and low concentration (12,600 cfu/g soil)
of pathogen inoculum. Each experiment was conducted twice.

Treatments included an uninfested control (untreated) mixed with sterile ground
millet seed, an infested control, and infested soil drenched with either benomyl (Benlate
50 DF at 0.6g/L), 2-(4-Thiazolyl)benzimidazole (Mertect 340-F at 2.34 ml/L), ﬂudioxonil
(Maxim 1.56 ml/L), or T. harzianum (Root Shield at 8.98 g/L). The fungicide 2-(4-
Thiazolyl)benzimidazole was excluded from the experiment with low pathogen
concentrations because the manufacturer does not plan to label this product for use on
asparagus (personal communication, Novartis Crop Protection Inc.). All drenches were
applied two days after planting.

Effectiveness of the treatments in the experiments with a high inoculum level was
evaluated by counting the number of dead plants weekly, and effectiveness of the
treatments in the experiment with a low inoculum level was evaluated by measuring the
fresh weight of roots and shoots, and visually estimating the percentage of the root system
covered with lesions.

Evaluation of nonpathogenic F. oxysporum isolates for control of Fusarium

96

Crown and Root Rot of asparagus. Four commercial asparagus ﬁelds in Oceana Co..
Michigan, and a research ﬁeld on the campus at the Michigan State University (M.S.U.)
Botany and Plant Pathology farm were sampled for F usarium spp. isolates between
October 1987 and April 1988. Isolates were collected from asparagus crown and root
tissue (17 isolates), the vascular tissue at the base of asparagus ferns (97 isolates), and
soil from asparagus ﬁelds (224 isolates). All plant tissue was rinsed in 20% household
bleach for 30-60 seconds, rinsed in sterile distilled water, and placed on Komada’s media
(Komada, 1975) which is selective for F usarium spp. Cultures were obtained by diluting
the soil onto Komada’s media and incubating for 5-7 days at 25°C. Fungal cultures
exhibiting vigorous ﬂuffy mycelial growth were transferred to carnation leaf agar (CLA)
and allowed to grow for 10-14 days under ﬂuorescent plant lights. CLA plates were
ﬂooded with sterile distilled water, scraped with a scalpel to dislodge spores, and poured
into sterile 100 ml Erlenmeyer ﬂasks through three layers of sterile cheesecloth. Spore
solutions were diluted to approximately 1x105 spores per ml, and ﬁve drops of each
solution were placed on tilted water agar plates and allowed to run to the bottom. Water
agar plates were incubated for 8 to 12 hours at room temperature, then single germinated
macroconidia were picked from plates with a ﬂattened sterile dissecting needle in a
laminar ﬂow hood with the use of a dissecting microscope and transferred to CLA. After
10 to 14 days, colonies identiﬁed as F oxysporum according to the criteria of Nelson et
al.(1 983) (227 isolates) were tested for pathogenicity in a method adapted from Stephens
and Elmer (1988). Spore solutions (1x106 spores/ml) were made from CLA plates, and

0.5 ml of spore solution from each isolate was added to each of ﬁve sterile, capped, 16 x

97

150 mm test tubes containing two-week-old asparagus seedlings grown in 6.5 ml of
Hoagland’s solution agar. Asparagus seeds (cv. ‘Mary Washington’, Rupp Seeds,
Wauseon, OH) were sterilized according to the methods of Damicone et al. (1981), and
germinated on water agar plates, then aseptically placed in the culture tubes once their
radicals had emerged. Cultures were allowed to infest the seedling tubes for 28 days in a
growth chamber under ﬂuorescent lights (16 hours of light/day) at temperatures of 25°C
(day), and 20°C (night). Disease symptoms were evaluated by visually estimating the
percentage of the root system exhibiting lesions or collapse. Cultures in which each of
the ﬁve seedlings exhibited disease symptoms on less than 5% of their roots were
considered non-pathogenic, and were put into long term storage on silica gel according to
the method described by Windels et al. (1988).

Forty-one fungal cultures isolated from Michigan asparagus ﬁelds and identiﬁed
as nonpathogenic forms of the fungus F oxysporum were tested for their potential to
control Fusarium crown and root. Two additional F oxysporum isolates found to be
nonpathogenic on asparagus, but pathogenic on celery and cyclamen, respectively, were
also included in this screen. Each of the isolates was added to 8 two-week-old ‘Mary
Washington’ asparagus seedlings grown in Hoagland’s solution agar in the manner
described previously. Cultures were allowed to colonize the tubes for 7 days, then a 0.5
ml of spore solution (1x106 spores/ml) of a virulent, pathogenic isolate of F oxysporum f.
sp. asparagi isolated from a diseased asparagus crown at the MSU. Botany and Plant
Pathology F arm was applied to four tubes colonized by each isolate, and 0.5 ml of a spore

solution (1x106 spores/ml) of a virulent, pathogenic isolate of F proliferatum isolated

98

from the base of a diseased asparagus stalk from a commercial asparagus production ﬁeld
in Oceana Co., MI was applied to the other four tubes. The pathogens were allowed to
infest the plants for 28 days, after which time the seedlings were evaluated for root rot
severity by a visual estimation of the percentage of the root system exhibiting lesions or
collapse.

Five isolates were chosen from this test for further study based on their potential
to control both F oxysporum f. sp. asparagi and F proliferatum. Each had an average
root rot rating of 25% or below when challenged with each F usarium pathogen, while
most isolates allowed a high level of disease in the presence of either one or both
pathogens. These isolates, designated D-l, S-13, E-12. F-21, and 2-5 were then evaluated
in a more rigorous experiment conducted in the growth chamber. Each isolate was added
to 15 two-week-old seedlings in tubes as 0.5 ml of a 1x10" spores/m1 solution in the
manner previously described, allowed to colonize for 7 days, then challenged with the
virulent isolate of F oxysporum f. sp. asparagi. A control was included in the
experiment by adding 0.5 ml of sterile distilled water to one set of tubes, instead of a
spore solution from a non-pathogenic isolate. The experiment was arranged in a
completely randomized design, and conducted twice in a growth chamber with
ﬂuorescent lights on a 16-hour cycle, and with temperatures of 25°C (day) and 20°C
(night). The experiment was conducted in the same manner for the pathogen F.
proliferatum. Results of all experiments were evaluated by visually estimating the
percentage of the root system that was discolored or collapsed. Dead plants were

considered to be 100% infected.

99

Greenhouse experiments were performed to test the effectiveness of non-
pathogenic F oxysporum isolates in a potting mixture. Seedlings of cultivar Mary
Washington (Rupp Seeds, Wauseon, OH) were disinfested by the method previously
described and planted into 96 cc plug trays ﬁlled with Bacto High Porosity Professional
Planting Mix (Michigan Peat Co., Houston, TX). Plants were grown for three weeks,
then clean, uniform seedlings were transfered to 512 cc plastic pots ﬁlled with steam
sterilized sandy loam soil and Bacto High Porosity Professional Planting Mix in a 5:1
ratio either infested with F oxysporum f. sp. asparagi and F proliferatum, infested with
these pathogens and non-pathogenic isolates of F oxysporum, or left uninfested.

F usarium spp. were added to the soil by mixing infested ground millet seed into the
potting mixture in a cement mixer for two hours. Pathogens were added to the potting
mixture as F oxysporum f. sp. asparagi isolate 2-7 and F proliferatum isolate P-67,
both from Michigan asparagus ﬁelds at 1 g/L of soil each. Non-pathogenic F. oxysporum
isolates (D-l, F-21) were added to pathogen-infested potting mixture. Eighteen plants
were used for each treatment and pots were arranged in a completely randomized design.
The plants were fertilized twice weekly with Peters 20-20-20 fertilizer (Grace Sierra
Horticultural Products Company, Milpitas, CA) and were grown for six weeks before the
number of dead plants in each treatment was counted. The experiment was conducted
twice.

Field experiments with fungicides and biological controls. Plots were
established in Oceana County, Michigan in two non-adjacent commercial ﬁelds

previously cropped to asparagus, and having a history of Fusarium seedling blight.

100

Asparagus seeds (Jersey variety) were sown into 1.22 m beds in three rows spaced 45.72
cm apart at a rate of one seed every 3.18 cm (8.97 kg seed/ha) using a Gaspardo vacuum
planter (Gaspardo Co., Morsano, Italy). Treatment plots consisted of a 3.05 m section of
the center row on each bed. Five feet of buffer were left between each treatment within a
row, and data were taken from a 2.13 m section in the center of each plot. Treatments
were arranged in a randomized complete block design with six blocks.

Treatments were applied as drench applications made to a 22.86 cm wide section
of the seedling row (except where noted), applied immediately following planting, and
consisted of the following: i) an untreated control (water only applied), ii) the fungicide
ﬂudioxonil at a high rate (42.06 l/ha Maxim 4FS, Novartis Crop Protection Inc.,
Greensboro, NC), iii) ﬂudioxonil at a low rate (2.03 kg/ha Medallion, Novartis Crop
Protection Inc., Greensboro,NC), iv) ﬂudioxonil at a high rate (42.06 l/ha Maxim 4FS)
applied every 30 days until harvest, v) the fungicide ﬂudioxonil at a low rate (2.03 kg/ha
Medallion) applied every 30 days until harvest, vi) the biological control fungus T.
harzianum (24.64 kg/ha RootShield, BioWorks Inc., Geneva, NY), vii) the fungicide
benomyl (1.17 l/ha Benlate 50 DF , E. I. du Pont de Nemours & Co. Inc., Wilmington,
DE), viii) non-pathogenic isolates of the fungus F oxysporum (infested, ground pearl
millet seed 429.25 kg/ha) broadcast over the treatment row and worked into the soil 20
cm deep, ix) uninfested ground pearl millet seed (429.25 kg/ha) broadcast over the
treatment row and worked into the soil, x) and ﬂudioxonil (0.11 ml/kg Maxim 4FS)
applied as a seed treatment.

At the end of the season (7 September, ﬁeld 1; 15 September, ﬁeld 2), plants were

101

dug, roots and shoots weighed, plant height measured, the number of shoots per plant
counted, and the root system visually evaluated for the total root area (%) exhibiting
lesions.

Statistical Analysis. All analyses were conducted using the Mixed Model
procedures of SAS statistical analysis software (SAS Institute Inc., Cary, NC), except for
those involving the number of dead plants, which were analyzed using the Exact
procedure of LogXact statistical analysis software (Cytel Software Corporation,
Cambridge, MA). Before the analyses were carried out, data on the percentage of root
system exhibiting lesions (growth chamber and greenhouse experiments) and fresh weight
of shoots (greenhouse experiments) were transformed using the arcsine square root and
log transformations respectively (Sokal and Rohlf, 1969). Differences between
treatments were determined by using Least Squared Means comparisons with Bonferroni
adjustment factor (P<0.05). Data from the two replications of each experiment were

presented together unless signiﬁcant experiment by treatment interactions were found.

RESULTS
Greenhouse experiments with fungicides and a biological control product. A
high level of pathogen inoculum resulted in plant death in all treatments except for the
uninfested control (no treatment) (Figure 10). By week ﬁve, all plants were dead in both
the infested control and T. harzianum biological control treatments, signiﬁcantly more
than in any other treatment. An average of two plants (20%) died over the ﬁve weeks in

the ﬂudioxonil treatment which was comparable to uninfested control and 2-(4-

102

 

+ infested control

--O-- Tn'choderma harzianum
+ benomyl

12 _ -v-- 2-(4 Thiazolyl) benzimidazole
+ fludioxonil

 

—I - - uninfested control

 

 

 

 

Average number dead plants

 

 

 

Weeks after treatment

Figure 10. Average number of asparagus plants killed in two
greenhouse experimemts when grown in soil infested with a
high level of F. proliferatum and F. oxysporum f. sp. asparagi
and treated with a chemical or biological control product.
Letter notation indicates signiﬁcant differences between
treatments.

103

Thiaziolyl)benzimidazole treatments. An average of 6.5 (65%) plants were dead after
ﬁve weeks with the benomyl treatment, signiﬁcantly more than either the ﬂudioxonil
treatment or the uninfested control.

When the lower pathogen inoculum level was used, all plants survived, regardless
of treatment. Fresh shoot weights for the uninfested control varied between the two low
inoculum experiments, resulting in a signiﬁcant experiment by treatment interaction. In
the ﬁrst experiment, the only signiﬁcant difference observed was in the infested control,
which exhibited lower mean shoot weights than the uninfested control (Figure 11). Fresh
shoot weight of the treatments did not differ signiﬁcantly from either control. In the
second low inoculum experiment, the shoot weight of the uninfested control was
signiﬁcantly greater than that of the ﬂudioxonil treatment and the infested control.
Signiﬁcant differences among other treatments were not noted. Because the analysis of
root weight and root system exhibiting lesions (%) showed no signiﬁcant experiment by
treatment interaction, the data for each of these parameters were combined. Root weight
was signiﬁcantly greater for the uninfested control than for any other treatment or the
infested control (Figure 12). Root weights for all treatments were signiﬁcantly greater
than the infested control, but were not signiﬁcantly different from each other. The
uninfested control plants had a lower percentage of root system with lesions than any
other treatment (Figure 13). All treatments resulted in reduction of root lesions compared
to the infested control, but did not differ signiﬁcantly from one another.

Growth chamber and greenhouse studies with non-pathogenic forms of F

oxysporum. All ﬁve of the selected non-pathogenic F oxysporum isolates signiﬁcantly

104

 

1.4 - _
Experiment 1
1.2 4
1.0 -
0.8 -
0.6 —

0.4 ~

0.2 4

 

 

 

0.0 -

 

1.4 ~ 3
Experiment 2

 

Shoot weight (g)

1.2 -
1.0 4
0.8 4
0.6 4
0.4 -

0.2 ~

 

 

   

0.0 -

unlnfested
control
control

Trlchoderma
harzianum
benomyl
ﬂudloxonil

Infested

Figure 11. Shoot weight of asparagus seedlings from two
greenhouse experiments when grown in soil infested with
a low level of F. proliferatum and F. oxysporum f. sp.
asparagi and treated with a chemical or biological control
product. Error bars represent standard error, and letter
notation represents signiﬁcant differences in L. S. means
comparisons.

105

 

Root weight (g)

 

 

 

'8- 3.
i9 5
'25 3
Eu 3
3

fludloxonll
Trlchoderme
harzianum
Infested
control

Figure 12. Root weight of asparagus seedlings from
two greenhouse experiments when grown in soil infested
with a low level of F. proliferatum and F. oxysporum f.
sp. asparagi and treated with a chemical or biological
control product. Error bars represent standard error,

and letter notation indicates signiﬁcant differences in L. 8.
means comparisons.

106

Root system exhibiting lesions (%)

30

 

 

 

 

benomyl
Infested
control

8
“3
as
£0
:0
3

fludioxonll
Trlchoderma
harzianum

Figure 13. Roots of asparagus seedlings exhibiting
lesions (%) in two greenhouse experiments when
grown in soil infested with a low level of F. proliferatum
and F. oxysporum f. sp. asparagi and treated with a
chemical or biological control product. Error bars
represent standard error, and letter notation indicates
signiﬁcant differences in L. S. means comparisons.

107

reduced root disease compared to the infested control in two growth chamber experiments
with F oxysporum f. sp. asparagi (Figure 14). Isolate D-l was signiﬁcantly more
effective at reducing root lesions than all other isolates in experiment 1, and signiﬁcantly
more effective than all isolates except for S-l3 in experiment 2. Isolate S-13 was
signiﬁcantly more effective than isolates 2-5 and E-12 in experiment 2 only.

No signiﬁcant treatment by experiment interaction occurred in trials with the
pathogen F proliferatum, thus the data from these experiments were combined (Figure
15). All isolates showed signiﬁcant disease reduction compared with the infested control,
although isolate D-l was signiﬁcantly less effective than the other isolates. Isolate F-21
showed the lowest mean disease rating of all the isolates.

In both greenhouse studies no signiﬁcant differences in plant death were found
between asparagus plants growing in media infested with F oxysporum f. sp. asparagi
and F proliferatum and left untreated, and those infested with pathogenic F usarium spp.
as well as non-pathogenic isolates. In experiment 1, 9 plants (50%) were killed in both
the infested control and the non-pathogen treatment, and in experiment 2. 12 plants
(66.7%) were killed in the infested control, while 13 (72.2%) were killed in the non-
pathogen treatment. No plants were killed in the uninfested control in either experiment.

Field trials using fungicides and biological controls. Although the trials were
conducted in seedling ﬁelds considered to be highly infested with pathogenic F usarium
spp., mean root lesions did not exceed 3% in the untreated control or the treatment plots
in either ﬁeld. Root infection (%) was not signiﬁcantly different among treatments, and

no consistent trends among the measured parameters were observed between the two

108

 

100
90 — 3 Experiment 1
80 -
70 ~
60 J
50 -
40 ~
30 -
20 -

10~

 

 

 

Water 2-5 S-13 E-12 F-21 D-1

 

100
90 g a Experiment 2
80 a

70‘

Root system exhibiting lesions (%)

60‘

50-

40-

30~

204

10~

 

 

 

Water 2-5 E-12 F-21 S-13 D-1

Figure 14. Root system of asparagus seedlings exhibiting
lesions (%) in two growth chamber experiments when
grown in culture tubes infested with F. oxysporum f. sp.
asparagi and treated with non-pathogenic F. oxysporum.
Error bars represent standard error, and different letter
notation indicates signiﬁcant differences in L. S. Means
comparisons.

109

 

 

 

Root system exhibiting lesions (%)

 

Water D-1 S-13 E-12 2-5 F-21

Figure 15. Root system of asparagus seedlings exhibiting
lesions (%) in two growth chamber experiments when
grown in culture tubes infested with F. proliferatum

and treated with non-pathogenic F. oxysporum. Error
bars represent standard error, and different letter

notation indicates signiﬁcant differences in L. 8. means
comparisons.

110

replicate ﬁelds (see appendix).

DISCUSSION

F usarium spp. pathogenic to asparagus are found in all Michigan soils, and
Fusarium decline has become a serious production limitation as pathogen populations
have built up in ﬁelds previously planted in the crop (Elmer et al., 1996; Hartung et al.,
1990). Our goal to develop an effective management strategy utilizing a commercially
available biological control product (T. harzianum), non-pathogenic isolates of F
oxysporum, and fungicides has met with limited success.

Non-pathogenic F oxysporum isolates were effective in limiting Fusarium disease
on asparagus seedlings grown in culture tubes, although isolates differed in their ability to
control disease caused by F proliferatum and F oxysporum f. sp. asparagi. Isolate D-l
was the most effective in controlling disease caused by F oxysporum f. sp. asparagi, but
was the least effective isolate when tested against F proliferatum. Vegetative
compatibility testing has shown that F oxysporum f. sp. asparagi is a diverse group. In
one experiment, 26 strains of this pathogen were placed into 8 vegtative compatibility
groups, while 34 strains appeared to belong to unique compatibility groups (Elmer and
Stephens, 1989). This kind of diversity is rare among pathogenic F usarium spp., and
non-pathogenic isolates may have varied activity against different isolates in a single
pathogenic formae.

When non-pathogenic isolates which effectively limited both F proliferatum and

F oxysporum f. sp. asparagi in culture tube tests were mixed, and tested in greenhouse

lll

studies with asparagus seedlings grown in soil, they were not effective in controlling
disease. It is likely that the high level of pathogen inoculum (180,000 cfu/ g soil) used in
this experiment overwhelmed and out-competed the non-pathogens. Hartung et. al.
(1990) found that levels of Fusarium in asparagus ﬁeld soils in Michigan were generally
much lower than this (between 0 and 60,000 cfu of F usarium spp. per gram of sieved
soil). Furthermore, Hartung et. al.’s (1990) numbers represent both pathogenic, and non-
pathogenic F usarium spp. found in soil, although F. proliferatum isolates were likely
underestimated because they are found more consistently on plant residue than in soil. It
is also possible that because the non-pathogenic isolates did not have a week to colonize
the asparagus seedlings, as in the grth chamber tubes, they could not provide adequate
protection. The added complexity of the soil system, or the method of inoculation could
have accounted for the failure of these isolates to control disease in the greenhouse
system as well.

Additional greenhouse studies with asparagus seedlings using fungicides and a
biological control product indicated that T. harzianum, benomyl, and ﬂudioxonil
increased root weight and decreased root disease compared to the infested control when a
low level of F proliferatum and F oxysporum f. sp. asparagi was used. Although shoot
weights did not differ signiﬁcantly among these treatments, the trends were similar to
those for the root data. The fungicide ﬂudioxonil limited disease caused by F usarium
spp. at both low and high inoculum levels. T. harzianum, however, was not effective
when a high rate of inoculum was used, indicating that the effectiveness of this product

may be dependent on the organism not being overwhelmed by the pathogen. Benomyl is

112

not currently labeled for use on asparagus in Michigan, and experiments by Lacy (1979)
indicated that it was not effective in reducing disease when applied as a dip to one-year-
old crowns from seedling nurseries.

Our attempt to test fungicides and biological control products under commercial
ﬁeld conditions was not successful. The low level of seedling infection in the ﬁelds
where the treatments were tested may be attributed to environmental conditions, and may
have been responsible for the absence of signiﬁcant or consistent differences between the
treatments. Previous research has suggested that dry conditions favor disease
development in seedlings (Graham, 1955; Elmer et al., 1997). In addition, Michigan
growers have observed that the critical time for seedling infection by F usarium is during
establishment (Jon Bakker, Director, Oceana County Asparagus Research Farm, Personal
Communication). Rainfall during the early part of the season was 13.35 cm (27 April
through 12 June), and total rainfall was 41.28 cm (27 April through 6 October).

Biocontrol activity of non-pathogenic F oxysporum isolates has been recognized
by others (Blok et al., 1997; Damicone and Manning, 1982; Tu et al., 1990). However,
only Damicone and Manning (1982) were successful in demonstrating disease control in
the ﬁeld using uninfested, greenhouse grown seedlings dipped in 0.4% methyl cellulose
containing non-pathogenic F oxysporum spores (4-6x10° spores/m1). Both Blok et. al.
and Tu et. al. failed to ﬁnd disease suppression in ﬁelds amended with soil containing
non-pathogenic F usarium spp., and an effective disease control product has not been
developed for the ﬁeld. Since studies by other researchers (Di Lenna and Foletto, 1990;
Lacy, 1979) have suggested that F usarium infection in seedling nurseries may be the
most important factor in predisposing asparagus plantings to decline, and have not

113

demonstrated disease control with fungicides when crowns were already infected, this
may be the best place to initiate root colonization by non-pathogenic species.

Future investigations on the use of non-pathogenic F oxysporum should consider
the optimal stage of plant development for non-pathogen colonization, different methods
for infesting plants with non-pathogenic isolates, and the level of inoculum needed to
obtain effective disease control. Using a mixture of avirulent strains may provide
enhanced disease suppression against a range of pathogenic F usarium isolates, and using
native isolates may insure that inoculum is able to survive and proliferate in the
conditions under which it may be used. New chemical and biological control products
should be tested further for their effectiveness against F usarium spp. pathogenic to
asparagus, and integrated strategies for disease control, such as those developed for other
F usarium pathogens (Beale and Pitt, 1990; Chakravarty et al., 1990; Sivan and Chet,

1993) may be explored as well.

114

 

LITERATURE CITED
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in relation to border cell production and Fusarium root rot of asparagus. M.S.
Thesis-Michigan State University, E. Lansing, 68 pp.
Beale, R. E., and Pitt, D. 1990. Biological and integrated control of F usarium
basal rot of Narcissus using Minimedusa polyspora and other micro-organisms.
Plant Pathology 392477-488.
Blok, W. J ., Zwankhuizen, M. J ., and Bollen G. J. 1997. Biological control of
F usarium oxysporum f. sp. asparagi by applying non-pathogenic isolates of F
oxysporum. Biocontrol Science and Technology 7:527-541.
Cassini, R. C., E1 Medawar, S., and Cassini, R. P. 1985. A biological control
technique to prevent F usariurn decline in the ﬁelds. Pages 228-234 in:
Proceedings of the Sixth lntemational Asparagus Symposium, Guelph, Ontario, E.
C. Lougheed and H. Tiessen eds.
Chakravarty, P., Peterson R. L., and Ellis B. E. 1990. Integrated control of
Fusarium damping-off in red pine seedlings with the ectomycorrhizal fungus
Paxillus involutus and fungicides. Canadian Journal of Forestry Research.
20:1283-1288.
Damicone, J .P., Cooley, D. R., and Manning, W. J. 1981. Benomyl in acetone
eradicates F usarium moniliforme and F usarium oxysporum from asparagus seed.
Plant Disease 65: 892-893.

Damicone, J .P., and Manning, W. J. 1982. Avirulent strains of F usarium

115

oxysporum protect asparagus seedlings from crown rot. Canadian Journal of Plant
Pathology 42143-146.

Davis, D. 1967. Cross-protection in F usarium wilt diseases. Phytopathology.
57:311-314.

Di Lenna, P., and Foletto, B. 1990. Effect of nursery management on subsequent
F usarium decline of asparagus in ﬁeld. Acta Horticulturae 271:299-303.

Elmer, W. H. 1989. Effects of chloride and nitrogen form on growth of
asparagus infected by F usarium spp. Plant Dis. 73: 736-740.

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.

Elmer, W. H., and LaMondia, J. A. 1997. Studies on the supression of Fusarium
crown and root rot of asparagus with NaCl. Pages 54-67 in: The IX international
Asparagus Symposium, Pasco, WA, B. Benson ed.

Elmer, W. H., LaMondia, J. A., and Taylor, G. S. 1997. Asparagus cultivar trials
in Connecticut. Pages 420-426 in: The IX international Asparagus Symposium,
Pasco, WA, B. Benson ed.

Elmer, W. H., and Stephens, C. T. 1989. Classiﬁcation of F usarium oxysporum

f. sp. asparagi into vegetatively compatible groups. Phytopathology 79:88-93.
Faloon, P. J ., and F auser-Kevem, H. A. 1996. Effect of thiabendazole (Tecto
20S) and metalaxyl (Ridomil MZ72) on asparagus establishment in replant soil.

Acta Horticulturae 415: 289-295.

116

Fantino, M. G. 1990. Research on asparagus decline in Italy. Acta Horticulturae
271:291-298.

Graham, K. M. 1955. Seedling blight, a Fusarial disease of asparagus. Canadian
Journal of Botany 33:374-404.

Grogan, R. G., and Kimble, K. A. 1959. The association of Fusarium wilt with
the asparagus decline and replant problem in California. Phytopathology 49:122-
125.

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.
Hervas, A., Trapero-Casas, J. L., and Jimenez-Diaz, R. M. 1995. Induced
resistance against Fusarium wilt of chickpea by nonpathogenic races of F usarium
oxysporum f. sp. ciceris and nonpathogenic isolates of F oxysporum. Plant
Disease 79:1110-1116.

Komada, H. 1975. Development of a selective medium for quantitative isolation
of F usarium oxysporum from natural soil. Rev. Plant Prot. Res. 8:1 14-124.

Lacy, M. L. 1979. Effects of chemicals on stand establishment and yields of
asparagus. Plant Disease Reporter 63:612-616.

Larkin, R. R, Hopkins D. L., and Martin, F. N. 1996. Suppression of Fusarium
wilt of watermelon by nonpathogenic F usarium oxysporum and other
microorganisms recovered from disease-suppressive soil. Phytopathology 86:812-
819.

Mandeel, Q., and Baker, R. 1991. Mechanisms involved in biological control of

117

Fusarium wilt of cucumber with strains of nonpathogenic F usarium oxysporum.
Phytopathology 81: 462-469.

Marois. J. J. 1993 Biological control of diseases caused by F usarium oxysporum.
Pages 77-81 in: F usarium Wilt of Banana. R. C. Ploetz ed., APS Press, St. Paul.
Nelson, P. E., Toussoun, T. A., and Marsas, W. F. O. 1983. F usarium Species:
An Illustrated Manual For Identiﬁcation. The Pennsylvania Stare University Press,
University Park.

Oyarzun, P. J ., Postma, J ., Luttikholt, A.J.G., and Hoogland, A. E. 1994.
Biological control of foot and root rot in pea caused by F uasarium solam' with
nonpathogenic F usarium oxysporum isolates. Canadian Journal of Botany 72:843-
852.

Sivan, A., and Chet, l. 1993. Integrated control of Fusarium crown and root rot
of tomato with Trichoderma harzianum in combination with methyl bromide or
soil solarization. Crop Protection 12:380-386.

Sokal, R. R., and Rohlf, J. F. 1969. Biomerty: The Principles and Practice of
Statistics in Biological Research. W. H. Freeman and Company, San Francisco.
Stephens, C. T., De Vries, R. M., and Sink, KC. 1989. Evaluation of Asparagus
species for resistance to F usarium oxysporum f. sp. asparagi and F moniliforme.
HortScience 24: 365-368.

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resistance in Asparagus spp. to Fusarium crown and root rot. Plant Disease

72:334-337.

118

Tu. C. C.. Cheng, Y. H., and Cheng, A. S. 1990. Recent advance in biological
control of Fusarium wilt of asparagus in Taiwan. Acta Horticulturae 271: 353-
357.

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asparagus (Asparagus ofﬁcinalis L.) agroecosystem. M. S. Thesis-Michigan State
University. 101 pp.

Windels, C. E., Burnes, P. M., and Kommedahl, T. 1988. Five-year preservation

of F usarium species on silica gel and soil. Phytopathology 78:107-109.

119

APPENDIX

SECTION I

120

Treatment

 

 

 

Measured 560 kg 1120 kg
Parameter Year Control 560 kg 1120 kg Lime & Lime & Lime
pH Top 1998 5.59 5.66 5.53 5.75 5.78 5.69

2000 6.28 abc 6.19 a 6.27 ab 6.55 c 6.58 be 6.51 bc
pH Bottom 1998 5.54 5.68 5.58 5.58 5.57 5.53
2000 5.87 a 6.00 a 6.1 lab 6.12 ab 6.16 ab 6.26 b
Potassium 1998 258 268 264 272 245 264
Top
2000 375 ns 374 ns 384 ns 343 ns 373 us 391 ns
Potassium 1998 223 229 233 232 254 233
Bottom
2000 301 ns 306 ns 315 ns 290 us 298 ns 293 ns
Calcium 1998 723 843 715 782 817 800
Top
2000 799 ab 755 a 826 ab 914 ab 970 b 979 b
Calcium 1998 71 l 798 744 730 671 71 1
Bottom
2000 587 ns 639 ns 641 ns 631 us 601 us 750 ns
Magnesium 1998 182 193 166 216 162 208
Top
(Healthy) 2000 239 us 196 us 213 ns 228 us 245 ns 234 ns
Magnesium 1998 52 60 50 65 69 66
Top
(Declined) 2000 80 a 93 ab 85 ab 168 b 162 b 151 ab
Magnesium 1998 78 93 83 89 85 92
Bottom
2000 l27ab 116a 109 ab 154b 141 ab 134 ab
Sodium 1998 3.8 4.8 5.8 8.8 7.5 7.8
Top
2000 26.9 ns 29.0 ns 35.7 ns 26.9 ns 27.4 ns 37.7 ns
Sodium 1998 9.1 9.0 6.6 3.7 6.0 6.3
Bottom
2000 24.1 a 35.7 b 51.4 c 23.8 a 29.3 a 55.6 c

Table 1. Mean data for soil samples from 4/98 (before any treatments were
applied) and 5/00 (after two years of treatment) in Oceana County ﬁeld
plots, tables show combined data for both ﬁelds unless signiﬁcant ﬁeld by
treatment effects were found, letter notation indicates signiﬁcant differences
between treatment means from 2000 with 1998 data used as a covariate.

121

 

NaCl No NaCl NaCl N0 NaCl

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

Year AppLeg AM Applied Appﬂeg
M pH Bottom

1998 5.38 5.34 5.74 5.62

2000 5.98 ns 5.72 ns 6.20 ns 5.84 ns
w No NaCl NaCl No NaCl

Year App_li_eg Appli_ed Applied Appﬂg

Potassium Top Potassium Bottom

1998 175 202 95 91

2000 230 ns 242 us 110 ns 224 ns
NaCl No NaCl NaCl No NaCl

Year Applje_d_ 5pm Applied M

Calcium Top Calcium Bottom

1998 2618 2455 2800 2609

2000 2609 ns 2590 us 2819 ns 2724 ns
Egg No NaCl NaCl No NaCl

Year Appljg AM Applied Appﬂeg

 

 

Magnesium Top

Magnesium Bottom

 

 

 

 

 

1998 250 220 254 207
2000 249 ns 290 ns 240 us 269 ns
NaCl No NaCl NaCl No NaCl
Year Appm Appﬁgg Applied mg
Sodium Top SthLn Botttmr
1998 10.2 10.0 6.0 5.8
2000 90.0 b 21.0 a 94.6 b 26.6 a

 

 

Table 2. Mean data for soil samples ﬁ'om 4/98 (before any treatments were
applied) and 5/00 (after two years of treatment) at MSU Botany and Plant
Pathology Farm, letter notation indicates signiﬁcant differences between
treatment means from 2000 with 1998 data used as a covariate for top and
bottom data separately.

122

Average sodium level in soil samples (ppm)

 

No NaCl

 

 

 

 

I

4198 4\ 7198 1 0l98 5/99/I\

 

80‘

O
O
I

.5
O
1

No NaCl

”ﬂ. ................. .. ................. .

N
O
r

 

 

 

 

o 'J r r r f
4I98 ¢ 7I98 10I98 5I99/I\

1 0I99

5/00

Figure 16. Mean sodium level in soil samples from MSU
Botany and Plant Pathology Farm ﬁeld 4/98-5/00 in parts
per million, arrows indicate treatment application times.

123

Na Levels in 0-15.2 cm of soil

Na Levels in 15.2-30.5 cm of Soil

9

140 -

 

 

 

 

 

 

 

 

 

 

 

 

f + Control
12° 1 /‘ \ + 500 lbs
, - —v-- 1000 lbs
100 - / .V\ \ + 500+L
. / a \ -I~- 1000+L
30 — . " \._-
E .
Q 60
Q
v
U) 40
2
Q- 20 J
E
8 0 1
'6
‘9 «use/F
IE
3 80 a
>
0
_l
m 60 ~
2
d}
m .1
e 40
0
>
<
20 -
0 4
4I98 /I\ 7I98 1 0I98 599/? 10I99

 

5l00

Figure 17. Mean sodium level in soil samples from healthy
ﬁeld in Oceana County between 4/98 and 5/00 in parts per
million, arrows indicate treatment application times.

124

Na Levels in 0-15.2 cm of Soil

——> Na Levels in 15.2-30.5 cm of Soil

 

140 ~

 

 

 

 

 

 

 

 

 

 

  

 

 

+ Control
. Lime =
120 1 f\ + 500 lbs 0
' - —v~- 1000 lbs {’3
100 - / V\ \ + 500+L o
// .. \ -I-- 1000+L E
A 80 4 ' .- \_ ' ,i,’
E 1‘.‘
a 60 :3
V .E
m 40 vi
a s
E 20 - 3
n: a
U) o - 2
g I T I I ‘1
C 4198 /I\ 7I98 10I98 5I99/I\ 10l99 5I00/I\
7, 70 -
r. =
.J 50 - (I)
10 . '6
Q)
E ..-——..——- 3.
20 - 5
1’
o
10 - z
.1
............. a
0 ~ 2
4/98 /1\ 7I98 10l98 5I99 I 10/99 5/00 /I\

Figure 18. Mean sodium level in soil samples from declined
ﬁeld in Oceana County between 4/98 and 5/00 in parts per

million, arrows

indicate treatment application times.

125

APPENDIX

SECTION II

126

Height of fern (Cm)

4O

30

20

30

20

 

 

 

 

: Field#1]
T c
_ T
l l
l T Field#2
- 4 r T
: T
L
l
t
l I l l l I
E E .3 a 2 a
E E i .E o B o o E w
o s 1: g E g g 5 E
0 g E S .c tn 3 'i
s -‘=’ s g
'— 2

Figure 19. Mean fern height in two seedling nursery trials
with chemical and biological controls. Error bars represent
standard error.

127

 

Field #1

 

ire

E L ‘Fl hi pLIrLIHIFlflkLl l

3

2

1

Field #2

 

06 5 4. 3 2 1

EaE 5n. «.095 we .mnEaz

l uwme_xms_
7
l «2:5.

‘ cmmoﬁancoz

. _>Eo:mm
,
V

l «stowage;
1 85:38.2
- cn—Exms.

,l :o=_wuws_

l .5me

l 85:00

LL LLLlFl LLL i, Flilclr Tr rlLLLL LF‘LLl ﬂ

0

trials with chemical and biological controls. Error bars represent

Figure 20. Number of shoots per plant in two seedling nursery
standard error.

128

Weight of shoots (g)

 

Field #1

  
    

i V‘ﬁ‘ﬁ—I‘Y‘I‘T’T‘I—I—fI‘I I VTTTW‘V 1*

 

 

Field #2

I i I 1 I I i I I I I

 
 

l I l I

 

 

Maxim
Millet

'3
I-
u
C
O
U

Medallion
Maxlm30
Medallion30
Trlchoderma
Benomyl
Nonpathogen
MaxlmSeed

Figure 21. Fresh weight of shoots in two seedling nursery
trials with chemical and biological controls. Error bars
represent standard error.

l
129

Weight of roots (9)

 

Field #1

 

 

 

 

 

8
- Field#2
_ i
6 T - l I i
i I
- T
4 i
2}
0

 

 

Medallion
Maxlm30
Benomyl .

MaximSeed

l
o
n
c
.2
E
1:
o
E

Trlchoderma »
Nonpathogen

Figure 22. Fresh weight of roots in two seedling nursery
trials with cheimcal and biological controls. Error bars
represent standard error.

130

Root system exhibiting lesions (%)

 

 

 

i Field #1 }

 

 

 

1’ Field #2

 

Maxhn
Benomyl!
Mﬂwte

“E
h
u
c
O
U

MedaMonv
MedaMon30
Tﬂchodenna
Nonpathogen

MaxhnSeed—

Figure 23. Percentage of root system exhibiting lesions
in two seedling nursery trials with chemical and biological
controls. Error bars represent standard error.

131