INVESTIGATING BIOLOGICAL INTERACTIONS AND SOURCES OF RESISTANCE TO PLANT PARASITIC NEMATODES IN A GRAFTED WATERMELON PRODUCTION SYSTEM
By
CODY L. SMITH
A THESIS PRESENTED TO THE GRADUATE SCHOOL OF THE UNIVERSITY OF FLORIDA IN PARTIAL FULFILLMENT OF THE REQUIREMENTS FOR THE DEGREE OF MASTER OF SCIENCE
UNIVERSITY OF FLORIDA
2017
© 2017 Cody L. Smith
To my parents and step-parents, without your love and support none of this would have been possible
ACKNOWLEDGMENTS
I’d first like to thank God for all that you have brought me through in this life. I’ve had more than my share of bumps in the road and without you, and the love and support of my family and friends I would still be in a much darker place.
I would also like to thank my advisor, Dr. Joshua Freeman for his patience with answering all my dumb questions and giving this kid from the middle of nowhere a shot at something better. In addition to Dr. Freeman I would like to thank my advisory committee, Dr. Nancy Burelle, Dr. Peter Dittmar, and Dr. Mathews Paret for their training and assistance with my research project.
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TABLE OF CONTENTS
page
ACKNOWLEDGMENTS ...... 4
LIST OF TABLES ...... 7
LIST OF FIGURES ...... 8
LIST OF ABBREVIATIONS ...... 9
ABSTRACT ...... 10
CHAPTER
1 INTRODUCTION ...... 11
Watermelon ...... 11 Significance and Value ...... 11 Florida Watermelon ...... 11 Methyl Bromide ...... 12 Efficacy ...... 12 Phaseout ...... 13 Grafting ...... 13 Background Information ...... 13 Positives ...... 14 Negatives ...... 16 Nematodes in Watermelon Production ...... 17 Root-knot Nematodes (Meloidogyne spp.) ...... 17 The Reniform Nematode (Rotylenchulus reniformis) ...... 18 Nematode Movement and Soil Depth ...... 20 Symptomology and Diagnosis ...... 22 Management ...... 24 Fusarium Wilt ...... 25 Biology and Background Information ...... 25 Fusarium Wilt of Watermelon ...... 26 Symptomology and Diagnosis ...... 27 Management ...... 29 Nematode/Pathogen Interactions and Disease Complexes ...... 30 Nematode interactions with Pathogens ...... 30 Nematode interactions with Fusarium ...... 30 Nematode interactions with other Nematodes ...... 31
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2 SCREENING CUCURBIT ROOTSTOCKS FOR VARIETAL RESISTANCE TO MELOIDOGYNE SPP. AND ROTYLENCHULUS RENIFORMIS ...... 34
Introduction ...... 34 Materials and Methods...... 37 Results ...... 39 Discussion ...... 40
3 ASSESSMENT OF WATERMELON ROOTSTOCK RESISTANCE TO FUSARIUM OXYSPORUM F.SP. NIVEUM WHEN EXPOSED TO MELOIDOGYNE INCOGNITA ...... 47
Introduction ...... 47 Materials and Methods...... 48 Results ...... 50 Discussion ...... 51
4 TRACKING SEASONAL MOVEMENT OF MELOIDOGYNE SPP. AND ROTYLENCHULUS RENIFORMIS THROUGH A NINETY DAY WATERMELON CROPPING SEASON ...... 61
Introduction ...... 61 Materials and Methods...... 63 Results ...... 64 Discussion ...... 65
5 CONCLUSION ...... 71
LIST OF REFERENCES ...... 73
BIOGRAPHICAL SKETCH ...... 86
6
LIST OF TABLES
Table page
3-1 2016 Fusarium Wilt Incidence...... 57
3-2 2016 Fusarium Wilt Severity ...... 57
3-3 2017 Fusarium Wilt Incidence ...... 60
3-4 2017 Fusarium Wilt Severity ...... 60
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LIST OF FIGURES
Figure page
2-1 60 DAP Root Gall Index ...... 44
2-2 90 DAP Root Gall Index ...... 44
2-3 90 DAP Meloidogyne spp. Soil Populations...... 45
2-4 90 DAP Meloidogyne spp. Root Tissue Populations ...... 45
2-5 90 DAP R. reniformis Soil Populations...... 46
2-6 90 DAP R. reniformis Root Tissue Populations ...... 46
3-1 90 DAP Root Gall Index ...... 55
3-2 2016 Total Daily Rainfall ...... 56
3-3 2016 Soil and Air Temperature Averages ...... 56
3-4 2017 Yield by FON Inoculation ...... 58
3-5 2017 Yield by Variety ...... 58
3-6 2017 Total Daily Rainfall ...... 59
3-7 2017 Soil and Air Temperature Averages ...... 59
4-1 Meloidogyne spp. Pre-plant Soil Populations ...... 68
4-2 R. reniformis Pre-plant Soil Populations ...... 68
4-3 Meloidogyne spp. Soil Populations 45 Days after Planting ...... 69
4-4 R. reniformis Soil Populations 45 Days after Planting ...... 69
4-5 Meloidogyne spp. Soil Populations 90 Days after Planting ...... 70
4-6 R. reniformis Soil Populations 90 Days after Planting ...... 70
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LIST OF ABBREVIATIONS
DAP Days after planting
FON Fusarium oxysporum f.sp. niveum
FW Fusarium wilt
MeBr Methyl bromide
MI Meloidogyne incognita
PPN Plant parasitic nematode
Pi Initial population
REN Rotylenchulus reniformis
Rf Reproductive factor
RKN Meloidogyne spp.
SAR Systemic acquired resistance
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Abstract of Thesis Presented to the Graduate School of the University of Florida in Partial Fulfillment of the Requirements for the Degree of Master of Science
INVESTIGATING BIOLOGICAL INTERACTIONS AND SOURCES OF RESISTANCE TO PLANT PARASITIC NEMATODES IN A GRAFTED WATERMELON PRODUCTION SYSTEM
By
Cody L. Smith
December 2017
Chair: Joshua H. Freeman Major: Horticultural Sciences
Incidence of the soil-borne plant pathogen Fusarium oxysporum f.sp niveum
(FON), and Meloidogyne spp. (RKN’s) in watermelon production systems is rising, due in part to the phase out of methyl bromide and reduced crop rotation. One method of management involves grafting a susceptible scion on a FON or RKN resistant rootstock.
A critical issue with some FON resistant rootstocks is lack of testing for RKN susceptibility. Both pathogens occur in similar locations and can interact synergistically.
Research has shown that many FON resistant rootstocks are highly susceptible to RKN damage. Studies have demonstrated that RKN’s have the ability to break host FON resistance, rendering these rootstocks non-viable. In addition to RKN’s, reniform nematode (Rotylenchulus reniformis) was observed on these FON resistant rootstocks.
Watermelons, are generally considered to be a non-host of R. reniformis. At the NFREC in Quincy, FL., the opposite was observed. The objectives of my research were to: (1) screen FON resistant rootstocks for resistance to RKN’s and R. reniformis, (2) assess
FON resistant rootstock resistance to FON when subjected to RKN’s, and (3) track migration of RKN’s and R. reniformis in the soil profile during a 90 day cropping season.
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CHAPTER 1 INTRODUCTION
Watermelon
Significance and Value
Cultivated watermelon (Citrullus lanatus) accounts for 46% of the world’s entire cucurbit production (MAGRAMA, 2013). The largest watermelon producer in the world is
China harvesting 75,054,330 metric tons which represents approximately 67% of global production. Globally, the United States ranks 8th in world production with 1,505,616
metric tons harvested in 2014 (FAOSTAT, 2014). Within the United States, Florida
ranks 1st in watermelon production, with 389,095,164 kg, followed closely by Texas
(368,317,004 kg) and California (342,915,832 kg) in 2016 (USDA NASS, 2017).
Average yields in Florida were 43,310 kg/ha with average values of $5,555/ha and
$0.13/kg in 2016 (USDA NASS, 2016). The market trend for watermelons has shifted
considerably in a twelve-year period from large seeded diploid fruit to smaller seedless
triploid fruit. Total U.S. market share for seedless watermelons increased from 51% in
2003 to 85% in 2014 (Naeve, 2015).
Florida Watermelon
Citrullus lanatus is considered a very tolerant crop on acidic soils with an
acceptable pH range of 5.0-6.8. The optimum yield to applied macronutrients for
watermelon occurs at a rate of 168 kg/ha nitrogen (N), 168 kg/ha phosphorus (P2O5),
and 135-168 kg/ha potassium (K2O) which can be applied both pre and post-planting
(Hochmuth and Hanlon, 2013). Northern regions of the state produce watermelons
almost exclusively in the spring due to late season cold temperatures, increased
whitefly pressure and high severity of gummy stem blight in fall planting (Newark et al.,
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2011; Webb et al., 2013). Weed management incorporates fumigants, pre and post-
emergence herbicides applied prior to planting or in row middles before vine overlap
(Dittmar and Stall, 2003). Disease management utilizes a rotation of fungicides and
bactericides with different modes of action to combat resistance. Fungicidal sprays are
best started one week post-transplant and continued on a 7-10 day interval or 5-7 day
interval during periods of heavy rainfall (Dufault and Paret, 2015). In north Florida
watermelons are planted between the months of February and April typically on raised
beds covered with black polyethylene mulch. Harvest for transplanted crops occurs
between April and July while direct seeded crops are harvested between May and
August depending on initial planting date and cultivar selected (Freeman et al., 2016).
Methyl Bromide
Efficacy
Plasticulture production systems once relied on the soil fumigant methyl bromide
(MeBr) for control of weeds, soil-borne pests and pathogens (USDA ERS, 2000). Methyl
bromide possessed a specific gravity and high vapor pressure that provided
unparalleled control and exceptional movement both laterally and vertically in the soil.
Downward movement was extensive and significant gas levels were discovered at a
depth of 9 feet (~274 cm) in the soil profile. In addition to superior soil movement, high
vapor pressure of MeBr led to rapid dissipation which allowed producers to fumigate
and plant crops in a matter of days (Kolbezen et al., 1974; Munnecke and Van Gundy,
1979). Despite its wide-scale usage over a roughly 40 year period no resistance in
soilborne pests or pathogens was documented and efficacy remained (Klein, 1996).
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Phaseout
In 1991 MeBr was found to be a causal agent in the degradation of the ozone
layer and motions were made to ban its use. Multi-national governing bodies assembled
to form the Montreal Protocol and MeBr was phased out in 2005, marking the last year
a producer could apply it without a critical use exemption (CUE) (US EPA, 2017). As of
2016, the only applicants who received CUE’s are strawberry growers in California and
U.S. cured pork producers (US EPA, 2015). The ban on methyl bromide was estimated
to have a significant economic impact, costing consumers and producers at least $1
billion annually (Ferguson and Padula, 1994). Alternatives to MeBr have lower vapor
pressure, water affinity and poorer soil distribution both vertically and laterally when
compared to the MeBr (Candole et al., 2007; Desaeger et al., 2004; Munnecke and Van
Gundy, 1979; Noling and Becker, 1994). In the absence of methyl bromide, producers
and researchers alike were pushed to look for alternative forms of pest and pathogen
management. Old chemistries once replaced by methyl bromide returned to use, novel
chemistries were trialed, a greater emphasis was placed on breeding for resistance,
cultural and mechanical practices received greater emphasis, and grafting vegetables
for disease management became a viable option (Martin, 2003).
Grafting
Background Information
The concept of grafting cucurbits originated as an early method to manage soil-
borne pathogens. In all cases regardless of crop, grafting involves the union of a scion
with desirable traits to a pathogen resistant rootstock. Cucurbit grafting techniques
include tongue approach grafting, hole insertion grafting, one cotyledon grafting, and
side grafting. Preferred grafting technique can vary by region, available equipment, or
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the selected rootstock species (Hassell et al., 2008). Cucurbit grafting for management of soil-borne pathogens is a relatively common practice in Europe and Asia where arable land is limited by encroaching urban areas and crop rotation may not be an option. Cucurbit grafting has also become a common practice in major watermelon producing Central American countries with specialized greenhouses developed by the
United Nations Industrial Development Organization (Amadio, 2004). Within the United
States, grafting is used extensively in perennial fruit and nut crops and, more recently, tomatoes (Kubota et al., 2008; Mudge et al., 2009; Reid and Hunt, 2000). Cucurbit grafting in the U.S. however, is still in its fledgling stages where only an estimated 250 hectares are in production (Davis et al., 2008b). Slow adoption in the U.S. market was due to methyl bromide, ample land for rotation, and significantly higher plant costs. As time has progressed methyl bromide has been lost, arable land built upon, gradual mechanization of the grafting process, and access to commercially grafted plants is reducing plant cost (Davis et al., 2008b; Lee et al., 2010).
Positives
While techniques and technology advance, the main objective of grafting remains unchanged; manage soil-borne pathogens and reduce overall inputs. Aside from added initial cost, a producer can save money by using reduced fumigant and pesticide rates for soil borne pest management. Reduced pesticide use lowers runoff potential and aids in shifting public opinion about modern agriculture. Capital investment on grafted plants should decline gradually as mechanization of the grafting procedure reduces labor and input costs (Kubota et al., 2008). Management of soil-borne pathogens with grafting provides organic producers with limited chemistries at their disposal as a viable solution to mitigate crop loss (Barrett et al., 2012). Grafting watermelon can provide added
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benefits like higher yields, harvest date flexibility, increased vigor, larger fruit, firmer
fruit, and added tolerance to abiotic stress (Alexopoulos et al., 2007; Davis et al., 2008a,
2008b; Lee et al., 2010). Aforementioned qualities are all traits watermelon breeders try
to incorporate into new cultivars. Breeding for disease resistance in watermelon can be
a lengthy process due to negative traits affiliated with the resistant line or accession
(Sitterly, 1972; Wehner, 2008). Current high-yielding cultivars are already marketable
with established reputations among producers. Grafting provides a method of achieving
disease resistance without the arduous process of creating a new cultivar.
Resistant cucurbit rootstocks are already commercially available with disease
resistance to powdery mildew (‘Marvel’ (Takii Seed, Salinas, CA)) , Rhizoctonia root rot
(‘RS-841’ (DeRuiter Seed / Seminis Seeds, St. Louis, MO)), Verticillium wilt (‘Kazako’
(Syngenta Seed, Boise, ID)), melon necrotic spot (‘Asisto’ (Takii Seed, Salinas, CA)) ,
root-knot nematodes (‘Dragon-2’ (BF Agritech, Kfar Yehoshua, Israel)) , and Fusarium
wilt races 0, 1, and 2 (‘Carnivor’ (Syngenta Seed, Boise, ID)) (USDA-NIFA, 2015).
Decreased rate of Fusarium wilt incidence and increased yields were observed when
watermelon scions were grafted on Fusarium wilt race 2 resistant hybrid rootstocks
(Keinath and Hassell, 2014). In addition, rootstocks of various lines and accessions
demonstrate significant ability in management of Meloidogyne spp. (Guan et al., 2014;
Pofu et al., 2011; Thies et al., 2015, 2010, 2007). Currently, resistance to both Fusarium
wilt of watermelon and root-knot nematodes is limited to only one commercial rootstock
‘RS-841’ (DeRuiter Seed / Seminis Seeds, St. Louis, MO) (USDA-NIFA, 2015). A wild
cucumber line (Cucumis pustulatus) demonstrated resistance to F. oxysporum f.sp. melonis and M. incognita, but resistance has not been demonstrated on F. oxysporum
15
f.sp. niveum thus far (Liu et al., 2014). In all, grafting cucurbits is an alternative for consideration in an integrated pest management plan or for use in a field with considerable Fusarium wilt pressure.
Negatives
Despite the multiple benefits of grafting as a management tool, it is not without flaws. An analysis conducted in 2008 assessed input costs of grafted watermelon plants versus non-grafted plants; grafting was estimated to increase initial production cost by
$1743 per hectare, with grafted plants costing almost three times that of a non-grafted
plant. Break-even points were set at $0.12/kg at a production rate of 50,000 kg/ha and
$0.19/kg at a production level of 40,000 kg/ha (Taylor et al., 2008). Last year, average
yields per hectare in Florida were 43,310 kg/ha with an average value of $0.13/kg
(USDA NASS, 2016). If a producer grew grafted watermelons and their crop performed
at the Florida state average they would not break even. Grafted seedlings are more
expensive than non-grafted seedlings because rootstock seeds are F1 (C. maxima x C.
moshcata) hybrids, the grafting procedure is labor intensive, and transplants are lost at
variable rates due to failed unions (estimated 65-95% survival rate) (Davis et al.,
2008b). Failed unions mean more scion and rootstock seedlings have to be produced
than the actual quantity needed. The majority of commercial rootstock lines available in
the United States are C. maxima x C. moschata hybrids. Characteristically these F1
hybrids are Fusarium wilt resistant but have been found to be susceptible to root-knot
nematodes (Thies et al., 2012, 2010). ‘RS-841’(DeRuiter Seed / Seminis Seeds, St.
Louis, MO), an F1 hybrid rootstock with marketed root-knot nematode resistance
performed similarly to other C. maxima x C. moschata hybrids in the presence of root-
knot nematodes (Giné et al., 2017; USDA-NIFA, 2015). Numerous citations in the
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literature describe the ability of plant parasitic nematodes to predispose resistant hosts to susceptibility (Bergeson, 1972; Golden and Van Gundy, 1975; Powell, 1971a). Plant cost and disease susceptibility are not the only negative aspects associated with grafting, various impacts to fruit quality are described in the literature. There are reports of grafted watermelons having low fruit weights, low Brix, yellow internal hard core, poor flesh texture, delayed maturity and increased rind thickness (Alexopoulos et al., 2007;
Davis et al., 2008a; Kyriacou et al., 2017; Lee et al., 2010; Turhan et al., 2012).
However the literature is contradictory on many issues involving fruit quality in grafted watermelons. Poor selection of rootstocks and scions can cause negative rootstock/scion interactions that may not be observed with a different combination.
Delayed maturity has been reported in grafted watermelon and muskmelon when compared to non-grafted counterparts. Trials containing both grafted and non-grafted plants, harvested at a uniform interval could be causing low Brix in experiments (Davis et al., 2008a; Xu et al., 2005)
Nematodes in Watermelon Production
Root-knot Nematodes (Meloidogyne spp.)
Within the genus Meloidogyne there are 97 species described in the literature
(Castagnone-Sereno et al., 2013). The most widespread and economically important species in regards to vegetable crops are M. incognita, M. javanica, M. arenaria, and M. hapla (Luc et al., 2005). Root-knot nematodes are primarily tropical to sub-tropical organisms excluding two species, (M. hapla and M. chitwoodi) which are more acclimated to temperate regions. Meloidogyne spp. are sedentary endoparasites that primarily reproduce parthenogenetically, although males are often observed under conditions of nematode stress. Root-knot nematodes possess a broad host range that
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encompasses many vegetable crops. Development of the second stage juvenile (J2) occurs within the egg prior to hatching. J2 is the infective stage for root-knot nematodes so once hatched the female locates a host, penetrates root tissue, and establishes a permanent feeding site in the root cortex (Taylor and Sasser, 1978). Three molts will
occur inside root tissue prior to becoming reproductive adults. Root-knot nematodes can
potentially lay up to 2000 eggs in a lifetime. Development is temperature dependent
with the ideal soil temperature ranging between 21 and 27°C. At 26°C most
Meloidogyne spp. will complete their lifecycle in 21 days (Taylor and Sasser, 1978).
Lifecycle completion is also affected by host, with more susceptible hosts producing
generations at an accelerated rate (Anwar et al., 1994).
Root-knot nematodes are estimated to inflict 12.3% yield loss on major crops
across the world (Sasser, 1987). Early research estimated Meloidogyne spp. cause
crop loss of 18-33% in subtropical climates annually (Sasser, 1979). Multiple
Meloidogyne spp. are capable of inflicting damage on C. lanatus. Watermelons are
susceptible to all 4 races of M. incognita, M. javanica, and both races of M. arenaria
(Sasser and Carter, 1982).
The Reniform Nematode (Rotylenchulus reniformis)
The reniform nematode (Rotylenchulus reniformis) is a semi-endoparasite and
one of the two parasitic members of the Rotylenchulus genus (A F Robinson et al.,
2005). The first juvenile stage (J2) molts within the egg then emerges. One to two
weeks post hatching the infective stage (J4) is reached and root penetration can occur.
Once the root is penetrated, one to two more weeks is required to reach reproductive
maturity where eggs are deposited in a gelatinous matrix (Wang, 2007). The reniform
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nematode lifecycle can be completed in 17-29 days and is greatly influenced by host plant and soil temperature (Chitambar, 1997).
On a global scale, R. reniformis is considered the second most important plant
parasitic nematode affecting vegetables crops, but is often overlooked or disregarded
where it occurs concomitantly in soil with Meloidogyne spp. (Luc et al., 2005).
Rotylenchulus reniformis has demonstrated infective capabilities on 314 plant species
including many cucurbits and other vegetable crops, making it a limiting factor in
vegetable production where populations are present (Robinson et al., 1997).
Populations of R. reniformis are at a minimum in the late spring and during the early
months of a summer cropping season. This however changes as soil temperatures rise
and reproductive rates increase, putting soil populations at a maximum as a spring
planted crop nears maturity (Luc et al., 2005). Research in Cuba during 1971
demonstrated that R. reniformis was able to survive 29 months in the absence of a host.
Long term survival without a host is due in large part to the reniform nematode’s ability
to undergo anhydrobiosis and preserve itself through desiccation (Stoyanov, 1971). The
ability to survive more than one year in the absence of a host makes it difficult to control
with rotation to a non-host crop.
R. reniformis is not a well-documented pathogen of the family Cucurbitaceae in
the United States, even with the earlier statement from Luc et. al (2005) which regarded
the nematode as the 2nd most important plant parasitic nematode (PPN) for vegetables.
In the Southeastern United States the reniform nematode is most commonly described
as a pest of cotton or soybeans (Jones et al., 1958; Robbins and Rakes, 1996). While
not as heavily researched as soybeans and cotton, literature involving cucurbits and the
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reniform nematode does exist. In a host differential screening R. reniformis demonstrated the ability to infect and reproduce on cultivated watermelon (Robinson
1997). An experiment in North Carolina demonstrated that pre-plant fumigation in summer squash with Telone (1,3-Dichloropropene) provided a 69% yield increase over the non-fumigated control when reniform nematode was present (Heald, 1978). Heald et al. (1975) later demonstrated that heavy infestation of R. reniformis on cantaloupe
(Cucumis melo) was capable of greatly reducing above ground biomass by
approximately 40%. In Brazil it was shown that R. reniformis can reproduce on the roots
of cucurbit species and does reduce above ground biomass as populations increase
(Torres, 2005). Citrullus lanatus cv. ‘Sugar Baby’ was included, but had a very low
reniform reproductive rate. Cucurbita moschata was observed to have significantly
greater nematode reproductive rate for R. reniformis (Torres 2005) and is one of the
genetic components in interspecific hybrid rootstocks which have documented
susceptibility to Meloidogyne spp. (Giné et al., 2017; Thies et al., 2012). Given the
ability to host on watermelon, reduce cucurbit yield, and reproduce extensively on C.
moschata, R. reniformis is a pest for further consideration in grafted melon systems.
Nematode Movement and Soil Depth
The majority of nematode movement is passive in nature and often carried out by
an external stimulus. While the aforementioned is true, nematodes are still capable of
migrating in soil independently, and sometimes travel great distances. Voluntary
nematode movement in the soil is almost always stimulus driven (root exudates,
temperature, moisture, mating). Work in tomato demonstrated movement of
Meloidogyne spp. 25 cm in nine days with 50% of the population migrating a distance of
50 cm in a sandy soil profile (Prot, 1978). Similarly R. reniformis migrated a lateral
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distance of 200 cm, and 91 cm downward at a rate of 0 – 3.3 cm a day from a point of
inoculation at soil surface (Moore et al., 2010). Movement can vary by soil temperature
and species. M. incognita and R. reniformis differ in temperature preference with root-
knot migrating toward simulated thermal sunny day temperatures and reniform moving
away. (Robinson, 1994)
Nematode management in a plasticulture system is accomplished by applying
nematicides using shanks or drip irrigation lines. This type of application may only
control nematodes in the top 30 cm of soil. Previous research in cotton indicated
substantial populations of R. reniformis occur below the 20 cm plow layer, with
substantial numbers deeper than 36 cm (A F Robinson et al., 2005; A. F. Robinson et
al., 2005; Robinson et al., 2000). Given the previous, actual depths where nematodes
survive and thrive is often much deeper than the top 30 cm of soil. Additional studies on
cotton demonstrated low reniform nematode populations at a depth of 1.75m (Heald
and Thames, 1980). Fumigation at 60-120 cm deep increased cotton yields by 68%
compared to the non-fumigated control (Westphal and Smart, 2003). Westphal also
noted that population densities in the 0-120 cm horizon provided a more accurate
perspective on potential crop damage than 0-30 cm. While the majority of depth
research for plant parasitic nematodes has been cotton/reniform oriented, a few studies
involve horticultural crops. Meloidogyne spp. collected from vineyard soil were
predominantly located at 60 cm in depth with similarity to the 120 cm depth and isolated
individuals detected at 330 cm deep (Ferris and McKenry, 1974). Florida strawberry
crops fumigated with a deep shank (40 cm) and deep drip applications of 1,3-
dichloropropene (Telone; 140-168 L/ha), saw reduced populations of sting and root-knot
21
nematodes at seasons end, and increased yields 9-29% when compared to the grower fumigation standard (Noling et al., 2016). Root-knot nematode populations inoculated at a depth of 120 – 135 cm from the soil surface reduced tomato yield by 11% and 59% in spring and fall experiments in sandy soil (Johnson and Mckeen, 1973). This movement
in sandy soils can contribute to the rapid recolonization of a fumigated area post
treatment when a susceptible host is planted (Prot, 1980; Yeates et al., 1991).
Symptomology and Diagnosis
Plant parasitic nematodes vary by species in mobility, speed, body shape, and
life cycle. Nematode extraction from soil and root tissue varies by technique depending on financial resources, species, or desired life stage. For comprehensive assessment of nematode root tissue populations, roots must first be macerated with a blender or finely reduced in some manner. Extraction methodology falls in two different categories: passive extraction or active extraction (McSorley and Walter, 1991). Passive nematode extraction is ideal when either motile and non-motile life stages or eggs are desired.
Passive techniques include various centrifuge flotation methods, filtration or sieving, and enzymatic digestion which can be used for soil or plant material. Active nematode extraction removes only motile, vermiform nematodes and is not applicable collecting eggs or sedentary adults. Active methods of extraction include Baermann funnels, incubation chambers, mist chambers or a Whitehead tray which effectively process soil or root samples (EPPO, 2013).
Meloidogyne spp. above ground symptomology is characteristic of many plant parasitic nematodes, including irregular damage distribution of stunted growth, chlorosis, and reduced yield. Below ground disease expression is characterized by gall formation on roots that are distinct to two genus of plant parasitic nematodes:
22
Meloidogyne and Naccobus (false root-knot nematode). Root-knot galling differs from false root-knot galling by pattern. Meloidogyne spp. galling occurs in a more sporadic distribution while Naccobus spp. produce galls in a bead-like chain, giving it the nickname “rosario” in Latin America (Luc et al., 2005).
Identification of root-knot nematodes by species can be accomplished by two
methods. The oldest method involves studying perineal patterns and morphological
features on enlarged sedentary females. Each Meloidogyne spp. possesses distinct
perineal rings, cuticular annulations, labia shapes, and different labial disk shapes
(Eisenback et al., 1980). This technique however is time consuming, intricate, uses
specialized equipment, and requires a highly trained individual. Even when performed
by an experienced individual, technical flaws, poor mounts and divergence within a
species may lead to an incorrect diagnosis. Polyacrylamide gel electrophoresis (PAGE)
is a more accurate form of species identification, which can encompass a broader range
of skill levels. The use of PAGE in 1971 created stable enzymatic profiles composed of
four dehydrogenases (lactate, malate, glucose-6-phosphate, and a-glycerophosphate)
and three hydrolases (acid and alkaline phosphatase and esterase) for Meloidogyne spp.: arenaria, hapla, incognita, and javanica (Dickson et al., 1971).
Symptomology of R. reniformis is often hard to discern. Like many other plant
parasitic nematodes above ground symptoms include stunted growth, chlorosis, and
yield reduction. Heavily infected roots may appear dirty even after rinsing due to soil
adhesion to the gelatinous egg masses. Root interveinal tissue may have a speckled
appearance or brown root cortex, but the most visible symptom is growth reduction of
the root system (Luc et al., 2005).
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Like root-knot nematodes, reniform nematodes can be speciated by morphological appearance but it requires extensive experience. Within R. reniformis, morphology however can vary greatly by regional populations within a single country and phenotypic polymorphism occurs when compared on a global scale (Agudelo et al.,
2005). Identification of reniform nematodes is best accomplished by using PCR with species specific primers developed for R. reniformis (Van Den Burg et al., 2015).
Management
Equipment sanitization reduces plant parasitic nematode spread from heavily
infected fields to nematode free fields (Collange et al., 2011). Rotation to non-host crops
aids in reducing most (excluding cyst and species capable of anhydrobiosis) plant
parasitic nematodes below damage threshold levels (Barker and Koenning, 1998).
Cover crops, green manures, and residue from Brassica spp. all demonstrate some
level of nematode management (Collange et al., 2011). Varietal resistance to root-knot
nematodes is not well documented in available watermelon cultivars, but grafting using
rootstocks ‘Dragon-2’ (BF Agritech, Kfar Yehoshua, Israel) ‘RS-841’ (DeRuiter Seed /
Seminis Seeds, St. Louis, MO), ‘Shelper’ (Takii Seed, Salinas, CA), or ‘Vita’ (Vilmorin,
Salinas, CA) remains an option (Guan et al., 2014; Thies et al., 2012, 2015, 2010, 2007;
USDA-NIFA, 2015).
Chemical control of plant parasitic nematodes is performed either pre-plant or
post plant depending on application method and active ingredient. Pre-plant control
generally utilizes soil fumigants applied by shank or through irrigation which varies in
efficacy due to fumigant distribution in the soil (Desaeger et al., 2017; J. Desaeger et
al., 2004). Post-plant chemical control uses systemic compounds or compounds with
little to no phytotoxicity (Haydock et al., 2006; Jones et al., 2017).
24
An economic threshold is the pest density where treatment for control provides
an economic return. For nematodes this relates populations to crop damage where the
value of destroyed crop exceeds the cost of nematode control (Hunt et al., 2009). A
damage threshold level is very similar to an economic threshold, but rates to plant
pathology and is rarely static (Nutter Jr. et al., 1993). A damage threshold for M.
incognita on watermelon was estimated at an initial population (Pi) equal to 122
eggs/100 cm3 of soil or 3.6 J2/100 cm3 of soil, and 1.6 galls on bioassay roots/100 cm3
of soil (Xing and Westphal, 2012). Management of plant parasitic nematodes at an
economic threshold is best achieved through a combination of methods and no method
should be relied on solely (Noling, 1997).
Fusarium Wilt
Biology and Background Information
Fusarium oxysporum has been described as one of the most economically impactful and adaptive fungal pathogens in the Ascomycota. Interestingly both pathogenic and non-pathogenic strains occur in agricultural soils and pathogenic formae specialis are non-pathogenic in the absence of a proper host (Gordon and Martyn,
1997). Non-pathogenic strains of F. oxysporum can be beneficial in the right settings,
providing biocontrol of pathogenic variants by direct resource competition (Larkin and
Fravel, 1999). Some endophytic variants act as systemic bio-controls and are capable
of providing significant resistance to plant parasitic nematodes (Vu et al., 2006). The
concept of formae specialis was first proposed to aid pathologists in the classification of
host specificity within F. oxysporum. Snyder and Hansen (1954), describes the first 25
forma specialis based on physiological and host capabilities without altering formal
taxonomy. As of 2009 there are 120 documented formae specialis for F. oxysporum that
25
infect a broad range of plant species (Michielse and Rep, 2009). To further divide the
Fusarium complex, variation in virulence can occur within a formae specialis, which are
designated as races. Races are defined by differential interaction with cultivars that may
or may not carry genes for pathogen resistance (Armstrong and Armstrong, 1978).
Fusarium Wilt of Watermelon
The market dominance of seedless watermelon, combined with the phase-out of
MeBr, and reduced available land for crop rotation has caused an increase of Fusarium
wilt in watermelon. Fusarium wilt in cultivated watermelon (Citrullus lanatus) is a
disease caused by the host specific fungal pathogen, Fusarium oxysporum f.sp.
niveum. Fusarium wilt of watermelons was first documented in 1894 in South Carolina
and Georgia by E.F. Smith (Smith, 1894). Fusarium wilt has been described as the most
economically important disease of cultivated watermelon on a global scale, with an
estimated 75 - 100% possible crop loss when susceptible varieties are grown in
combination with heavily infested fields (Bruton et al., 1999; Egel, 2007). To date, four
races are documented: 0, 1, 2, and 3. Varietal resistance to races 0 and 1 were
identified and incorporated in commercial cultivars but no reliable resistance has been
found to race 2 and 3 (Everts and Hochmuth, 2010; Zhou et al., 2010). One of the
greatest issues with managing this pathogen is the longevity and viability of
chlamydospores. Fusarium oxysporum prefers acidic, light, sandy soils which is the
predominant texture of most soils found in the southeastern U.S. Soil factors impact
symptom development of Fusarium wilt in watermelon. Fusarium wilt prefers cool
season temperatures with an optimum range at 25-27°C, temperatures greater than
27°C significantly reduce activity (Egel, 2007). In addition, disease severity is often
increased with low soil pH, reduced soil moisture, and high soil nitrogen levels. Surveys
26
were conducted in Delaware and Maryland in 2004 to determine F. oxysporum f.sp.
niveum levels in each field and the amount of wilt observed in the race 1 and 2
susceptible cultivar ‘Sangria’ at harvest. The conclusion drawn was that the minimum
amount of inoculum to cause wilt was 166 CFU/g soil and that 367 CFU/g was enough
to cause wilt in 50% of the plants (ID50). In the majority of the fields examined (73%)
the CFU/g soil values ranged between 100 and 1,200 (Zhou and Everts, 2004).
Symptomology and Diagnosis
Symptoms of F. oxysporum f.sp. niveum (FON) is first observed in the field by
wilting or “flagging” of leaves. Following flagging, insipient wilt, necrosis, and eventually
vine termination or plant death results. Heavy infestations on young plants can lead to
damping off and plant death. Under wet field conditions pinkish-white fungal hyphae can
be observed at the junction of stem base and soil line (Egel, 2007). Wilt symptomology
is caused by tyloses which are vascular blockages created by the plant to prevent
pathogen advancement. In susceptible plants the formation is delayed and does not
form quickly enough to block the advancement of F. oxysporum (Beckman, 1964). The
characteristic field diagnostic symptom for Fusarium wilt in the field is the discoloration
of the xylem. Vascular discoloration can be observed by using a sharp object to make a
parallel cut at the stem base to observe the root cortex. The other method includes
cutting a cross section revealing a brown ring of necrotic xylem tissue. Heavily infected
plants may die in a matter of ten days while plants with a lesser inoculum load may take
several weeks (Kleczewski and Egel, 2011).
If Fusarium wilt is suspected on watermelon then an isolation test should be
performed. Tissue should be harvested near the plant base on freshly infected tissue to
reduce contamination risk with another fungi or bacteria. Remove soil and debris rinsing
27
with room temperature water then cut 1-2 cm pieces. Place pieces in a 0.6-1.2% bleach solution for a time period of 1-5 minutes for surface disinfection (Leslie and Summerell,
2006). Rinse sections 2-3 times with deionized water to remove excess bleach solution then plate, preferably on Komada’s media. If Komada’s media is not available select
Malachite-green, streptomycin amended PDA plate or a pentachloronitrobenzene
(PCNB) selective media (Castella et al., 1997). Incubate under florescent lighting for twelve hours at room temperature and in five days a colony should grow. Fusarium colonies can vary widely in both in morphology and in color. Hyphae can range from white to pale violet, macroconidia may be violet or orange and coloration of agar can range from deep violet to no color in some isolates (Kleczewski and Egel, 2011; Leslie and Summerell, 2006). F. oxysporum forms three types of reproductive spores: microconidia, macroconida, and chlamydospores (Egel, 2007). Microconida are reniform shaped and form on false heads. Macroconidia are banana or canoe shaped with 3-5 separated cells produced in large clumps called sporodochia. Chlamydospores are thick walled, circular reproductive structures forming either terminally or intercalary in long chains during periods of duress(Leslie and Summerell, 2006). Identification in culture can be determined by shape of the macroconidium, the structure of the microconidiophore, and the formation orientation of chlamydospores (Gordon and
Martyn, 1997). Confirmed F. oxysporum does not necessarily indicate pathogenicity as non-pathogenic species can also be cultured (Gordon and Martyn, 1997). Pathogen status can be determined with further plating on carnation leaf agar for a period of weeks. Macroconida formation in the center on this agar plate indicate pathogenicity.
Further classification can be accomplished by race testing. Race determination of FON
28
is performed with a host susceptibility test using cultivars of varying resistance: ‘Black
Diamond’ (susceptible), ‘Charleston Grey’ (0), ‘Calhoun Grey’ (1), and PI-296341-FR
(2). Race 0 causes disease in ‘Black Diamond’, race 1 causes disease in ‘Charleston
Gray’ and ‘Black Diamond’, race 2 causes disease in ‘Calhoun Grey’ and aforementioned lines, and race 3 causes disease in all cultivars present (Martyn and
Netzer, 1991).
Management
A major concern with Fusarium wilt of watermelons is the longevity and viability of reproductive structures. Chlamydospores formed within the mycelium can persist in soil without a viable host for a maximum of 15-20 years (Egel, 2007). Prolonged survival in the absence of a host reduces efficacy of rotation with a non-host crop but rotation every 5-7 years can significantly reduce viable fruiting bodies (Zhou and Everts, 2004).
When the United States grew predominantly diploid watermelon, resistance to FON was prevalent and the disease was better controlled (Elmstrom and Hopkins, 1981). Within the past 20 years the US market has gone to predominantly triploid seedless watermelon production which does not possess comparable varietal resistance (Bruton et al., 1999).
The best management strategy for Fusarium wilt is to avoid introduction of inoculum in to a field. Sanitation of equipment and clean seed or transplants can greatly reduce chances of introduction (Bruggen and Finckh, 2015). FON and other Fusarium spp. prefer cooler soil temperatures so planting date adjustment can reduce disease impact (Matheron et al., 2005). Pre-plant fungicidal treatments can also reduce
Fusarium wilt symptomology in young transplants (Egel and Hoke, 2007). If FON is already present in a field then soil fumigation in combination with varietal resistance or
29
grafted transplants provides the greatest management for Fusarium wilt of watermelon
(Egel, 2007; Elwakil et al., 2008; Hopkins and Elmstrom, 1979; Keinath and Hassell,
2014).
Nematode/Pathogen Interactions and Disease Complexes
Nematode interactions with Pathogens
Plant parasitic nematodes interact with numerous disease causing agents including bacteria, fungi, and viruses (Al-hazmi et al., 2002; McGuire, 1972; Moura et al., 1975). Pathogen interactions result from virus transmission, plant wounding, or host modification and vary in severity from synergistic to antagonistic (Bergeson, 1972).
Rotylenchulus reniformis interacts with Rhizoctonia solani to create a synergistic disease impact on cotton (Sankaralingam and McGawley, 1994). Pythium ultimum and
Rhizoctonia solani demonstrate affinity for root-knot nematode feeding sites. Both pathogens colonize giant cells in the root tissue and the host plant succumbs to root rot disease, antagonistically displacing M. incognita (Golden and Van Gundy, 1975;
Melendez and Powell, 1970). Pathogen relationships are not incurred intentionally and in some cases nematode survival is compromised by the invader.
Nematode interactions with Fusarium
A point of concern arises when both F. oxysporum and plant parasitic nematodes occur in the same cropping system, which is not uncommon in Florida soils. Synergism
between pathogens can inflict greater disease in a susceptible host or potentially
predispose a resistant host to pathogen susceptibility. Fusarium spp. exhibit possible
attraction to root-knot nematodes in the context of root wounding. Gomes et al., (2013)
documented Fusarium solani mycelium increasing propagule counts and greater
disease virulence when exudates from M. enterolobii parasitized roots were applied.
30
Previous literature demonstrated when both Meloidogyne spp. and F. oxysporum occur as co-contaminates, Fusarium wilt severity increases (Garber et al., 1979; Mai and
Abawi, 1987; Powell, 1971a, 1971b; Taylor, 1979). Pathogenicity of nematodes varies within genus by species, with some species causing a greater synergistic effect with F. oxysporum than others (Jenkins and Coursen, 1957). Conflict within literature informally addresses the intricacy and variability associated with fungal disease complexes.
Similar research in tomato demonstrated no degradation or synergism in host
resistance when subjected to a co-inoculation of F. oxysporum and M. incognita (Abawi
and Barker, 1984). Not all forma speciales of F. oxysporum are pathogenic to plants
and not all nematode Fusarium interactions produce disease complexes or synergism.
Antagonism has also been documented between nematodes and Fusarium spp.
Damage caused by the sugar beet cyst nematode (Heterodera schachtii) was greatly
reduced when co-inoculated with F. oxysporum due to disruption of feeding sites
(Jorgenson, 1970). Endophytic variants of F. oxysporum demonstrated significant
reduction of burrowing nematode (Radopholus similus), creating an antagonistic
interaction.(Vu et al., 2006)
Nematode interactions with other Nematodes
By definition a plant parasitic nematode can be classified as either a parasite or
pathogen due to its feeding habit and its ability to cause disease. Plant parasitic
nematode interactions with different plant parasitic species can be competitive,
synergistic or antagonistic. A classic example of nematode competition is that of M.
incognita and R. reniformis. Competition between sedentary endoparasites is based
largely on environmental conditions and viable sites for reproduction, and are
considered weak interactions (Eisenback, 1985). When both species are present in a
31
field, the species with greater populations tends to dominate the microcosm (Diez et al.,
2003; Thomas and Clark, 1983). Variability of competition outcomes involving two species could be influenced by systemic acquired resistance (SAR). SAR is a form of induced resistance to a pathogen caused by previous infection of an alternate pathogen
(Sticher et al., 1997). In context of reniform and root-knot species, SAR forms to whichever nematode establishes secondarily on the root system (Aryal et al., 2011). R. reniformis may however be more competitive in monocultural settings than M. incognita.
Reniform nematode demonstrated greater competitiveness in cotton than M. incognita
(Robinson, 2007) and progressively replaced M. incognita across multiple cotton fields in a 10 year period (Spurlock et al., 2014). Species replacement in an agricultural field could be influenced by both crop and soil texture. Cotton has proven to be a better host for R. reniformis than M. incognita, and R. reniformis prefers fine textured soils while M. incognita prefers sandy or coarse textured ones (Koenning et al., 1996).
With all nematode/pathogen interactions it is important to remember combinations of only two pathogens do not often occur in nature. In many cases pathogenic interactions involve more than one species of plant parasitic nematode (Freckman and
Caswell, 1985). As a broad overview, migratory endoparasites are antagonistic to sedentary endoparasites and ectoparasites, and ectoparasites can be suppressive or experience mutual antagonism with endoparasites (Eisenback, 1985). Cumulative plant damage however is typically synergistic with multiple species of plant parasitic nematodes present in the same system. Overall root reduction increased in six species of Bermudagrass as nematode species: ring nematode (Criconemoides ornatus), stunt nematode (Tylenchorhynchus martini), and sting nematode (Belonolaimus
32
Iongicaudatus) were introduced together (Johnson, 1970). Observations made in
Chrysanthemum involving Pythium aphanidermatum, Belonolaimus longicaudatus (sting nematode), and Meloidogyne incognita demonstrated greatest disease severity when both nematode species were co-inoculated with the oomycete versus individually
(Johnson and Littrell, 1970)
33
CHAPTER 2 SCREENING CUCURBIT ROOTSTOCKS FOR VARIETAL RESISTANCE TO MELOIDOGYNE SPP. AND ROTYLENCHULUS RENIFORMIS
Introduction
Damage caused by soil-borne pathogens, like Fusarium oxysporum f.sp. niveum and plant parasitic nematodes (Meloidogyne spp. and Rotylenchulus reniformis) has increased since the phase out of methyl bromide. The ban on methyl bromide was estimated to have a significant economic impact, costing consumers and producers at least $1 billion annually (USDA ERS, 2000). The loss of methyl bromide has pushed research toward identifying viable alternatives for pathogen management.
A growing method for soil-borne pathogen management in U.S. cucurbit production is grafting. In all cases regardless of crop, grafting involves the fusion of a high yielding, susceptible scion to a pathogen resistant rootstock. Currently cucurbit grafting is most common in European and Asian countries where intensive land use and inability to rotate crops make it a viable option. Cucurbit grafting techniques include tongue approach grafting, hole insertion grafting, one cotyledon grafting, and side grafting. Preferred grafting technique can vary by region, available equipment, or the selected rootstock species (Hassell et al., 2008). While resistance to both Meloidogyne spp. and F. oxysporum f.sp. niveum have been identified independently, no commercial rootstocks or cultivars currently possess resistance to both (Keinath and Hassell, 2014;
Pofu et al., 2011; Thies et al., 2010, 2007). Research conducted in cantaloupe
(Cucumis melo) demonstrated significant reduction in root gall index (RGI), reproductive factor (Rf), and J2 recovered from soil when grafted on root-knot nematode resistant
Cucumis metulifer (Guan et al., 2014). Decreased rate of Fusarium incidence and increased yields were observed when watermelon scions were grafted on Fusarium wilt
34
race 2 resistant hybrid rootstocks (Keinath and Hassell, 2014). Previous research shows interspecific (C. maxima X C. moschata) hybrids are susceptible to root-knot
nematodes and can sometimes perform comparably to the susceptible control (Giné et al., 2017; Guan et al., 2015; Thies et al., 2012, 2010). Resistance to both F. oxysporum
f.sp. melonis and M. incognita has recently been documented in China in wild cucumber
(Cucumis pustulatus), but the research did not involve F. oxysporum f.sp. niveum (Liu et
al., 2014).
Fusarium wilt of watermelon, caused by F. oxysporum f.sp. niveum (FON) has
four current documented races: 0, 1, 2, and 3. Currently, varietal resistance to races 0
and 1 are found in commercial watermelon cultivars but no reliable resistance has been
found to race 2 and 3 (Everts and Hochmuth, 2010; Zhou et al., 2010). Fusarium wilt of
watermelon, when uncontrolled has the potential to cause 100% crop loss in a field
planted with a susceptible variety (Egel, 2007). A major concern with Fusarium wilt of watermelon is the longevity and viability of reproductive structures. Chlamydospores formed within the mycelium can persist in soil without a viable host for a maximum of
15-20 years (Egel, 2007). Extensive survival in the absence of a host removes potential management options like rotation with a non-host crop.
Multiple Meloidogyne spp. (RKN’s) are capable of inflicting damage on C.
lanatus. Watermelon is susceptible to all 4 races of M. incognita, M. javanica, and both
races of M. arenaria (Sasser and Carter, 1982). Members of the Cucurbitaceae family
differ in susceptibility to root-knot nematodes and resistant cultivars are not commercially available (Guan et al., 2014; Levi et al., 2009; Thies et al., 2007). Early
literature estimated root-knot nematodes cause crop loss of 18-33% in subtropical
35
climates annually (Sasser, 1979). The M. incognita damage threshold level of the
watermelon cultivar ‘Royal Sweet’ was estimated with an initial population (Pi) equal to
122 eggs/100 cm3 of soil, 1.6 galls on bioassay roots/100 cm3 of soil, or 3.6 J2/100
cm3 of soil (Xing and Westphal, 2012). Meloidogyne incognita is a common pest in
sandy soil which is the dominant component of many soils in Florida.
Historically, Rotylenchulus reniformis is not a well-documented pathogen of the
family Cucurbitaceae in the United States. In the Southeastern United States it is most
commonly known as a pest of cotton (Robinson et al., 2005), and has demonstrated
host ability on cultivated watermelon (Robinson 1997). Early literature stated that pre-
plant fumigation in summer squash with 1,3-dichloropropene provided a 69% yield
increase over the non-fumigated control (Heald, 1978). In Brazil it was shown that R.
reniformis can reproduce on the roots of cucurbit species and does reduce above
ground biomass as populations increase (Torres 2005). Given the ability to host on
watermelon and reduce yield on other cucurbits, R. reniformis is a pest for
consideration.
A point of concern arises when both F. oxysporum and PPN’s occur within the
same cropping system, which is not uncommon in sandy soils. A synergistic interaction
between pathogens can potentially predispose a resistant host to pathogen
susceptibility. Previous literature has demonstrated that when both RKN’s and F.
oxysporum are present in the same area, Fusarium symptomology increases (Mai and
Abawi, 1987; Powell, 1971a, 1971b; Taylor, 1979). However, the literature is unclear
regarding this interaction and a disease complex does not always form. A study
conducted in tomato demonstrated no degradation in host resistance when subjected to
36
a co-inoculation of F. oxysporum and M. incognita (Abawi and Barker, 1984). The
primary objective of this experiment was to screen cucurbit rootstocks resistant to FON
for additional resistance to RKN’s and other plant parasitic nematodes.
Materials and Methods
Experiments were conducted during spring and fall of 2015 and 2016 at the
North Florida Research and Education Center (NFREC) located in Quincy, Florida. The
soil type was Dothan-Fuquay fine, sandy loam (Fine-loamy, kaolinitic, thermic Plinthic
Kandiudults - Loamy, kaolinitic, thermic Arenic Plinthic Kandiudults). The experiment was arranged as a randomized complete block design with four replications. Soil was cultivated to a depth of 25 cm prior to bed formation. Rows were spaced 2.44 meters apart, and beds were 76.2 cm wide and 20.3 cm tall. The field used had a history of R.
reniformis and RKN infestation. White on black polyethylene mulch was used in fall and
black polyethylene mulch was used in spring with irrigation being provided by a single
off center drip tape. Plants were started from seed in 128 cell transplant trays. Twelve,
three-week-old seedlings were transplanted in each experimental plot and spaced 91
centimeters apart. The rootstock ‘Carnivor’ (Syngenta Seed, Boise, ID) was used as the
Fusarium wilt resistant, root-knot nematode susceptible control. Interspecific hybrid (C.
maxima x C. moschata) rootstocks like ‘Carnivor’ have documented susceptibility to
RKN’s (Thies et al., 2012, 2010). Spring 2015 treatments included ‘Carnivor’, ‘Bulldog’
(United States Vegetable Lab (USVL), Charleston, SC), ‘USVL-482351’ (United States
Vegetable Lab, Charleston, SC), ‘USVL-246’ (United States Vegetable Lab, Charleston,
SC), ‘USVL-252’ (United States Vegetable Lab, Charleston, SC), and ‘USVL-360’
(United States Vegetable Lab, Charleston, SC). Fall 2015 treatments included
‘Carnivor’, ‘Bulldog’, ‘USVL-482351’, ‘USVL-246’, and ‘USVL-252’. Spring 2016
37
treatments included ‘Carnivor’, ‘Bulldog’, ‘USVL-482351’, ‘USVL-246’, ‘USVL-252’, and
‘SP-6’ (Syngenta Seed, Boise, ID). Fall 2016 treatments included ‘Carnivor’, ‘Bulldog’,
‘USVL-246’, ‘USVL-252’, ‘USVL-360’, and ‘SP-6’. Rootstock use varied by experiment due to limited seed availability on the non-commercial lines.
Root gall index is a rating system utilized to quantify damage caused by RKN’s on a plant root system. A rating scale of 0 – 10 is used where 0 indicates no galling and
10 is a completely dead plant (Bridge and Page, 1980). RGI was collected at 30-day intervals beginning 30 days after planting (DAP) in 2016 spring and fall experiments.
Subsequent ratings were collected at 60 DAP for fall of 2015 and spring and fall of
2016. Final ratings were collected at 90 DAP for all four experiments. Three plants were evaluated at 30 and 60 DAP, and five or all remaining plants were collected and evaluated at 90 DAP. Root systems were excavated and triple washed prior to root gall evaluation.
Soil and root tissue samples were collected at 90 DAP for each plot in all but one experiment (Spring 2015). Ten 15 cm soil samples were collected with a hand probe from the individual plots, then homogenized prior to collecting a sub sample of 100 cc of soil for extraction. For root tissue samples, ten grams of root tissue was collected for each plot and chopped finely to aid in extraction. Nematode extraction from both soil and root tissue was conducted in the same manner. Soil or root samples were first placed in a large coffee filter then put in a Baermann funnel with water (EPPO, 2013).
After a period of 48 hours nematodes were removed from the bottom of the funnel and
plant parasitic species were counted.
38
For data analysis individual plant RGI ratings were first averaged by plot. Once plot averages were obtained the data was subjected to one way ANOVA, then mean separation was performed by Fisher’s LSD test at P = 0.05 in the SAS Program version
9.4. Nematode soil population data was first separated by parasitic species (RKN’s and
R. reniformis) to be analyzed independent of one another. With the data separated, each species was subjected to a one way ANOVA with mean separation determined by
Fisher’s LSD at P = 0.05 in the SAS Program version 9.4. Nematode root tissue
population data was first separated by parasitic species (M. incognita and R. reniformis)
to be analyzed independent of one another. The species data separated was subjected
to a one way ANOVA with mean separation determined by Fisher’s LSD at P = 0.05 in
the SAS Program version 9.4.
Results
In spring of 2015, ‘Bulldog’, and USVL lines ‘246’, ’252’, ‘360’ had significantly
less galling than the control and ‘USVL-482351’ at the 90 DAP sampling interval.
In fall of 2015, ‘Bulldog’ and USVL lines ‘246’ ‘252’ showed resistance in terms of
less root galling than ‘Carnivor’ in both 60 and 90 DAP sampling intervals. ‘USVL-
482351’ was galled less at both 60 and 90 DAP when compared to the control but had
greater galling than the three previously mentioned lines. 90 DAP soil populations for
both plant parasitic nematode species were found at the greatest density in the ‘USVL-
482351’ rootstock, while the control, ‘Carnivor’, and resistant lines ‘Bulldog, ‘USVL-246’
and ‘USVL-252’ had significantly less. Root tissue populations at 90 DAP for M.
incognita were greatest in ‘Carnivor’ and separated from all other lines present in the
trial. However, R. reniformis root tissue populations however were numerically highest
in ‘USVL-482351’ and comparable to the control.
39
During spring 2016 at the 60 DAP RGI, ‘Carnivor’ exhibited significantly greater root galling than all other lines in this trial. At the 90 DAP interval however only ‘Bulldog’,
‘SP-6’, and USVL lines ‘246’ and ‘252’ had less galling than ‘Carnivor’, ‘USVL-482351’ was similar. 90 DAP RKN populations from root tissue were greatest in ‘Carnivor’ with
all other varieties having significantly lower populations. No significance was observed
in populations collected from the soil for either species and no significance was found
for R. reniformis from the root tissue.
During fall 2016 at both the 60 and 90 DAP RGI, ‘Bulldog’, ‘SP-6’, and USVL
lines ‘246’, ‘252’, and ‘360’ all had significantly less root galling than ‘Carnivor’. At 90
DAP, R. reniformis soil populations were greater on ‘Carnivor’ than any of the selected
entries. Root tissue populations for RKN’s at 90 DAP were observed to be highest in the
control with similarity to ‘SP-6’. ‘Bulldog’, and USVL lines ‘246’, ‘252’, and ‘360’ all had
less RKN’s present in the root tissue. R. reniformis root tissue populations at 90 DAP
were greatest in ‘SP-6’, with all other experimental entries including ‘Carnivor’ having
lower populations. No significance was observed in 90 DAP soil population data for
RKN’s.
Discussion
The primary objective of this study was to screen Fusarium wilt resistant cucurbit
rootstocks for resistance to RKN’s and R. reniformis. Through all four experiments the
RKN susceptible control ‘Carnivor’ maintained a higher RGI than all treatments
excluding ‘USVL-482351’ at the 2015 and 2016 spring 90 DAP sampling. Entries
‘Bulldog’, ‘SP-6’, ‘USVL-246’, and ‘USVL-252’ consistently exhibited less root galling
than ‘Carnivor’ at both 60 and 90 DAP sampling intervals in all seasons. While not
always significant, separation also occurred between ‘USVL-482351’ and other USVL
40
lines. 30 DAP RGI data was not reported due to occurrence of galling in only two experiments and no significant differences.
Soil populations were not consistently different between rootstocks for
Meloidogyne species. For RKN’s, the fall 2015 experiment observed the greatest populations in ‘USVL-482351’ instead of the control. Populations extracted from soil for
R. reniformis however demonstrated significance in both 2015 and 2016 fall experiments. Fall 2015 R. reniformis soil data had a similar trend to the RKN data with
‘USVL-482351’ having greatest density of nematodes in soil affiliated with the rootstock.
Fall of 2016 data found the highest observed populations of R. reniformis on ‘Carnivor’,
which may correspond with ‘USVL-482351’ being absent in that experiment.
Nematode numbers extracted from 10 grams of root tissue was far more
important for RKN’s than for R. reniformis. All three experiments where root tissue was
collected demonstrated a very similar trend where the root-knot susceptible cultivar
‘Carnivor’ contained a greater number of RKN’s than other lines. Minor variation only
occurred in this trend during the fall 2016 trial where ‘SP-6’ demonstrated similarity to
both the control and other resistant entries. R. reniformis root population data was
significant in both fall experiments but did not demonstrate any discernable trend. Fall
2015 reniform root data demonstrated the highest populations in ‘USVL-482351’ which
was similar the control, and ‘SP-6’ had the highest observed populations in fall 2016
with no reniform nematodes present in the control. We believe the spatial and overall
low presence of R. reniformis in the root tissue is due largely to its feeding habit. The
reniform nematode is a semi-endoparasite meaning that only the anterior body portion
is inside the root cortex and the posterior section of its body is exposed (Wang, 2007).
41
The triple wash procedure performed on roots prior to collecting root samples may have incidentally removed both sedentary reproductive females and vermiform juveniles.
Experimental data may indicate tolerance or vertical resistance in one variety
(Rohde, 1972). During fall 2016, ‘SP-6’ had similar populations of RKN’s and the
highest populations of R. reniformis when compared to ‘Carnivor’. Even with nematode
presence in the root tissue it maintained little to no galling. While the nematodes were
able to establish feeding sites and penetrate the root tissue some stage of their lifecycle
was interrupted where gall formation could not occur.
‘SP-6’ which is a Fusarium wilt resistant pollenizer for seedless watermelon
production, demonstrated low root gall index and some degree of tolerance or
resistance. ‘SP-6’ is not currently utilized as a rootstock but the resistance
demonstrated in this experiment makes it a viable candidate for rootstock use when
both Fusarium wilt and RKN’s are present in a field. Another advantage to ‘SP-6’ over
the other lines in this experiment is its’ commercially availability. While strong resistance
has been isolated within the USVL lines, only the variety ‘Bulldog’ is close to being
commercially available.
One of the more interesting results from this experiment is the presence of
Rotylenchulus reniformis in the root tissue and surrounding soil affiliated with the
rootstocks. In the southeastern United States it is generally thought of as a major
economic pest of cotton or soybeans. The literature is deficient on the economic impact
of R. reniformis on watermelon and other cucurbit crops. Given the high populations in
the soil affiliated with the rootstocks further research and a possible reconsideration of
its pathogenicity on cucurbit rootstocks should be made.
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Grafting has potential to be another management tool for soil-borne diseases available to growers. Two of the biggest hindrances to widespread utilization of grafting are cost (around three times as much as a non-grafted plant) and industrial production of grafted plants. Selecting rootstocks for resistance to Fusarium wilt and plant parasitic nematodes will not only reduce the cost of additional control measures but will help ensure host resistance does not degrade. Literature states numerous nematode/disease interactions and synergistic disease complexes, which sometimes cannot be replicated. The possibility of comparable losses to non-grafted transplants due to degradation of Fusarium wilt resistance signifies the importance of incorporating plant parasitic nematode resistance. For grafting to be an economically viable option, multiple forms of resistance and other added benefits need to be included to ensure greater return on invested capital.
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60 Day Root Gall Index 4 A a 3.5 Fall 2015 3 Z Spring 2016 2.5
10) Fall 2016 - 2 B
RGI (0 RGI 1.5 b 1 b b C 0.5 A C b * Y * Y Y b Y * * Y 0
Figure 2-1. Root gall index (RGI) collected 60 days post planting for selected cucurbit rootstocks from research conducted during fall of 2015 and spring and fall of 2016 in Quincy, FL.. Means not followed by the same letter are significantly different at P=0.05 by Fisher’s LSD. Means were compared within the season. * used when rootstock was not represented in that experiment.
90 Day Root Gall Index 7 Z 6 a Spring 2015 Fall 2015 z 5 Spring 2016 10)
- 4 Fall 2016 3
RGI (0 RGI A b 2 A Y 1 Y Y B * B c y * * Y y B c y B c Y y * * y 0
Figure 2-2. Root gall index (RGI) collected 90 days post planting for selected cucurbit rootstocks from research conducted during spring and fall of 2015 and 2016 in Quincy, FL. Means not followed by the same letter are significantly different at P=0.05 by Fisher’s LSD. Means were compared within the season. * used when rootstock was not represented in that experiment.
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90 Day Meloidogyne spp. Soil Populations 120 A 100 Fall 2015 NS Spring 2016 80 Fall 2016 60 B NS 40
20 Nematodes / 100 cc soil cc of / 100 Nematodes * B * B B * * 0
Figure 2-3. Nematode populations from soil associated with selected cucurbit rootstocks from research conducted in Quincy, FL during the fall of 2015 and spring and fall of 2016. Meloidogyne spp. J2 populations recovered from 100 cc of soil at 90 DAP. Means not followed by the same letter are significantly different at P=0.05 by Fisher’s LSD. Means were compared within season. * used when rootstock was not represented in that experiment.
90 Day Meloidogyne spp. Root Tissue
250 Populations A Fall 2015 200 Spring 2016 150 Fall 2016 Z B 100 a ZY 50 b b Y * B b * B b Y B b Y * * Y 0 Nematodes / 10 g tissue of root Nematodes
Figure 2-4. Nematode populations from roots of selected cucurbit rootstocks from research conducted in Quincy, FL during fall of 2015 and spring and fall of 2016. Meloidogyne spp. J2 populations recovered from 10 g. of root tissue at 90 DAP. Means not followed by the same letter are significantly different at P=0.05 by Fisher’s LSD. Means were compared within season. * used when rootstock was not represented in that experiment.
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90 Day Rotylenchulus reniformis Soil Populations 700 A 600 Fall 2015 500 Spring 2016 a 400 Fall 2016 B 300
200 b b b b 100 NS B Nematodes / 100 cc soil cc of / 100 Nematodes * B * B b * * 0
Figure 2-5. Nematode populations from soil and roots of selected cucurbit rootstocks from research conducted in Quincy, FL during fall of 2015 and spring and fall of 2016. Vermiform R. reniformis populations recovered from 100 cc of soil at 90 DAP. Means not followed by the same letter are significantly different at P=0.05 by Fisher’s LSD. Means were compared within the season. * used when rootstock was not represented in that experiment.
90 Day Rotylenchulus reniformis Root Tissue Populations 60 A Fall 2015 50 AB 40 Spring 2016 a Fall 2016 30 20 10 NSb * B b * B b B b * * b 0 Nematodes / 10 g tissue of root Nematodes
Figure 2-6. Nematode populations from soil and roots of selected cucurbit rootstocks from research conducted in Quincy, FL during fall of 2015 and spring and fall of 2016. Vermiform R. reniformis populations recovered from 10 g. of root tissue at 90 DAP. Means not followed by the same letter are significantly different at P=0.05 by Fisher’s LSD. Means were compared within the season. * used when rootstock was not represented in that experiment.
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CHAPTER 3 ASSESSMENT OF WATERMELON ROOTSTOCK RESISTANCE TO FUSARIUM OXYSPORUM F.SP. NIVEUM WHEN EXPOSED TO MELOIDOGYNE INCOGNITA
Introduction
Control of soil-borne pathogens like Fusarium oxysporum f.sp. niveum (FON) and southern root-knot nematode (Meloidogyne incognita) in vegetable plasticulture was historically achieved largely through methyl bromide application. Methyl bromide was found to be a causal agent in the depletion of the ozone layer and was phased out of general use in 2005 (United States Environmental Protection Agency, 2017). The ban on methyl bromide was estimated to have a significant economic impact, costing consumers and producers at least $1 billion annually (USDA ERS, 2000). The loss of this efficacious fumigant has directed research towards alternate methods of soil-borne pathogen control.
One alternative control method involves grafting a high yielding susceptible scion onto a rootstock with appropriate resistance. Studies in pepper has shown that grafting a susceptible bell pepper scion on to root-knot nematode resistant rootstocks effectively reduced root gall index (RGI) ratings and demonstrated increased fruit weight over the control (Kokalis-Burelle et al., 2009). Similar work conducted in cantaloupe (Cucumis melo) demonstrated significant reduction in RGI, reproductive factor (Rf), and J2 recovered from soil when grafted on a resistant rootstock (Guan et al., 2014). Grafting watermelon onto resistant rootstocks has shown reduction of Fusarium wilt incidence and increased yields (Keinath and Hassell, 2014).
A major point of concern involving FON and root-knot nematodes is that these pathogens often occur as co-contaminates in the soil. Previous research reported C. maxima X C. moschata hybrids have susceptibility to root-knot nematodes and can
47
sometimes perform comparably to the non-grafted control with respect to root galling
(Giné et al., 2017; Thies et al., 2010). Research has demonstrated that when both
Meloidogyne spp. and Fusarium oxysporum are present in the same area, Fusarium severity increases (Mai and Abawi, 1987; Powell, 1971a, 1971b; Taylor, 1979). A
Synergistic effect between both pathogens can potentially compromise host plant resistance. Research in tomato demonstrated that a co-inoculation of F. oxysporum f.sp. lycopersici and M. incognita resulted in 100% wilt symptoms in Fusarium resistant cv. ‘Chesapeake’, and F. oxysporum f.sp. lycopersici with M. hapla produced 66% wilt symptoms (Jenkins and Coursen, 1957). Results do however vary between experiments. Similar work conducted in tomato demonstrated no decrease in host resistance when subjected to a co-inoculation of F. oxysporum and M. incognita (Abawi and Barker, 1984). Additional research in cotton observed no pronounced synergistic effect when F. oxysporum and M. incognita were co-inoculated (Starr et al., 1989). Little to no work has been conducted in watermelon (Citrullus lanatus) concerning the impact on host resistance to FON. The purpose of this research was to determine if rootstock resistance to FON is compromised when co-inoculation with Meloidogyne incognita occurs in a watermelon cropping system.
Materials and Methods
Experiments were conducted in the spring seasons of 2016 and 2017 at the
North Florida Research and Education Center (NFREC) located in Quincy, Florida. The soil type at NFREC is a Dothan-Fuquay complex, which is a fine, sandy loam (Fine- loamy, kaolinitic, thermic Plinthic Kandiudults - Loamy, kaolinitic, thermic Arenic Plinthic
Kandiudults). Soil was cultivated to a depth of 25 cm. prior to bed formation, rows were spaced 2.44 meters apart, and beds were 20.3 cm tall and 76.2 cm wide. Raised beds
48
were covered with black polyethylene mulch and drip tape was deployed concurrently
with the mulch. Non-grafted plants were started from seed in 72 cell transplant trays in a peat based potting media. Grafted plants were produced using the one cotyledon method (Hassell et al., 2008). The seedless watermelon cultivar ‘Fascination’ (Syngenta
Seeds, Boise, ID) (C. lanatus) was the scion used in these experiments. ‘Fascination’
has intermediate resistance to FON race 1, but no marketed resistance to RKN’s or
FON race 2 (Syngenta, 2015) . Two rootstocks were selected for this experiment.
‘Bulldog’ (C. lanatus var. citroides) (United States Vegetable Lab (USVL), Charleston,
SC), demonstrating resistance to both RKN’s and FON races 1 and 2, and ‘Carnivor’ (C.
maxima x C. moschata) (Syngenta Seeds, Boise, ID) with susceptibility to RKN’s and
resistance to FON races 1 and 2. Grafting treatments included non-grafted
‘Fascination’, ‘Fascination’ grafted onto ‘Bulldog’, and ‘Fascination’ grafted onto
‘Carnivor’. Ten plants were transplanted into each plot on 8 April, 2016 and 21 March,
2017 and spaced 91 cm apart. Three ‘SP-6’ (Syngenta Seeds, Boise, ID) pollenizer
plants were included in each plot.
Inoculation treatments used for this experiment consisted of a non-inoculated control
(NI), inoculations of FON race 2 (FW), M. incognita (MI), and a co-inoculation of both M.
incognita and FON race 2 (FWMI). All scion and rootstock combinations were
subjected to the treatments. In both 2016 and 2017 spring seasons each treatment was replicated four times. Inoculation of FON was conducted at planting with 1.5 g of
infected wheat kernels per plant hole. M. incognita inoculations were applied in a liquid
solution to seedlings just prior to planting with an inoculation rate of 2500 eggs per
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transplant. The experiment was arranged as a split-plot design with grafting treatment as main plot and inoculation treatment as sub plot.
Incidence and disease severity were quantified for Fusarium wilt with a visual rating system. Rate of incidence was recorded as a percentage and determined by number of infected plants out of 10 total plants. Percent severity was collected for each
infected plant in the plot then the average was recorded. Incidence and severity ratings
were taken 21 and 28 DAP in spring 2016. In 2017 incidence and severity was collected
at 21, 28, 35, 42 49, 56, 63, 70, and 77 DAP.
The root gall index (RGI) rating was used to quantify root system damage caused by root-knot nematodes. RGI uses a 0 – 10 scale where a 0 indicates no galling and 10 represents complete plant death (Bridge and Page, 1980). The use of 10 in the rating
system is somewhat subjective because without proper testing the causal agent in plant
death cannot be determined. To indicate prolific galling 9 is the more appropriate
number to use. Five plants per plot were collected for RGI. The plant populations in the
individual plots was crucial for Fusarium wilt ratings so RGI was collected only at the
end of the season (90 DAP).
Watermelon fruits were harvested one time at the end of the 2017 season. Fruit
were individually weighed and a subsample of five fruits per plot were selected for fruit
quality analysis. 2016 yield data was not collected.
Data were subjected to analysis of variance using SAS version 9.4. When
appropriate, means were separated using Duncan’s Multiple Range Test with a P= 0.05.
Results
In spring of 2016, ‘Bulldog’ and ‘Fascination’ demonstrated resistance in terms of
less root galling than ‘Carnivor’ at the 90 DAP sampling interval. Fusarium wilt incidence
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and severity on 4/29/2016 was greater in ‘Fascination’ and ‘Bulldog’ than ‘Carnivor’.
Sampling on 5/5/2016 observed ‘Carnivor’ and ‘Bulldog’ with lower Fusarium wilt incidence and severity than ‘Fascination’.
In spring of 2017, ‘Carnivor’ demonstrated greater root galling at 90 DAP when compared to ‘Fascination’ and ‘Bulldog’. No significant difference in total yield was observed when separated by variety. Total yield was greatly reduced in plots inoculated with FON compared to non-inoculated plots. On 4/11/2017 Fusarium incidence and severity was greatest in ‘Bulldog’ and lower in ‘Fascination’ and ‘Carnivor’. Percent incidence on 4/18/2017 occurred least in ‘Carnivor’ and similarly in ‘Fascination’ and
‘Bulldog’. Fusarium wilt incidence and severity was greatest in ‘Fascination’ at the
4/25/2017 sampling interval compared to ‘Carnivor’ and ‘Bulldog’. On 5/23/2017
Fusarium incidence and severity was least in ‘Bulldog’ with similarity between ‘Carnivor’ and ‘Fascination’. At the 6/6/2017 sampling interval the greatest Fusarium incidence and severity was demonstrated on ‘Fascination’ with lesser rates on ‘Bulldog’ and
‘Carnivor’. No significance for incidence or severity occurred on sampling intervals 5/2,
5/8, 5/16 and 5/31 in spring 2017.
Discussion
The primary objective of this experiment was to test the retention of FON
resistance when both M. incognita and FON race 2 were both present in the same crop.
Both seasons saw Fusarium wilt symptomology present in plots where FON race 2
inoculum was not applied as a treatment. Root galling caused by Meloidogyne spp. also
appeared in plots where no root-knot egg inoculant was applied. However, both occurred at very low incidence. We believe this was due to use of a non-sterilized field.
Given the field conditions and presentation of both Meloidogyne spp. and FON in plots
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that did not receive those treatments, all treatment and rootstock combinations potentially contain varying levels of both pathogens. ANOVA revealed that there were no significant interactions between nematode inoculation and fusarium inoculation for fusarium wilt incidence. Fusarium wilt symptomology was observed on ‘Carnivor’ and
‘Bulldog’ which have resistance to both FON race 1 and 2. While not true at every
sampling interval, as a general trend ‘Carnivor’ demonstrated greater FON resistance
than ‘Bulldog’.
Fusarium wilt incidence and severity were not observed heavily in non-grafted,
susceptible, ‘Fascination’ plots and overall field pressure was low in both experiments.
A potential cause could be that our inoculum load was not high enough to cause
consistent and successful infection. Additionally both spring seasons experienced
prolonged periods of drought and drip irrigation may not have maintained an adequate
soil moisture level for the pathogen to thrive.
2017 yield data demonstrated no significant difference when separated by
variety. Concerning yield, no significant difference was exhibited with MI or FWMI
inoculations. Additionally, no interaction between FW and MI was observed with
ANOVA. Yield was greater in treatments that did not receive FW inoculations at P=.10,
making FW the only significant factor involving yield reduction.
‘Fascination’, the FON race 2 susceptible cultivar selected for this trial
demonstrated significantly lower RGI ratings than the M. incognita susceptible rootstock
‘Carnivor’ in both 2016 and 2017 seasons. RGI ratings for ‘Fascination’ are comparable
to the root-knot resistant rootstock, ‘Bulldog’. Separation in both experiments
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demonstrates ‘Fascination’ possesses some degree of non-marketed resistance to root- knot nematodes.
An interesting phenomenon was observed in plots that were planted with transplants inoculated with 2500 M. incognita eggs. When root systems were collected for rating RGI the observation was made that root galling only occurred in a 10-15 cm diameter area in the root system’s central point. Outside of the galled region, roots were clean and free of nematode damage. While this was not observed on every inoculated root system, frequency of occurrence could not go unnoticed. In a field setting, cucurbit root growth may be too aggressive for this inoculation method. Another potential cause
could be movement restriction due to water applied by drip irrigation. Water applied
from emitters covers only a portion of the root zone which could prevent further
advancement in the root system.
The potential for disease complex formation and degradation of host resistance
to FON races 1 and 2 on cucurbit rootstocks is still a possibility. Complex formation can
sometimes be circumstantial and vary as previously demonstrated in the literature
(Abawi and Barker, 1984; Mai and Abawi, 1987; Powell, 1971a). No interaction between
MI and FW occurred in this experiment for all three varieties. During both experiments
‘Carnivor’, the FON resistant and root-knot nematode susceptible rootstock demonstrated the greatest root gall index ratings and low expression of Fusarium wilt symptoms. ‘Carnivor’ is not representative of all cucurbit rootstocks, but performed similarly to other C. maxima x C. moschata varieties with root-knot nematode susceptibility (López-Gómez et al., 2016; Thies et al., 2012, 2010). In terms of this
53
experiment, no definite or significant degradation of host resistance to FON race 2 was exhibited.
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90 Day Root Gall Index 4.5 A 4 3.5 a 2016
3 2017 10)
- 2.5 2 RGI (1 RGI 1.5 1 B 0.5 B b b 0 Carnivor Fascination Bulldog
Figure 3-1. Root gall index (RGI) for selected varieties for spring 2016 and 2017 in Quincy, FL at 90 days after planting (DAP). Means not followed by the same letter are significantly different at P=0.05 by Duncan’s Multiple Range Test. Means were compared within a year.
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2016 Total Daily Rainfall 12.00
10.00
8.00
6.00
4.00 Rainfall (cm)Rainfall
2.00
0.00
Figure 3-2. 2016 daily rainfall recorded in centimeters from Quincy, FL
2016 Soil and Air Temperature Averages 30.0 C) ° 25.0
20.0 Temperature ( Temperature
15.0
Average Air Temperature 60 cm Average Soil Temperature 10 cm
Figure 3-3. 2016 average daily soil temperatures at a depth of 10 cm below and air temperatures 60 cm above ground level recorded in degrees Celsius from Quincy, FL.
56
Table 3-1. Fusarium wilt incidence expressed as percent incidence collected in 2016 in Quincy, FL. Data were compared within sampling date. Means not followed by the same letter are significantly different at P=0.05 by Duncan’s Multiple Range Test.
2016 Fusarium Wilt Incidence
Sampling Interval Variety 4/29 5/5 Carnivor 1.397 B 2.582 B Fascination 13.062 A 27.995 A Bulldog 11.365 A 8.427 B
Table 3-2. Fusarium wilt severity expressed as percent severity collected in 2016 in Quincy, FL. Data were compared within sampling date. Means not followed by the same letter are significantly different at P=0.05 by Duncan’s Multiple Range Test.
2016 Fusarium Wilt Severity
Sampling Interval Variety 4/29 5/5 Carnivor 1.128 B 1.247 B Fascination 6.262 A 5.643 A Bulldog 5.890 A 2.318 B
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2017 Yield Weight by FON Inoculation 38000 A 36000 34000 32000 B 30000
Kg/Ha 28000 26000 24000 22000 20000 Non-Inoculated FON Inoculated
Figure 3-4. Watermelon fruit yield as affected by F. oxysporum f.sp. niveum (FON) inoculation in 2017 in Quincy, FL. Means not followed by the same letter are significantly different at P=0.10 by Duncan’s Multiple Range Test.
2017 Yield Weight by Variety 36000 34000 NS 32000 30000 28000 Kg/Ha 26000 24000 22000 20000 Carnivor Fascination Bulldog
Figure 3-5. Watermelon fruit yield as affected by variety, 2017 in Quincy, FL. Means not followed by the same letter are significantly different at P=0.05 by Duncan’s Multiple Range Test.
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2017 Total Daily Rainfall 8.00 7.00 6.00 5.00 4.00 3.00 Rainfall (cm)Rainfall 2.00 1.00 0.00
Figure 3-6. 2017 daily rainfall recorded in centimeters from Quincy, FL.
2017 Soil and Air Temperature Averages 30.0
C) 25.0 ° 20.0 15.0 10.0
Temperature ( Temperature 5.0 0.0 4-Apr 11- 18- 25- 2-May 9-May 16- 23- 30- 6-Jun Apr Apr Apr May May May
Average Air Temperature 60 cm Average Soil Temperature 10 cm
Figure 3-7. 2017 average daily soil temperatures at a depth of 10 cm below and air temperatures 60 cm above ground level recorded in degrees Celsius from Quincy, FL.
59
Table 3-3. Fusarium wilt incidence expressed as percent incidence collected in 2017 in Quincy, FL. Data were compared within sample date. Means not followed by the same letter are significantly different at P=0.05 by Duncan’s Multiple Range Test.
2017 Fusarium Wilt Incidence
Sampling Interval Variety 4/11 4/18 4/25 5/2 5/9 5/16 5/23 5/31 6/6 ‘Carnivor’ 0.625 B 2.5 B 0 B 0 NS 1.875 NS 1.875 NS 2.5 AB 4.375 NS 0 B ‘Fascination’ 4.375 B 6.25 AB 3.75 A 1.875 2.5 1.25 5.625 A 11.25 8.75 A ‘Bulldog’ 8.75 A 11.25 A 0 B 1.25 0 0 0 B 3.125 1.875 B
Table 3-4. Fusarium wilt severity expressed as percent severity collected in 2016 in Quincy, FL. Data were compared within sampling date. Means not followed by the same letter are significantly different at P=0.05 by Duncan’s Multiple Range Test.
2017 Fusarium Wilt Severity
Sampling Interval Variety 4/11 4/18 4/25 5/2 5/9 5/16 5/23 5/31 6/6 ‘Carnivor’ 0.625 B 8.44 NS 0 B 0 NS 6.875 NS 6.25 NS 6.25 AB 4.375 NS 0 B ‘Fascination’ 3.125 B 15.94 3.75 A 3.125 5.313 2.5 9.688 A 8.438 8.438 A ‘Bulldog’ 15.250 A 23.13 0 B 1.25 0 0 0 B 4.063 1.25 B
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CHAPTER 4 TRACKING SEASONAL MOVEMENT OF MELOIDOGYNE SPP. AND ROTYLENCHULUS RENIFORMIS THROUGH A NINETY DAY WATERMELON CROPPING SEASON
Introduction
Intensive vegetable production the United States involves the use of plastic mulch and soil fumigants. Fumigation in a plasticulture system can manage multiple crop pests including weeds, insects, nematodes, and soil-borne diseases. A point of concern when making a pre-plant application is the depth at which the fumigant is applied. Fumigation shanks used in United States vegetable production under
plasticulture almost exclusively apply fumigant at a depth of 30 cm. Available fumigants have lower vapor pressure, water affinity and poorer soil distribution both vertically and laterally when compared to the previous industry standard, methyl bromide (Candole et al., 2007; Desaeger et al., 2004; Munnecke and Van Gundy, 1979; Noling and Becker,
1994).
Previous research in cotton has shown that substantial populations of plant
parasitic nematode species can occur below the 20 cm plow layer. In Louisiana, Texas,
and Arkansas, 10 cotton fields surveyed were found to have more than half of the
Rotylenchulus reniformis population deeper than 36 cm. (Robinson et al., 2000).
Additional research on cotton (Robinson et al., 2005; Robinson et al., 2005)
demonstrated the actual depths where these nematodes survive and potentially thrive is
often much deeper than the top 30 cm of soil. Nematode populations at a depth of 120 –
135 cm from the soil surface reduced tomato yield by 11% in a spring experiment and
59% that following fall in Canadian greenhouse soil (Johnson and Mckeen, 1973).
Fumigation controls nematodes in the top 30 cm of soil which provides young plants
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with a “disease free” window, but does not provide full season protection (Yeates et al.,
1991). Work in tomato demonstrated that entire populations of Meloidogyne spp. moved at least 25 cm in a sandy soil profile in nine days and 50% of the population moved a distance of 50 cm in nine days (Prot, 1978). This movement in sandy soils can contribute to the rapid re-colonization of a fumigated area post treatment when a susceptible host is planted (Prot, 1980).
In 1927 a root study was conducted on dryland watermelon using the cultivar
‘Kleckly Sweet’. It was determined that root growth extended 1.2 m deep, with most roots occurring in the first 30 cm of soil (Weaver and Bruner, 1927). Research conducted in 2013 demonstrated similar results with rootstocks under polyethylene mulch producing the majority of root biomass in the first 30 cm of soil with roots still deeper in the profile (Miller et al., 2013). The majority of tomato root biomass when
grown under drip irrigation occurred at a depth range of 30-40 cm with some roots still
reported at a depth of 100 cm (Oliveira et al., 1996). The damage threshold level for
above ground dry weight of the watermelon cultivar ‘Royal Sweet’ for M. incognita was
estimated with an initial population (Pi) equal to 122 eggs/100 cm3 of soil, 1.6 galls on
bioassay roots/100 cm3 of soil, or 3.6 J2/100 cm3 of soil (Xing and Westphal, 2012). A
damage threshold for tomatoes is mentioned for three Meloidogyne spp. represented as
Juveniles/100 cc of soil: M. javanica (<0.2 J2/100 cc soil), M. hapla (<4 J2/100 cc soil)
and M. incognita (<4 J2/100 cc soil) (Barker and Olthof, 1976).
Control of Rotylenchulus reniformis with fumigation has shown increased yields
in okra, tomatoes, lettuce, and squash by 19%, 13%, 57% and 69%, respectively
(Heald, 1978). R. reniformis historically is not viewed as a major watermelon crop pest
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so a damage threshold has yet to be established. Watermelon has however been a proven host for R. reniformis (Robinson et al., 1997) and research in Brazil has shown reduced watermelon shoot biomass in fields infested with R. reniformis (Torres et al.,
2005). The objective of these experiments was to determine seasonal movement of
Meloidogyne spp. and Rotylenchulus reniformis within the soil profile during a typical 90 day watermelon cropping season.
Materials and Methods
Experiments were conducted in the spring seasons of 2016 and 2017 and fall of
2016 at the North Florida Research and Education Center (NFREC) located in Quincy,
Florida. The soil type in this experiment was Dothan-Fuquay, which is a fine, sandy loam (Fine-loamy, kaolinitic, thermic Plinthic Kandiudults - Loamy, kaolinitic, thermic
Arenic Plinthic Kandiudults). Soil was cultivated to a depth of 25 cm prior to bed formation. Rows were spaced 2.44 meters apart, and beds were 76.2 cm wide and 20.3 cm tall. Spring experiments were planted on black polyethylene mulch, while the fall experiment was planted on white-on-black polyethylene mulch. Irrigation water was delivered to the trial through one offset drip tape per bed. The experiment was located in a field with historically high populations of Meloidogyne spp. (RKN) and
Rotylenchulus reniformis. The interspecific hybrid squash (C. maxima x C. moschata)
‘Carnivor’ (Syngenta Seed, Boise, ID) was chosen as planting material. ‘Carnivor’ is a cucurbit rootstock with marketed resistance to Fusarium oxysporum f.sp niveum races 1 and 2. C. maxima X C. moschata hybrids like ‘Carnivor’ have documented susceptibility to root-knot nematodes (López-Gómez et al., 2016; Thies et al., 2012, 2010). Seedlings were grown in 128 cell transplant trays and transplanted to their plots on 4/22/16 (spring
2016), 8/19/16 (fall 2016), and 5/19/17 (spring 2017), three weeks post seeding at a
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spacing of 91 centimeters. Fifteen seedlings were planted in each 18.3 m. plot and the
experiment was arranged as a randomized complete block design (RCBD) with four
replications. Sampling was conducted at three different intervals; pre-plant, 45 days
after planting (DAP), and 90 DAP.
Three 5 cm diameter, 120 cm long core samples were collected from each plot at
each collection date using a model 5096 Geoprobe system (supplier information). Pre-
plant cores were taken from the bed centers while mid and season’s end cores were
taken adjacent to the plant base to collect root zone soil. Once collected, the cores were
segmented in four 30 cm sections (0-30, 30-60, 60-90, 90-120 cm). Each sampling date
had a collection of 12 samples for each depth, for a total of 48 samples for all depths.
From each 30 cm sample, a sub-sample of 100 cc of soil was collected and nematodes
were extracted using the Baermann funnel technique (EPPO, 2013). After 48 hours
nematodes were identified and counted.
Data was first log transformed and subjected to a one way ANOVA. Populations
were analyzed independent of each other to produce final averages for RKN’s and R.
reniformis at all four depth intervals (0-30, 30-60, 60-90, 90-120 cm). Once averaged
the data was subjected to a Fischer’s LSD Test with a P-value of 0.05 in SAS.
Results
During spring of 2016 at the pre-plant sampling the highest concentration of R.
reniformis occurred at the 30-60 and 60-90 cm depths. The numerically highest
occurrence of RKN’s occurred at the 30-60 cm depths with similarity at the 0-30 and 60-
90 cm depths. Mid-season R. reniformis had a homogenous distribution with no
significance difference at depth, while populations of RKN’s were greater in the 0-30 cm
range than below 60 cm. End of season sampling showed RKN’s numerically highest at
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the 60-90 cm depth but statistically similar to 0-90 cm. End of season R. reniformis populations were the greatest in the 0-90 cm range.
During fall of 2016 at the pre-plant sampling RKN’s were similar in the 0-90 cm range with population in the 30-60 and 60-90 cm range being statistically greater than
90-120 cm. The numerically highest population of R. reniformis was found in the 30-60 cm section as well at pre-plant with similarity to both the 0-30 and 60-90 cm sections. At the mid-season sampling both species of plant parasitic nematodes had greater densities in the 30-90 cm depth range than 0-30 cm. End of season R. reniformis populations were greater at the 30-60 cm range than 0-30 cm with similarities between
60-90 and 90-120 depths and the 0-30 depth. RKN’s were found in greater concentrations at depths 30-60 and 60-90 cm than 90-120 cm. Populations were similar at 0-30 cm and 90-120 cm at season’s end.
During spring of 2017 at the pre-plant sampling interval both species of plant parasitic nematodes were found in greater concentrations at depths of 30-60 and 60-90
cm than 0-30 cm. Mid-season sampling showed greatest populations of RKN’s
occurring in the 30-60 cm depth while R. reniformis was numerically highest at 30-60
cm with similarity to 60-90 cm. At the end of season, RKN’s were at their highest
concentration in the 0-30 cm range while R. reniformis populations were stratified
between 0-90 cm with the highest numerical occurrence at 30-60 cm.
Discussion
The interspecific hybrid rootstock ‘Carnivor’ was chosen for this experiment
based on its commercial availability and susceptibility of C. maxima x C. moschata
hybrids to plant parasitic nematodes (Thies et al., 2010). ‘Carnivor’ is currently marketed
as resistant to Fusarium wilt (F.oxysporum f.sp. niveum) races 1 and 2 and Verticillium
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wilt (Verticillium dahliae) but does not have marketed resistance to root-knot nematodes
(Meloidogyne spp.) (USDA-NIFA, 2015). Given the possibility of use in a commercial watermelon production system it fit well as a model plant. The primary objective of this experiment was to determine where plant parasitic nematode species were present before, during, and at the end of a watermelon season.
In many areas of the United States, soil fumigants are applied in plasticulture production with 30 cm shanks. This aids in the management of soil-borne pests and pathogens in the top 30 cm. of soil. Current fumigant alternatives do not control pathogens vertically or laterally as well as methyl bromide (MeBr). Research in
California demonstrated the downward movement of MeBr by finding significant detectable levels at depths of 7-9 feet (~213-274 cm) below the point of application
(Kolbezen et al., 1974). The depth achieved by MeBr is lacking in modern fumigants, which places many soil-borne pathogens outside the zone of control. Many researchers historically have not considered the spatial distribution of nematode populations in the soil profile for this very reason. Given that the populations of both RKN’s and R. reniformis at 30-90 cm were similar to or greater than those at 0-30 cm range prior to planting, the inference could be made that applying fumigant at a depth of 30 cm is only providing marginal nematode control. In Florida strawberry, it was demonstrated that with deep shank (40 cm) and deep drip applications of 1,3-dichloropropene (Telone;
140-168 L/ha), soil population densities of sting and root-knot nematode were significantly lower at seasons end and increased yield 9-29% when compared to grower fumigation standard (Noling et al., 2016). In cotton, fumigation at 60-120 cm. deep increased cotton yields by 68% compared to the non-fumigated control (Westphal and
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Smart, 2003). Westphal also noted that population densities in the 0-120 cm. horizon provided a more accurate estimation of potential crop damage than 0-30 cm.
The southern root-knot nematode (Meloidogyne incognita) in addition to other
RKN’s are known parasites of watermelon. Conversely, far less is known concerning the
reniform nematode (Rotylenchulus reniformis) and watermelon. On a global scale, R.
reniformis is considered the second most important nematode affecting vegetables
crops, but is often overlooked where it occurs concomitantly in soil with RKN’s (Luc et
al., 2005). R. reniformis demonstrated a pathogen/host relationship with C. lanatus
(Robinson et al., 1997), but the experiment was only a host test. Additionally, reduction
of watermelon shoot biomass has been documented in Brazil (Torres et al., 2005), but
minimal research has occurred in the United States. Given that in both 2016 and 2017
spring seasons in this experiment, R. reniformis populations exponentially increased
between the 0 and 90 day sample dates at all soil profile depths demonstrates
reproductive capabilities. While not currently viewed as a major soil-borne pest of
cucurbits in the United States, it deserve further research. While a watermelon rootstock
was used as the model crop for this experiment, the data collected is more than
applicable to other fruits or vegetables grown in a plasticulture system where nematode pressure inhibits yield.
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Meloidogyne spp. Pre-plant Soil Populations 1400 A 1200 1000 800 600 A AB 400 200 A AB Nematodes / 100 cc Soil cc of / 100 Nematodes AB A B B AB C BC 0 Spring 2016 Fall 2016 Spring 2017
0-30 cm 30-60 cm 60-90 cm 90-120 cm
Figure 4-1. Meloidogyne spp. populations from soil pre-planting during spring and fall of 2016 and spring of 2017 from experiments conducted in Quincy, FL. Means are to be compared within season.
R. reniformis Pre-plant Soil Populations 1000 A
800
600 AB 400 A 200 AB B B A A A
Nematodes / 100 cc Soil cc of / 100 Nematodes B B B 0 Spring 2016 Fall 2016 Spring 2017
0-30 cm 30-60 cm 60-90 cm 90-120 cm
Figure 4-2. R. reniformis populations from soil pre-planting during spring and fall of 2016 and spring of 2017 from experiments conducted in Quincy, FL. Means are to be compared within season.
68
Meloidogyne spp. Soil Populations 45 Days after Planting 1200 A 1000
800 A AB 600
400 BC A AB 200 BC
Nematodes / 100 cc Soil cc of / 100 Nematodes B C C B C 0 Spring 2016 Fall 2016 Spring 2017
0-30 cm 30-60 cm 60-90 cm 90-120 cm
Figure 4-3. Meloidogyne spp. populations from soil 45 days after planting during spring and fall of 2016 and spring of 2017 from experiments conducted in Quincy, FL. Means are to be compared within season.
R. reniformis Soil Populations 45 Days after Planting 1200 A
1000
800
600 A
400 AB
200 AB B Nematodes / 100 cc Soil cc of / 100 Nematodes NS C BC C 0 Spring 2016 Fall 2016 Spring 2017
0-30 cm 30-60 cm 60-90 cm 90-120 cm
Figure 4-4. R. reniformis populations from soil 45 days after planting during spring and fall of 2016 and spring of 2017 from experiments conducted in Quincy, FL. Means are to be compared within season.
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Meloidogyne spp. Soil Populations 90 Days after Planting 400 A 350 300 A
250 A 200 150 A A B 100 AB AB
Nematodes / 100 cc Soil cc of / 100 Nematodes 50 B B B B 0 Spring 2016 Fall 2016 Spring 2017
0-30 cm 30-60 cm 60-90 cm 90-120 cm
Figure 4-5. Meloidogyne spp. populations from soil 90 days after planting during spring and fall of 2016 and spring of 2017 from experiments conducted in Quincy, FL. Means are to be compared within season.
R. reniformis Soil Populations 90 Days after Planting 6000 A 5000
4000
3000 A A A 2000 A A 1000 A Nematodes / 100 cc Soil cc of / 100 Nematodes B B AB AB B 0 Spring 2016 Fall 2016 Spring 2017
0-30 cm 30-60 cm 60-90 cm 90-120 cm
Figure 4-6. R. reniformis populations from soil 90 days after planting during spring and fall of 2016 and spring of 2017 from experiments conducted in Quincy, FL. Means are to be compared within season
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CHAPTER 5 CONCLUSION
The phase out of methyl bromide left many watermelon and cucurbit producers without a viable option to control soilborne pathogens like F. oxysporum f.sp. niveum and plant parasitic nematodes. Variable efficacy and soil distribution, combined with off-
putting odors and various issues associated with available fumigants indicate the need
for further incorporation of additional management tactics.
A concern with using grafted plants is absence of resistance to both Fusarium
wilt and plant parasitic nematodes. Parasitic nematodes including Meloidogyne spp. and
Rotylenchulus reniformis threaten host resistance to Fusarium wilt through a
nematode/disease complex. No perceived degradation of host resistance was observed
in Fusarium resistant, root-knot nematode susceptible cultivar ‘Carnivor’ in this experiment. Fusarium wilt resistant rootstock lines and varieties trialed in this experiment demonstrated strong resistance to plant parasitic nematodes. The greatest concern for a producer interested in using grafted plants for soilborne pest management is the cost. Currently grafted transplants cost almost three times that of a non-grafted plant. Grafting technology in recent years has become more automated and plant cost is falling, meaning grafting may become a more common management tool in the future.
Plant parasitic nematodes are commonly managed in a watermelon system with
1, 3-dichloropropene, a drip applied or shank applied fumigant nematicide, or a different
fumigant. Many of the nematicidal compounds on the market today do not possess the soil mobility of methyl bromide. Poor soil movement both vertically and laterally reduces compound efficacy against nematodes. This research and previous experiments on other crops demonstrates significant nematode population at depths far out of reach of
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currently available methyl bromide alternatives. Given the high populations at depth and nematode migration capabilities, a reassessment of fumigant application depth for plasticulture systems should be addressed.
In order for grafting to become an economically viable option for soilborne pathogen management two aspects must improve. Overall cost of grafted plants needs to decrease to a point where implementation applies to more producers than just those with high Fusarium wilt pressure. Further incorporation of multiple disease resistances to create better disease resistant packages will also improve value, lower planting cost, and provide growers with better return on invested capital. Grafting as a supplement or alternative to fumigation offers a glimpse at a promising future.
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BIOGRAPHICAL SKETCH
Cody Smith was born in 1991 to Steve Smith and Brenda Macrides and is a
Florida native from the town of Fort White. He graduated from Santa Fe High School in
Alachua, FL in 2009 and later received a Bachelor of Science in plant science at the
University of Florida in the spring of 2015. During his time as an undergraduate he worked for U.F.’s Agronomy Department, a landscape architect, a pecan/timber producer, and managed his own nuisance animal removal business. In the fall of 2015 he began his master’s program with the University of Florida’s Horticultural Sciences
Department under Dr. Joshua Freeman and graduated in the fall of 2017.
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