Factors affecting the efficacy of 1,3-Dichloropropene for plant-parasitic nematode management in Florida deep sandy soils

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Factors affecting the efficacy of 1,3-Dichloropropene for plant-parasitic nematode management in Florida deep sandy soils
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FACTORS AFFECTING THE EFFICACY OF 1,3-DICHLOROPROPENE FOR
PLANT-PARASITIC NEMATODE MANAGEMENT IN FLORIDA DEEP SANDY
SOILS












By

CLAUDIA RIEGEL


A DISSERTATION PRESENTED TO THE GRADUATE SCHOOL
OF THE UNIVERSITY OF FLORIDA IN PARTIAL FULFIL LNIENT
OF THE REQUIREMENTS FOR THE DEGREE OF
DOCTOR OF PHILOSOPHY

UNIVERSITY OF FLORIDA


2001




























To My Parents,

Jodo Antonio and Wilma Sanchez Riegel














ACKNOWLEDGE NI ENTS


Many wonderful people have crossed my path since I started college and I am

grateful to the many friends who have stood by my side during these years. I would like

to first to thank my advisor, Dr. Donald W. Dickson, for all the support, encouragement,

and advice he has given me the past several years. He has devoted many hours to

discussing my research, working in the field, and reading my papers. In turn, I have given

him more grey hair and spent his supply of green and purple pens. Dr. Dickson has

opened many doors for me and I will forever be grateful.

Thanks are expressed to my committee members Drs. R. D. Berger, L. T. Ou,

L.G. Peterson, G. C. Smart, Jr. and E. B. Whitty for their guidance and suggestions.

Special recognition goes to Dr. Peterson who has been enormously generous with his time

and efforts to move my project forward. He has gone beyond the call of duty to promote

and maintain interest in my project so that it would be funded by Dow AgroSciences LLC

each year. I thank him also for his support and encouragement professionally and for

providing me with the contacts to move successfully into the job market. I very much

value his advice and friendship. I thank Dow AgroSciences LLC for financially supporting

my assistantship and research for 4 years.

I thank Dr. Smart for spending enormous amounts of time teaching me nematode

anatomy and morphology and reading my dissertation. I thank Dr. K. Nguyen for









teaching me taxonomy and nematode identification. Dr. Nguyen always took time out of

his busy schedule to help me identify a nematode or answer my questions.

I thank Dr. Dickson's technical support staff and friends, Tom Hewlett, Reggie

Wilcox, and Tim Sheffield. They have been enormously helpful with my experiments; in

fact, I would not have been able to complete the work without them. They worked many

hours in the hot sun taking samples, transplanting seedlings, spraying, and worst of all

weeding. I appreciate all the times they stayed after hours to help me finish my field work.

Many friends have supported me throughout these past few years. I acknowledge

Janete Brito, Billy Crow, Socorro Hall, Hye-Rim Han, Liz Herrera, the McNelis family,

the Powers family, the Swienton family, and Marco Toapanta for their help. Finally, I

thank my family for all the love, encouragement, and support they have given me the past

four years: my parents, Joao and Wilma, my brother, Christian, and my relatives in Brazil,

Vera Lucia, Luiz Renato, Marcus, and Thais Cerchi and Chico, Dilma, Isis, and Kaue

Sanchez.














TABLE OF CONTENTS

Page

ACKNOW LEDGM ENTS ............................................... iii

LIST O F TA BLES ..................................... .. ............. viii

LIST OF FIGURES .................................................... xi

ABSTRACT ...................................................... xiii

CHAPTERS

1 IN TRO DU CTION ..................................................... 1

History of 1,3-Dichloropropene ....................................... 1
Biological Activity of 1,3-Dichloropropene ............................. 2
M ode of Action .................................................... 3
Chemical and Physical Properties of 1,3-Dichloropropene .................. 5
Soil Texture ................................................ 7
Soil Temperature ............................................ 9
Soil M oisture .............................................. 11
Time of Application, Seeding, or Transplanting Restrictions ............... 13
Enhanced Degradation ............................................ 15


2 THE EFFICACY OF 1,3-DICHLOROPROPENE FOR CONTROL OF
ROOT-KNOT NEMATODES WHEN APPLIED IN SOIL
ENHANCED FOR DEGRADING 1,3-DICHLOROPROPENE ............ 20

Introduction ...................................................... 20
M materials and M ethods ............................................. 22
Results and Discussion ............................................ 33









3 RATE RESPONSE OF 1,3-DICHLOROPROPENE FOR NEMATODE
CONTROL IN SPRING SQUASH IN DEEP SANDY SOILS ............. 45

Introduction .................................. .. .................. 45
M materials and M ethods ............................................. 46
R results ......................................................... 49
D discussion ...................................... ................. 56


4 COMPARISON OF CHISEL TYPE, SEALING, AND APPLICATION
DEPTH ON THE EFFICACY OF 1,3-DICHLOROPROPENE IN
DEEP SANDY SOILS ............................................. 60

Introduction ..................................................... 60
M materials and M ethods ............................................. 62
Results and Discussion ............................................ 73


5 COMPARISON OF DIFFERENT CHISEL TYPES FOR
1,3-DICHLOROPROPENE FUMIGATION IN DEEP SANDY SOILS ...... 82

Introduction ...................................................... 82
M materials and M ethods ............................................. 83
Results and Discussion ............................................ 85


6 THE EFFICACY OF 1,3-DICHLOROPROPENE IN COMPOSTED AND
NONCOMPOSTED SOIL .......................................... 90

Introduction ..................................................... 90
M materials and M ethods ............................................. 88
R results ......................................................... 9 1
D discussion ..................................................... 10 1

7 SUM M ARY ....................................................... 104








8 APPENDIX A. EFFICACY OF SEVERAL RATES OF
1,3-DICHLOROPROPENE, DIFFERENT METHODS OF
APPLICATION, AND THE USE OF PLASTIC MULCH FOR
SEALING AFTER FUMIGATION TO CONTROL ROOT-KNOT
NEMATODES IN FLORIDA DEEP SANDY SOILS

Introduction .................................................... 107
M material and M ethods ............................................ 109
Results and Discussion ........................................... 113


LIST OF REFERENCES ................................................ 119

BIOGRAPHICAL SKETCH ............................................ 131














LIST OF TABLES


Table Page
1-1. Chemical and physical properties of 1,3-dichloropropene .................... 6

2-1. Number of galls on tomato roots grown in soil with Meloidogyne
spp. in microplots amended with enhanced soil (previously
identified to degrade 1,3-dichloropropene at an
accelerated rate, autoclaved enhanced soil, and nonenhanced soil ........... .35

2-2. Number of eggs on tomato roots grown in soil infested with
Meloidogyne spp. in microplots amended with enhanced soil (previously
identified to degrade 1,3-dichloropropene at an accelerated rate,
autoclaved enhanced soil, and nonenhanced soil ......................... 36

2-3. Regression analysis used to interpret the effects of 1,3-dichloropropene
exposure periods on Meloidogyne spp. gall production on tomato
grown in microplots that were amended with enhanced soil (previously
identified to degrade 1,3-dichloropropene at an accelerated rate,
autoclaved enhanced soil, and nonenhanced soil ......................... 37

2-4. Regression analysis used to interpret the effects of 1,3-dichloropropene
exposure periods on Meloidogyne spp. egg production on tomato
grown in microplots that were amended with enhanced soil (previously
identified to degrade 1,3-dichloropropene at an accelerated rate
[Ou et al., 1995]), autoclaved enhanced soil, and nonenhanced soil ........ .38

2-5. Galls induced by Meloidogyne spp. second-stage juveniles (J2) that were
extracted from infested soil from microplots amended with enhanced
soil (previously identified to degrade 1,3-dichloropropene at
an accelerated rate, autoclaved enhanced soil, and nonenhanced soil. ........ 39

2-6. Comparisons of the number of Meloidogyne incognita second-stage
juveniles in tomato roots grown in 1,3-dichloropropene (1,3-D,
112 liters/ha) treated or untreated field soil previously identified as
enhanced for degrading 1,3-D at an accelerated rate,
and in soil without a history of 1,3-D fumigation........................ 43








2-7. Comparisons of the number of Meloidogyne incognita second-stage
juveniles in tomato roots grown in PCV tubes filled with enhanced
soil known for degrading 1,3-dichloropropene (1,3-D) at an
accelerated rate and with soil without a history of fumigation,
both of which were treated with 1,3-D (84 liters/ha). ..................... 44

3-1. Contrasts to compare gall indices of root-knot nematode on squash
and rates of 1,3-dichloropropene in trials 1 and 2......................... 54

4-1. Number of Meloidogyne spp. second-stage juveniles (J2) per 100 cm3 of
soil at harvest following fumigation with 1,3-dichloropropene
applied with three chisel types in three peanut trials conducted in 1998
(trials 1 and 2) and 1999 (trial 3) at the University of Florida Green
Acres Agronomy Research Farm located in Alachua County, Florida. ....... 75

4-2. Yield and incidence of root-knot nematode galls on peanut pods at
harvest in soil fumigated with 1,3-dichloropropene applied
with three chisel types in three peanut trials conducted in 1998 (trials 1
and 2) and 1999 (trial 3) at the University of Florida Green Acres
Agronomy Research Farm located in Alachua County, Florida .............. 76

4-3. Effects of broadcast or in-row fumigation of 1,3-dichloropropene
applied at 84 liters/ha with conventional chisels or parachisels on
fruit production and yield of squash (Cucurbita pepo), number of
second-stage juveniles of Meloidogyne spp. per 100 cm3, soil and
reproductive factors 34 days and 65 days after planting at the
University of Florida Green Acres Agronomy Farm located in
Alachua County, Florida in 1999.................................... 78

5-1. The number Meloidogyne spp. second-stage juveniles (J2) that penetrated
4-week-old tomato seedlings after in-row fumigation with
1,3-dichloropropene applied at 84 liters/ha with conventional chisels
(with and without disking), parachisels (with and without disking),
and subsurface hooded sweep chisels in 1998 and 1999. .................. 86

5-2. Orthogonal contrasts to compare the number of Meloidogyne spp.
per tomato root system from plants grown in untreated soil and in
soil fumigated with 84 liters of 1,3-dichloropropene/ha using
conventional chisels (with and without disking), parachisels (with
and without disking), and subsurface hooded sweep chisels in 1998 .......... 87








6-1. Physical and chemical properties of the noncomposted and composted
soil used in two trials testing the efficacy of 1,3-dichloropropene
in the control of root-knot nematodes.................................. 96

6-2. Gravimetric soil water content at the time of fumigation with
1,3-dichloropropene for soils taken from 15, 30, and 45 cm deep
in noncomposted and composted microplot soils for two trials. ............. 97


6-3. The effect of 1,3-dichloropropene (applied broadcast at the rate of
112 liters/ha) on root-knot nematodes as indicated by a tomato
root bioassay 7 weeks after transplanting in noncomposted and
com posted soil ...........................................

Al-1. Comparison of plant-parasitic nematode population densities per
100 cm3 soil and yield from soil broadcast fumigated with 56, 84,
112, and 168 liters per hectare of 1,3-dichloropropene. ...........

A 1-2. Comparison of plant-parasitic nematodes and yield following soil
fumigation with 1,3-dichloropropene applied by different methods
and at different depths ......................................


........ 98



....... 116



....... 117


A1-3. Comparison of plant-parasitic nematodes and yield from soil fumigated
with 1,3-dichloropropene and covered with industry standard
black polyethylene plastic film or clear very impermeable polyethylene
plastic film (VIF), disked and covered with plastic film, and disked only .... 118














LIST OF FIGURES


Figure page

2-1. Schematic of field plot layout used to evaluating the efficacy of
1,3-dichloropropene in enhanced and nonenhanced soil ................... 28

2-2. Schematic of screened mesh template used for placement of
1,3-dichloropropene in the enhanced and nonenhanced soil at two
different field sites ................................................ 29

2-3. Schematic of a plot with six tomato seedlings transplanted in a row
surrounded by a border of fumigated soil ............................... 30

2-4. PVC tubes constructed for the greenhouse study had an upper chamber
filled with fumigated soil and a lower chamber filled with heat-
pasteurized soil ................................................... 31

3-1. Effects of rates of 1,3-dichloropropene for control of Meloidogyne spp.
on the marketable yield and growth of squash produced in two trials
conducted at the University of Florida Green Acres Agronomy Farm
in 1999 ........................................................ 5 1

3-2. Effects of rates of 1,3-dichloropropene on root galling of squash induced
by Meloidogyne spp. 65 days after planting in two trials conducted at
the University of Florida Green Acres Agronomy Farm in 1999 ............. 52

3-3. Effects of rates of 1,3-dichloropropene on the density of Meloidogyne spp.
second-stage juveniles (J2) per 100 cm3 soil in two trials conducted at
the University of Florida Green Acres Agronomy Farm in 1999. ........... 55

3-4. Degree days for Meloidogyne spp. were based on maximum and
minimum soil temperatures 10 cm deep recorded from 23 April
to 23 June 1999 ................................................... 57

4-1. The conventional chisel used for in-row and broadcast fumigation of 1,3-
dichloropropene ............................................... 66








4-2. Parachisels used for in-row and broadcast fumigation of 1,3-dichloropropene. 67

4-3. A schematic of the parachisel used for fumigation with 1,3-dichloropropene .... 68

4-4. Schematic drawings of the parachisel in side and plan views................. 69

4-5. The subsurface hooded sweep chisel used for fumigation with
1,3-dichloropropene .............................................. 70

4-6. Root-knot nematode galling in squash, Cucurbitapepo, grown
in soil fumigated with 84 liters of 1,3-dichloropropene/ha applied
in-row and broadcast with conventional chisels and parachisels with
and w without disking.. .............................................. 81

6-1. The effects of increasing the exposure period of Meloidogyne incognita
to 112 liters of 1,3-dichloropropene/ha on the number of galls produced
per root system of tomato grown for 7 weeks in containers in soil
sampled from 0 to 15 cm deep...................................... 99

6-2. The effects of increasing the exposure period ofMeloidogyne incognita
to 112 liters of 1,3-dichloropropene/ha on the formation of galls
produced in tomato roots grown in soil sampled 15 to 30 cm deep. ........ 100








Abstract of Dissertation Presented to the Graduate School
of the University of Florida in Partial Fulfillment of the
Requirements for the Degree of Doctor of Philosophy


FACTORS AFFECTING THE EFFICACY OF 1,3-DICHLOROPROPENE FOR
PLANT-PARASITIC NEMATODE MANAGEMENT IN FLORIDA DEEP SANDY
SOILS

By

Claudia Riegel

May 2001

Chairman: Don W. Dickson
Major Department: Entomology and Nematology

1,3-Dichloropropene (1,3-D) has been used for many years in Florida to manage

plant-parasitic nematodes on many crops. 1,3-Dichloropropene is a fumigant nematicide

available to growers and it will likely be among the most important alternative

replacements for methyl bromide, which is proposed for complete phase-out in 2005.

Rate, application depth, method of application, abiotic degradation, and microbial

degradation are all factors that contribute to the performance of 1,3-D. Experiments were

conducted in the field, microplots, and greenhouse with the goal to improve knowledge of

the relative performance of 1,3-D in Florida deep sandy soils and soils capable of microbial

degradation of 1,3-D at an accelerated rate (enhanced soil). Field and greenhouse

experiments were performed with soil infested with root-knot nematodes enhanced for

degrading 1,3-D. The number of second-stage juveniles (J2) that penetrated tomato

seedlings that were transplanted into fumigated soil were evaluated to determine the

efficacy of 1,3-D against root-knot nematodes in enhanced and nonenhanced soils. There








were no differences in the number of J2 that entered the roots grown in fumigated

enhanced and nonenhanced soils. No decrease in efficacy of 1,3-D could be detected.

Field studies were performed to determine efficacious rates and methods of applications

for plant-parasitic nematode control in deep sandy soils. Rates tested included 56, 84,

112, and 168 liters/ha of 1,3-D applied broadcast with conventional chisels followed by

disking. Soil fumigated with 1,3-D at broadcast rates of 84, 112, and 168 liters/ha offered

the best control of root-knot nematodes on squash, whereas 56 liters of 1,3-D/ha provided

marginal control. Methods tested included in-row and broadcast fumigation of 1,3-D

using conventional and parachisels followed by disking and subsurface hooded chisels.

Number and yield, plant-parasitic nematode population density, root-knot nematodes gall

indicies were determined. The highest number of fruit were from plots treated with 1,3-D

applied broadcast followed by disking (P < 0.1). The highest yield was from plots

fumigated with 1,3-D applied in row with conventional chisels followed by disking (P <

0.1). At midseason and at harvest, the number of J2 was lower in all fumigated plots

except in 1,3-D applied in row with parachisels and the untreated control (P < 0.1). The

performance of 1,3-D applied by subsurface hooded sweep chisels was poor. Fumigation

with 1,3-D, regardless of rate or methods, resulted in a decrease in root-knot nematodes

that infected indicator crops.














CHAPTER 1
INTRODUCTION



History of 1,3-Dichloropropene



Carter (1943) was the first to report the effectiveness of a mixture of 1,3-

dichloropropene (1,3-D) and 1,2-dichloropropane (1,2-D) for management of root-knot

nematodes in Hawaiian pineapple fields. The original material, sold under the trade name

D-D, was produced by Shell Development Company and was available in two grades, a

50-50 mixture of 1,3-D and 1,2-D, and a crude form containing about 75% of the mixture

and 25% impurities (Carter, 1943).

The 1,3-D fraction, which was identified as the nematicidal fraction (Carter, 1945),

was increased in marketed products thereafter. The D-D mixture was purified in 1944 and

beginning in the 1950's, Dow Chemical Company marketed the nematicide as Telone,

which was generally composed of 85% 1,3-D and 15% 1,2-D (McKenry and Thomason,

1974a). Vidden D, formulated with 85% to 93% 1,3-D and the rest 1,2-D, also was

marketed by Dow Chemical Company. In 1977, a new formulation with essentially all the

1,2-D component removed was produced and named Telone II (Dow AgroSciences LLC,

Indianapolis, IN. This product was composed of 92% 1,3-D, and 8% inert ingredients.

Over the years, efforts have continued to improve the purity Telone II and the product is








2

currently composed of 94% 1,3-D (Anonymous, 1999b), but it can be as high as 99% and

as low as 93% (Eger, pers. comm.).



Biological Activity of 1,3-Dichloropropene



1,3-Dichloropropene has a biological spectrum of activity that includes nematodes,

some fungi, and some soil-dwelling insects (Anonymous, 1996b; Carter, 1945; McKenry,

1981; Wright, 1981). The product is used widely as a soil fumigant to control

economically important plant-parasitic nematodes such as the burrowing, citrus, cyst

(carrot, golden, soybean, sugar beet, and wheat), dagger, lance, lesion, pin, needle,

reniform, ring, root-knot, spiral, sting, and stubby root (Anonymous, 1996a; Dunn and

Noling, 1995; Fletcher, 1956; Youngson and Goring, 1970).

1,3-Dichloropropene also is labeled to control several plant diseases. These

include bacterial canker of peach (Botryosphaeria dothidea), Fusarium wilt of cotton,

which is suppressed by the control of the cotton root-knot nematode (Meloidogyne

incognita race 3), Verticillium wilt of potato (Verticillium dahliae), and wilt of mint

(Verticillium albo-atrum f. menthae). 1,3-Dichloropropene can suppress the effects of

sugar beet Rhizomania disease (Anonymous, 1996b). The fumigant is believed to reduce

the activity of Polymyxa beta that vectors the Rhizomania virus. The fumigant also is

effective for control of garden centipedes (symphylans) (Anonymous, 1996b).










Mode of Action



1,3-Dichloropropene has two isomers, cis and trans, which have similar chemical

and physical properties but the cis isomer is slightly more volatile than the trans isomer

(Yang, 1986). Cis- and trans-1,3-D are organic molecules that are categorized as alkyl

halides. Alkyl halides are compounds in which a halogen atom (chlorine, fluorine,

bromine, iodine) replaces a hydrogen atom of an alkane (C-C) or alkene (C=C)

(Solomons, 1988a). The chemical penetrates the nematode cuticle directly and promotes

dysfunction of the biochemical processes within the nematode. Organisms exposed to cis-

or trans-1,3-D become hyperactive with increased respiration, defecation, and mobility

(McKenry, 1981). Bimolecular nucleophilic substitution reactions occur on protein

surfaces. A nucleophile is a reagent that "seeks" a positive center in an organic molecule.

In an alkyl halide, the positive center is the carbon atom to which the halogen is attached

(Solomons, 1988a). This carbon atom bears a positive charge because the electronegative

halogen pulls the electrons of the carbon-halogen in its direction (Solomons, 1988a). In a

nucleophilic substitution, reactions occur with the carbon-halogen bond of the molecule

and with the substrate. Heterolysis occurs and the unshared pair of the nucleophile is used

to form a new bond to the carbon atom (Solomons, 1988b). Any molecule or negative ion

that has an unshared pair of electrons is a potential nucleophile. Nucleophilic substitution

reactions occur with NH2, SH, and OH groups on protein surfaces.

The cis isomer, which has been shown to be more toxic to nematodes than the

trans isomer (Moje et al., 1957), is more reactive in nucleophilic displacement reactions








4

than the trans isomer (Hatch et al., 1952). These reactions are slow and the second-order

rate constants for the solvolysis ofcis- and trans-l,3-D are in the order of 0.1 to 0.3

liter'/mole2/hour (Castro and Thomason, 1971). Castro and Thomason (1971) suggested

that the nucleophilic displacement reactions were too slow to explain the mortality of

nematodes, thus they suggested that oxidation of the iron Fe (II) at the center of the

hemeproteins in the cytochrome chain blocks respiration thereby causing cause death of

nematodes.

Rapid action of this toxicant could be explained by the oxidation of iron porphyrins

and hemeproteins. Porphyrins are molecules that have a complex planar structure that

contain substituted pyroles covalently linked in a ring containing a central metal atom.

Iron porphyrins are components of hemeprotein molecules that are found in terminal

oxidases or mitochondrial oxidase and are widely found in biological systems (Van Gundy

and McKenry, 1977). Alkyl halides react rapidly with Fe (II) porphyrins and result in iron

oxidation in the cytochrome chain and a breakdown in respiration. For example, cis-1,3-D

oxidizes Fe(II) porphyrins at a constant rate of approximately 4x 104 liter2/mole2/minute

(Castro and Thomason, 1971).

Castro and Thomason (1971) hypothesized that nematodes exposed to low doses

of alkyl halide nematicides in the range of 1 to 200 ppm undergo a quiescent period before

death. Death could be the result of direct oxidative flooding of iron centers in the

respiratory sequences (Castro and Thomason, 1971). If the nematodes are removed from

contact with the nematicide, the process may be reversed. Nematodes within

nondecomposed root tissue will not necessarily be killed. These nematodes may be








5

exposed to sublethal doses of 1,3-D that have been shown to have a stimulatory effect on

Meloidogyne spp. present within the roots of fig or grape (Van Gundy and McKenry,

1977). Certain low sub-lethal dosages of the trans isomer of 1,3-D or its derivatives can

have a stimulatory effect on hatching of Globodera rostochiensis eggs in addition to the

desired decrease in hatching (Shomaker and Been, 1999). At higher concentrations of the

1,3-D, alyklation of many enzymes, or metabolic sites are simultaneously inhibited, thus

several vital processes stop and lead to a rapid and violent death (Castro and Thomason,

1971).



Chemical and Physical Properties of 1,3-Dichloropropene



The chemical and physical properties of 1,3-D (Table 1-1) and properties of the

soil, climatic conditions, and methods of application regulate the mobility of 1,3-D in soils,

and therefore, control of target organisms (Goring, 1957; Goring, 1962). Movement of

the cis and trans isomers of 1,3-D in soil occurs mainly as a result of diffusion in the vapor

phase (Goring, 1957; Leistra, 1970). During diffusion there is a tendency of the fumigant

to establish an equilibrium of the concentration in vapor, water, and adsorbing phases

(Goring, 1957; Leistra, 1970).

To be effective, 1,3-D must penetrate an envelope of water surrounding soil

particles in order to reach target organisms. The vaporized molecules of the fumigant will

tend to dissolve in the soil water to establish an equilibrium distribution ratio between the

soil-air and the soil-water governed by Henry's Law (Goring, 1957; Leistra, 1970). Water








6

Table 1-1. Chemical and physical properties of 1,3-dichloropropene (Anonymous,
1996a).


Property


Description


Chemical name

Trade name

Molecular weight

Empirical formula

State

Color

Odor

Melting point

Boiling point


Vapor pressure

Saturated vapor pressure


Solubility
in water

in organic solvents

Density


Henry's Law constant
(water/vapor)


1,3-Dichloropropene (1,3-D)

Telone II (Dow AgroSciences LLC, Indianapolis, IN)

11.98 g/mole

C3H4C12

Liquid

Colorless to straw colored

Sweet penetrating

The melting point is lower than -50 C

The boiling point of the cis-isomer is 104.1 C
The boiling point for the trans-isomer is 112.6 C

28 mmHg at 20 C

34.3 mmHg at 25 C (cis)
23.0 mmHg at 25 C (trans)


The solubility of the cis-isomer in water is 2.18 g/liter
The solubility of the trans-isomer in water is 2.32 g/liter
Probably soluble in most organic solvents

1.205 to 1.219 at 20 C
1.2 to 1.214 at 25 C

Cis-isomer 1.8 x 10-3 m3 atm/gmol (25 C )
Trans-isomer 1.05 x 10-3 m3 atm/gmol (25 C)










solubility and vapor pressure are two properties that are related in Henry's Law that

influence the rate of a fumigant's movement. Changes in conditions that tend to shift this

equilibrium will tend to increase or decrease the rate of diffusion of the fumigant through

the soil and modify the relative concentrations of the fumigant in the various phases

(Goring, 1957). The diffusion constant of 1,3-D in the gas phase is 10,000 times greater

than in the water phase (Leistra, 1972) and diffusion in water is considered negligible.

Any decomposition of the fumigant in the soil will affect its concentration in the

various phases (Goring, 1957). The cis- and trans-1,3-D can be transformed rapidly in soil

by abiotic hydrolysis. Hydrolysis rates vary from 1% per day (Williams, 1968) to 3.5%

per day (Castro and Belser, 1966). Microbial transformation is thought to be responsible

for the subsequent steps in the degradation of cis- and trans-3-chloroallyl alcohol (3-CAA)

(Van Dijk, 1974). The parent compound, 1,3-D, is the most toxic to nematodes (Moje et

al, 1957) but the hydrolysis products (cis- and trans-chloroallyl alcohol) also are toxic to

nematodes. Degradation of the hydrolysis products occurs quickly and serves to remove

the toxicant from the soil.



Soil Texture

Soil texture itself does not appear to be the primary limiting factor in the diffusion

of chemicals because it has an indirect influence (Goring, 1957; Thomason and McKenry,

1974). Coarser-textured soils generally allow maximum diffusion of 1,3-D through the

soil. In fine textured soils clay loam and clay soils, the moisture-holding capacity, cation

exchange capacity, organic matter content, and percentage of blocked pore spaces are










generally increased (Thomason and McKenry, 1974). Blocked pore spaces occur when

soil moisture is high or soil structure is destroyed through compaction or deflocculation

(Thomason and McKenry, 1974). Blocked pore spaces restricts gaseous diffusion thus

decreasing the chance that nematodes will come in contact with a lethal dose of the

fumigant (Thomason and McKenry, 1974).

1,3-Dichloropropene has a low vapor pressure and diffusion through the soil

occurs on a random molecular basis (Goring, 1957). The compound is a liquid and the

first process that occurs after application of the fumigant to the soil is initial vaporization.

The rate at which this occurs will be influenced by the inherent volatility of the fumigant,

the extent of dilution of the fumigant with solvents, and the restraint on diffusion caused

by the porous medium through which the fumigant is diffusing (Goring, 1957).

If the air space of the soil is limited and the pore spaces discontinuous, the

diffusion pattern will be restricted (Thomason and McKenry, 1974). As a general rule, a

sandy loam soil should not be fumigated if the soil moisture tension is less than 0.5 bars

suction at a depth of 30.5 cm (Thomason and McKenry, 1974). A clay loam soil should

have a 0.6 bar suction. It is possible to fumigate sandy or clay type soils when the

moisture level is at the permanent wilting point (15 bars tension) (Thomason and

McKenry, 1974). For optimum movement of the toxicant, soils should be fumigated when

soils are in a drying condition (Thomason and McKenry, 1974).

When a marginal dose of 1,3-D is used, greater control occurs when soil has low

porosity rather than high porosity (Goring, 1957). Soil texture will often vary with depth










in the soil profile and make it difficult to predict fumigant dispersion, and compacted

layers will limit diffusion of the fumigant (Munnecke and Van Gundy, 1979).

Diffusion of 1,3-D is greatly affected by organic matter because the sorption of the

compound is due primarily to the colloidal organic matter in soils (Siegel et al., 1951).

Reliable fumigation of soils is difficult to obtain when organic matter content is high

(Goring, 1957; Leistra, 1970). If soil with high organic matter are to be fumigated, the

soil should be moist. When fumigating, it is best to aim for a high soil porosity throughout

to give the fumigant the best chance to diffuse to considerable distances before extensive

sorption or decomposition occurs (Goring, 1957). A soil with high porosity and low soil

moisture will generally give the best results.



Temperature

Temperature of soils is important when fumigating. However, nematode control

was shown to be independent of temperature when there is a range from 10 to 32 C

(Youngson and Goring, 1962). Temperature can affect the diffusion and activity of a

fumigant in many ways. Generally a rise in soil temperature increases the volatility

because the vapor pressure of the fumigant is increased (Goring, 1957). For alkyl halides

(e. g. 1,3-D), solubility in water tends to decrease with increased temperatures. There is a

shift in Henry's Law constant that predicts a greater proportion of the fumigant in the

water and the rate of diffusion through soil decreases.

Theoretically, better nematode control is achieved at higher temperatures when

concentrations of the toxicant is low but well distributed throughout the soil profile










(Thomason and McKenry, 1974). At lower temperatures, diffusion of the chemical

throughout the soil profile is restricted and concentrations in the water are higher than in

air spaces (Thomason and McKenry, 1974). In laboratory and field studies, Thomason

and McKenry (1974) stated that the best control of nematodes would be attained between

15 and 20 C. Below 15 C, the solubility of 1,3-D is greatly increased and seriously

limits diffusion through soil because the concentration of the fumigant increases

dramatically with a decrease in temperature, therefore, the diffusion rate in the soil is

limited.

Hydrolysis rates for 1,3-D are influenced most by temperature and soil moisture.

In experiments conducted in the field (Thomason and McKenry, 1974) and laboratory

(Castro and Belser, 1966; Van Dijk, 1974), hydrolysis of 1,3-D at significant rates

occurred at higher temperatures of 15 to 25 C. Roberts and Stoydin (1976) showed that

both cis- and trans-1,3-D are hydrolyzed initially to respective cis- and trans-3-chloroallyl

alcohols, which are in turn, rapidly oxidized to corresponding 3-chloroacrylic acids.

Temperature has a pronounced affect on the disappearance of 3-chloroallyl alcohol (Van

Dijk, 1980). In the laboratory, 3-CAA disappeared at a faster rate at a higher temperature

and the trans isomer degraded faster than the cis isomer in unsterilized soils. Under these

conditions, the alcohol degraded faster than the 1,3-D. There was a minimal effect of

temperature in the sterile soil and the disappearance of the alcohols was much slower than

in unsterilized soil.










Soil Moisture

Soil moisture is a limiting factor in the diffusion of fumigants such as 1,3-D and D-

D (Thomason and McKenry, 1974). If the moisture is held constant and the air space is

decreased by 2.5 times, the proportion of the fumigant in water is increased. The result

will be a longer exposure of the nematode to the fumigant but at a considerably lower

concentration (Goring, 1957).

Maximum fumigant diffusion occurs in soils with a soil moisture tension of 0.6 to

15 bars (Munnecke and Van Gundy, 1979). Soils that are wetter than 0.6 bars (10 to 12%

moisture) will have a number of air passageways blocked by water thereby restricting

fumigant movement (McKenry and Thomason, 1974a). Soil fumigation at high soil

moisture levels results in a slow movement of the 1,3-D fumigant vapors and uneven

distribution in soils (McKenry and Thomason, 1974a). Lateral diffusion also is affected.

The higher the soil moisture, the more the fumigant is diluted and the total pattern of

diffusion will be more restricted (Thomason and McKenry, 1974a). Excessively dry soils

(15 bars) that exceed the wilting point for vegetation have few molecules of water

surrounding soil particles. Under these conditions, the sorption of the fumigant molecules

may occur directly on the surface of the soil particle (McKenry and Thomason, 1974a;

Munnecke and Van Gundy, 1979).

Hydrolysis rates will vary depending on the soil type due to differences in diffusion

rate and sorption capacity. Physical characteristics of the adsorbent surface, surface

acidity, nature and availability of the catalytic sites, soil pH, and redox potential (Bailey et

al., 1974) are all factors that may influence hydrolysis. However, no consistent correlation








12

between 1,3-D hydrolysis and soil organic matter, clay content, or pH has been observed

(Van Dijk, 1974).

Castro and Belser (1966) showed that hydrolysis and degradation of 1,3-D occurs

in moist soil. They applied high concentrations (10-2 M) of 1,3-D to wet soil and observed

a hydrolysis rate of 3.4% per day, however this dissipation rate was not collaborated by

Thomason and McKenry (1974) when they showed 1,3-D to persist in a wet, warm soil

for 2 to 4 months. Van Dijk (1974) observed 1,3-D to have a longer half-life in a buffer

solution than in soil. In the buffer solution, 1,3-D dissipated faster with increased

temperature. The dissipation rates of cis- and trans-isomers were not significantly

different from each other in either soil slurry or buffer solution (Castro and Belser, 1966;

Van Dijk, 1974).

Microbial and chemical degradation are two ways that 1,3-D disappears from soil.

The other major way is the diffusion of 1,3-D vapor to the soil surface and subsequent

evaporation into the atmosphere (Yang, 1986). In addition, downward movement via

leaching into the groundwater also may occur. Van der Pas and Leistra (1987) reported

that low concentrations (
beneath commercial flower-production fields in The Netherlands. Detection of trace

amounts or no 1,3-D residues in groundwater beneath treated fields was not unusual.

However, 1,3-D in water is subject to rapid chemical hydrolysis (McCall, 1987), and for

this reason, there is little or no chance that 1,3-D residues would leached into the

groundwater.










Soil moisture also affects the susceptibility of nematodes to the fumigant. The

amount of 1,3-D required to control sugar beet cyst nematode in sealed containers in soil

at low and relatively high moisture contents were different (Goring, 1957). Higher doses

are required to kill nematodes at low soil moisture than at high soil moisture.



Time of Application, Seeding, or Transplanting Restrictions



Applications of 1,3-D in winter and spring in north Florida equally effective in the

management of Meloidogyne arenaria race 1 on peanut (Kinloch and Dickson, 1991).

Yields of peanut were increased 2.5 fold as compared to banded plus applications at

pegging applications ofaldicarb (Kinloch and Dickson, 1991).

Preplant applications of 1,3-D are specified on the label for Telone II. A 7 day

waiting period before planting is specified (Anonymous, 1999b). There should be no odor

of 1,3-D at planting and for determining the presence if in doubt a seed test is an

inexpensive method that indicates the presence of the chemical (Anonymous, 1996b).

Lettuce and radish seeds are sensitive to 1,3-D vapors and can be used as indicators of the

presence of the fumigant. A composite sample from soil collected randomly from treated

areas 15.2 to 45.7 cm deep is placed in sealed bottles that contain the seeds. If 1,3-D is

not present, the seeds should geminate in less than 1 week and the seedlings should be

green and healthy. If the seedlings are discolored (brown or chlorotic) or stunted, 1,3-D

probably is still present in the soil. If the concentration of 1,3-D is high, the seeds may not








14

germinate or the seedlings may be killed. Soil from an untreated area also should be tested

to check the viability of the seeds.

Aeration of the soil may be done several ways depending on the planting site

(Anonymous, 1996b). Any equipment moved into a treated area should be

decontaminated to prevent reinfestation of the planting site. The soil may be worked to

the depth of the treated zone. In the case of row treatment, a narrow shank may be pulled

through the bed without turning the soil. Following a broadcast treatment, plowing or

deep cultivation to the depth of the treatment zone can be performed. This is especially

desirable in northern climates after fall fumigation of muck soil (Anonymous, 1996b). If

planting sites have deep-rooted trees and shrubs that have been fumigated, then an

aeration period of 3 to 6 months may be required.

Seeds, seed pieces, and transplants used must be free of pathogens so that the

planting site is not infested or reinfested with pathogens. When possible, one should use

certified seed to ensure the quality of the plant material. Seeds or transplants should be

placed 8 to 10 cm to one side of the furrow left by the applicator shank (off the treatment

line) (Anonymous, 1996b). When two shanks are used, the center area between shanks

should be planted.

Certain growing stages of plants are more sensitive to 1,3-D than others. Dormant

seeds are more tolerant to 1,3-D exposure than are seedling transplants (Anonymous,

1996b). In unbedded and bedded rows, the seeds should be placed at a depth proper for

germination. In a use pattern violating the Telone II label, Rich and Kinloch (1998; 2000)

conducted field trials in northern Florida that showed 1,3-D applied in-row at planting at










16, 32, and 48 liters/ha caused limited phytotoxicity and produced comparable yields to

the standard preplant application.



Enhanced Degradation



The duration of the availability of soil-applied pesticides can be influenced by a

range of biotic and abiotic processes that interact extensively so that the contribution of

any one variable is usually difficult to discern (Suett et al., 1996). However, it has been

long accepted that the degradative capacity of the soil microbial community is a significant

factor to determine residue stability in soil (Suett et al., 1996). Microorganisms have been

shown to degrade several pesticides in soil including 1,3-D (Chung, et al., 1999; Jones and

Norris, 1998; Oldenhuis et al., 1989; Ou and Sharma, 1989; Ou et al., 1993; Ou et al.,

1995; Rack and Coats, 1990; Vannelli et al., 1990).

Cis- and trans-1,3-D are hydrolyzed to cis- and trans-3-CAA and oxidized to

corresponding 3-chloroacrylic acids (Roberts and Stoydin, 1976). The acids are further

degraded to aliphatic carboxylic acids such as acetic acid, propionic acid, and succinic

acids (Anonymous, 1996a; Roberts and Stoydin, 1976). These acids are then degraded to

CO2 and H20 (Ou et al., 1989). The initial step in hydrolysis from 1,3-D to 3-CAA was

considered to be an abiotic process and the subsequent steps were considered biological

(Roberts and Stoydin, 1976).

The two isomers are similar kinetically (Leistra et al., 1991; Van der Pas and

Leistra, 1987; Van Dijk, 1974, Van Dijk, 1980). Half-life values for cis-1,3-D in soils








16

under laboratory conditions at 10, 15, and 20 C ranged from 17 to 47, 4 to 32, and 3 to

15 days, respectively (Leistra et al., 1991; Smelt et al., 1989; Van Dijk, 1974; Van Dijk,

1980). Due to the volatile nature of 1,3-D, it is difficult to determine true degradation

rates of 1,3-D in soils under field conditions. Unlike cis- and trans-l,3-D, cis- and trans-3-

CAA appear to have different degradation rates in soil with the trans-isomer being more

rapidly degraded than the cis-isomer (Leistra et al., 1991; Van Dijk. 1974).

1,3-Dichloropropene has been reported by many scientists to degrade at a faster

rate in soil after repeated applications (Chung et al, 1999; Lebbink et al., 1989; Ou et al.,

1995; Smelt et al., 1989; 1996; Verhagen et al., 1995; 1996). Smelt et al. (1989) and

Lebbink et al. (1989) were the first to report that enhanced degradation could occur with

1,3-D. Smelt et al. (1989) reported that enhanced degradation of 1,3-D occurred in some

loamy soils, however, in some soils that had been treated up to six times enhanced

degradation of 1,3-D was not observed. Smelt et al. (1989) did not attempt to determine

if enhanced degradation was isomer specific. They attributed the enhancement to the prior

presence of microorganisms capable of degrading 1,3-D in those soils that exhibited rapid

degradation of 1,3-D. However, the enhanced degradation reported by Smelt et al. (1989)

really was not chemically measured in corresponding control soils. Lebbink et al. (1989)

reported lab studies using soil collected from a field in The Netherlands that had been

fumigated with 1,3-D annually for 12 years degraded 1,3-D at an accelerated rate. The

field from which the soil was collected had a 70% reduction in efficacy over the course of

the 12 years (Lebbink et al., 1989).










The first documented case of enhanced degradation of 1,3-D in Florida was

observed by Ou et al. (1995). In subsequent research conducted at the reported enhanced

sites, Chung et al. (1999) and Ou et al. (1995) found that degradation oftrans 1,3-D in

surface and subsurface sandy soils (Arredondo fine sand) was faster than for cis-1,3-D.

Degradation rates of both isomers were similar in untreated soil. This site had been

treated with 1,3-D at least seven times over the past 13 years, and had been planted with

peanut or tomato. Both cis- and trans- isomers of 1,3-D in surface and subsurface soil

samples collected from the treated plot were degraded significantly faster than in the

corresponding soil samples collected from the untreated plots. Half-life values for cis- and

trans-l,3-D in the fumigated soil 0 to 15 cm deep were 8 and 3 days, respectively. Half-

life values for cis- and trans-1,3-D in corresponding untreated surface soil were 20 and 17

days, which were not different. None, or trace amounts, of trans-3-CAA were detected in

fumigated and soil nonfumigated soil, whereas substantial amounts of cis-3-CAA were

detected in these soils. Cis- and trans-CAA are the hydrolysis products of cis- and trans-

1,3-D, respectively. Based on results of the degradation rates of cis- and trans-1,3-D in

fumigated and soil nonfumigated soils, Ou et al. (1995) concluded that differential

enhanced degradation occurred in the treated soil, with trans-1,3-D being degraded faster

than cis-1,3-D. They termed this phenomenon "differential enhanced degradation" to

indicate that these two isomers are degraded at different rates. They also suggested that

biological hydrolysis is the main factor for hydrolysis of trans-1,3-D in treated soil,

whereas, chemical hydrolysis was the main factor for hydrolysis of cis- and trans-1,3-D in

untreated soil.










Microorganisms capable of hydrolyzing 1,3-D have been isolated from soil and

water. Lebbink et al., (1989) isolated a fluorescent Pseudomonas sp. from soil enhanced

to degrade 1,3-D. This strain used 1,3-D as the sole source of carbon for growth

(Lebbink et al., 1989). Verhagen et al. (1995) isolated 15 bacterial isolates that were

capable of degrading 1,3-D when grown in a mineral medium containing yeast extract. In

addition, six strains of a Gram negative bacterium that were isolated contained the plasmid

dhlA (haloalkane dehalogenase)-like gene that was thought to be involved in degradation

of 1,3-D (Verhagen et al., 1995). Two different strainsof Pseudomonas spp. isolated from

soil were found to have the capacity to use 3-CAA or 3-chloroacrylic acid as a sole source

carbon for growth (Belser and Castro, 1971). The Psuedomonas sp. isolated by Van

Hylckama and Janssen (1992) produced two dehalogenases, one specific for cis-3-

chloroacrylic acid and the other specific for trans-3-chloroacrylic acid.

Currently, it is unknown whether one, two, or three enzymes are responsible for

the hydrolysis of cis- and trans-1,3-D and whether axenic cultures isolated or consortia are

responsible for degradation of 1,3-D in soil and water. Psuedomonaspavonaceae was

capable of producing at least three different dehalogenases (Verhagen et al., 1995). In

addition, Ou et al. (1995) isolated a enriched a mixed bacterial culture from a Florida

sandy soil that had limited capacity to mineralize 4C-labeled 1,3-D cometabolically. A

bacterial consortium from a Florida sandy soil that was capable of enhanced degradation

of 1,3-D was isolated (Ou et al., 2001). From that bacterial consortium, an axenic culture

of Rhodococcus sp. capable of degrading cis- and trans-1,3-D was isolated after

continuous subculturing of the bacterial consortium (Ou et al., 2001). This isolated culture








19

was capable of degrading both 1,3-D isomers in the presence of a suitable carbon source

(Ou et al., 2001).














CHAPTER 2
THE EFFICACY OF 1,3-DICHLOROPROPENE FOR CONTROL OF ROOT-KNOT
NEMATODES WHEN APPLIED IN SOIL ENHANCED FOR DEGRADING 1,3-
DICHLOROPROPENE



Introduction



A near equal ratio of cis-and trans-1,3-dichloropropene (1,3-D) is the main active

ingredients of the fumigant nematicide Telone II (Dow AgroSciences LLC, Indianapolis,

IN) (Ou et al., 1995). The commercial formulation is composed of 94% 1,3-D and 6%

inert ingredients (Anonymous, 1999a). In Florida, fumigation with 1,3-D is recommended

for a variety of crops including vegetable and agronomic crops (Dunn and Noling, 1995),

and 1,3-D formulated with chloropicrin (17% and 35%) is the likely alternative for methyl

bromide for nematode control in the near future when methyl bromide is no longer

available to growers (Holman, 1999).

Although the fumigant has been used successfully to control plant-parasitic

nematodes on many crops, in some cases the efficacy of the fumigant has been less than

expected when used under varying conditions. One factor thought to be involved in

reducing the efficacy of 1,3-D was microbial degradation. Microorganisms have been








21

shown to degrade pesticides in soil (Jones and Norris, 1998; Oldenhuis et al., 1989; Ou et

al., 1993; Ou et al., 1995; Rack and Coats, 1990; Vannelli et al., 1990).

Enhanced degradation of 1,3-D has been reported following repeated applications

of the fumigant (Chung et al, 1999; Lebbink et al., 1989; Ou et al., 1995; Smelt et al.,

1989; 1996; Verhagen et al., 1995; 1996). The first documented case of enhanced

degradation of 1,3-D in Florida was reported by Ou et al. (1995). They found that

degradation oftrans 1,3-D was faster than cis-1,3-D in surface and subsurface sandy soils

collected from a 1,3-D treated site. Degradation rates of both isomers were similar in

untreated soil (Ou et al., 1995). This field site located at the University of Florida Green

Acres Agronomy Farm in Alachua County, had been fumigated repeatedly and bacteria in

the soil were capable of degrading 1,3-D to CO2 and H20 in 14, 7, and 5 days in

experiments conducted during 1994, 1995, and 1996, respectively. In 1997, both cis- and

trans-1,3-D were degraded completely in less than 5 days (Chung et al., 1999). Half-life

values in 1997 for 1,3-D in the surface soil (0 to 15 cm deep) was 1 day for cis-l,3-D and

0.8 days for trans-1,3-D, and in subsurface soil (15 to 30 cm deep) it was 1.4 days for the

cis-1,3-D and 0.7 days for the trans-l,3-D (Chung et al., 1999). A consortium of soil

bacteria and a strain of Rhodococcus sp. have been isolated and shown to degrade 1,3-D

at an accelerated rate in the presence of a second organic substrate (Ou et al., 2001).

The objective of this study was to determine whether this enhanced soil affected

the efficacy of 1,3-D against root-knot nematodes. Experiments were designed to

ascertain the impact of the enhanced soil on 1,3-D and root-knot nematodes.










Materials and Methods



Microplot Study

Microplots infested with a mixed population of Meloidogyne arenaria (Neal)

Chitwood race 1 and M incognita (Kofoid and White) Chitwood race 1 were used to

evaluate the efficacy of 1,3-D. The microplots were located at the University of Florida

Green Acres Agronomy Research Farm in Alachua county, Florida. A total of 54

microplots (76-cm-diam. encircled with 60-cm-wide fiberglass sheets inserted 50 cm deep

into the soil) (Johnson et al., 1981) spaced in rows 1.5 m apart were used. The soil was

Arredondo fine sand (93% sand, 4% silt, and 3% clay, and 3% organic matter). The

microplots were divided into three groups of 18 and designated as low, medium, and high

based on an initial sampling to determine the density of second-stage juveniles (J2) of the

two Meloidogyne species. Low density was 1 to 28 J2/100 cm3 of soil, intermediate was

29 to 88 J2/100 cm3 of soil, and high was 89 to 648 J2/100 cm3 of soil.

The soil in the top 45 cm, estimated at 0.53 m3, was amended with 4 kg of one of

three soil types: enhanced to degrade 1,3-D, enhanced to degrade 1,3-D and autoclaved

twice, each time at 121 C at 103.4 kilopascal for 30 minutes, and soil not enhanced to

degrade 1,3-D. Microplots containing untreated enhanced soil and nonamended soil were

included as controls. The experimental design was a randomized complete block with 10

replicates. A 30-cm-wide by 30-cm-deep soil core was removed from the center of each

microplot and replaced by one of the three soil types mentioned above, and the soil for










each treatment was thoroughly mixed with soil in the microplot. In nonamended

microplots, an equal portion of soil was removed, placed back into the hole, and mixed

with the soil present in the microplot. The plots were irrigated by microjets ca. 5 minutes

daily for 1 month.

The microplots were broadcast fumigated with 1,3-D at a rate of 168 liters/ha 1

month (19 May 1996) after incorporating the enhanced soil into the microplots. Soil

moisture at the time of fumigation was 13.1% at 30 cm deep. A marked circular template

was used to aid in placement of the fumigant. A glass pipette was used to inject 1,3-D 30

cm deep on 30.5 cm centers five times in each microplot and the soil around the injection

points was pressed firmly with a wooden stake. Two liters of water were sprinkled over

the surface of each microplot to form a water barrier seal because it has been shown

effective in reducing volatilization losses in the field (Gan et al., 1998).

Soil samples were taken from the microplots at three depths (0 to 15 cm, 15 to 30

cm, and 30 to 45 cm) 24 hours before fumigation, and 24,48, 72, 96, and 120 hours after

fumigation. Five soil cores per each depth (ca. 250 cm3 soil) were removed with a soil

sampling tube (2.5-cm-diam.) and placed in a 10x15x20 cm, 0.002 cm thick polyethylene

bag and immediately taken to the laboratory for processing. The soil samples at each

depth within each microplot were combined and mixed thoroughly. Two bioassays with

tomato seedlings were done with these samples in a shadehouse, and a third bioassay was

done directly in the microplots. A bioassay was used because it is the most reliable

method to determine the number of nematodes that survive exposure to a toxicant and

successfully infect a host plant (Thomason et al., 1964).










Bioassay 1: A 100 cm3 subsample was transferred from each composite soil

sample and then placed into a 164 ml ultraviolet-stabilized container (Stewe and Sons,

Corvallis, OR). Before soil was added, 40 cm3 of heat-pasteurized sandy-loam soil was

added to the bottom of each container. A 3-week-old tomato seedling, Lycopersicon

esculentum Mill. cv. Rutgers, was transplanted into each container. The number of galls

was estimated by counting those that were visible on the root system 6 weeks later. Root-

knot nematode eggs were extracted using a 0.5% sodium hypochlorite solution (Hussey

and Barker, 1973) and the number of eggs per root system was determined.

Bioassay 2: Two microplots from the three categories (low, intermediate, and

high densities of J2) from each treatment were chosen arbitrarily for the second bioassay.

Second-stage juveniles were extracted by a centrifugal-flotation method (Jenkins, 1964)

from a 100 cm3 subsample at depths of 0 to 15 cm, 15 to 30 cm, and 30 to 45 cm. Soil

samples were removed from the same microplots 24 hours before fumigation and 24 and

48 hours after fumigation. The number of J2 per 100 cm3 of soil was counted and these

nematodes were placed in containers filled with 140 cm3 of heat-pasteurized soil and a

tomato seedling was transplanted into the soil. The number of galls was estimated by

counting those that were visible on the root system 6 weeks later.

Bioassay 3: Three tomato seedlings, cv. Summerset, were transplanted into each

microplot 48 hours after fumigation. The tomato plants were grown for 6 weeks before

they were removed and the roots indexed for galling caused by root-knot nematodes on a

scale of 0 to 10 (0 = 0 galls, 1 = 10% of the roots galled, 2 = 20% of the roots galled,

..... 10 = 100% of the roots galled) (Barker et al., 1986).










Five cores (2.5-cm-diam., 20-cm-deep) of soil from each microplot were taken

with a cone-shaped sampling tube before fumigation (19 May) and 7 weeks after

fumigation. Soil cores from each microplot were combined and nematodes were extracted

from a 100-cm3 subsample by a centrifugal-flotation method (Jenkins, 1964). All plant-

parasitic nematodes were counted.

The number of galls and eggs per root system from bioassay 1 and the number of

galls per root system from bioassay 2 were subjected to regression analysis to determine

the nematode response to length of 1,3-D exposure in the soil. Gall indices from bioassay

3 were subjected to analysis of variance.

1,3-D Degradation: Rate of 1,3-D degradation for soil collected from the three

treatments was determined by Dr. L.-T. Ou in the Soil and Water Science Department,

University of Florida with the use of a previously described method (Chung et al., 1999).

To make the determination, 50 cm3 of soil 30 cm deep was collected from each microplot

and combined according to treatment.



Field Study

Two field sites in close proximity (150 m apart) were used for the study. The site

(162 m2) with soil enhanced for degrading 1,3-D was located on the west side of a 1.02 ha

field. During the past 20 years, the site had been planted mainly with peanut (Arachis

hypogaea L.), but with some plantings of corn (Zea mays L.), and one crop of tomato

(Lycopersicon esculentum Mill.) in 1993. The second site (nonenhanced soil) had the

same dimensions and was located 144 m east of site one. This site located within a fenced










area, was planted with bahiagrass (Paspalum notatum Flugge) for the past 25 years and

had no known history of exposure to 1,3-D. Soil from both sites was collected from the

top 45 cm and combined for soil texture analysis. The soil texture in the top 45 cm was

88% sand, 8% silt, and 4% clay with 3% organic matter, and pH 6.

Both sites were reinfested with root-knot nematodes to ensure a uniform and large

population density by incorporating soil and roots containing Meloidogyne incognita

(Kofoid & White) Chitwood race 1 on 15 March and 10 May 1997. The inoculum was

prepared in the greenhouse by infecting 300 tomato plants cv. Rutgers each with 4,000

eggs of M incognita race 1. The plants were maintained in the greenhouse until the roots

were heavily galled and egg masses were abundant. After each site was infested, the root-

knot nematode population was increased further by growing a crop of hairy vetch (Vicia

villosa Roth) through the winter and soybean (Glycine max L.) cv. Davis in the summer

(planted on 12 July 1997). The soybean plants were cut at the soil line and the roots were

tilled into the soil 3 weeks before fumigation. Before 1,3-D was applied, three 0.91 m

wide beds were formed at both field sites.

Twenty, 0.91 m by 1.5 m plots were prepared on the raised beds at each site (Fig.

2-1). The untreated control and fumigated plots were completely randomized within each

site. The treatments, which were applied at both sites (25 August 1997), consisted of 1,3-

D applied broadcast at 112 liters/ha and an untreated control. Paraquat at a rate of 0.19

liters of formulation/ha was applied broadcast 5 days before fumigation. A template of

mesh screen on a wooden frame (1.6 m x 1.1 m) was constructed to ensure precise








27

placement of 1,3-D in each plot (Fig. 2-2). The center of each square was marked as the

point of injection. A glass and teflon syringe with a stainless steel needle 30 cm long was

used to inject the fumigant. The soil around the injection point was firmly pressed to seal

in the fumigant. The soil temperature taken (average of 5 samples) at 20 cm deep at the

time of fumigation was 31 C and the soil moisture was 6.4% at 0 to 15 cm deep, 6.5% at

15 to 30 cm deep, and 7.9% at 30 to 45 cm deep. One week after fumigation, 10-10-10

NPK mineral fertilizer was applied broadcast at the rate of 448 kg/ha and was

incorporated in the top 8 cm of soil. Three drip tubes with emitters spaced 30 cm apart

and a flow rate of 62 ml/30.5 m was placed 8 cm from the plant stems for application of

water. The drip tubes were spaced 0.3 m apart over each bed. Six tomato seedlings cv.

Solarset were transplanted in the three central squares of each plot (Fig. 2-3). The plants

were removed from the soil 11 days after planting. The roots were stained (Byrd, et al.,

1983) and the J2 that penetrated the root systems were counted with the aid of a

stereomicroscope. The data were subjected to the Kruskal-Wallis test for completely

randomized nonparametric data (SAS Institute, Cary, NC).



Greenhouse Study

Soil from the enhanced and nonenhanced sites at the University of Florida Green

Acres Agronomy Farm in Alachua county, Florida was brought from the field and used for

this study. Thirty-six 12.7-cm-diameter PVC tubes were constructed to provide two

chambers, an upper chamber (42 cm long) to be filled with fumigated soil and a lower

chamber (30.5 cm long) to be filled with pasteurized soil (Fig. 2-4). The upper fumigation









































Fig. 2-1. Schematic of field site plot layout used to evaluate the efficacy of 1,3-
dichloropropene in enhanced and nonenhanced soil. Site one soil had been identified previously to
degrade 1,3-D (Ou et al., 1995); site two, located 144 m from site one, was planted in bahiagrass
for the past 25 years and had no known history of exposure to 1,3-D. The two field sites covered
an area of 7.6 mn wide by 1.5 m long.

















Injection points


1.5m


Wooden frame


Screen mesh





Fig. 2-2. Schematic of screened template used for the placement of 1,3-
dichloropropene in enhanced and nonenhanced soil at two different field sites. Injection
points located on 30 cm centers.


-0.3m-


i-----
*> S S




S S Si
-t -
_______ ______ _
















Two tomato seedlings were transplanted per square.


1 r m


Soil samples were taken from the three center squares.


Fig. 2-3. Schematic of a plot with six tomato seedlings transplanted in a row
surrounded by a border of fumigated soil.

















Nitex mesh


12.5 cam-diam.






Greenhouse bench


.,-Two tomato seedlings



- Fumigated soil chamber






.----- Pasteurized soil chamber


Fig 2-4. PVC tubes constructed for the greenhouse study had an upper chamber filled
with fumigated soil and a lower chamber filled with heat-pasteurized soil. The two chambers were
separated by a single layer of Nitex mesh (45 pm openings) that was glued to the bottom of the
upper chamber. Two tomato seedlings were transplanted into the upper chamber 5 days after
fumigation with 1,3-dichloropropene at a rate of 84 liters/ha.








32

chamber held 836 cm3 of soil and the lower pasteurized chamber held 240 cm3 of soil. A

Nitex (Tetko, Inc., New York, NY) screen mesh (45 ,m openings) was glued to the

bottom of the upper chamber thereby allowing for movement of water and nematodes but

holding the soil in the chamber. The lower chambers of all 36 PVC tubes were filled with

heat pasteurized field soil (Arredondo fine sand) and of the upper chambers, 18 were filled

with nonenhanced soil and 18 were filled with enhanced soil. The tubes were placed in a

greenhouse and secured in place with wooden stakes.

Meloidogyne incognita race 1 eggs were extracted from egg masses on tomato

roots using a 0.1% NaOCI solution (Hussey and Barker, 1973). Each upper chamber was

infested with 27,000 eggs ofMeloidogyne incognita race 1 the same day the eggs were

extracted. Three holes, each at different depths (5, 15, and 30 cm deep), were made to

ensure that the nematode eggs were distributed throughout the soil profile of the upper

chamber. Each tube was then watered with 100 ml of tap water.

The experimental design was a randomized complete block with four treatments,

1,3-D treated and nontreated enhanced and nonenhanced soils, each replicated 9 times. A

broadcast rate of 84 liters of 1,3-D was injected into the upper chamber of the appropriate

treatments 7 days after infestation with nematodes. A glass syringe outfitted with a 30-

cm-long, stainless steel needle was used to inject the fumigant 25 cm deep into the soil on

6 June 1998. Each tube was draped with a plastic bag and covered with water to a depth

of 2.5 cm after the fumigant was injected. The soil temperature was 33 C and the soil

moisture was 10% at the time of fumigation.








33

The plastic bags and water were removed 5 days after fumigation and two tomato

seedlings cv. Rutgers were transplanted into each upper chamber. The plants were

removed from the soil 5 days after transplantation and the roots were washed carefully and

stained (Byrd et al., 1983). The number of root-knot nematodes that penetrated the root

systems was counted using a stereomicropscope. Data were subjected to analysis of

variance.



Results and Discussion



Microplot Study

The microplots with low, intermediate, and high densities of J2 of Meloidogyne

spp. were not different 7 weeks after fumigation when densities were compared among

individual treatments (P > 0.05), thus they were combined for data analysis (Table 2-1).

Bioassay 1: Dosage response of plant-parasitic nematodes to soil fumigants have

been determined using a variety of methods in the laboratory, greenhouse, and field

(Goring, 1957; McKenry and Thomason, 1977a; 1977b; 1977c; McKenry et al., 1977;

Rich and Kinloch, 2000; Schomaker and Been, 1999). In this study, the number of galls

and eggs per root system decreased with increased periods of exposure to 1,3-D for all

treatments (Tables 2-1, 2-2), however, there were no statistical differences among the

three treatments (amended enhanced soil, amended autoclaved enhanced soil, and the

nonenhanced soil). Apparently, transfer of soil shown to degrade 1,3-D was not sufficient

to affect soil response in the microplots to 1,3-D, or perhaps our test procedures were








34

insufficient to detect differences if they existed. The reaction of 1,3-D was similar among

all three treatments and gave a typified reaction to that expected against root-knot

nematodes. There were no galls or eggs produced after 96 hours of exposure. Regression

analysis was used to interpret the relationship between the number of galls and eggs per

root system and exposure period of Meloidogyne spp. to 1,3-D. A decrease in galls and

eggs was observed in all treatments and at all three depths with increased exposure

periods to 1,3-D (Tables 2-3, 2-4).

Bioassay 2: The continuing fumigation action of 1,3-D was stopped when the

nematodes were extracted from the soil. This approach was used by Thomason et al.

(1968) because the most critical assessments of the effects of exposure time are made

when the nematodes are exposed to the fumigant and then removed from the solution of

the toxicant. Second-stage juveniles extracted from 1,3-D-treated soil 24 hours after

fumigation were motionless, but some recovered and were infective (Table 2-5). A similar

response has been observed with the fumigant nematicide EDB and the plant-parasitic

nematode Aphelenchus avenae (Castro and Thomason, 1971). Motility and reproduction

of the nematode recovered after 8 hours of in vitro exposure to ethylene dibromide and a

96 hour exposure study was adequate to assess mortality (Castro and Thomason, 1971).

In the current study, after fumigation, the number of galls induced by the treated J2 was

low and only appeared from samples collected 24 hours following fumigation. No viable

J2 were found in samples collected 48 hours later. Galling caused by J2 at each sample

depth for all three treatments showed a nonlinear response to exposure periods to 1,3-D

(P 0.05).










Table 2-1. Number of galls on tomato roots grown in soil infested with Meloidogyne spp. in microplots amended with enhanced soil
(previously identified to degrade 1,3-dichloropropene [1,3-D] at an accelerated rate [Ou et al., 1995]), autoclaved enhanced soil, and nonenhanced
soil.


Number of galls per root system

Before fumigation After fumigation

Treatmentb Depth (cm) 24 hr 24 hr 48 hr 72 hr 96 hr 120 hr

Amended with enhanced soil 0 to 15 98 28 8 4 0 0
15 to 30 95 8 0 0 0 0
30 to 45 12 0.2 0.1 0 0 0

Amended with autoclaved enhanced soil 0 to 15 94 20 11 8 0 0
15 to 30 140 3 0.1 0 0 0
30 to 45 160 2 3 0 0 0

Nonenhanced soil 0 to 15 60 39 0 2 0 0
15 to 30 125 0.3 0 0 0 0
30 to 45 97 1 0.4 0 0 0


Microplots were broadcast fumigated with 1,3-D at 168 liters/ha, and data are means of 18 replications.

'Enhanced and autoclaved enhanced soils (4 kg) were placed in the center of each microplot and mixed with the remaining soil to 30 cm
deep.

bA 100 cm3 subsample was transferred from each composite soil sample and then placed into a 164 ml ultraviolet-stabilized container
(Stewe and Sons, Corvallis, OR). Each container had 40 cm3 heat-pasteurized sandy-loam soil added at the bottom.

CA 3-week-old tomato seedling, Lycopersicon esculentum Mill. cv. Rutgers, was transplanted into each container. The number of galls was
estimated by counting those that were visible on the root system 6 weeks later.










Table 2-2. Number of eggs on tomato roots grown in soil infested with Meloidogyne spp. in microplots amended with enhanced soil
(previously identified to degrade 1,3-dichloropropene [1,3-D] at an accelerated rate [Ou et al., 1995]), autoclaved enhanced soil, and nonenhanced
soil.

Number of eggs per root system

Before fumigation After fumigation

Treatment Depth (cm) 24 hr 24 hr 48 hr 72 hr 96 hr 120 hr

Amended with enhanced soil 0 to 15 761 235 767 0 0 0
15 to 30 4,994 913 0 0 0 0
30 to 45 8,460 25 18 0 0 0

Amended with autoclaved enhanced soil 0 to 15 1,456 810 211 27 0 0
15 to 30 4,913 27 0 0 0 0
30 to 45 2,103 36 84 0 0 0

Nonenhanced soil 0 to 15 1,321 808 0 2 0 0
15 to 30 6,343 42 0 0 0 0
30 to 45 755 66 1 0 0 0


Microplots were broadcast fumigated with 1,3-D at 168 liters/ha, and data are means of 18 replications.


'Enhanced and autoclaved enhanced soils (4 kg) were placed in the center of each microplot and mixed with the remaining soil to 30 cm


deep.


bA 100 cm3 subsample was transferred from each composite soil sample and then placed into a 164 ml ultraviolet-stabilized container
(Stewe and Sons, Corvallis, OR). Each container had 40 cm3 heat-pasteurized sandy-loam soil at the bottom.

CA 3-week-old tomato seedling, Lycopersicon esculentum Mill. cv. Rutgers, was transplanted into each container. Root-knot nematode
eggs were extracted from galled roots with a 0.5% sodium hypochlorite solution (Hussey and Barker, 1973).










Table 2-3. Regression analysis used to interpret the effects of 1,3-dichloropropene" exposure periods on Meloidogyne spp. gall production
on tomato grown in microplots that were amended with enhanced soil (previously identified to degrade 1,3-dichloropropene [1,3-D] at an
accelerated rate [Ou et al., 1995]), autoclaved enhanced soil, and nonenhanced soil.



Treatments Depth (cm) Response R2 P value

Amended with enhanced soil 0 to 15 Y= 89- 2.2X+ 0.0 IX2 0.93 0.05
15 to 30 Y= 80 2.3X+0.01X2 0.84 0.1
30 to 45 Y= 92 2.7X+ 0.02X2 0.79 0.1

Amended with autoclaved enhanced soil 0 to 15 Y=83 -2.OX+0.01X2 0.89 0.05
15 to 30 Y= 116 3.4X+0.02X2 0.80 0.1
30 to 45 Y= 132 3.9X+ 0.02X2 0.79 0.1

Nonenhanced soil 0 to 15 Y=61 1.4X+0.01X2 0.94 0.05
15 to 30 Y= 103 3.1X+0.02X2 0.79 0.1
30 to 45 Y=23 -2.4X+0.01IX2 0.79 0.1


aMicroplots were broadcast fumigated with 1,3-D at 168 liters/ha and data are means of 18 replications.


bEnhanced and autoclaved enhanced soils (4 kg) were placed in the center of each microplot and mixed with the remaining soil to 30 cm


deep.


CA 100 cm3 subsample was transferred from each composite soil sample and then placed into a 164 ml ultraviolet-stabilized container
(Stewe and Sons, Corvallis, OR). Each container had 40 cm3 heat-pasteurized sandy-loam soil at the bottom.

dA 3-week-old tomato seedling, Lycopersicon esculentum Mill. cv. Rutgers, was transplanted into each container. The number of galls was
estimated by counting those that were visible on the root system 6 weeks later.










Table 2-4. Regression analysis used to interpret the effects of 1,3-dichloropropenea exposure periods on Meloidogyne spp. egg production
on tomato grown in microplots that were amended with enhanced soil (previously identified to degrade 1,3-dichloropropene [1,3-D] at an
accelerated rate [Ou et al., 1995]), autoclaved enhanced soil, and nonenhanced soil.



Treatment Depth (cm) Response R P value

Amended with enhanced soil 0 to 15 ns
15 to 30 Y=6,860- 196X+0.01X2 0.86 0.1
30 to 45 Y= 6,957 -207X+0.02X2 0.79 0.1

Amended with autoclaved enhanced soil 0 to 15 Y= 1,452 32X+ 0.2X2 0.99 0.001
15 to 30 Y= 4,044- 121X+ 0.8X2 0.79 0.1
30 to 45 Y= 1,739 5 IX+0.3X2 0.79 0.1

Nonenhanced soil 0 to 15 Y= 1,345 32X+ 0.2X2 0.95 0.05
15 to 30 Y= 5,223 156X+X2 0.79 0.1
30 to 45 Y= 641 18X+0.1X2 0.84 0.1


aMicroplots were broadcast fumigated with 1,3-D at 168 liters/ha and data are means of 18 replications.


bEnhanced and autoclaved enhanced soils (4 kg) were placed in the center of each microplot and mixed with the remaining soil to 30 cm


deep.


cA 100 cm3 subsample was transferred from each composite soil sample and then placed into a 164 ml ultraviolet-stabilized container
(Stewe and Sons, Corvallis, OR). Each container had 40 cm3 heat-pasteurized sandy-loam soil at the bottom.

dA 3-week-old tomato seedling, Lycopersicon esculentum Mill. cv. Rutgers, was transplanted into each container. Root-knot nematode
eggs were extracted from galled roots with a 0.5% sodium hypochlorite solution (Hussey and Barker, 1973).










Table 2-5. Galls induced by Meloidogyne spp. second-stage juveniles (J2) that were extracted from infested soil from microplots amendedb
with enhanced soil (previously identified to degrade 1,3-dichloropropene [1,3-D] at an accelerated rate [Ou et al., 1995]), autoclaved enhanced soil,
and nonenhanced soil.

Number of galls per root system

Before fumigation After fumigation

Treatment Depth (cm) 24 hr 24 hr 48 hr

Amended with enhanced soil 0 to 15 101 12 0
15 to 30 95 0 0
30 to 45 122 0 0

Amended with autoclaved enhanced soil 0 to 15 67 2 0
15 to 30 129 0 0
30 to 45 43 0 0

Nonenhanced soil 0 to 15 0 0 0
15 to 30 72 63 0
30 to 45 44 0 0

Microplots were broadcast fumigated with 1,3-D at 168 liters/ha, and data are means of six replications.

aSoil samples were removed from six microplots 24 hours before fumigation and 24 and 48 hours after fumigation.

bEnhanced and autoclaved enhanced soils (4 kg) were placed in the center of each microplot and mixed with the remaining soil to 30 cm
deep.

'The number of J2 per 100 cm3 of soil was counted and these nematodes were placed in containers filled with 140 cm3 of heat-pasteurized
soil and a tomato seedling was transplanted into the soil. The number of galls was determined by counting those that were visible on the root system
6 weeks later.










Bioassay 3: Galls on tomato roots were evaluated 7 weeks after fumigation

determine if the enhanced soil that was amended to the microplots had a negative impact

on root-knot nematode control. All plots were fumigated and no differences in gall

indicies were noted on tomato transplanted directly into the microplots 48 hours following

fumigation (P 0.05). The average gall indices for soil amended with enhanced soil, soil

amended with autoclaved enhanced soil, and nonenhanced soil were 0.8, 1.2, and 0.9,

respectively. The gall indices ranged from 0 to a high of 7, which is 70% of the root

system being galled. There was a low level of root-knot nematodes that escaped

fumigation (Alphey and Boag, 1987; Been and Schomaker, 1999; McKenry et al., 1977).

A variety of factors, e.g., tolerant life stages (cysts and cryptobiotic stages), position of

the nematode in the soil profile, such as, depth of the nematode in the soil profile or in a

clumps of soil, survival in plant tissue (Schroeder et al., 2000), differences in soil moisture

or the presence of restrictive layers in the soil play a role in the survival of a low level of

plant-parasitic nematodes after fumigation with 1,3-D (McKenry and Thomason, 1976b).

1,3-D Degradation: Regardless of treatment, soil collected from each of the

microplots did not degrade 1,3-D at an accelerated rate (Ou, pers. comm.). Consequently,

the addition of enhanced soil to the microplots did not result in an enhanced degradation

of 1,3-D over the course of this experiment. There was no detectable effects of the

enhanced soil on the efficacy of 1,3-D. The effect of 1,3-D on root-knot nematodes over

time, however, provided useful information on the rapidity of its killing action.










Field Study

Nematode numbers that entered roots were highly variable among samples

collected from each of the two field sites. In the enhanced field soil, the number of J2 that

entered roots in the untreated soil was greater than occurred in fumigated soil (P < 0.05)

(Table 2-6), but when fumigated enhanced soil and nonenhanced soil were compared the

number of nematodes entering tomato roots was not different (P 0.5). In the

nonenhanced soil, the number of J2 that entered the roots in the untreated soil was higher

than in the fumigated soil (P < 0.05). Soil from both sites did not differ in the number of

J2 per root system (P > 0.05) (Table 2-6).

The number of infective juveniles that penetrated the root systems of tomato

seedling was reduced with fumigation. There were no differences in the number of

juveniles that entered the roots of tomato from fumigated enhanced and nonenhanced soil,

suggesting that the enhanced soil did not affect the efficacy of 1,3-D.



Greenhouse Study

Fumigation with 1,3-D lowered the number of infective juveniles that penetrated

tomato roots (Table 2-7) (P < 0.1). More root-knot nematodes were detected in tomato

roots from both the untreated enhanced and nonenhanced soils (P < 0.1). The number of

J2 per root system in tomato roots from both enhanced and nonenhanced soils did not

differ (P 0.1). The number of J2 per root systems from the 1,3-D fumigated soils in the

enhanced and the nonenhanced soil also were not different (P > 0.05). Less than two

nematodes per root system were detected in the fumigated enhanced soil and an average








42

of 0.2 nematodes penetrated the root systems grown in nonenhanced soil after fumigation.

Although 1,3-D greatly reduced the root-knot nematode densities, not all J2 were killed or

rendered noninfective. A low level of root-knot nematodes escaped fumigation which

confirms previous reports (Alphey and Boag, 1987; Been and Schomaker, 1999; McKenry

et al., 1977). These data also support the field data that shows soils identified to degrade

1,3-D does not decrease the efficacy of 1,3-D on root-knot nematodes. The effects of

1,3-D on plant-parasitic juveniles is rapid, with 95% plus nematodes killed within 48 to 72

hours, whereas Ou et al. (1995) reported that degradation of 1,3-D occurred in 5 days.











Table 2-6. Comparisons of the number of Meloidogyne incognita second-stage juveniles
in tomato roots grown in 1,3-dichloropropene (1,3-D, 112 liters/ha) treated or untreated field soil
previously identified as enhanced for degrading 1,3-D (Ou et al., 1995) at an accelerated rate, and
in soil without a history of 1,3-D fumigation.

Treatment Siteb J2/root system'


Untreated Enhanced 30.0
Fumigated Enhanced 0.3*

Untreated Nonenhanced 118.7
Fumigated Nonenhanced 1.7*

Untreated Nonenhanced 118.7
Untreated Enhanced 30.0

Fumigated Nonenhanced 1.7
Fumigated Enhanced 0.3


aThe enhanced soil was fumigated at least seven times at a rate of 55 to 110 liters/ha over
the past 13 years (Ou et al., 1995).

bThe nonenhanced site had been planted with bahiagrass for the past 25 years and had no
known history of exposure to 1,3-D.

'Data are means of 10 replicates. The data were subjected to analysis by the Kruskal-
Wallis test for completely randomized nonparametric data. Asterisks indicate differences from
corresponding control at P < 0.05.









44

Table 2-7. Comparisons of the number of Meloidogyne incognita second-stage juveniles
in tomato roots grown in PCV tubes filled with enhanced soil known for degrading 1,3-
dichloropropene (1,3-D) (Ou et al., 1995) at an accelerated rate and with soil without a history of
fumigation', both of which were treated with 1,3-D (84 liters/ha).

Treatment Sitec J2/root system

Untreated Nonenhanced 14.5 a

Untreated Enhanced 25.6 a

Fumigated Nonenhanced 0.2 b

Fumigated Enhanced 1.8 b


Data are means of nine replicates. The data were subjected to analysis by Duncan's
multiple-range test (P < 0.1).

aThe enhanced soil was fumigated at least seven times at a rate of 55 to 110 liters/ha over
the past 13 years (Ou et al., 1995).

bThe nonenhanced site had been planted with bahiagrass for the past 25 years and had no
known exposure to 1,3-D.

cSoil was collected from two sites (enhanced for degrading 1,3-D and nonenhanced) at the
University of Florida Green Acres Agronomy Research Farm in Alachua county, Florida.














CHAPTER 3
RATE RESPONSE OF 1,3-DICHLOROPROPENE FOR NEMATODE CONTROL IN
SPRING SQUASH IN DEEP SANDY SOILS


Introduction



The phase-out of methyl bromide by 2005 (Anonymous, 1999a) creates a need for

effective and reliable alternative chemicals to control pests and pathogens of high-valued

vegetables, nursery crops, some ornamentals, and turfgrass renovations or new

installations. The soil fumigant, 1,3-dichloropropene (1,3-D) plus chloropicrin, is

considered a strong candidate to replace methyl bromide for the vegetable industry in

Florida (Stephans, 1996). It is one of the few registered fumigants available for broad-

spectrum management of nematodes.

1,3-Dichloropropene is toxic to plant-parasitic nematodes (Fletcher, 1956;

Youngson and Goring, 1970), and is currently recommended in Florida for a variety of

vegetable and agronomic crops (Dunn and Noling, 1995). Although the fumigant has

been used successfully to control plant-parasitic nematodes on many crops (Dickson,

1985; Weingartner and Shumaker, 1990), its performance can vary (Dickson and Hewlett,

1988; Melton, 1996). Factors that may affect its efficacy in deep, sandy soils include

depth of placement, sealing after injection, soil moisture and temperature, and dosage.









Recommended application rates for field crops in Florida range from 56 to 112

liters/ha (Dunn and Noling, 1995), but there is a tendency among agriculturalists to lower

rates to reduce crop production cost. The objective of this study was to evaluate the

efficacy of four rates of 1,3-D for the management of root-knot nematodes in deep sandy

soils of Florida.



Materials and Methods



Two experiments (trials 1 and 2) were conducted during 1999 in fields at the

University of Florida Green Acres Agronomy Research Farm in Alachua County, Florida.

The soil was an Arredondo fine sand (92% sand, 5% silt, 3% clay; 4% organic matter; pH

6.1). The sites were naturally infested with Meloidogyne spp., Criconemoides spp.,

Pratylenchus brachyurus (Godfrey) Filipjev and Stekhoven, and Paratrichodorus minor

(Colbran) Siddiqi. Because root-knot nematodes were patchy in distribution, both sites

were infested in fall 1997 with Meloidogyne arenaria (Neal) Chitwood race 1 and M

incognita (Kofoid and White) Chitwood race 1, each cultured on tomato (Lycopersicon

esculentum Mill. cv. Rutgers). Heavily galled, chopped tomato roots (ca. 600 g of galled

roots/1 2.2 m row) were placed in 15-cm-deep open furrows spaced 0.9m apart. Hairy

vetch (Vicia villosa Roth) was planted to both sites in fall 1997. Peanut (Arachis

hypogaea L. cv. Sunrunner) was planted in the summer of 1998, and hairy vetch was

again planted in the fall of the same year as a winter cover crop. The hairy vetch was

plowed under 30 March 1999.








47
The experimental design was a randomized complete block with six replications.

Plots were 12.2 m long with a row spacing of 90 cm, and a 2.4-m alley separated each

block. An untreated fallow plot on either side of each treated plot served as a border.

1,3-D was broadcast-injected 13 April 1999 at 0, 56, 84, 112, or 168 liters/ha 30 cm deep

with six conventional chisels (two forward and four rear inclined double-beveled blades)

spaced 30 cm apart. The two outer chisels were forward-swept and the four inner chisels

were backward-swept. A 1.8-m-wide, two-gang disk was run through each plot

immediately after fumigant injection cutting the soil surface ca. 15 cm deep to form a

surface seal and break the continuity of chisel traces (1 to 1.3-cm openings in the upper

soil profile left by conventional chisels). Soil moisture at both sites at the time of

fumigation was 13%, and the soil temperature at 10 cm deep on the day of fumigation

increased from a minimum of 25 C to a maximum of 28 C.

Twelve cores (2.5-cm-diam., 20-cm-deep) of soil from each plot were taken with a

cone-shaped sampling tube at pretreatment (Pi) on 12 April, mid-season (Pm) on 16 May,

and final harvest (Pf) on 23 June. Soil cores from each plot were combined and

nematodes were extracted from a 100-cm3 subsample by a centrifugal-flotation method

(Jenkins, 1964). All plant-parasitic nematodes were counted.

Summer squash, Cucurbitapepo L. cv. Sunex 9602, was planted on 20 April

1999, 7 days after fumigation. Three weeks after planting, seedlings were thinned to one

plant every 45 cm of row. The center 12 plants per plot were designated for yield.

Preplant, 55 kg/ha of 10-10-10 fertilizer were applied broadcast and incorporated.

Postplant, a total of 16.5 kg/ha of nitrogen in the form ofNH4NO3 and 16.5 kg/ha of









potassium in the form of K20 was divided into six weekly applications applied by drip

irrigation beginning 2 weeks after seedling emergence. Double-wall drip tubing (Chapin

Twinwall, Watertown, NY), with emitters spaced 30 cm apart and a flow rate of 62

ml/minute/30.5 m, was placed 7.5 to 10 cm from the center of the bed for application of

water, fertilizer, and calcium. Calcium (8.4% CaO) at a rate of 0.19 liters/ha was applied

weekly beginning at flowering to reduce the incidence of blossom end rot. One hive of

honey bees (Apis mellifera L.) per 2.5 ha was placed 50 m from the fields at the time of

flowering to aid in pollination (Hochmuth et al., 1998).

Broadleaf weeds were managed with applications of naptalam and bensulide, and

foliar pathogens were managed with chlorothalonil and manzate (Hochmuth et al., 1998).

Methomyl was applied to manage insect pests. The plots were cultivated on 19 May for

weed control.

Squash was first harvested on 30 May 1999, and harvesting continued three times

a week for 3 weeks. The number and weight of marketable squash per plot were

recorded. Sixty-five days after planting an overall rating for plant growth was made for

each plot based on a subjective scale of 1 to 10 (1 = stunted, chlorotic, or dead plants, 10

= full, lush, green growth). Root-knot nematode galling was determined on 23 June for all

12 plants in each plot based on an index scale of 0 to 10 (0 = no galls on roots and 10 =

100% of the root system galled) (Barker et al., 1986).

Reproductive factor for the Meloidogyne spp. was determined by Pm/Pi and Pf/Pi

ratios, where Pi = initial population density, Pm = population density 34 days after

planting, and Pf= population density 65 days after planting. The means for each









treatment at the three sampling times (Pi, Pm, and Pf) were calculated and used for the

proportion calculations.

Daily minimum and maximum soil temperatures between planting and harvest were

obtained from the Agronomy Department climatological data collected from a NOAA-

approved weather station located within 100 to 150 m of both trials. The mean daily soil

temperature 10 cm deep was calculated by averaging the daily minimum and maximum

temperatures. Degree days were calculated by temperature summations (Arnold, 1960)

with the formula, degree day = (Tmax + Tmin)/2 z, where Tmax, was the maximum daily soil

temperature, Tmin was the minimum daily soil temperature, and z was the lower

developmental threshold of 10 C.

Quadratic regression analyses were used to estimate 1,3-D dosage responses (SAS

Institute, Cary, NC). Orthogonal contrasts were performed on gall indices and rates of

1,3-D. All nematode data were transformed with loge (x + 1) before analysis.



Results



The results for each trial are presented separately because of a significant trial x

treatment interaction for most variables. The R2 values were low for most models because

of variability in estimates of population densities of Meloidogyne spp. and fruit

production, but the overall model effects were significant (P 0.1).

An increase in total fruit production was observed in both trials when the plants

were grown in soil fumigated with 1,3-D (trial 1 :Y = 42.30 + 0.47X 0.002X2, R2 = 0.28,









P < 0.05; trial 2: Y = 13.49 + 0.17X 0.0006X2, R2 = 0.2, P < 0.1). In trial 1, the mean

number of fruit increased 52, 54, 78, and 66% compared to the untreated control for 56,

84, 112, and 168 liters of 1,3-D/ha, respectively. In trial 2, the mean number of fruit

increased 48, 113, 64, and 96% compared to the untreated control for 56, 84, 112, and

168 liters of 1,3-D/ha, respectively.

Marketable yield of squash increased in soil fumigated with 1,3-D compared to

plants grown in untreated soil (Fig. 3-1A). In trial 1, the greatest increase in yield (a 2.1-

fold increase compared to the untreated control) was from plants grown in soil that

received 1,3-D at 112 liters/ha. In trial 2, the greatest yield increase was from plants

grown in soil fumigated with 84 liters of 1,3-D/ha (Fig. 3-1A).

Overall plant growth in both trials was improved with fumigation (Fig. 3-1B), with

nearly a 1.9-fold improvement in the growth ratings in the fumigated plots compared to

the growth ratings in the untreated control. Plants from the untreated control were

stunted, whereas plants in plots fumigated with 1,3-D had a growth rating of 6 to 7 (Fig.

3-1B).

Fumigation decreased the severity of root galling caused by Meloidogyne spp. in

both trials (Fig. 3-2). The untreated controls in trial 1 had a mean gall rating of 71% of

the roots being galled and in trial 2, the untreated control has a rating of 34%. A decrease

in the galling index was observed for all fumigated plots compared to the untreated

control. The mean galling indices for the untreated control and fumigated plots were

lower in trial 2 than in trial 1. Gall indices from fumigated plots from both trials combined

had a lower mean gall index than the untreated control (P < 0.0001) (Table 3-1). More












25

S20

15

10
--- -- -
5 --- Trial 1 Trial 2
0 -----------------------
0
10
B
3-- 8

-0


S4



| 2 Trial 1 Trial 2
0-- ---
0
0 56 84 112 168
1,3-Dichloropropene (liters/ha)


Fig. 3-1. Effects of rates of 1,3-dichloropropene for control ofMeloidogyne spp.
on the marketable yield and growth of squash produced in two trials conducted at the
University of Florida Green Acres Agronomy Farm in 1999. Each data point is a mean of
six replicates. A) Quadratic models described a positive response between squash yield
and the rate of 1,3-dichloropropene applied in both trials. Trial 1: Y = 14.54 + 0.24X -
0.001X2, R2 = 0.28, P < 0.05; trial 2: Y = 3.78 + 0.06X 0.0002X2, R2 = 0.21, P < 0.05.
B) Effects of rates of 1,3-dichloropropene on the growth of squash 65 days after planting.
The plants were rated on a scale of 1 to 10 (1 = stunted, chlorotic, or dead plants, 10 =
full, green, lush growth). Quadratic models described a positive response between the
growth of the plants and the rate of 1,3-dichloropropene applied in both trials. Trial 1: Y
= 3.17 + 0.06X 0.0003X2, R2 = 0.28, P < 0.05; trial 2: Y= 3.68 + 0.04X 0.0001X2, R2
= 0.24, P < 0.05.
















2



1.5 -
+ Ik
\ Trial 1
S1\ \ -
= 1 \Trial 2
"-" \ \-.--
0
Q. 0.5 \
s.


o -
0 56 84 112 168
1,3-Dichloropropene (liters/ha)



Fig. 3-2. Effects of rates of 1,3-dichloropropene on root galling of squash induced
by Meloidogyne spp. 65 days after planting in two trials conducted at the University of
Florida Green Acres Agronomy Farm in 1999. Each data point is a mean of six replicates.
The data were transformed with log, (x + 1) before analysis. Trial 1: Y = 1.84 0.02X +
0.00008X2, R2 = 0.53, P = 0.0001; trial 2: Y = 1.37 0.02X + 0.00007X2, R2 = 0.81, P =
0.0001.








53
galls were produced on plants grown in soil fumigated with 56 liters of 1,3-D/ha in trial 2

compared with plots fumigated with 84, 112, and 168 liters/ha (Table 3-1).

Population densities of Meloidogyne spp. at 34 DAP (Pm) in trial 1 and 65 DAP

(Pf) in both trials were lower in plots fumigated with 1,3-D than in untreated plots (Fig.3-

3A,B). At 34 DAP in trial 1 (Fig. 3-3A), a mean of 35 J2/100 cm3 of soil was detected in

the untreated control compared to 0.3 J2/100 cm3 of soil in plots fumigated with 84 liters

of 1,3-D/ha. At 65 DAP, the untreated control had 77 J2/100 cm3 of soil, whereas

population densities remained low in fumigated plots. At this time in trial 2, the

population density of J2 decreased with soil fumigation with 15, 23, 70, and 23-fold

decreases in numbers of J2 compared with the control (Fig. 3-3B). The reproductive

factor (Pm/Pi and Pf/Pi ratios) for the untreated control and all four 1,3-D rates in both

trials were less than 1 (data not shown). The reproductive factor in the fumigated plots

were lower than the untreated control.

The population densities of Paratrichodorus minor, Pratylenchus brachyurus, and

Criconemoides spp. were low at 34 and 65 DAP in both trials; therefore, the results are

not reported. The population density of Criconemoides spp. at both sampling times in

both trials decreased after fumigation. Criconemoides spp. at 34 DAP decreased with

increased rates of 1,3-D in trial 1, Y = 3.7 0.007X 0.00004X2 (R2 = 0.41; P = 0.001),

whereas at 65 DAP, their numbers were reduced from 156/100 cm3 soil in the untreated

control to 71, 37, 30, and 4/100 cm3 of soil in plots fumigated with 56, 84, 112, and 168

liters of 1,3-D/ha, respectively. Transformed data fit the quadratic model Y = 1.3 0.01X

- 0.0004X2 (R2 = 0.24; P = 0.05). The response of Criconemoides spp. to rates of 1,3-D









Table 3-1. Contrasts to compare gall indices of root-knot nematode on squash
and rates of 1,3-dichloropropene in trials 1 and 2b.
Contrast Mean F value P

Trial 1

Untreated 1.85 22.6 0.0001

Fumigated 0.35

Trial 2

Untreated 1.37 91.1 0.0001

Fumigated 0.16

56 Liters/ha 0.50

vs. 84 Liters/ha 0.07 7.2 0.05

vs. 112 Liters/ha 0.07 7.0 0.05

vs. 168 Liters/ha 0 9.6 0.01


aRoot galling was determined 65 days after planting based on a scale of 0 to 10 (0
= no galls and 10 = 100% of the root system galled) (Barker et al., 1986). The data were
transformed with loge (x + 1) before analysis.

bTwo experiments (trials 1 and 2) were conducted during 1999 in fields of close
proximity at the University of Florida Green Acres Agronomy Research Farm in Alachua
County, Florida.

c1,3-Dichloropropene was applied-broadcast at 0, 56, 84, 112, and 168 liters/ha.
Data for the untreated control and each fumigant rate are means of six replications.
Pooled data for the 1,3-dichloropropene fumigated plots are the means of 24 plots.











4

3.5 A Pm f
M
g 3 \
1',
S2.5

2
^ \

+ 1.5


0 N
", l

0.5 -. ,

n2 I I I ---.....----- I
2.5

B Pf



S1.5 "


+ 1
01.5
I-*




I0
\
\*
0 i--- "- I ---
0 56 84 112 168
Rate 1,3-di chloropropene (liters/ha)




Fig. 3-3. Effects of rates of 1,3-dichloropropene on the density of Meloidogyne
spp. second-stage juveniles (J2) per 100 cm3 soil in two trials conducted at the University
of Florida Green Acres Agronomy Farm in 1999. Sampling occurred at 34 (Pm) and 65
(Pf) days after planting. Each data point is a mean of six replicates. Data were
transformed with loge (x + 1) before analysis. A) Trial 1: Y = 2.0 0.03X + 0.0002X2, R2
= 0.28, P < 0.05 (Pm), and Y = 3.74 0.05X + 0.0002X2, R2 = 0.67, P < 0.0001 (Pf). B)
Trial 2: Y = 1.98 0.03X + 0.0001X2, R2 = 0.41, P < 0.01 (Pf).









in trial 2 were similar to those in trial 1 (data not shown) except that the number of

Criconemoides spp. recovered from untreated plots and from plots that were fumigated

with 56 liters of 1,3-D/ha was higher than from plots treated with 84 and 168 liters of 1,3-

D/ha (P 0.05).

The daily degree days from the day of planting until the completion of the study

ranged from 11.9 to 19.7 (Fig. 3-4A). The cumulative degree days from 20 April to 23

June 1999 was 1,098 (Fig 3-4B) therefore, it is estimated that there were sufficient heat

units for development of at least one generation of root-knot nematodes during the

interval of these two trials. Egg masses were observed on the squash roots at harvest.



Discussion



The number of fruit and total yield of squash increased when soil was fumigated

with 1,3-D. The overall number of fruit and total fruit weight were greater in trial 1 than

in trial 2, despite higher numbers ofMeloidogyne spp. in the trial 2 site. Increases in yield

after fumigation are commonly reported with many kinds of crops (Kinloch and Rich,

1998; Schenk, 1990; Weingartner et al., 1993). In these trials, no phytotoxicity was

observed in the squash seedlings at any of the rates of 1,3-D applied. Plants that were

grown in fumigated soil were uniform in growth, consistent in plant stand, and larger than

plants grown in untreated soil.

Fumigation with 1,3-D decreased the number of second-stage juveniles of

Meloidogyne spp. in soil and the number of galls on squash roots. In trial 2, the higher










20
A

U18
0

A
16


S14


S12


10 -------------

1,200 ........

1,000 B
A
800

600
-o/
> 400 /


S200

0 -
110 116 122 128 134 140 146 152 158 164 170
Calender date (20 April to 23 June 1999)




Fig. 3-4. Degree days for Meloidogyne spp. were based on maximum and
minimum soil temperatures 10 cm deep recorded from 23 April to 23 June 1999.
Temperatures were obtained from the Agronomy Department climatological data
collected from a NOAA approved weather station. The lower developmental threshold
was 10 C. A) Daily degree days for the duration of both trials. B) Degree day
accumulations for the duration of both trials.









rates (84, 112, and 168 liters of 1,3-D/ha) were more effective in reducing root-gall

severity than the low rate of 56 liters/ha. However, in plots fumigated with 1,3-D at a rate

of 56 liters/ha only a low level of root galling was observed on roots from fumigated plots

at final harvest with 15% of roots galled in trial 1 and 7% in trial 2. Squash, which is an

excellent host for Meloidogyne spp., is a short-season crop (Robinson and Decker-

Walters, 1997) that is normally planted in late winter and early spring (February to April)

and late summer and early fall (August to September) in north Florida (Hochmuth et al.,

1998). In the former case, soil temperature may remain relatively low, thereby slowing the

rate of development of root-knot nematodes.

Temperature influences the development of root-knot nematodes (Vrain and

Barker, 1978; Wong and Mai, 1973), and degree days have been used as a means to

measure the rates of development of several nematode species including root-knot

nematodes (Arnold, 1960). Tyler (1933) found that the minimum temperature for

development of Meloidogyne spp. was between 9 C and 10 C and it took between

6,000 and 8,000 heat units for the nematode to develop from second-stage juveniles to

egg-laying females in tomato roots. In the field, approximately 9,000 heat units were

necessary for the development of M javanica in tobacco roots (Milne and DuPlessis,

1964). Under the conditions of this study, the soil temperature remained below 20 C at

10 cm deep. According to the relatively low number of heat units accumulated, root-knot

nematodes would have just reached reproductive maturity at the conclusion of these trials.

Unfortunately, we did not measure egg production; however, abundant root-knot

nematode galls formed on roots, and egg masses were observed, especially in the








59
untreated control. The extracted second-stage juveniles in soil at 34 and 65 DAP in both

trials were lower than the initial estimations of population density and the rate of

reproduction was below 1 and in a few cases, the nematode reproductive rate was zero.

Therefore, we conclude there was only one generation of the nematode developed during

the period of these experiments.

For most crops planted in Florida, whether in the northern or southern regions,

there is a relatively long growing season and root-knot nematodes in soil even at low

densities generally have sufficient heat units to complete three, four, or more generations

per year thereby the population densities substantially increased. For long-season crops

such as peanut, 1,3-D applied-broadcast at low rates (28, 47, and 65 liters of 1,3-D/ha)

provided only marginal control of M arenaria race 1 (Dickson and Hewlett, 1988).

However, for a short-season crop such as squash grown in early spring months in north

Florida, low rates of 1,3-D may be all that is necessary for satisfactory management of

root knot nematodes.














CHAPTER 4
COMPARISON OF CHISEL TYPE, SEALING, AND APPLICATION DEPTH ON
THE EFFICACY OF 1,3-DICHLOROPROPENE IN DEEP SANDY SOILS


Introduction



The fumigant 1,3-dichloropropene (1,3-D) is recommended broadly to manage

plant-parasitic nematodes for a wide variety of vegetable and agronomic crops (Dickson,

1985; Dunn and Noling, 1995; Weingartner and Schumaker, 1991). 1,3-Dichloropropene

volatilizes in soil (Gan et al., 1995; 1998; Goring, 1957; Lembright, 1990) and the

maintenance of the fumigant in the soil at a lethal dose after application is of crucial

importance. Rate (Kinloch and Rich, 1998; Schenk, 1990), methods of application, e. g.

row vs. broadcast (Sipes et al, 1993; Weingartner and Shumaker, 1991), placement depth

(Rodriguez-Kdbana and Robertson, 1987), and application equipment (Lembright, 1990,

Schneider et al., 1996) are all key factors affecting the efficacy of 1,3-D. However,

proper sealing of the soil surface after application may be one of the most important

factors to prevent premature losses of the fumigant from the soil.

1,3-Dichloropropene is generally applied with conventional chisels (forward or

backward-swept) that leaves an open slit in the soil 1 to 1.3 cm wide called a chisel trace

(Anonymous, 1996b; Lembright, 1990). The fumigant moves quickly into the air space

left by the chisel, thus allowing it to escape rapidly into the atmosphere (Goring, 1957;








61
Lembright, 1990). Disking immediately after injection of 1,3-D is one method to eliminate

chisel traces (Anonymous, 1996b), thereby the fumigant is sealed in the soil. Other

methods that have been suggested to close chisel traces includes the use of fluted colters,

press wheels, a cultipacker, a roller, a spike-tooth harrow, and irrigation (Dallimore, 1955;

Lembright, 1990; Rhoades et al., 1962). Unfortunately, none of these methods work well

in all situations and there is little or no data to support the use of any one of these

methods.

The necessity of disking after fumigation to break up chisel traces adds an

additional cost and time factor. To eliminate this step a different type of chisel for

placement of fumigants that does not leave a chisel trace was developed. This chisel, was

called a parachisel. This type of chisel minimally disturbs the soil surface because the

blade runs through the soil on a 45 plane from vertical and the soil is raised slightly and

falls back as the blade passes. Consequently, no chisel trace is left behind. In-row or

broadcast applications can be made with parachisels. The design of the parachisel is

similar to that of a soil injection chisel developed by Dow Chemical Co. (Midland, MI) to

provide a suitable means to inject or place chemicals into bedded land with minimal

disturbance of the beds to the degree that would occur with conventional chisels (Miller et

al., 1967).

Depth of application may be a factor that increases the efficacy of 1,3-D. Thomas

(1994) reported that an increase in the injection depth for in-row fumigation with 1,3-D

from 28 cm to 48 cm increased yields of red chili pepper. In contrast, Rodriguez-Kdbana

and Robertson (1987) reported that in-row fumigation with 56 liters of 1,3-D/ha in sandy








62
loam soil was effective to control J2 of M arenaria race 1 when the fumigant was applied

25 cm deep compared to 8, 15, and 36 cm. Most researchers evaluated shallow and deep

placement in-row, bed, or broadcast treatments. Direct comparisons of in-row and

broadcast fumigation and depth of application with 1,3-D in deep sandy soils needs to be

understood better.

Soil fumigation is costly and placement of the fumigant in-row will reduce the

amount of material used and therefore reduce the cost. The crop response may be equal

or superior to soil that received broadcast applications (Lear and Thomason, 1956;

Lembright, 1990; Rhoades et al., 1962; Zehr and Golden, 1986). However, others have

reported in-row fumigation to be less effective than broadcast fumigation (Weingartner

and Shumaker, 1991). Our objectives were to compare parachisels versus conventional

chisels for in-row and broadcast fumigation with 1,3-D, with and without disking, and to

compare the efficacy of deep and shallow applications of 1,3-D with different equipment

(conventional, parachisel, and subsurface hooded sweep chisel) to manage of root-knot

nematodes in a deep sandy soil.



Materials and Methods



Two experiments were conducted during 1998 and 1999 at the University of

Florida Green Acres Agronomy Research Farm located in Alachua County, Florida.

Experiment 1 was conducted to evaluate different types of chisels and depth of placement

on the efficacy of 1,3-D for management ofMeloidogyne arenaria (Neal) Chitwood race








63
1 in peanut. Two trials of experiment 1 were conducted concurrently in 1998 (trials 1 and

2) in fields of close proximity and a third trial (trial 3) was conducted in 1999. Experiment

2 (spring of 1999) was conducted to compare in-row and broadcast fumigation of 1,3-D

with conventional chisels and parachisels to manage Meloidogyne spp. in spring planted,

straight-neck yellow squash.

The soil was an Arredondo fine sand (91.5% sand, 5.5% silt, 3% clay; and 1.5%

organic matter; pH 6.3). The sites for both experiments were naturally infested with

Meloidogyne spp., Criconemoides spp., Pratylenchus brachyurus (Godfrey) Filipjev and

Stekhoven, and Paratrichodorus minor (Colbran) Siddiqi. Because the population of

root-knot nematodes was patchy in distribution, the sites were infested in the fall of 1997

with Meloidogyne arenaria race 1 and M incognita (Kofoid and White) Chitwood race 1,

each grown on tomato (Lycopersicon esculentum Mill. cv. Rutgers). Heavily galled,

chopped tomato roots of both species (600 g of galled roots per 12.2 m of row) were

placed in 15-cm-deep open furrows spaced 90 cm apart. Hairy vetch (Vicia villosa Roth)

was planted to the sites in fall 1997 and 1998 as a winter cover crop. The hairy vetch was

plowed under 8 May 1998 and 30 March 1999.

The experimental design for both experiments was a randomized complete block

with six replications. Plots were 12.2 m long with a row spacing of 91 cm and a 2.4 m

alley separated each block. Peanut plots consisted of four planted rows with the outer

rows serving as borders. Squash plots also consisted of four rows; however, the outer

two border rows were fallow and squash was planted in the center two rows. The soil

temperature at 10 cm deep on the day of fumigation in 1998 (experiment 1) reached a









minimum of 28 C and a maximum of 34 C and soil moisture was 11%. The soil

temperature at 10 cm deep on the day of fumigation in 1999 (experiment 2) reached a

minimum of 25 C, and a maximum of 28 C, and soil moisture was 13%.

To determine nematode densities, 12 cores (2.5-cm-diam., 20 cm deep) of soil

from each plot were taken with a cone-shaped sampling tube at pretreatment (Pi)

(experiment 1 on 19 May 1998 and 4 April 1999; experiment 2 on 12 April 1999), mid-

season (Pm) (experiment 2 on 16 May 1999), and final harvest (Pf) (Experiment 1 on 23

October 1998 and 19 September 1999; Experiment 2 on 23 June 1999). Soil cores from

each plot were combined and nematodes were extracted from a 100 cm3 subsample by a

centrifugal-flotation method (Jenkins, 1964). All plant-parasitic nematodes were counted.

The reproductive factor for root-knot nematodes was determined by Pf/Pi ratios in

experiment 1, where Pi = the initial population density and Pf = final population density at

146 days after planting (DAP) in trials 1 and 2, and 150 DAP in trial 3; and Pm/Pi and

Pf/Pi ratios in experiment 2, where Pi = initial population density, Pm = population density

34 DAP, and Pf= population density 65 DAP, respectively. The means for each treatment

at the three sampling times (Pi, Pm, and Pf) were calculated and used to determine the

ratios.

The equipment used for soil injection of 1,3-D in these experiments was

conventional chisels, parachisels, and subsurface hooded sweep chisels. The conventional

chisels were forward swept or backward swept, straight, double-beveled blades (Fig. 4-1).

The prototype of the parachisel was designed and manufactured in the machine shop of

the University of Florida Agricultural and Biological Engineering Department (Fig. 4-2,4-








65
3,4-4A). This chisel is a rearward inclined, side-wise sloped on a 450 plane, single-beveled

blade that was manufactured from C1080 steel. A stainless steel tube (1/8th inch thread)

was welded onto the blade for chemical delivery (Fig. 4-4B). These chisels are

manufactured currently by Chemical Containers (Lake Wales, FL). The subsurface

hooded sweep chisel was an experimental prototype developed by Dow Chemical Co.

(Midland, MI). Each unit had a flat spray tip 8003 TeeJet (Spraying Systems Co.,

Wheaton, IL) spray nozzle mounted under each subsurface hooded sweep chisel (Fig. 4-

5).

All fumigation was done on a flat bed surface. For in-row fumigation with

conventional chisels and parachisels, two chisels per row spaced 25 cm apart were

centered over each of the two plot rows. One subsurface hooded sweep chisel per row

was used for in-row fumigation. For broadcast fumigation with conventional and

parachisels, six chisels, each spaced 30 cm apart, were used. The conventional chisels

were arranged as follows: two outer swept-forward chisels and four inner swept-back

chisels. Three subsurface hooded sweep chisels spaced 30 cm apart (0.9 m wide) were

used for bed fumigation. If a plot was disked after fumigation, a 1.8 m two gang disk was

run through it immediately after fumigation to cut the soil surface ca. 15 cm deep.

Experiment 1: In trials 1 and 2, 1,3-D was applied either broadcast, in bed, or in-

row at a rate of 84 liters/ha (78 ml/chisel/30.5 m of row). The broadcast applications

were done with conventional chisels set at either an injection depth of 20 cm or 30 cm

(both followed with or without disking). Bed treatment was done with three subsurface

hooded sweep chisels set at an injection depth of 30 cm. In-row applications were made








































Fig. 4-1. The conventional chisel used for in-row and broadcast fumigation of 1,3-
dichloropropene. The chisel was a forward or rear inclined, double beveled blade.


MIBman

NIP, -P ? :









67

























.. C



-.--
4W












Fig. 4-2. Parachisels used for in-row and broadcast fumigation of 1,3-
dichloropropene. The parachisel was a rearward inclined, side-wise sloped, single beveled
blade.








































PARACHISEL PATTERN PLAN VIEW
Not To Scale
Material: C1080 Steel




Fig. 4-3. A schematic of the parachisel used for fumigation with 1,3-dichloropropene.

00








69








A










4 304.8m.,
Side View


U l/Oth Inch Stainless Steel Pipe
-~~' ~for Chemical Deliven/ -- ^'/
OO O0118






254mm E [

0sB Plan View
25.4mm N. TO 5.





Fig. 4-4. Schematic drawings of the parachisel in side and plan views. A) Side
view of the parachisel. B) Plan view of the parachisel showing the stainless steel tube
used for delivery of the fumigant. The width of the tube is given in inches because of
industry standards for tube threading.












































Fig. 4-5. The subsurface hooded sweep chisel used for fumigation with 1,3-
dichloropropene.








71
with conventional chisels, parachisels, and subsurface hooded sweep chisels each set at an

injection depth of 30 cm. In trial 3, treatments were the same except that the subsurface

hooded sweep chisels were omitted and broadcast treatments of 1,3-D at 84 and 112

liters/ha (104 ml/chisel/30.5 m of row) applied with parachisels were added. Gauge

wheels were added to the fumigation rig to help control depth of placement.

Peanut cvs. Sunrunner and Georgia Green were planted 7 days after fumigation in

1998 and 1999, respectively. Broadleaf weeds were managed with a preplant application

of pendimethalin, and purple nutsedge (Cyperus rotundus L.) and yellow nutsedge

(Cyperus esculentus L.) were managed with postplant applications of imazamethipyr.

Foliar plant pathogens were managed by broadcast applications of chlorothalonil and

tubuconazol on a 10 to 14-day schedule. All pesticides were applied at labeled rates.

Gypsum was applied broadcast at a rate of 895 kg/ha at initiation of pegging.

Peanuts in trials 1 and 2 were dug on 22 October 1998 (146 DAP) and trial 3 was

dug 17 September 1999 (150 DAP). Three to five days later the pods from the center two

rows were combined and dried at 60 C until they reached 9.6% moisture content in 1998,

and 8.6% moisture content in 1999. The pods were weighed to determine yield. The

incidence of root-knot nematode galls on the pods was determined by taking a subsample

of 200 pods per plot from trials 1 and 2, and 100 pods per plot from trial 3. The number

of pods per subsample with root-knot nematode galls was counted.

Experiment 2: Treatments consisted of 1,3-D applied at a rate of 84 liters/ha in-

row and broadcast with conventional chisels and parachisels (both with or without

disking), and an untreated control. Plots that were broadcast fumigated had one row of









squash seeded down its center and plots that were in-row fumigated had two rows of

squash seeded on 91.4 cm centers. An untreated, fallow plot on either side of each treated

plot served as a border. Straight-neck yellow squash, Cucurbita pepo L. cv. Sunex 9602,

was seeded on 20 April 1999, 7 days after fumigation. Three weeks after planting,

seedlings were thinned to one plant every 45 cm of row. Twelve plants from the center of

each plot (either broadcast or in-row fumigated) were designated for yield. Fifty-five

kilograms per hectare of 10-10-10 (N-P-K) fertilizer was applied broadcast and

incorporated into the top 10 cm of soil before planting. A total of 16.5 kg/ha of nitrogen

in the form of NH4NO3 and 16.5 kg/ha of potassium in the form of K20 was divided into 6

weekly applications. Calcium (8.4% CaO) at a rate of 0.2 liters/ha was applied weekly

beginning at flowering to reduce the incidence of blossom-end rot. Double-wall drip

tubing (Chapin Twinwall, Watertown, NY) with emitters spaced 30 cm apart and a flow

rate of 62 ml/minute/30.5 m was placed 7.5 to 10 cm from the plant stems for application

of water, fertilizer, and calcium. One hive of honey bees (Apis mellifera L.) per 2.5 ha

was placed 50 m from the field at flowering to aid in pollination (Hochmuth et al., 1998).

Broadcast applications ofnaptalam and bensulide were incorporated 10 cm deep

for management of broadleaf weeds before planting (Hochmuth et al., 1998). The plots

were cultivated on 19 May for weed control. Foliar plant pathogens were managed with

applications ofchlorothalonil and manzate on a 10-day schedule (Hochmuth et al., 1998).

Methomyl was applied to manage insect pests. All pesticides were applied at labeled

rates.








73
Squash was first harvested on 30 May 1999 and harvesting continued three times a

week for 3 weeks. The number and yield of marketable squash per plot was recorded.

Root-knot nematode galling was determined on 23 June 1999 based on a scale of 0 to 10

(0 = 0 galls, 1 = 10% of the roots galled, 2 = 20% of the roots galled,...... 10 = 100% of

the roots galled) (Barker et al., 1986). Data were subjected to analysis by ANOVA and

means separated by Duncan's multiple-range test (SAS Institute, Cary, NC).



Results and Discussion



Experiment 1: The number ofMeloidogyne spp. J2 at harvest in trial 1 was lower

in all treatments except for bed fumigation with the subsurface hooded sweep chisel, than

in the untreated control (P _< 0.1) (Table 4-1). No differences were observed among

treatments in trials 2 and 3 (P > 0.1), except for the high number of J2 where parachisels

were used to apply 1,3-D at 112 liters/ha (Table 4-1).

In this study we evaluated in-row and broadcast fumigation at a shallow (20cm)

and deep (30 cm) placement of 1,3-D. In experiment 1, deeper placement did not increase

efficacy. In-row and broadcast fumigation with 1,3-D at 20 cm without disking following

injection was not different when the number of J2 at the end of the season were evaluated

(Table 4-1). Disking, following a shallow or deep broadcast application of 1,3-D, did not

improve any of the parameters tested for all three trials of experiment 1 (Tables 4-1,4-2).

The peanut fields had a mixed population of root-knot nematodes, but since the

fields were maintained nearly weed free throughout the season, peanut was the only








74
available host. The reproductive factor (Pf/Pi) in trial 1 was lower in treatments in which

1,3-D was applied in-row with conventional chisels and broadcast at 20 cm deep with

conventional chisels (with and without disking), and at 30 cm deep with disking than that

for the untreated control (P 0.1) (Table 4-1). The reproductive factor in the untreated

control in Trials 2 and 3 was lower or was not different than that in the fumigated

treatments (P < 0.1).

In trial 1, the lowest yields recorded were in plots treated with 1,3-D applied in-

row with conventional chisels (Table 4-2), whereas yields in all other treatments were

similar. However, in trial 2, plants grown in fumigated plots, regardless of treatment, had

higher yields than plants grown in untreated soil (P 0.1) (Table 4-2). No differences in

yield were observed among the untreated control and fumigated plots in trial 3 (P > 0.1)

(Table 4-2). However, in that trial 70% of the peanut plants were heavily infected with

Diplodia gossypina Cooke (collar rot of peanut) and that disease probably masked

differences among treatments. The incidence of root-knot nematode galling was not

different among treatments in trial 1 except for plots treated with 1,3-D applied with

subsurface hooded sweep chisels placed in bed (P < 0.1) (Table 4-2). In Trial 2, there was

a higher incidence of galling on pods from the untreated control (91%) on pods from soil

fumigated with 1,3-D (P 0.1) (Table 4-2). 1,3-Dichloropropene applied broadcast with

conventional chisels at 30 cm deep (with and without disking) had the lowest incidence of

galls on pods. In trial 3, no differences in incidence of root-knot nematode galls on pods

were observed (P >2 0.1) (Table 4-2). In these trials, broadcast fumigation with

conventional chisels and parachisels followed by disking were equally effective for the











Table 4-1. Number of Meloidogyne spp. second-stage juveniles (J2) per 100 cm3 of soil at harvest following fumigation with 1,3-dichloropropene
(1,3-D) applied with three chisel types in three peanut trials conducted in 1998 (trials I and 2) and 1999 (trial 3) at the University of Florida Green Acres
Agronomy Research Farm located in Alachua County, Florida.

Trial 1a Trial 2' Trial 3b

Treatment No. of J2 Rf No. of J2 Rf No. of J2 Rf

Untreated 779 a 93 ab 1,746 17 b 238 b l b

In-row
Conventional chisel 120 b 5c 1,977 211 a
Conventional chisel, shallow 237 b 2 b
Parachisel 194b 16 bc 1,924 14b 328 b I b
Subsurface hooded sweep chisel 79 b 25 bc 2,379 98 b

Broadcast
Conventional chisel, shallow 1 b 0.04 c 1,383 32 b 381 b 3 ab
Conventional chisel, shallow, disking 14 b 2 c 1,570 87 ab 288 b I b
Conventional chisel 45 b 20 bc 1,096 32 b 489 ab 3 ab
Conventional chisel, disking 13 b 0.2 c 791 8 b 328 b 3 ab
Parachisel 241 b 2b
Parachisel, 112 liters of 1,3-D/ha 647 a 4 a

Bed
Subsurface hooded sweep chisel 690 a 108 a 1,941 46.0 b
'In trials 1 and 2, 1,3-D was applied at a rate of 84 liters/ha. Data for the untreated control and fumigated plots were each the mean of six replicates.
Means within columns followed by the same letter are not significantly different according to the Duncan's multiple-range test (P < 0.1).
bin trial 3, 1,3-D was applied with parachisels at 112 liters/ha. Data for the untreated control and fumigated plots were each the mean of six
replicates. Means within columns followed by the same letter are not significantly different according to the Duncan's multiple-range test (P 0.1).

The reproductive factor (Rf) for Meloidogyne spp. was determined by Pf/Pi ratio where Pi = the initial population density and Pf = final population
density 146 to 150 days after planting. Data for Pi and Pfare each the mean of six replicates.


dThe shallow application of 1,3-D was injected at 20 cm deep and the application depth for all other treatments was 30 cm deep.










Table 4-2. Yield and incidence of root-knot nematode galls on peanut pods at harvest in soil fumigated with 1,3-dichloropropene (1,3-D) applied with
three chisel types in three peanut trials conducted in 1998 (trials 1 and 2) and 1999 (trial 3) at the University of Florida Green Acres Agronomy Research Farm
located in Alachua County, Florida.

Trial 1P Trial 2' Trial 3b

Treatment Yield (kg/ha) Incidence' Yield (kg/ha) Incidence' Yield (kg/ha) Incidence'

Untreated 4,709 ab 1.5 b 2,735 d 91.2 a 3,543 12

In-row
Conventional chisel, shallow 4,170 15
Conventional chisel 3,767 c 1.3 b 3,901 ab 61.5 b
Parachisel 4,888 a 1.1 b 3,722 abc 67.0 b 3,767 11
Subsurface hooded sweep chisel 4,081 bc 1.2 b 3,677 bc 51.0 b

Broadcast
Conventional chisel, shallow 4,978 a 2.0 b 3,318 abc 64.3 b 4,395 7
Conventional chisel, shallow, disking 4,574 ab 8.7 b 3,857 abc 41.8 bc 4,036 8
Conventional chisel 4,843 a 0.8 b 4,170 ab 20.3 c 4,170 11
Conventional chisel, disking 4,709 ab 5.8 b 4,359 a 20.7 c 4,170 5
Parachisel 4,574 8
Parachisel, 112 liters 1,3-D/ha 4,126 11

Bed
Subsurface hooded sweep chisel 4,619 ab 25 a 3,991 ab 50.2 b
'In trials 1 and 2, 1,3-D was applied at a rate of 84 liters/ha. Data for the untreated control and fumigated plots were each the mean of six replicates.
Means within columns followed by the same letter are not significantly different according to the Duncan's multiple-range test (P 0.1).
bIn trial 3, 1,3-D was applied with parachisels at 112 liters/ha. Data for the untreated control and fumigated plots were each the mean of six
replicates. Means within columns followed by the same letter are not significantly different according to the Duncan's multiple-range test (yield = P 0.1 and
incidence = P 0.5).

'Incidence of root-knot nematode galls on peanut was calculated as the percentage of 200 peanut pods with root-knot nematode galls.
'The shallow application of 1,3-D was injected at 20 cm deep and the application depth for all other treatments was 30 cm deep.








77
parameters tested. However, in a similar study, disking after in-row fumigation with 1,3-

D with conventional chisels increased the efficacy of the fumigant (see Chapter 3).

Experiment 2: Fruit production and yield of squash were higher in plots broadcast

fumigated with 1,3-D using conventional chisels followed by disking compared with the

untreated control (P 0.1) (Table 4-3). All other treatments were similar to that of the

control. At 34 and 65 days after planting, the number of Meloidogyne spp. J2 per 100 cm3

of soil was lower in all the fumigated plots except for those that were fumigated with 1,3-

D in-row with parachisels compared with untreated plots (P < 0.1) (Table 4-1). At 34

days after planting, disking after in-row and broadcast fumigation using conventional

chisels or parachisels did not impact the number of J2 detected in the soil (P > 0.1).

Fewer J2 were recovered from soil broadcast fumigated using parachisels without disking

than from in-row fumigation with parachisels without disking. No differences were

observed in the number of J2 recovered from in-row or broadcast fumigation with

conventional chisels. The population densities of Criconemoides spp., P. minor, and P.

brachyurus were variable, but were generally lower in numbers following broadcast

fumigation compared with the untreated control (data not shown). The reproductive

factors (Pm/Pi and Pf/Pi) for the untreated control and all fumigated plots at 34 and 65

days after planting were less than one (Table 4-3). The untreated control had a higher

reproductive factor than fumigated treatments for each sampling date. Cool soil

temperatures throughout the growing season likely contributed to this low level of root-

knot nematode reproduction.










Table 4-3. Effects of broadcast or in-row fumigation of 1,3-dichloropropene applied at 84 liters/ha with conventional chisels or parachisels on fruit
production and yield of squash (Cucurbita pepo), number of second-stage juveniles of Meloidogyne spp. per 100 cm3, soil and reproductive factors 34 days and
65 days after planting (DAP) at the University of Florida Green Acres Agronomy Farm located in Alachua County, Florida in 1999.
Treatment No. of fruit/ha (x104) Yield (metric ton/ha) No. of J2' Rmb Rf

34 DAP 65 DAP

Untreated 8.4 bc 14.6 bc 2.7 a 3.1 a 0.1 0.3

In-row

Conventional chisel 1.8 ab 21.1 ab 0.5 bc 1.6 bc 0.01 0.07
Conventional chisel, disking 9.9 bc 17.5 bc 1.4 bc 1.5 bc 0.04 0.03
Parachisel 7.9 bc 16.1 bc 1.7 ab 3.2 a 0.04 0.22
Parachisel, disking 7.2 c 12.8 c 0.9 bc 1.1 bc 0.03 0.07

Broadcast

Conventional chisel 7.2 c 13.6 bc 1.0 bc 0.4 c 0.01 0.003
Conventional chisel, disking 13.8 a 22.1 a 0.7 bc 1.4 bc 0.08 0.06
Parachisel 8.5 bc 15.3 bc 0.2 c 0.2 c 0.002 0.002
Parachisel, disking 8.8 bc 16.5 bc 0.9 bc 1.8 b 0.1 0.05
aData were transformed with loge (x + 1) before analysis.

b'Rm = Midseason reproductive factors for Meloidogyne spp. were determined by the Pm/Pi ratio, where Pm = population density 34 days after
planting and Pi = the pretreatment population density.

CRf = Final reproductive factors for Meloidogyne spp. were determined by the Pf/Pi ratio, where Pf = population density at 65 days after planting.

Data for the untreated control and fumigated plots were each the mean of six replicates. Means within columns followed by the same letter are not
significantly different according to the Duncan's multiple-range test (P < 0.1). The means for each treatment at the three sampling times (Pi, Pm, and Pf) were
calculated and used for determining the ratios.








79
Squash grown in soil fumigated with 1,3-D had fewer galls on the roots compared

to roots from the untreated control (P 0.1) (Fig. 4-6). Plants from broadcast treatments

had overall lower gall indicies compared to plants from in-row treatments. Fumigation

with 1,3-D reduced the number of J2 in soil and the number of root galls, which has been

reported previously (see Chapter 3; Weingartner and Schumaker, 1991). Row treatment

can be a cost-effective method for nematode management because only the soil in which

the crop will be grown is treated (Lembright, 1990). Less material per hectare is used as

compared to broadcast fumigation in which the entire field is treated. The crop response

may be equal to or superior to soil receiving a broadcast treatment (Lear and Thomason,

1956; Lembright, 1990; and Rhoades et al., 1962). In deep sandy or sandy loam soils,

deep application with a single chisel has been shown to be equal to or better than two

chisels in performance and emissions reduction when the same rate of fumigant per hectare

is used (Lembright, 1990; Schneider et al., 1995; Sipes et al., 1993). However, in our

experiment, in-row applications were less effective for root-knot nematode control on

squash plantings in the deep sandy soil of Florida than broadcast applications.

Disking following in-row and broadcast fumigation with conventional chisels or

parachisels did not result in a reduction in the amount of galling on roots (Fig. 4-6).

However, in this experiment, 4 hours after fumigation, a light rain totaling 2.5 cm fell on

the field over an 8 hour period. The deep placement of the fumigant and the light rain may

have sealed in the fumigant thereby masking any differences that may have occurred

among treatments.










Regardless of the method of application, fumigation with 1,3-D at 84 or 112

liters/ha reduced the number of infective juveniles. The few nematodes that escape

fumigation may reproduce rapidly thereby increasing the population density by the end of

the season. In the case of peanut, the roots, pods, and pegs are susceptible to root-knot

nematode infection until the day of harvest. During this relatively long period, root-knot

nematodes can undergo several life cycles in a season. Under these conditions, the small

number of root-knot nematodes that survived fumigation could result in crop losses later

in the season. On the other hand, in experiment 2 and in a previous study on squash, root-

knot nematodes were managed well in the spring with broadcast applications of 84 liters

of 1,3-D/ha (see Chapter 3). Fumigation with higher rates such as 112 and 168 liters of

1,3-D need to be evaluated for control of root-knot nematodes for long season crops such

as peanut and tobacco.















S-1 Parachisels
7 Conventional chisels
a D = Disked plots


0
"- .5
I

bU b
bc 3 bcjb
cl 2
bcd
D d cd


Untreated In-row Broadcast







Fig. 4-6. Root-knot nematode galling in squash, Cucurbita pepo, grown in soil
fumigated with 84 liters of 1,3-dichloropropene/ha applied in-row and broadcast with
conventional chisels and parachisels with and without disking. The galls were indexed on
a scale of 0 to 10 (0 = 0 galls, 1 = 10% of the roots galled, 2 = 20% of the roots galled.
..... 10 = 100% of the roots galled) (Barker et al., 1986). Data for the untreated control
and fumigated plots were each the mean of six replicates. Means within columns with the
same letter are not significantly different according to the Duncan's multiple-range test (P
< 0.1).













CHAPTER 5
COMPARISON OF DIFFERENT CHISEL TYPES FOR 1,3-DICHLOROPROPENE
FUMIGATION IN DEEP SANDY SOILS


Introduction



A major concern with the use of 1,3-D in Florida deep sandy soil is retaining an

adequate dosage to provide suitable efficacy while preventing premature emissions. The

fumigant is generally applied with conventional chisels, 15 to 20 cm deep, which generally

leave a 1 to 1.3 cm chisel traces in the soil (Lembright, 1990). Intact chisel traces allow

the fumigant to escape rapidly into the atmosphere (Goring, 1957; Lembright, 1990).

Rollers, cultipackers, press wheels, and other similar equipment may not be adequate to

these chisel traces. The label for Telone II (Dow AgroSciences LLC, Indianapolis, IN)

was changed to address this concern. The new label requires that 1,3-D be injected at a

depth of 30 cm deep below the final soil surface and that the chisel traces be broken and

filled with soil bedding disks following application.

Although 1,3-D has been used successfully to control plant-parasitic nematodes on

many crops, its performance is known to vary (Dickson and Hewlett, 1988; Melton,

1996). One factor which may affect its efficacy in deep sandy soils is the method of

application. Modification of application equipment and method of fumigating has been

shown to reduce 1,3-D emissions. Methods, such as, elimination of chisel traces







83
(Anonymous, 1996b; Dallimore, 1955), reduction of number of chisels (Schneider et al.,

1995; Sipes et al., 1993), fumigation at different depths (Gan et al., 1998; Rodriguez-

Kdbana and D. G. Robertson, 1987), and application of water to the soil surface after

fumigation (Gan et al., 1998) have reduced 1,3-D emissions. The objective of this study

was to compare the efficacy of in-row fumigation of 1,3-D with conventional chisels and

parachisels (with and without disking), and subsurface hooded sweep chisels to manage

Meloidogyne spp. in deep sandy soils.



Materials and Methods



Two field trials were conducted at the same site at the University of Florida Green

Acres Agronomy Research Farm in Alachua County, Florida in 1998 and 1999. The site

was infested with M incognita (Kofoid and White) Chitwood, and M. javanica (Treub.)

Chitwood. Tobacco (Nicotiana tabacum L.) was planted in the field in spring 1997. Rye,

Secale cereale L. cv. Wrens Abruzzi, was planted as a winter cover crop in the fall of both

1997 and 1998 and plowed under in the spring. The soil was an Arredondo fine sand

(91% sand, 6% silt, 3% clay, and 2.5% organic matter; pH 6.5).

The experimental design was a randomized complete block with six replications.

The two-row plots were 9.1 m long with a row spacing of 91 cm. 1,3-Dichloropropene

was applied 8 October 1998 and 15 June 1999. Applications with conventional chisels

(with and without disking) or parachisels (with and without disking) were made with two

chisels per row spaced 22 cm apart centered over each row. 1,3-Dichloropropene was

injected 30 cm deep at 84 liter/ha (78 ml/chisel/30.5 m of row) into the soil and a 1.8-m-








wide, two-gang disk set to cut 15 to 18 cm deep was run through the plots immediately

following 1,3-D application. The conventional chisels were forward or rear inclined,

double beveled blades, and were arranged with one outer swept-forward chisel and one

inner swept-back chisel per row (see Chapter 4). Parachisels were rearward inclined, side-

wise sloped on a 450 plane, single beveled blades (see Chapter 4). The subsurface hooded

sweep chisels (Dow Chemical Co., Midland, MI) had a flat tip 8003 TeeJet (Spraying

Systems Co., Wheaton, IL) spray nozzle mounted under each subsurface hooded sweep

(see Chapter 4). The soil moisture at time of fumigation was 13% in 1998 and 11% in

1999 and the soil temperature at 10 cm deep reached a maximum of 29 C and a minimum

of 27 C in both years.

Twelve soil cores (2.5-cm-diam., 20 cm deep) were taken from each plot x% ith a

cone-shaped sampling tube at pretreatment (5 October 1998 and 12 June 1999) and at

termination of the study (25 November 1998 and 29 July 1999). Soil cores from each plot

were combined and nematodes were extracted from a 100 cm3 subsample by a centrifugal-

flotation method (Jenkins, 1964). All plant-parasitic nematodes were counted.

Glyphosate at the labeled rate was applied broadcast on the soil surface 1 week

before planting to manage weeds, and halosulfuron was sprayed at the labeled rate 1 week

after planting to manage purple and yellow nutsedge. A rate of 318 kg of 10-10-10 N-P-

K fertilizer/ha was applied broadcast and incorporated into the top 10 cm of soil on the

day of planting. Supplemental water was applied through overhead irrigation. Tomato

seedlings, Lycopersicon esculentum Mill. cv. Solarset, were transplanted 31 and 38 cm

apart in all plots on 15 October 1998 and 25 June 1999, respectively.








Root systems of the center 10 plants in each plot were removed 35 days after

transplanting (DAT) in 1998 and 37 DAT in 1999. Root-knot nematodes that penetrated

the tomato root systems were stained (Byrd et al., 1983) and counted.

Data were subjected to analysis by ANOVA and means separated by Duncan's

multiple-range test (SAS Institute, Cary, NC). Differences among the number of root-

knot nematodes per tomato root system per treatment were compared by orthogonal

contrasts. All data were transformed with loge (x + 1) before analyses, and all differences

reported were significant at P < 0.05.



Results and Discussion



No differences among treatments were observed in final soil population densities

of Meloidogyne spp. (P 0.1). Fumigation in 1998 with 1,3-D regardless of application

methods, reduced the number of infective juveniles (J2) (Table 5-1). Fewer numbers of

second-stage juveniles (J2) entered the root system of plants grown in fumigated soil

compared to plants grown in untreated soil. Plants from untreated soil had an average of

87 J2/root system, whereas those from fumigated soil showed an average of 19 J2/root

system. The most effective 1,3-D application methods were with conventional chisels

followed by disking and parachisels (with and without disking). Fumigation with

parachisels with and without disking were equally effective in reducing the number of J2

that penetrated tomato roots (Table 5-1). Fewer J2 were in tomato roots from soil

fumigated with 1,3-D using conventional chisels followed by disking compared to roots

from soil fumigated with conventional chisels without disking (Tables 5-1,5-2).









Table 5-1. The number Meloidogyne spp. second-stage juveniles (J2) that
penetrated 4-week-old tomato seedlings after in-row fumigation with 1,3-dichloropropene
applied at 84 liters/ha with conventional chisels (with and without disking), parachisels
(with and without disking), and subsurface hooded sweep chisels in 1998 and 1999.

Loge (J2/root system + 1)

Treatment 1998 1999a

Untreated 4.3 a 2.5

Conventional chisel 2.0 bc 1.8

Conventional chisel + disking 1.0 d 3.1

Parachisel 1.4cd 3.3

Parachisel + disking 0.9 d 2.6

Subsurface hooded sweep chisel 2.7 b 3.1

Data for all treatments were each the mean of 60 replicates and were transformed
with loge (x + 1) before analysis. Means within columns followed by the same letter are
not significantly different according to Duncan's multiple-range test (P < 0.05).


aLack of a letter denotes nonsignificance (P 0.1)