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Evaluation of Resistance to Meloidogyne Arenaria in the Peanut (Arachis Hypogaea. L) Cv. Tifguard

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Title:
Evaluation of Resistance to Meloidogyne Arenaria in the Peanut (Arachis Hypogaea. L) Cv. Tifguard
Creator:
Yuan, Weimin
Place of Publication:
[Gainesville, Fla.]
Florida
Publisher:
University of Florida
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Language:
english
Physical Description:
1 online resource (113 p.)

Thesis/Dissertation Information

Degree:
Doctorate ( Ph.D.)
Degree Grantor:
University of Florida
Degree Disciplines:
Entomology and Nematology
Committee Chair:
DICKSON,DONALD W
Committee Co-Chair:
TILLMAN,BARRY
Committee Members:
BRITO,JANETE A
PRESTON,JAMES F,III

Subjects

Subjects / Keywords:
resistance -- tifguard
Entomology and Nematology -- Dissertations, Academic -- UF
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bibliography ( marcgt )
theses ( marcgt )
government publication (state, provincial, terriorial, dependent) ( marcgt )
born-digital ( sobekcm )
Electronic Thesis or Dissertation
Entomology and Nematology thesis, Ph.D.

Notes

Abstract:
Tifguard, which was released as a peanut cultivar resistant to root-knot nematode and tomato spotted wilt virus in 2007, was found to be heavily infected by Meloidogyne arenaria in several peanut production fields in Florida in 2012. The goal of this project was to determine why the cultivar that was reported to be highly resistant succumbed to root-knot nematode infection. The objectives were to evaluate the resistance of three different sources of Tifguard seeds; to determine the seasonal population changes and vertical population densities of M. arenaria collected from resistant and susceptible peanut rhizospheres in two different soil types; to determine the effects of high temperature on the resistance in Tifguard, and to compare the yield of Tifguard, isogenic Tifguard and Georgia-06G treated vs. nontreated with 1,3-dichloropropene. In three M. arenaria infested field sites, a comparison of Tifguard seed obtained from three sources showed that 2.5, 28, and 39.5% of plants that were infected by the nematode were negative for the nematode resistance gene. The seasonal distribution of second-stage juvenile (J2) of M. arenaria followed similar trends in two different soil types, with a peak occurring during late summer and early fall at harvest. Number of J2 dropped following harvest and reached a density less than 10 J2/200 cm3 of soil in February. Comparison of vertical population densities of J2 collected from Georgia-06G rhizosphere in two different soil types showed that greater numbers occurred in the upper 60 cm of soil during the growing season in a Candler sand, whereas in a Norfolk loamy sand greater densities were found only in the top 45 cm. The population densities of J2 collected from Georgia-06G rhizosphere at all depths were much greater in Norfolk loamy sand than that in the Candler sand. Tifguard reduced the nematode population to 1 and 28 in the Candler sand and the Norfolk loamy sand, respectively at harvest. Comparison of nematode numbers from different developmental stages at different temperatures demonstrated that the high soil temperature increased nematode infection rate and accelerated nematode development in Georgia-06G. No further development of J2 occurred in Tifguard roots at 28 or 31 C, however at 34 C a few J3-J4, females, egg laying females, and males of M. arenaria were observed. Comparisons indicated that there was no effect of 1,3-dichloropropene on root-knot nematode damage and yield of Georgia-06G, Tifguard, or isogenic Tifguard. ( en )
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In the series University of Florida Digital Collections.
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Includes vita.
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Includes bibliographical references.
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Description based on online resource; title from PDF title page.
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This bibliographic record is available under the Creative Commons CC0 public domain dedication. The University of Florida Libraries, as creator of this bibliographic record, has waived all rights to it worldwide under copyright law, including all related and neighboring rights, to the extent allowed by law.
Thesis:
Thesis (Ph.D.)--University of Florida, 2017.
Local:
Adviser: DICKSON,DONALD W.
Local:
Co-adviser: TILLMAN,BARRY.
Statement of Responsibility:
by Weimin Yuan.

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EVALUATION OF RESIST ANCE TO Meloidogyne arenaria IN THE PEANUT ( A rachis hypogaea L) cv TIFGUARD By WEIMIN YUAN A DISSERTATION PRESENTED TO THE GRADUATE SCHOOL OF THE UNIVERSITY OF FLORIDA IN PARTIAL FULFILLMENT OF THE REQUIREMENTS FOR THE DEGREE OF DOCTOR OF PHILOSOPHY UNIVERSITY OF FLORIDA 2017

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2017 Weimin Yuan

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This work is dedicated to my parents for bringing me up and educating me well

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4 ACKNOWLEDGMENTS This project was completed with the contribution s of many people and headed by Professor Donald W. Dicks on. Since 2012 when I joined his research team, my scientific perspectives and skills in professional studies were acquired with contributions from a number of scientists and peers. Foremost, I would like to express my great gratitude to Professor Dickson, my adviser, for giving me finan cial support and the opportunity to work in his lab with patient guidance and consideration throughout the duration of my entire program. Importantly, I also appreciated th at he was confident and trusted that I could do and finish this research. His resear ch passion in nematology is truly worth emulating, which was influencing and encouraging for me through four and half years. It is an honor for me to have worked with him. I extend my sincere appreciation to the members of my advisory committee, Dr s James F. Preston, Janet e A. Brito and Barry Tillman for their instructions and encouragement This research was also aided scientifically by the University of Georgia Tifton campus and Nespal (National Environmentally Sound Production Agriculture Laborator y, GA) I give heartfelt thanks to the biological scientist Dr. Maria de Lourdes Mendes who was working in the Nematod e M anagement Lab, for her technical support for laboratory experiment s, greenhouse management, and field work. I would like to mention al so Dr s Billy Crow, Robert McSorley, and Tesfamariam Mengistu for unselfishly sharing their vast knowledge and wisdom gained through their long history of working with nematodes. I also thank my colleagues in the lab, Silvia Vau, Kanan Kustuwa, and Sai Qi u for the mentoring, encouragement and meaningful conversations. I acknowledge

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5 especially Silvia Vau for her wonderful friendship and her spiritual g uidance and support through my years in Gainesville. I also give special thanks to K anan Kustuwa who was a lways standing by my side taking care of my life and giving me strong encouragement. Many Chinese friends have supported me throughout these pa st years. My earnest thanks to R uohan Liu who was a graduate student working in the same department for not only offering me many favors when I was a beginning student but most importantly, bringing out the best and strongest in me. Through her, I realized that being independent is the only way you c an become stronger It is impossible for you to change the world b ut it is possible to change yourself to adapt to the world. Thanks go to Dr. Jianan Wang for his help with takin g care of my life for 2 years. Above all, I would like to express my gratitude to my mother, for her unconditional love, and for being with me all the time. Other sincere acknowledgments are to my fellow nematology majors, dearest friends and roommates in the 4 years for imp acting my life in one way or other. Through them, I learned how to productively deal with work and problems in diverse setti ngs, smartly take risks, and courageously attack the most difficult situations. Lastly, I am indebted to the University of Florida for sponsoring my study and stay in the United States. I would like to express my appreciation to the Florida Peanut Produce rs Association for partial funding of my research project These institutions were instrumental to making this research possible.

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6 TABLE OF CONTENTS page ACKNOWLEDGMENTS ................................ ................................ ................................ .. 4 LIST OF TABLES ................................ ................................ ................................ ............ 9 LIST OF FIGURES ................................ ................................ ................................ ........ 11 ABSTRACT ................................ ................................ ................................ ................... 13 CHAPTER 1 INTRODUCTION ................................ ................................ ................................ .... 15 Root knot Nematode ................................ ................................ ............................... 15 Economic Impact ................................ ................................ .............................. 16 History of the Genus ................................ ................................ ......................... 16 The Distribution of Meloidogyne S pecies ................................ ......................... 17 Peanut Production ................................ ................................ ................................ .. 18 Plant parasitic Nematodes Associated with Peanut ................................ ......... 20 Species of Meloidogyne Parasitizing Peanut ................................ .................... 21 Other Species of Nematodes Associated with Peanut ................................ ..... 22 Root knot Nematode on Peanut ................................ ................................ ............. 22 Disease Cycl e ................................ ................................ ................................ .. 23 Parasitic Mechanism ................................ ................................ ........................ 25 Populati on Dynamics ................................ ................................ ........................ 27 Temperature Effects on Root knot Nematode ................................ .................. 30 Root knot Nematode Management ................................ ................................ .. 31 Development of Resistant Peanut Cultivars ................................ ..................... 31 Identification of Resistant Gene to Root knot Nematode ................................ .. 33 The Application of Host Resistance ................................ ................................ ........ 35 2 FIELD EVALUATION OF RESISTANCE TO Meloidogyne arenaria IN BREEDER, FOUNDATION AND GROWER KEPT TIFGUARD ............................. 37 Introduction ................................ ................................ ................................ ............. 37 Materials and Me thods ................................ ................................ ............................ 38 Field Sites and Experimental Designs ................................ .............................. 38 Determination of Root knot Nematode Species ................................ ............... 40 Analysis of Resistance Gene in Infected Tifguard Plants ................................ 40 Results and Discussion ................................ ................................ ........................... 41 464 Farm 2013 2015 ................................ ................................ ....................... 41 Brown Fa rm and AREC 2014 2015 ................................ ................................ 41 Determination of Root knot Nematode Species ................................ ............... 41 Analysis of Resistance Gene in Infected Tifguard Plants ................................ 42

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7 3 THE SEASONAL AND VERTICAL DISTRIBUTION OF Meloidogyne arenaria FROM RESISTANT AND SUSCEPTIBLE PEANUT GROWN IN TWO SOIL TYPES ................................ ................................ ................................ .................... 46 Introduction ................................ ................................ ................................ ............. 4 6 Materials and Methods ................................ ................................ ............................ 48 Field Locations and Experimental Designs ................................ ....................... 48 Seasonal and Vertical Distribution of Meloidogyne arenaria ............................ 49 Statistical Analysis ................................ ................................ ............................ 50 Determination of Effects of Soil Texture on Nematode Population ................... 50 Results ................................ ................................ ................................ .................... 51 Above and Belowground Symptoms of Root knot Disease .............................. 51 Seasonal Distribution of Meloidogyne arenaria Recovered from Georgia 06G Rhizosphere ................................ ................................ .......................... 51 Vertical Distribution of Meloidogyne arenaria Recovered from Georgia 06G Rhizosphere ................................ ................................ ................................ .. 52 Seasonal and Vertical Distribution of Meloidogyne arenaria Recovered from Tifguard Rhizosphere ................................ ................................ .................... 54 Distribution of Nematodes Associated with Soil Temperature .......................... 54 Effects of Soil Texture on Nematode Population ................................ .............. 54 Discussion ................................ ................................ ................................ .............. 54 Occurrence of Meloidogyne arenaria in the Soil ................................ ............... 54 Seasonal and Vertical Distribution of Meloidogyne arenaria ............................ 56 The Influence of Soil Temperature on Nematode Population ........................... 57 Effect of Soil Texture on Nematode Population ................................ ................ 57 4 THE INFLUENCE OF TEMPERATURE ON THE SUSCEPTIBILITY OF CVS. TIFGUARD AND GEORGIA 06G PEANUT TO Meloidogyne arenaria .................. 67 Introduction ................................ ................................ ................................ ............. 67 Materials and Methods ................................ ................................ ............................ 70 Nematode Origin ................................ ................................ .............................. 70 Penetration and Development of Meloidogyne arenaria ................................ ... 70 Data Collection ................................ ................................ ................................ 71 Statistical Analysis ................................ ................................ ............................ 71 Resistance Gene Marker Analysis of Tifguard ................................ ................. 71 Results ................................ ................................ ................................ .................... 72 Penetration of Meloidogyne arenaria in the Resistant and Susceptible Peanut Cultivars ................................ ................................ ............................ 72 Development of Meloidogyne arenaria in the Resistant and Susceptible Peanut Cultivars ................................ ................................ ............................ 73 Reproduction of Meloidogyne arenaria in Different Peanut Genotypes ............ 74 Discussion ................................ ................................ ................................ .............. 74 Effect of Peanut Genotypes and Temperatures on Meloidogyne arenaria ....... 74 The Function of Resistance Gene ................................ ................................ .... 75 Hypersensitive Reaction ................................ ................................ ................... 75

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8 5 COMPARISON OF YIELD FROM BREEDER TIFGUARD, ISOGENIC TIFGUARD AND GEORGIA 06G TREATED AND NONTREATED WITH 1,3 DICHLOROPROPENE ................................ ................................ ........................... 90 Introduction ................................ ................................ ................................ ............. 90 Materials and Methods ................................ ................................ ............................ 91 Field Design and Treatments ................................ ................................ ........... 91 Peanut Harvest and Data Collection ................................ ................................ 92 Statistical Analysis ................................ ................................ ............................ 92 Results ................................ ................................ ................................ .................... 92 Peanut Growth and Root knot Nematode Infection ................................ .......... 92 Yield Comparison ................................ ................................ ............................. 93 Soil Type, Texture, Analysis ................................ ................................ ............. 93 Discussion ................................ ................................ ................................ .............. 93 LIST OF REFERENCES ................................ ................................ ............................... 96 BIOGRAPHICAL SKETCH ................................ ................................ .......................... 113

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9 LIST OF TABLES Table page 2 1 The average percentages of three different sources of Tifguard seed infected by Meloidogyne arenaria at the 464 Farm, Levy Co., FL in 2013, 2014, and 2015 ................................ ................................ ................................ .................. 43 3 1 Significance of main and interactive effects of both fixed and random variables for the vertical distribution of Meloidogyne arenaria extracted from 200 cm 3 of soil collected from plots planted Georgia 06G at the Brown Farm. ... 61 3 2 Significance of main and interactive effects of both fixed and random variables for the vertical distribution of Meloidogyne arenaria extracted from 200 cm 3 of soil collected from plots planted Georgia 06G at the Attapulgus ..... 61 3 3 Number of Meloidogyne arenaria second stage juveniles extracted from 200 cm 3 of soil collected at four depths 0 30, 31 60, 61 90, and 91 120 cm deep from plots planted Georgia 06G at the Brown Farm (2014 2015). ...................... 61 3 4 Number of Meloidogyne arenaria second stage juveniles extracted from 200 cm 3 of soil collected at four depths 0 15, 16 30, and 31 45 cm deep from plots pl anted Georgia 06G at the Attapulgus Research and Education ............. 62 3 5 Significance of main and interactive effects of both fixed and random variables for the vertical distribution of Meloidogyne arenaria extracted from 200 cm 3 of soil collected from plots planted with Tifguard at the Brown Farm ... 64 3 6 Significance of main and interactive effects of both fixed and random variables for the vertical distribution of Meloidogyne arenaria extracted from 200 cm 3 of soil collected from plots planted with Tifguard at the Attapulgus ...... 64 3 7 Number of Meloidogyne arenaria second stage juveniles extracted from 200 cm 3 of soil collected at four depths 0 30, 31 60, 61 90, and 91 120 cm from plots planted with Tifguard at the Brown Farm 2015 ................................ ......... 64 3 8 Number of Meloidogyne arenaria second stage juveniles extracted from 200 cm 3 of soil collected at four depths 0 15, 16 30, and 31 45 cm from plots planted with Tifguard at the Attapulgus Research and Education Center ........... 65 3 9 Sand, silt, and clay distribution at different soil depths collected from Brown Farm, Levy County, FL and University of Ge orgia, Attapulgus Research and Education Center, Attapulgus, GA. ................................ ................................ ..... 66 4 1 Analysis of variance of root weight, number of Meloidogyne arenaria per gram root system of two peanut cultivars. ................................ .......................... 77

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10 5 1 Effect of different treatments on plant growth, galling induced by root knot nematodes and yield of peanut in a field trial at Plant Science Research and Education Unit, Citra, FL, Spring Summer 2015. ................................ ................ 95

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11 LIST OF FIGURES Figure page 2 1 The average percentages of Tifguard breeder seed infected by Meloidogyne arenaria was compared with Georgia 06G at two locations in 2014 and 2015. .. 44 2 2 The average percentages of different Tifguard seed sources that were negative for the resistance gene marker for Meloidogyne arenaria ................... 45 3 1 Seasonal fluctuation of Meloidogyne arenaria second stage juveniles in the soil profile of Georgia 06G at four depths in a Candler sand at the Brown Farm, Levy County, FL. ................................ ................................ ...................... 59 3 2 Seasonal fluctuation of Meloidogyne arenaria second stage juveniles in the soil profile of Georgia 06G at three depths in a Norfolk loamy sand soil at University of Georgia, Attapulgus Research and Education Center, .................. 60 3 3 Seasonal fluctuation of Meloidogyne arenaria second stage juveniles in the soil profile of Tifguard at four depths in a Candler sand at the Brown Farm, Levy County, FL and at three depths in a Norfolk loamy sand soil at,. ............... 63 4 1 The effect of temperature on the number of different developmental stages of Meloidogyne arenaria in roots of two peanut cultivars and tomato recorded at 5 day intervals over a 30 day period at 28 (Experiment 1). ............................ 78 4 2 The effect of temperature on the number of different developmental stages of Meloidogyne arenaria in roots of two peanut cultivars and tomato recorded at 5 day inte rvals over a 30 day period at 31 (Experiment 1). ............................ 79 4 3 The effect of temperature on the number of different developmental stages of Meloidogyne arenaria in roots of two peanut cultivars and tomato recorded at 5 day intervals over a 30 day period at 34 (Experiment 1). ............................ 80 4 4 The effect of temperature on the number of different developmental stages of Meloidogyne arenaria in roots of two peanut cultivars and tomato recorded at 5 day inte rvals over a 40 day period at 28 (Experiment 2). ............................ 81 4 5 The effect of temperature on the number of different developmental stages of Meloidogyne arenaria in roots of two peanut cultivars and tomato recorded at 5 day intervals over a 40 day period at 31 (Experiment 2). ............................ 82 4 6 The effect of temperature on the number of different developmental stages of Meloidogyne arenaria in roots of two peanut cultivars and tomato recorded at 5 day inte rvals over a 40 day period at 34 (Experiment 2). ............................ 83

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12 4 7 The mean number of Meloidogyne arenaria of all development stages in roots of Tifguard and Georgia 06G recorded at 5 day intervals over a 30 day period in experiment 1. ................................ ................................ ....................... 84 4 8 The mean number of Meloidogyne arenaria of all development stages in roots of Tifguard and Georgia 06G recorded at 5 day intervals over a 40 day period in experiment 2. ................................ ................................ ....................... 85 4 9 The number of second stage juveniles (J2) per gram of root system of the resistant cultivar Tifguard and susceptible cultivar Georgia 06G 5 days after inoculation in two experiments. ................................ ................................ .......... 86 4 10 The number of egg laying females per gram of root system of the resistant cultivar Tifguard and susceptible cultivar Georgia 06G 30 or 40 days after in oculation in two experiments. ................................ ................................ ........... 87 4 11 Different developmental stages of Meloidogyne are naria in peanut and tomato roots at 34 ................................ ................................ .......................... 88 4 12 Arrows point to necrotic lesions formed around the root knot nematode infection sites in Tifguard roots at 5 and 40 days after inoculation (DAI). ........... 89

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13 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 EVALUATION OF RESIST ANCE TO Meloidogyne arenaria IN THE PEANUT ( A rachis hypogaea L) cv TIFGUARD B y Weimin Yuan May 2017 Chair: Donald W. Dickson Major : Entomology and Nematology Tifguard, which was released as a peanut cultivar resistant to root knot nematode and t omato spotted wilt virus in 2007 was found to be heavily infected by Meloidogyne arenaria in several peanut production fields in Florida in 2012. The go al of this project was to determine why the cultivar that was reported to be highly resistant succumbed to root knot nematode infection. The objectives were to evaluate the resistance of three dif ferent sources of Tifguard seed s ; to determine the seasonal population changes and vertical population densities of M. arenaria co llected from resistant and susceptible peanut rhizospheres in two different soil types ; to determine the effects of high temperature on the resistance in Tifguard, and to compare the yield of Tifguard, isogenic Tifguard and Georgia 06G treated vs. nontreated with 1,3 dichloropropene In three M. arenaria infested field sites a c omparison of Tifg uard seed obtained from three sources showed that 2.5 28 and 39.5 % of plants that were infected by the nematode were negative for the nematode resistance gene. The seasonal distribution of second stage juvenile (J2) of M. arenaria followed similar trends in two different soil types with a peak occurring during late summer and early fall at

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14 harvest. Number of J2 dropped following harvest and reached a dens ity less than 10 J2/200 cm 3 of soil in February. Comparison of vertical population densities of J2 collected from Georgia 06G rhizosphere in two different soil types showed that greater numbers occu r r ed in the upper 60 cm of soil during the growing season in a Candler sand, whereas in a Norfolk loamy sand greater densities were found only in the top 45 cm. The population densities of J2 collected from Georgia 06G rhizosphere at all depths were much greater in Norfolk loamy sand tha n that in the Candler sand. Tifguard reduced the nematode population to 1 and 28 in the Candler sand and the Norfolk loamy sand respectively at harvest Comparison of nematode number s from different developmental stages at different temperatures demonstrated that the high soil temperature increased nematode infection rate and accelerated nematode development in Georgia 06G No further development of J2 occurred in Tifguard roots at 28 or 31 however at 34 a few J3 J4, females, egg laying females, and males of M. arenaria were observed. Comparisons indicated that there was no effect of 1,3 dichloropropene on root knot nematode damage and yield of Georgia 0 6G, Tifguard, or isogenic Ti fgua rd ( P

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15 CHAPTER 1 INTRODUCTION Root k not Nematode Root knot nematode is the common name of members from the genus Meloidogyne (Go e ldi, 1892), which is derived from a Greek word meaning of apple shaped female. It is an economically important polyphagous group of obligate plant parasites with advanced parasitic adaptation s. Root knot nematode s have worldwide distribution and a b roa d host range They are capable of infecting nearly every species of higher plant s ( Karssen, 2006) Typically they parasitize internally within plant roots and feed on modified vascular plant cells, inducing small to larg e galls or knots on roots. This is the common underground symptom produced by the nema tode ( Christie, 1936 ). The abovegrou nd symptoms are usually defined as a patchy distribution pattern with chlorotic and stunted plants Both symptoms may be similar to nutrition deficiency or malfunction of root system s ( Bird, 1970 ). These severe abovegrou nd symptoms are explained by Meloidogyne infection that disrupts water and nutrient uptake and upward translocation by the root system (Bird, 1970). Hosts may be heavily infected without showing external symptoms on folia r tissue, e.g. symptomless plant le aves but galled roots, however more commonly the aboveground symptoms may be severe, especially under drought conditions The rapid development and reproduction on good hosts result in several generations during one crop season, which lead s to severe crop damage. Secondary infection by other pathogens usually causes extensive decay of nematode infected tissues ( Christie, 195 0 ). Because of nematode infection, not only crop yield is sup p ressed but also product quality is often not acceptable. Th us this genus is of great economic and social importance.

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16 Economic I mpact Generally, tropical regions provide more favorable environmental conditions for pathogen development, colonization, reproduction and dispersal, and as a result damage and yield l osses caused by plant pathogens, including plant parasitic nematodes are greater in tropical than in temperate regions (De Waele and Elsen, 2007). Severity of root knot nematode damage is usually affected by specific Meloidogyne species. Their effects dep end on susceptibility of hosts, crop history, season and soil type (Greco et al ., 1992; Potter and Olthof, 1993). Similarly, economic thresholds are primarily dependent on these same factors. Based on damage thresholds established for several crops, on ave rage, approximately 0.5 to 2 second stage juveniles per gram of soil is enough to cause damage (Di Vito et al., 1991). There is not only a direct cost to root knot nematode infestations, but also an indirect cost such as that cause by regulations and quara ntines imposed on some species of Meloidogyne. For example M. chitwoodi is increasingly regulated because of its effects as a serious pathogen of potato and other economically important crops. This nematode is on the list of prohibited pathogens of many countries namely (Canada, the EU, Mexico and other countries in Latin America, and the Far East) (Hockland et al ., 2006). M. fallax is another quarantine nematode that is being subjected to regulation. Also, the highly virulent M. enterolobii has been adde d to the EPPO alert list. Additional root knot nematodes that might be added to the list of quarantine species include recently described M. minor and M. citri in the USA (Viaene et al., 2007). History of the G enus The first root knot disease was reported during the middle of the 19th century by a clergyman, Miles Joseph Berkeley (1855), who first attributed galls extracted from

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17 glasshouse cucumber roots to the nematode. The nematode was first described by Cornu (187 9). He based his description on nematodes found in root galls of sainfoin ( Onobrychis sativus Lam.) in the Loire valley, France. In 1887, a brief description of a root knot nematode was made by Goeldi. He illustrated a root knot nematode extracted from cof fee plants in Brazil. He named the nematode M. exigua Up until 1949 all root knot nematodes were designated as Heterodera marioni. Benjamin Chitwood is credited with revising the genus Meloidogyne (Chitwood, 1949) He placed five species and in one genus, naming them M. arenaria M. exigua M. incognita, M. javanica M. hapla and M. incognita var. acrita He based their identification on morphological characters, such as perineal pattern, stylet knob shape, length of stylet, and the distance of the dorsal esophageal gland orifice from the base of the stylet knob. Currently there are over 100 Meloidogyne spp. described (Karssen, 2002). In addition to traditional morphological identification, more modern tools are being used such as biochemical and molecular biology (Hartman and Sasser, 1985). The D istribution of Meloidogyne S pecies It is difficult to determine the origin of most Meloidogyne spp. Their broad host range and the worldwide movement of vegetative planting stock infected with root knot nematodes makes it difficult to distinguish the nematode site of origin or whether the nematode adapt ed to the new site (Sasser, 1977). Some species may adapt to their new environment s whereas others may not (Sasser, 1977). M. incognita (Kofoid and White, 1919) the southern root knot nematode, is one of the most ubiquitous species in the genus. It has a wider geographi c range than any other species. The nematode is found from approximately 40 N lati tude to 33 S (Taylor et al., 1982). The average annual temperature reported in this geographic range lies between 18 to 30

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18 However, most populations were found where the annual temperature range s between 24 to 30 The optimum temperature development is 27 M. incognita rarely occurs in regions where the average lowest temperature is below 3 The second most common species, M. javanica (Treud, 1885) is also distributed throughout the world. The latitude range of M. javanica is from approximately 33 N to 33 S (Tayler et al., 1982). It is rarely found in cold regions where monthly temperature average below 3 Dryer soil conditions usually enhance survivability of M. javanica and as a result it is the dominant species in regions where monthly precipitation is 5 mm for at least 3 months. Al t hough M. arenaria (Neal, 1889) is not as common as the previous ly mentioned species, it is the most important plant parasitic nematode worldwide on peanut. The distribution of M. arenaria is app roximately 40 N latitude to 33 S latitude T he annual temperature range is similar to M. incognita M. hapla (Chitwood, 1949) is less prevalent than the other three Meloidogyne sp p. I t is usually found in cool regions with latitudes between 34 N and 43 N (Taylor et al., 1982). In tropical or subtropical regions, M. hapla often occurs at high altitudes that are more than 1 000 m (Brown, 1955). Peanut Production The peanut, or groundnut ( Arachis hypogaea ), is an annual herbaceous plant and a member of the legume or the "bean" family ( Fabaceae ). This family is characterized by symbiosis with rhizobium which leads to production of nitrogen needed for plant growth (Nwokolo, 1996). Growth of pe anut begins after the germination of a seed. T he plant produces a flower above ground and ends in maturation of a fruit underground. It has at least two distinct growth habits: erect and trailing (Dickson,

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19 1998). Appearance of flowers begins 4 to 6 weeks a fter planting with formation of yellow petaled, pea like inflorescence borne in the axillary clusters aboveground. The flowers wither responding to self pollination and the stalks called gynophores or pegs elongate downward into the soil. Subsequently they turn horizontally followed by formation of a swollen ovary that develops to become a pod. The pods of most cultivated peanut contain two or three seeds and they mature at 125 to 145 days after planting (Dickson, 1998). Peanut grows well in tropical a nd subtropical climates, optimally at temperatures ranging between 30 and 34 C, although they will tolerate a range between 15 and 45 C ( Williams et al., 1978 ). High temperatures (above 34C) may cause damage to peanut flowers. Plants grow best in a ligh t, well drain ed sandy loam soil with an optimal pH of 6.0 6.5 ( Smith, 1950 ). Variable yield losses caused by drought are dependent on different factors such as duration and local environment stress (Kambiranda et al., 2011). A well distributed rainfall pat tern between 50 and 60 c m over the course of the growing season is ideal for optimal peanut production ( Nicholaides, 1969 ). Peanut originated in South America where this species is believed to have existed for thousands of years (Hammons, 1982). Peanut pla yed an important role in the diet of the Aztecs and other Native Indians in South America and Mexico. The Portugese explorers may have brought peanut to Africa whose people later introduced the plant into what became the United States. On the other hand, t he Spanish have been credited for bringing peanut from South America to Asia through t he former colony, the Philippines (Kaprovickas, 1969).

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20 Today, peanut is a crop of global importance to both smallholder and large commercial producers. It has been widely grown in the tropics and subtropics, where it has been classified as both a grain legume and oil crop (Hymowitz, 1990) World annual production is about 46 million tons per year. China has the largest production of peanut, a share of about 45% of the overall world production, followed by India (16%) and the United States (5%). Peanut is the 12 th most valuable crop grown in the United States with a farm value of over one billion U.S. dollars. The United States produced m ore than 2.0 mmt from about 1 354, 00 0 planted acres, largely from the stat es of Georgia, Alabama, Florida, and Texas ( A nonymous 2015 ). I n 2014 Florida produce d about 13% of the USA peanut supply, mostly in nine counties. While there are many varieties of pe anut grown in the USA, Runner is by far the most commonly grown among the four market types ( Tillman et al., 2015 ). Because of their high protein content and their chemical profile, peanut is a good food source for the human diet (Grosso et al. 1994 ; Nelson and Guzman 1995). Worldwide a large percentage of peanut seed is used for the production of edible oil, whereas in the USA about 60% of the production is processed in a variety of ways and eaten as food (Allen, 1981). Plant parasitic Nematodes Associated with P eanut Diseases pose a major threat to the production of peanut each year, and prevention of disease on peanut is a major concern Plant parasitic nematodes are among the most important soil borne constraints in successful peanut production. Worldwide plant parasitic nematodes considered important on peanut include th ree species of Meloidogyne Pratylenchus brachyurus, Belonolaimus longicaudatus, Criconemoides ornatus, Aphelenchoides arachidis, Scutellonema cavanessi, Tylenchorynchus brevilineatus, and Ditylenchus africanus (Dickson, 2005).

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21 Species of Meloidogyne P arasitizing P eanut Three Meloidogyne spp. that caus e significant sup p ression of peanut yields and fruit quality are M arenaria (peanut root knot nematode) race 1, M. javanica (Javanese root knot nematode) races 3 & 4 and M. hapla (northern root knot nema tode). In the United States, M arenaria is the most damaging species on peanut in the more southern states, whereas M. hapla is the damaging in North Carolina, Virginia, and Oklahoma ( H irunsalee et al., 1995 ). M. javanica is sometimes found to be mixed wi th M. arenaria in the field (Cetintas, et al., 2003 ) M. haplanaria (Eisenback et al., 2003) was recently described as a new species parasitizing pea nut in Texas, however little is known regarding its effect s on peanut outside of Texas Cliff and Hirschmann (1985) described the morphological variation s among seven populations of M. arenaria representing both physiological races. R ace 1 infects and reproduces on groundnut, whereas race 2 does not (Taylor and Sasser, 1978). In a recent study, a race 3 of M. arenaria was reported. This race infect s and reproduce s on both resistant tobacco and pepper but not on cotton or peanut in Spain (Robertson et al ., 2009). Most recently, M. arenaria race 3 has been found infecting peach in Florida (Sai Qiu, personal communication, 2017). The loss of peanut yield caused by the peanut root knot disease can reach up to 50% or greater in severely infested fields (Proite et al., 2008). Peanut has been reported to be infected by M. javanica in Egypt (Ibrahim and EI Seady, 1976), Georgia (Minton et al., 1969) and Texas (Tomas zewski et al., 1994). Other reports of M. javanica on peanut are from India (Patel et al., 1988; Sakhuja and Sethi, 1985), and Brazil (Lordello and Gerin, 1981). The first observation of M. javanica parasiting peanut in Florida was found in a commercial production field in 2002 (Lima et

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22 al., 2002). Later, Cetintas et al. (2003) confirmed the occurrence of M. javanica on peanut in Florida Their investigation showed the ratio of M. javanica to M. arenaria race 1 in the peanut crop was approximately 29% vs. 71% respectively M. hapla has been reported damaging peanut North Carolina, Virginia, and Oklahoma ( Hirunsalee et al., 1995; Machmer 1951 ; M iller and Duke, 1961 ) and it has also been f ound on peanut in some s outhern s tates ( Christie, 1959 ). Other S pecies of N ematodes A ssociated with P eanut Pratylenchus brachyurus is commonly found associated with peanut grown in Alabama (Steiner, 1945), Georgia (Boyle, 1950) and Florida The extent of damage caused by this pathogen is not clear (Minton, 1984). Belonolaimus longicaudatus is also reported to occur on peanut depending on location (Ab u G harbieh et al., 1969). It was reported in Florida causing extensive field damage to peanut in Levy County (Kustuwa et al ., 2015). Damage on peanut by Criconemoides ornatus occurs especially in the southeastern United States, but only when they reach a high density (Sasser et al., 1968; Minton and Bell, 1969). The occurrence s of other nematode pa rasites causing disease of peanut are limited in their distribution (Dickson, 2005). Root k not N ematode on P eanut M eloidogyne arenaria race 1 is the dominant root knot nematode species infecting and causing damage on peanut. The infected roots pegs and pods have typical galls with egg mass es (Sasser, 1954). Galling of the root systems inhibits plant absorbing n utrients on that leads to slow growth, stunting, and yellowish leaves. Because s oil particles often adhere to egg masses, the infected roots have a crusty appearance. These diseased pe anut plants are often infected by second ary soil borne pathogens such as Sclerotium rolfsii and Sclerotinia minor (Starr et al., 1996, 2002).

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23 The nematode causes stunting of the plants and suppresses yield. Galled pods have a distorted appearance which is not acceptable in the market. Additionally, galling severity is positively correlated to aflatoxin contamination of peanut kernels (Timper and Wilson, 2002). Disease C ycle The disease cycle of root knot nematode i n peanut starts with the penetration of roots by the second stage juveniles (J2), which is the infective stage (Karssen, 2002). The J2 in the soil penetrates a suitable root by using its stylet to thrust repeated ly into cells at the surface. After forcing its way into the root, the juvenile move s between and through cells eventually positioning its head adjacent to the vascular tissues. Within 2 to 3 days, the juvenile becomes sedentary, with their head embedded in the vascular cylinder to establish a feeding site. The nematode then begins to grow in diameter, loses its ability to move, and matures. On the process of establishing a feeding site the female injects secretory proteins (from the esopha geal gland) that stimulate physiological changes within the parasitized cells known as "giant cells". When these specialized cells increas e in size changes occur that allows the giant cell s to produce large amounts of proteins as food resources for the nem atode (Proite et al., 2008) Because of the nematode esophageal gland cell secretions, the plant responds producing more growth regulator s which result in increasing cell size (hypertrophy) and number of cells within the adjacent cortex (hyperplasia) (B arker et al., 1998). Root cells around the giant cells enlarge and divide rapidly, leading to gall formation (Hussey and Mims, 1991). T he nematode goes through two additional juvenile stages interspersed by molts. T he mature female is much wider but not much longer than the original second stage juvenile. The f emale body becomes spherical or pear shaped, with a

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24 diameter of about 63.5 and a narrow neck embedded in the vascular tissue. The life cycle is completed wh en the female begins laying eggs. One single egg mass normally bears 500 to 800 eggs but the number varies from almost none under unfavorable conditions to as many as 2,000 under highly favorable conditions ( Taylor, 1978 ) The development of the nematode is dependent on e nvironmental conditions and hosts suitability ( Proite et al., 2008 ). Eggs are deposited in a gelatinous matrix that becomes visible outside or near the galled surface of the roots. Secondary infection of pe anut roots, pegs and pods occur a s newly hatched J2 move to initiate new feeding sites (Dickson and De Waele, 2005). Pa r th en ogenesis is the main mode of reproduction of M. arenaria Males are not required for reproduction and they are normally found in small numbers but may increase under adverse conditions such as food shortage, crowded population densities, and high soil temperature (Triantaphyllou, 1960 ) The length of the life cycle and the population densities depend upon several factors. S oil temperature, host suitability, and soil type are considered to be among the most important At 27C which is usually optimum for most Meloidogyne spp one generation on susceptible peanut requires approximately 25 to 35 days, whereas at 19C at least 87 days are necessary ( Tyler, 1933 ) Thus, three to five generations are possible in the field. The life cycle is longer on a less suitable host. Sandy, organic muck and peat soils favors population build up more than heavier clay soils ( Van, 1985 ). The nematode can overwinter inside or out sid e of galled roots or pods in soils ( Nusbaum, 1962 ) The second stage juveniles (J2) are hatched from eggs when soil temperature s are around to 12 These J2 serve as initial inocu l um for a following susceptible crop ( Wallace, 1963 ). Soil temperature, humidity, soil type and cultivar

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25 management are considered as main factors determining severity of root knot disease (Wallace, 1963). The egg mass index (EI), which was introduced in the literature by Taylor and Sasser (1978), was origin ally developed by members of the Southern Regional Technical Committee on Nematodes USA It gives an approximate measurement of root knot nematode reproduction based on numbers of egg masses/plant instead of actual numbers of eggs and juveniles/plant. Sin ce the number of eggs per egg mass varies, the El is not a quantitative measurement of reproduction; still, it has been used as a basis for estimates of host efficiency. Likewise, the gall index (GI), which is based on the same scale as the El, has been us ed as an indicator of plant damage even though little work has been used to correlate GI with crop yield. In the differential host test an average El and an average GI of 2 or less are interpreted as indicating host resistance and those greater than 2 as i ndicating host susceptibility. Parasitic M echanism For plant parasitic nematodes to develop it is necessary that they feed on certain hosts, absorbing nutrients to comple te their life cycle. T wo morphological characters a stylet and an esophagus, ma ke it possible for a nematode to be parasit ic (Blaxter, 1997). Root knot nematode is a sedentary plant parasite with an advanced evolution of parasitism. This nematode has a typical T ylenchoida stylet and an esophagus with strong metacorpus as a pump chamber and three secretory gland cells, one dorsal (DG) and two subventral (SvG). These glands are the major source of the secretions involved in plant parasitism (Hussey, 1990). Root knot nematodes are able to secret proteins to modify host root cells into very specialized feeding cells by the alteration of gene expression of host cells (Gheysen et al., 1997). During different

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26 stages in the life cycle of root knot nematodes, the morphology and function of esophageal gland cells is altered The SvG are most funct ional at initial parasitism by infective J2 whereas the DG is the predominant gland in the parasitic stages (Bird, 1983). Infective J2 penetrate the host root and migrate through the cortex cells to the vascular tissue. Meanwhile, degrading enzymes for ce ll wall penetration are secreted from the esophageal glands through the stylet of root knot nematodes to facilitate migration by weakening or rupturing cell walls. Giant cells induced by root knot nematode are a result of an abnormal increase in cell numbe r and size following repeated mitosis without wall degradation (Bird, 1996; Fenoll et al., 1997). When J2 undergo further development, the cuticle structure is altered by decreasing layers with the basement membrane left in order to absorb more nutrients f rom host. The nuclear region of the gland cell s is where synthesis of secretory proteins occurs. The mature enzymes are stored in Golgi derived membrane bounded granules that are connected with ampullae by microtubules in the gland cell (Burgess and Kelly 1987). The number of different secretory proteins may change according to gland cell type and parasitic stages (Burgess and Kelly, 1987). After a plant parasitic nematode penetrat es the host with its stylet, enzyme s are secreted from the esophageal glands and injected into plant cells through the stylet ( Carneiro et al., 1996 ). These enzymes are able to stimulate their host to produce abnormal growth regulators that aid in the development of feeding cells thereby providing nutrients ( Jone s and Northcote, 1972 ). S tudies on the esophageal glands and stylet secretions obtained from root knot nematodes confirmed that a group of proteins (e.g. glycosylated proteins ), but not nucleic acids, were involved in the secretions ( Davis et al., 2000). The composition of

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27 secreted proteins via the nematode stylet was analyzed by designed chemical stimulation of the production of stylet secretions in a vitro system Esophageal gland antigens have been iso lated by monoclonal antibodies and used for direct a nalyses or screen ing cDNA expression libraries corresponding to secretion genes (Davis et al., 2000) Population D ynamics Plant parasitic n e matode population dynamics play an important r ole in plant nematology because of the relationship with the incidence of crop disease These provide the basis for advisory programs in agriculture (Wallace 1963 ). Vertical and seasonal distribution s of nematode s are highly variable but they are generally related to the distribution of host roots (Wallac e 1963 ). In most cases, plant nematode s in soil seem to have favorable zone s where they concentrate most likely related to the ir survivability It has been suggested that host distribution, root depth, height of water table, soil moisture, soil type, dep th of subsoil and temperature affect vertical distribution patterns (Ferris, 1971; Miller, 1960; 1972; O'B annon 1972; Potter, 1967 ). There are several observations on the distribution of nematode in different soil types and different seasons ( Ferris and Mc Kenry 1974; Norton et al 1971 ). The t op 45 cm of soil seems to be the optimum zone for nematode s to congregate ( Barker and Nusbaum, 1971 ). However, different nematode species have diversity in their vertical distribution pattern s According to Godfre y (1924), root knot nematodes tend to concentrate at a depth to 20 cm deep) for determining prep lant nematode population densities Root knot ne matode populations overwi nter ing at a deeper soil depth may be considered as a

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28 strategy for survival A t this deeper zone they escape detection, fumigation and are less likely to be affected by other control management tactics. This ensures their survival to serve as an initial inoculum for the next crop season (Potter, 1967). After planting a crop, nematode damage increases at a ra te inconsistent with the number of nematodes found in the preplant sample. Cobb (1914) stated that nematodes are often very numerous near the surface and gradually drop in number s as the soil depth increases. H igher population densities are reported to be found from 20 to 50 cm deep with few being detected at 86 cm deep or greater (Godfrey, 1924). In extreme environmental conditions, nematode p opulation densities are low or variable in the upper 15 cm of soil with greater numbers being found at 30 to 45 cm deep. I n some instances, numbers may remain relatively high down to 120 cm deep There is much information about the influence of temperatur e on nematodes development; however there are no reports of nematode reactions to temperature fluctuation in the field (Ferris and McKenry, 1974). Nematodes move in films of water that surround soil particle. Dropkin (1980) stressed that many nematode genera a nd species have particular soil types and climatic requirements and that certain species of nemato des prefer to live in sandy soil, whereas others prefer clay or loamy soil. Most plant parasitic nematodes are reported to survive well in a coarse te xtured soil (Wallace, 1968; 1971). The relationship between nematode movement and soil texture is a function of the ratio of nematode size to pore and particle size. There are marked positive correlation s between length and diameter of the nematode and opt imum pore and particle sizes that allows for maximum movement (Brodie, 1975).

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29 In the presence of host plant s root knot nematodes can migrate towards roots of a host plant while moving up or down in the soil layers. Johnson and McKeen (1973) found that a population of M. incognita present in the top 15 cm can move down to a depth of 120 125 cm According to Pinkerton and McIntyre (1987), M. chitwoodi J2 occurring in soil can migrate 30 cm upwards to penetrate tomato plants. H igher population numbers were reported from 20 to 50 cm deep than at shallower depth s, whereas very few were detected at 86 cm deep or greater (Godfrey, 1924). Under extreme environmental conditions, the nematode population densities were low or variable in the upper 15 cm whereas the largest numbers of nematodes were found at 30 to 45 cm deep with decreasing numbers at deeper levels and relatively high number s down to 120 cm. However, Ferries and McKenry detected Meloidogyne spp in a vineyard in California 120 cm below the soil surfa ce and could be detected at 330 cm deep (Ferris and McKenry 1974). Juveniles and eggs are supposed to be survival stages of nematode s in soil Different nematode species have different survival stages that are affected by temperature. Daulton (1961) and N usbaum (1962) stated that based on their field experiment s Meloidogyne javanica and M. hapla overwinter as eggs which is the same survival form for Xiphinema americanum According to Baunacke (1922), with enough stored food, the juveniles of sugar beet c yst nematode can survive for months in the lab. However, their activities decreased at 15 or lower. As the temperature increases to 25 to 28 nematode activity increases so much so that the infective stage must find a suitable host or else it depletes its stored foods (Dropkin, 1963). It was reported that the higher the soil temperature, the shorter the survival period of juvenile s under

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30 fallow conditions because of increasing activity at higher temperature s (Bergeson, 1959). A ccording to these findings, Van Gundy et al. (1976 ) summarized that the nematode s internal food reserves were the main factor determining the survival of many plant parasitic nematodes Moreover, they concluded that root searching activity causes greater use of food reserves based on the fact that food reserves were consumed by juveniles faster in the presence of roots than in their absence. Temperature Effects on Root knot N ematode Most Meloidogyne spp. are important pathogens in tropical, subtropical and temperate regions with relatively mild winters. Thus temperature is considered as one of the most important factor s that govern s the occurrence and severity of root knot disease (Godfrey, 1926). Temperature has great bearing upon geographical distrib ution, population development and survival. I n southern states, long summers combined with favorable soil temperature and moisture conditions allow multiple generation s of root knot nematode s in one single growing season thereby le ading to high population densities in the soil (Godfrey, 1926) In contrast, in northern areas with a shorter summer season fewer generations of nematode s and low er population densities occur Meloidogyne sp p. that have different climate ranges have been reported in several studi es (Bird and Wallace, 1965; Olthof, 1967; Thomason and Lear, 1961). Temperature effects on the life cycl e of the root knot nematode have been well documented. Each nematode species has its own optimum temperature range for all phases of development from th e egg stage through the egg laying stage. Susceptible plant roots may be infected when the temperature range s from 12 to 40.5 ( Tyler, 1933) In addition, it pointed out that the temperature threshold for nematode maturity and reproduction seems to be higher than that for early stages of the life cycle, but that

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31 no egg s were found above 31.5 Bergeso n (1959) indicated that most egg s hatched at a temperature of 15.5 and above wh ereas below 10 surviv al and egg hatch decreased rapidly. Similar reports show ed that temperatures below 15.6 and above 35 limit ed the reproduction of M incognita ( Thomason and Lear, 1961). Root knot Nematode M anagement Cultural and physical practice s biological organism s host resistance, and nematicide s are traditional tactics used to manage the peanut root knot nematode Crop r otations are one of the most effective cultural management methods For example, rotation of peanut with cotton, velvet bean, or bahiagrass can m itigate M. arenaria population densities and increase peanut yield s ( Dickson and Hewlett, 1989; Johnson et al., 2000; Rodrguez Kbana et al., 1986, 1987, 1991a, 1991b ). However, long periods of growing low value crop s and the nematodes ability for long term survival in soil without host s compromise the usefulness of crop rotation. Both fumigant and nonfumigant nematicides have been proven effective for managing root knot disease (Dickson, 1988). The fumigant nematicide 1, 3 D i s highly effective, however because of measured production costs associated with its use and other environmental concerns many growers seek other options. Though biological antagonists are effective in suppressing root knot nematode, there are limitations to their use (Bale et al., 2008). Therefore, resistant cultivar s are thought to provide a more sustainable method for managing the endoparasitic nematode pathogen s such as Meloidogyne spp. Develo pment of Resistant Peanut C ultivar s Utilization of resistan t cultivar s is considered an effective component in pest management (Williamson, 1999). By definition, with plant resistance the reproduction of a nematode species should be suppressed up to 90% or more (Cook and Evans, 1987).

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32 In response to infection by a nematode, the plant resistance gene reduces the parasitic Moreover, compared to a susceptible plant, the galls formed on roots of a resistant cultivar may be reduced or completely lacking because of the incompatible inte raction between the parasites and the resistant host ( Williamson 1999 ). Numerous attempts were made to find root knot n ematode r esistance in cultivated peanut germplasm Researcher s were aware that a number of other Arachis spp. were highly resistant to the peanut root knot nematode ( Simpson, 1991 ). In 1995 a breakthrough was made when resistance genes from wild peanut were transferred into cultivated peanut. Resistant genes to root knot nematode from A. cardenasii were successful introgressed into A. hypogaea by a backcross breeding pat hway (Starr and Morgan, 2002). Garcia et al. (1996) reported that a germplasm line called TxAG 6 with resistance was generated by interspecific hybridization [ A. batizocoi x ( A. cardenasii x A. diogoi )]. The germplasm li ne TxAG 7 was obtained from the first backcross generation of TxAG 6 and resulted in release of the first root knot nematode resistant peanut cultivar COAN (Simpson and Starr 2001). In 2003, NemaTAM was the second named peanut cultivar released with resis tance to root knot nematodes. Both were released by the Texas Agriculture Experiment Station (Simpson et al 2003). NemaTAM had the same level of resistance as COAN, but greater yield potential. Both COAN and NemaTAM are derived from the same introgressio n but selected from a different backcross generation (Simpson, 1991). However, neither is resistant to M. hapla Moreover, neither of these cultivars was considered for planting in the southeastern USA because of the problems with tomato spotted wilt virus (TSWV) and

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33 other fungal diseases endemic to the area (Rich and Tillman, 2009). In 2007, a runner type peanut cultivar ( Arachis hypogaea L. subsp. hypogaea var. hypogaea ) Tifguard was developed that was resistant to both root knot nematodes and TSWV. Tifg uard was released by the USDA ARS and the Georgia Agricultural Experiment Stations (Holbrook et al exhibite d a high level of resistance to TSWV, M. arenaria and M. javanica (Holbrook et al 2008). Identification of Resistant G ene to Root knot N ematode It has long been a question of interest on how a single gene governs resistance to a nematode by interfering with the establishment of the elaborate changes caused by the parasite in host root s (Williamson, 1994). After being attracted to and penetrating roots, root knot nematode s migrate to the vascular tissue in a similar manner in both resistant and susceptib le plants (Dropkin, 1969; Ho et al., 1992; Paulson, 1972). However, in resistant plants, the nematode is not able to establish feeding sites Instead, a hypersensitive response (HR) formed by a localized region of necrotic cells are induced near the anteri or portion of the invading J2 (Dropkin, 1969; Ho et al., 1992; Paulson, 1972). The J2, surrounded by necrotic cells, usually die and others that fail to establish feeding sites may egress from the roots. The earliest indication of HR is visible about 12 h after inoculation (Paulson and Webster, 1972). HR does not occur wh en J2 are migrating through the root tissue, but do es so when they attempt to establish a feeding site. The resistance from wild Arachis spp. was identified and reported to be controlled by two dominant genes Mag and Mae. The former inhibits galls formation and

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34 the latter suppresses egg production (Choi et al 1999). The advanced germplasm line TxAG 6 mentioned above is an F1 from the com plex hybrid of [ A. batizocoi x ( A. cardenasii x A. diogoi )], with each one having resistant genes to M. arenaria (Choi et al 1999) The resistance to M. arenaria in A. cardenasii results in the inhibit ion of development and is considered to be a hypersen sitive host reaction that occurs near the anterior end of the nematode within a few days of infection (Starr et al 1995). The resistant mechanism to M. arenaria in A. batizocoi was considered to involve a different mechanism Though the J2 invade d into A batizocoi roots, few of them remain ed and develop ed inside the roots (Choi et al. 1999). In addition, A. batizocoi resistance is reported to increase the time required for M. arenaria to finish its life cycle. However, no hypersensitive reaction was obs erved in this Arachis spp Another germplasm line TxAG 7 is derived from the first backcross generation of A. hypogaea TxAG 6 (Starr et al 1995). The resistance to M. arenaria in TxAG 7 was similar to that of A. cardenasii in whic h an apparent necrotic HR occurred. But the occurrence of host cell necrosis of a HR reaction was not observed when J2 invaded into the root system (Starr et al., 1990) The resistance in COAN generated from TxAG 7 is contro lled by a single dominant gene. Most invasive J2 emigrate d from roots whereas the few remaining J2 develop ed to reproductive females (Choi et al 1999). T hree different experiments were designed to evaluate the mechanism of resistance to M. arenaria in COAN (Bendezu and Starr, 2003) Th e rare necrosis that occurred in host tissue in roots of COAN suggested that resistance to the peanut root knot nematode does not involve a necrotic, hypersensitive reaction. Furthermore, resistance in COAN block s most of the J2 from penetrating roots. Onl y 1 of 90 was observ ed within the vascular

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35 cylinder, whereas more than 70% of J2 were observed in the vascular tissue of the susceptible cultivar (Bendezu and Starr, 2003). A gene named Rma is assumed to be a dominant root knot nematode resistance gene in troduced from other Arachis species. Identifying and mapping of alien DNA sequence by codominant DNA markers, most of clones are leucine rich repeat family of plant genes. In addition, a four genome equivalent fosmid library was developed from the resistan t cultivar, which assembled into two long contigs. NT946 has about 32,923 bp whereas NT344 is about 30,462 bp. By finer mapping with marker assisted selection, the R gene of wild peanut was found to be located on chromosome 9A and 9B. Mag and Mae were disc overed to relate to Rma. The function of Mag is reported to inhibit formation of galls, whereas Mae suppresses egg production of the root knot nematode (Nagy et al., 2010). The Application of Host R esistance Long term use of nematode resistant cultivars ca n cause extensive selection pressure on the target species and affect nontarget nematodes as well (Lawrence, 1992). Problems from planting resistant cultivars successively include shifts in nematode races or species. An example of shifts in species of root knot nematode in response to planting resistant cultivars occurred with M. incognita on tobacco in North Carolina. There was a species shift from M. incognita with an increase finding of M. arenaria and M. javanica infecting resistant cultivars of tobacco in North and South Carolina (Barker, 1989; Johnson, 1989). This was believed to have resulted from long term use of resistant cultivars to combat M. incognita problems. In California there have been reports of resistant tomato cultivars being overwhelmed by resistance breaking nematodes within field populations. Although the Mi gene in tomato cultivars has proven

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36 to confer effective resistance against several root knot nematode specie s ( Williamson, 1998 ), numerous reports of root knot nematode have been identified worldwide parasitizing resistant tomato plants (Tzortzakakis et al., 1996; 2005; Carvalho et al., 2015). Severe root knot nematode infection on the resistant peanut cultiv ar Tifguard was reported in Levy County, Florida in 2012. Root knot disease was found on Tifguard in a total of 14 sites over the summer. Tifguard had previously been reported as nearly immune to M. arenaria and M. javanica. The reason for the brea k down i n Tifguard was unknown.

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37 CHAPTER 2 FIELD EVALUATION OF RESISTANCE TO M eloidogyne arenaria IN BREEDER, FOUNDATION AND GROWER KEPT TIFGUARD Introduction Tomato spotted wilt virus and root knot nematodes cause damage and yield suppression on peanut grown in Florida (Holbrook et al., 2008). Because of nematode problem, the US Department of Agriculture (USDA) released a runner cu ltivar in 2007 named Tifguard. The cultivar was reported as highly resistant to root knot nematode. It was the first peanut cultivar w ith a combination of resistance to TSWV and root knot nematodes. This cultivar provided growers with another option to manage nematode and TSWV diseases in the southeastern United States. Although, Georgia 06G became the dominant cultivar planted Tifguard gained a larger proportion of the acreage where root knot nematode was prob l ematic (Tillman et al., 2013). With Tifguard, root knot nematode infested fields could be put back into production without large increases in production costs. There are five sourc es of Tifguard seed breeder, Foundation, Certified, Registered, and grower kept. Breeders first produce a relatively small quantity of seed of any new cultivar. These seeds are called Breeder seed in the seed certification system and are directly controlled by the plant breeding institution, firm or individual that is the source for production for the certified classes (USDA, 2002). Foundation seed is the progeny of breeder seed, handl ed so as to maintain specific genetic purity and identity, production of which must be acceptable by the Department of Agriculture and Forestry (USDA, 2009). Registered and Certified seed are the progeny of breeder or Foundation seed, handled under the sam e procedures as Foundation seed to maintain satisfactory genetic purity and identity. Grower kept seed is the progeny of peanut

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38 produced from the previous season and used as a seed source for the following crop season. Although Tifguard was reported to be highly resistant to Meloidogyne arenaria and M. javanica root knot nematode damage was observed on grower kept Tifguard peanut in summer 2012 in production fields located primarily in Levy County, FL. Typical above ground symptoms of root knot disease we re noted. The emergence of a resistance breaking race of M. arenaria was considered as a possible reason. This possibility was highlighted because of the long term monoculture of peanut in some production fields in Levy County. The practice of continuous p lanting of resistant cultivars leads to the increased possibility of resistance breaking by the soi l borne pathogens ( Frisvold, 2010 ). In addition to the possible developme n t of resistance breakings strains of the nematode, higher soil temperature also was considered as a factor that could impact root knot disease on Tifguard. This has been demonstrated with the Mi gene in tomato that confers resistance to several Meloidogyne sp p ( Williamson 1998 ) The objective was to evaluate Tifguard seed obtained from three sources to determine the cause for root knot nematode infection on the resistant cultivar. Materials and Methods Field Sites and E xperim ental D esigns Resistance to M. arenaria in Foundation seed and grower kept Tifguard seed was evaluated in the same field site (464 Farm) where a large number of Tifguard plants infected with root knot nematodes were found in 2012. The field design was a randomized complete block with four replicates. In each block two rows of Foundation Tifguard and two ro ws of grower kept Tifguard, were com pared with two rows of Florida 07 (susceptible) ( Gorbet and Tillman, 2009 ). The plots were 3.0 m long and rows were

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39 spaced 0.9 m apart, with each row seeded with 16 seeds. The experiment was conducted in 2013, 2014 and 2 015 in the same blocks with the following exceptions. In 2014 six rows of breeder Tifguard were compared with two rows of FloRun 107 (susceptible) (Tillman and Gorbet, 2015) In 2015 Georgia 06G ( Branch, 2007b ) seed replaced Florida 07. The plots were spra yed with a back pack sprayer every 14 to 20 days with c hlorothalonil. Weeds were removed by hand pulling. After 140 days, all the peanut plants were dug to remove roots, pegs and pods. The roots were examined for galling and egg masses immediately after d igging. Red food coloring solution (0.5 L/10 L H 2 O) was used as an aid to visualize egg masses (Thies et al., 2002 ) The assessment for peanut plants was recorded as positive for infected roots and negative for roots without infection. In 2014 and 2015, r esistance to M. arenaria in breeder Tifguard seed was compared with Georgia 06G in two separated locations both known to be heavily infested with M. arenaria the Brown Farm, Levy County, Florida and the University of Georgia, Attapulgus Research and Education Center (AREC), Attapulgus, Georgia. The field design was a randomized complete block with five replicates at the Brown Farm and six replicates at AREC. The row length was 6 m and the row spacing was 0.9 m. The production methods followed the IFAS, University of Florida and University of Georgia Peanut Production Guides (Stephens, 1994; Lee and John, 2014). At harvest, the peanut plants were examined for root knot nematode galling and egg masses of roots, pegs and pods Red food coloring solution (0.5 L/10 L H 2 O) was prepared to stain egg masses. Each peanut plant was recorded as positive if infected by RKN and

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40 negative for plants without infection. Both f ields were overseeded with red crimson clover ( Trifolium cv Dixie) during autumn and winter seasons. Determination of Root knot Nematode S pecies A total of 250 (100 from Brown Farm, 100 from AREC and 50 from 464 Farm) females were extracted from infected roots and assessed by PAGE. Each female was placed in a microtube in a 5 l of deionized water and stored at 20 Identification of the root knot nematode species infecting Tifguard was made by analyses of isozyme phenotype following their resolution via the polyacrylamide gel electrophoresis protocol (Dickson, 1971 ; Ebenshade and Triantaphyllou, 1985 ). Electrophoresis was performed with the use of a Bio Rad mini PROTEAN II unit (Bio Rad, Philadelphia, PA). Single females were homogenized individually in the microtubes. Before homogenizing 5 l of loading dye (Bio Rad) was added. Each gel contained 15 wells, with each one loaded separately with a female extract. A single M. javanica female homogenate was placed into wells 1 and 15 to serve as a control. Electrophoresis was conducted in a discontinuous buffer system with 8% acrylamide running gel, pH 8.8, and 4% acrylamide stacking gel, pH 6.8. The voltage was maintained at 80 volts for the first 15 minutes and i ncrease to 200 volts for 35 minutes. After electrophoresis, the gels were removed and placed into an enzyme reaction mixture to resolve esterase and malate dehydrogenase isozyme patterns. Analysis of Resistance Gene in I nfected Tifguard P lants A young lea f was removed from Tifguard plants displaying root knot nematode galling or egg masses. The leaf was placed in a microtube for overnight delivery to the National Environmentally Sound Production Agriculture Laboratory (NESPAL), Tifton,

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41 GA. They were exami ned for the presence of the root knot nematode resistance gene marker (Chu et al., 2011). Results and Discussion 464 Farm 2013 2015 In 2013, 30% of the plants grown from Tifguard Foundation seed were found infected by M. arenaria whereas 40% of peanut g rown from Tifguard grower kept seeds were infected ( Table 2 1). All the peanut plants of Florida 07 were heavily galled. In the 2014 and 2015 season, no Tifguard breeder plants had visible galls or egg masses, whereas all Georgia 06G plants were heavily ga lled and with numerous egg masses ( Table 2 1). Brown Farm and AREC 2014 2015 In 2014 and 2015, 12% and 3.9% of Tifguard breeder plants were galled at the Brown Farm field site, respectively (Figure 2 2 A), whereas at the AREC field site 7% and 1.8% Tif guard breeder plants were galled, respectively (Figure 2 2 B). In both trials all the Georgia 06G susceptible plants were heavily galled and with numerous egg masses (Figure 2 2). Determination of Root knot Nematode Species Based on esterase (Est ) and malate dehydrogenase (Mdh) phenotypes. All the nematode samples were identified as Meloidogyne arenaria. These phenotypes are the most commonly found in populations of M. arenaria around the world infecting not only peanut but also other crops ( Brito et al., 2008 ).

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42 Analysis of Resistance Gene in I nf ected Tifguard P lants Based on the detection of the resistance gene ma r ker by the NESPAL, 2.5%, 28% and 39.5% of the Tifguard breeder, Tifguard Foundation and the Tifguard grower kept seed, respectively were negative for the resistant gene marker (Figure 2 2 ). The breeder seeds had the highest proportion of plants with the resistance gene. It appears that the seed purity declined after multiple increases of peanut seed production. It was surprising tha t the foundation seed had a relatively high percentage of plants with the resistance gene. The grower kept seed had the highest proportion of Tifguard plants negative for the resistance gene. Under the field condition where different peanut cultivars are p lanted in close proximity, it is possible that outcrossing occurred within Tifguard or that volunteers of a susceptible cultivar from the previous crop remained in the field and were mixed with Tifguard pods at harvest. The re were some false positive diagnoses in the field for galling on Tifguard. It is difficult to visualize root knot nematode galls on peanut roots, especially when there are only a few. The large number of rhizobium nodules on peanut roots interferes with t he determination of small root knot nematode galls. It is not possible to say whether some plants categorized as positive were not actually infected. It is also possible that some Tifguard plants that had the resistance gene could have been infected by a s train of the nematode capable of infecting the resistan t peanut

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43 Table 2 1. The average percentages of three different sources of Tifguard seed infected by Meloidogyne arenaria at the 464 Farm, Levy Co., FL in 2013, 2014, and 2015. P eanut seed sources 1 R oot knot nematode infection (%) 2013 2014 2015 Tifguard breeder N/A 0 0 Tifguard Foundation 30 N/A N/A Tifguard grower kept 40 N/A N/A Florida 07 100 N/A N/A FloRun 107 N/A 100 N/A Georgia 06G N/A N/A 100 1 Tifguard Foundation and grower kept seed were compared with Florida 07 in 2013 and Tifguard breeder seed was compared with FloRun 107 in 2014 and Georgia 06G in 2015.

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44 Figure 2 1 The average percentage s of Tifguard breeder seed infected by Meloidogyne arenaria was compared with Georgia 06G at two locations in 2014 and 2015 A ) the Br own Farm, Levy Co., FL. B) University of Georgia, Attapulgus Research and Education Cente r Attapulgus GA. 0% 20% 40% 60% 80% 100% 120% Georgia-06G Tifguard breeder Georgia-06G Tifguard breeder 2014 2015 Average percentage of peanut infected by Meloidogyne arenaria 100% 100% 0% 20% 40% 60% 80% 100% 120% Georgia-06G Tifguard breeder Georgia-06G Tifguard breeder 2014 2015 Average percentage of peanut infected by Meloidogyne arenaria A B 100% 100% 12% 3.9% 7% 1.8%

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45 Figure 2 2 The average percentage s of different Tifguard seed sources that were negative for the resistance gene marker for Meloidogyne arenaria 0.00% 5.00% 10.00% 15.00% 20.00% 25.00% 30.00% 35.00% 40.00% 45.00% Tifguard Breeder Tifguard Foundation Tifguard grower-kept Average percentage of different Tifguard seed sources without resistance gene 28% 39.5% 2.5%

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46 CHAPTER 3 THE SEASONAL AND VERTICAL DISTRIBUTION OF M eloidogyne arenaria FROM RESISTANT AND SUSCEPTIBLE PEANUT GROWN IN TWO SOIL TYPES Introduction Nematode population dynamics plays an important role in plant nematology because of its association with the incidence of crop dise ases and its utility as the basis for advisory service s (Wallace 1963 ). Vertical distribution of nematodes is highly variable but is generally related to the distribution of host roots (Wallace 1963 ). In most cases, plant nematodes concentrate in a favor able soil zone. It has been suggested that host distribution, root depth, height of water table, soil moisture, soil type, depth of subsoil, and temperature affect vertical distribution patterns (Ferris, 1971; Miller, 1960; 1972; O'B annon 1972; Potter, 19 67). There are several observations on spatial distribution of plant parasitic nematodes in different soil types, different host crop and seasons ( Ferris and Mc Kenry 1974; Norton et al 1971 ). The top 45 cm of soil seems to be the optimum zone for most nematodes to congregate ( Barker and Nusbaum, 1971). However, nematode species vary in their vertical distribution patterns. Meloidogyne arenaria is a highly virulent soilborne pathogen of peanut that has the ability to produce multiple generations during the relatively long peanut growing season. Therefore, the population density of this nematode often builds to high levels. The nematode remains nearly undetectable and symptoms on peanut may be difficult to observe until 75 to 80 days after planting (Rodr guez Kbana et al., 1986). But as the growing season progresses, stunted and yellow plants are often accompanied with increasing nematode populations in infested sites.

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47 The pre plant threshold for this nematode was determined to be one egg or second stage juvenile /100 cm 3 of soil (McSorley et al., 1992). It is difficult for farmers to detect this low preplant threshold density by conventional extraction techniques (McSorley et al., 1992; Rodrguez Kbana et al., 1982). Population densities of this nematod e can increase exponentially as the host crop develops, driven by multiple reproduction cycles by harvest time (Rodrguez Kbana et al., 1986). At harvest, damage is most easily observed on roots, however pegs and pods may be galled. A carrying capacity of 5,000 to 6,000 J2/100 cm 3 of soil was reported in microplot experiments (McSorley et al., 1992), however in field soil numbers were about 10 fold lower (Rodrguez Kbana and Ivey, 1986). In Alabama the highest and most variable population density of J2 w as found in the top 30 to 40 cm in a vertical distribution study (Rodrguez Kbana and Robertson, 1987). The fluctuation of the J2 densities in the top soil layers was associated with the abundance of host roots, and a peak was observed before or at harves t. Deeper in the soil, the population densities were lower but fairly constant no matter at what time of year sampling occurred. However, this vertical distribution of J2 density observed in Alabama may different in deep sandy soil, such as those that exis t in Florida. In such conditions, high densities of J2 may be recovered deeper in the soil (Dickson, 1985). The vertical distribution of M. chitwoodi to a depth of 70 cm was examined for 2 years under summer barley, carrot, fodder beet, bean, marigold and black salsify in two fields with sandy soils (Wesemael and Moens, 2008). The results indicated that the vertical distribution was not significantly influenced by degree of susceptibility of

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48 different crops. More important ly the population density was closely related to the different growing periods of the crops. In microplots studies, M. incognita moved rapidly in both directions to a soil depth of 150 cm (Johnson and McKeen, 1973). High n ematode population densities and root gall indices on tomato roots were recorded to depths of 150 cm in soil inoculated with 4,400 M. incognita kg 1 of soil. The high population density in the top 30 cm of soil reduced tomato yields by 20% in the first cro p and 70% in the following crop. Greenhouse evaluations demonstrated that high population densities of M. incognita could be recovered as deep as where the crop roots reach (Ward, 1964; Bird, 1969). Preplant population density is a predictor for root knot disease damage, so data from soil samples provide guidance for nematode management tactics (Been and Schomaker, 2006). The nematode distribution both horizontally and vertically over the period of a crop season or year usually fluctuates based on food sour ce and environmental conditions (Mojtahedi et al., 1991). To develop adequate sampling procedures, data is required regarding the vertical distribution to determine the approximate sampling depths for estimating population densities (Verschoor et al., 2001 ). The objective of this study was to quantify the vertical and seasonal distribution of M. arenaria from a susceptible and resistant peanut cultivar grown in two different soil types. Materials and Methods Field L ocations and Experimental D esigns Two pea nut field s that were previously heavily damaged by root knot nematodes were selected for comparison of M arenaria distribution over two crop seasons, 2014 and 2015. Site one, the Brown Farm, consists of a Candler deep sand and was located

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49 in Levy County, Florida Site two consisted of a Norfolk loamy sand stratified over a dense clay layer and was located at the University of Georgia, Attapulgus Research and Education Center (AREC), Attapulgus, GA. Breeder seed of Tifguard and Georgia 06G peanut cultivars were planted in paired rows with a 0.9 m row spacing. Rows were 6.1 m in length and each plot was replicated five and six times at the Brown Farm and the AREC for 2 years in a randomized complete block design The plots were carefully marked so as to repea t the planting of Tifguard and Georgia 06G in as near as possible the same spot in both crop seasons. The Brown farm was planted on 10 April, 2014 and h arvested on 8 September, 2014. In 2015, the Brown farm was planted on 1 May and harvested on 28 October. The AREC was planted on 5 June, 2014 and h arvested on 30 October in 2014. In 2015, the AREC was planted on 19 May and harvested on 04 October. The production methods followed the IFAS, University of Florida and University of Georgia Peanut Production Guid es (Stephens, 1994; Lee and John, 2014). During the winter season, both sites were overseeded with red clover ( Trifolium cv Dixie). Seasonal and Vertical D istribution of M eloid o gyne arenaria During the peanut growing season, samples were taken in the rhizosphere of Tifguard and Georgia 06G peanut cultivars beginning in spring 2014 and ending in fall 2015. After peanut harvest samples continued to be collected within the former peanut plots marked by flags. At the Brown Farm, soil samples were collected from four depths at 30 cm increments with a 10 cm diameter bucket auger to a depth of 120 cm. At the AREC, soil was collected from three depths at 15 cm increments to a depth of 45 cm. Sampling at the AREC was restricted to the top 45 cm depth b ecause of a dense clay layer. The soil from each depth sample was mixed thoroughly placed in a plastic bag

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50 and stored at 10 (Hooper, 1986) until processed. At each sampling period, a flag was placed to mark the spot to ensure a new site was chosen for th e next sampling date. Nematodes were extracted from 200 cm 3 of soil via the centrifuge flotation technique (Jenkins, 1964). The root knot nematode J2 and other plant parasitic nematodes were counted under an opt ical microscopy Temperature sensors (HOBO T idbit Temperature Data Logger, Onset Computer Corporation, MA ) were installed to measure soil temperature of each depth at both sites in 2015. A hole was dug and the sensors were inserted in a horizontal position at 18, 36, 54 and 72 cm de e p at the Brown Farm and at 6, 18, 30 cm de e p at the AREC. The sensors were set to record s oil temperatures at 5 min intervals. Statistical A nalysis The effects of soil depth, host genotype, and days after planting (DAP) on the number of J2 extracted from 200 cm 3 of soi l were subjected to analysis of variance using the following model: nematode number = constant + depth + date + (depth date ) HSD (honest significant difference) test. Det ermination of Effects of Soil T extu re on Nematode P opulation Soil chemical component analyses and pH were completed by University of Florida, Institute of Food and Agricultural Sciences Analytical Service Laboratories and soil texture analyses were performed by the Bouyoucos Hydrometer Method (Day, 1965).

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51 Results Above and B elow ground Symptoms of Root knot D isease In 2014, the aboveground symptom of root knot ne matode damage on Georgia 06G was more readily observed at the Brown Farm than that at the AREC. Symptoms inclu ded stunted, chlorotic plants at the Brown Farm, whereas at the AREC there was hardly any visible symptom of stunting or chlorosis. However, in contrasts to the aboveground symptoms, the below ground symptoms of galls and egg masses wer e more readily apparent on Georgia 06G at the AREC than that at the Brown Farm. There were no above ground symptoms on Tifguard regarding stunting or chlorosis at either site. In the 2015 season, there was a marked decrease in the aboveground root knot nem atode symptoms at the Brown Farm, whereas at the AREC the symptoms were unchanged from that observed in 2014. Seasonal D istribution of Meloid o gyne arenaria R ecovered from Georgia 06G R hizosphere I nitial population densities of J2 from all replicates and depths average d 18 and 156/200 cm 3 of soil at the Brown farm and the AREC respectively The trend lines of the nematode populat ion density over the sampling period are shown in Figure 3 1. At the AREC 2014 harvest, the average popu lation of J2 had increased up to 7,000/200 cm 3 of soil, whereas at the Brown Farm the number of J2 detected at harvest was only 289/200 cm 3 of soil. Similarity at the second harvest 2015 the population density of M. arenaria occurred much greater in the N orfolk loamy sand soil (AREC) than that in the Candler sandy soil (Brown F arm). However, at both sites the numbers of J2 recovered

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52 in the second season was much lower than that which occurred during the 2014 crop season. At the Brown Farm 2014 population densit ies of M. arenaria at the four depths increased slowly during the first 86 days until mid June after which there was a rapid increase in their numbers, peaking in mid August. This was followed with a small decrease in numbers immediately before harvest. The population numbers continued to decrease reaching a low in mid February through April 2015. There was only a small increase in numbers in late May at time the clover was turned under in preparation for planting peanut. Aft er peanut w as planted in 2015, the population increased and reached the highest number at harvest. Similar to the Brown Farm, population densit ies of M. arenaria at the three depths increased slowly during the first 77 days until late August after which there was a rapi d increase in their numbers, peaking at harvest at the end of October (Figure 3 2). This was followed with a rapid decrea se in numbers immediately after harvest. The population numbers continued to decrease reaching a low in mid February through April 2015 There was only a small increase in numbers following the planting of clover in early April. After peanut s were planted in 2015, the population increased and reached the highest number at harvest. Vertical D istribution of Meloid o gyne arenaria R e covered fr om Georgia 06G R hizosphere M ultifactor ial ANOVA performed on nematode data showed significant effect s of DAP, depth and interactions among these factors at both locations ( P (Table 3 1 2).

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53 At the Brown Farm, the highest numbers throughout the two seasons at most sampling dates were recorded from the 31 60 cm depth, and the lowest numbers were recorded from the 91 120 cm depth ( P 3). There was only one difference in the numbers recorded at the deepest depths among the 15 samplin g dates and that was the highest number recorded in September 2014 compared wit h the lowest number recorded July 2015 ( P 3). Between January and July 2015 there were no differences in numbers recorded among the four sampling depths ( P 0. 05) The highest numbers recorded were similar in the upper two depths between mid July and early October 2014 ( P 3). However, the increase in the soil population in the upper two depths during 2015 season, between mid August and mid Octob er, was much less than that occurred in 2014 ( P 3). The vertical distribution of J2 in the Georgia 06G rhizosphere was quite different at the AREC from that at the Brown farm. The number of J2 fluctuated among the three soil layers over th e entire sampling period (Table 3 4 ). However, the highest numbers of J2 w ere found at the depth of 0 15 cm at the first harvest whereas greatest number of J2 w ere observed at the depth of 16 30 cm at the second harvest. There was no difference of J2 numb er s occur ring from the end of July to late August among the three sampling depths ( P 4). Also from December 2014 to February 2015 and between May and September 2015 there were no differences in numbers recorded among the three sampling depths ( P 4). The highest numbers recorded were similar in the upper two depths in September 2014 and 2015 ( P 4).

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54 Seasonal and Vertical D istribu tion of Meloid o gyne arenaria R e covered from Tifguard R hizosphere M ultifactor ANOVA performed on nematode data showed significant effect s of DAP, depth and interactions among these factors at both locations ( P (Table 3 5 6). The number of nemato des extracted from Tifguard roots at harvest was 1/200 cm 3 of soil at the Brown Farm and 28/200 cm 3 of soil at the AREC. The highest number of nematode s was found at the soil depth of 0 30 cm over the whole season at the Brown Farm and at the soil depth of 31 45 cm at the AREC except at harvest (Figure 3 7 8). The resistant peanut cultivar indeed reduced the population density in the soil. Distribution of Nematodes A s sociated with Soil T emperature Based on statistica l analysis, soil temperature had no significant effect on nematode distribution. Most J2 w ere found within the soil temperature range s of 28 30 at the Brown Farm and within 22 24 at the AREC. E ffects of Soil Texture on Nematode P opulation The soil at the Brown farm was classified as a Candler sand and was composed of 97.5% sand, 2% silt and 0.5% clay, whereas the soil at the AREC was a Norfolk loamy sand comprised of 84% sand, 5.5% silt and 10. 5% clay (Table 3 9). Discussion Occurrence of Meloidogyne arenaria in the S oil As an obligate parasite, the population density of M. arenaria is strongly influenced by the availability of a suitable host ( Barker, 1976 ). The second stage juvenile is the infective stage that penetrates roots in the presence of a host. About 16 33 days are required for M. arenaria to complete one life cycle on a host, which usually

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55 depend s on host species and environmental conditions ( Machmer, 1977 ). Th e po pulation density of M. arenaria at both field sites was low following the winter season, preceding the spring season for pl anting peanut. S ubsequently there was a slow but progressive increase during the first 75 to 90 days following the planting of peanut A s the crop grows nematode infection increases and appears to reach the highest population densities during the remaining 45 days until harvest ( Dickson and Hewlett, 1988 ). Most peanut cultivars mature around 135 after planting. The increase of nematode number s on peanut is relatively slow compared to that which occurs on tomato ( see chapter 4 ). Root penetration on tomato us ually occurs in 2 days for M arenaria and galls are detectable after 10 15 days. In contrast galls in field planting os peanut can hardly be observed on until ca. 70 days after p lanting. The highest population density of M. arenaria in the soil was observed at harvest at the AREC whereas the highest population of M. arenaria was detected 105 days after planting at the Brown Farm. A t time of harvest at the Brown F arm in 2014 the plants were in a state of decline from what was believed to be caused by root knot disease however at the AREC site the gr owth of peanut was much better relatively to what occurred at the Brown Farm Vine growth of the susceptible Georgia 06G was ca. 38 to 45 cm tall at AREC, whereas at Brown Farm peanut height was less than 20 cm tall. The disparity in growth of the heavily infected peanuts may have been because of differences in soil type that result ed in much better peanut growth and development at the AREC. At the AREC site peanut roots grew down into the underpinning of clay, which provided harbor from the nematode and provided moisture and nutrients. In contrast, peanut roots at the Brown Farm wer e vulnerable to infection regardless of soil

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56 depth in th at J2 and root knot nematode eggs may be widely distributed through the soil profile. In the 2015 season the growth of peanut at the Brown Farm was very poor however peanut growth at the AREC was sim ilar to that that occurred in 2014 Seasonal and Vertical D istribution of Meloidogyne arenaria The seasonal distribution of J2 at both field sites followed similar trends, with a peak occurring during late summer a nd early fall around harvest Numbers of J2 d ecreased following harvest and this decrease continue during the winter season. There was only a small increase following the winter cover crop of clover. This period was followed by a significant and sharp rise in J2 numbers in September and October. Alt hough the population density of M. arenaria is affected by soil t emperature which impact s hatching and development, the host plant susceptibility is another factor that strongly i nfluence s the dynamics ( Blair, 1989 ). At the Brown Farm, during the crop season there was the largest increase of J2 occurring at the soil depth s of 0 to 30 and 31 60 cm which suggest that the nematode follows the roots as they move down into the soil p rofile, perhaps seeking a more favorable temperature for development and survival. There was also an increase of J2 at all soil depths sampled beginning in mid July and continuing up to harvest. Starr and Jeger (1985) and Sohlenius and Sandor (1987) both concluded that root abundance or root biomass largely determine the vertical distribution of Meloidogyne spp It appears that as roots grow deeper in soil the J2 follow, increasing in numbers as the season progress es T he large decrease in numbers at all depths that beg a n in January 2015 and continue until late May was not expected. However, the seeding of crimson clover in November most likely affected the number of nematodes detected in that viable J2 would penetrate the clover roots. Surprisingly, t he increase on clover was only slight

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57 over the 5 month period of growth Clover roots that were observed for galls in early April showed only minimum galling. T he highest population density of J2 at the AREC was obtained in the top 15 cm depth but numbers were only slightly less in the deeper depths of 16 30 and 31 45 cm deep. The numbers reached the highest numbers at harvest followed by a gradual decrease reaching a very low number at al l soil depths in mid February. Again, this drop was likely associate d with the planti ng of clover in November 2014. There was an increase in J2 recovered from samples taking in early April and this decline continued until early September when they spiked at all depths. The I nfluence of Soil Temperature on Nematode P opulation The soil temperature is dependent on geographic locations and seasonal fluctuations. Different Meloidogyne sp p. have different survival temperature ranges Bannon and Santo, 1984 ). Franklin (19 37) revealed that Meloidogyne spp. could survive 16 months ou t of doors in the absence of a host and could withstand some freezing. Kincaid (1946) found that root knot nematodes found in the southern states of the U.S.A. failed to survive the winter in Ind iana and Wisconsin. E xistence of physiological races responding differently to low temp er atures could be an explanation for these contradictory findings. Generally, the higher soil temperature has positive effects on nematode reproduction within the limita tion of high temperature extremes. Nevertheless, in this study, temperatures at different soil depths did not appear to impact nematode numbers. Effect of Soil Texture on Nematode P opulation Soil texture is considered as one of the important factors for nematode survival and movement. Sandy soil is prefe r red for vital act ivities of root knot nematode (Prot

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58 and Van Gundy, 1981 ). The soil at the Brown Farm had more than 97% sand wh ereas at AREC sand content was only 85%. However, much higher numbers of nematodes were observed at the AREC than at the Brown farm. Although contradictory these results might be explained by the heavy soil clay layer underpinning that occurred only 45 cm deep. As has been pointed out the i nitial population is usually correlated with the final population and root damage (McSorley, 1998). Garcia (2012) also suggested a strong nonlinear relationship existed between Pi and Pf as well as percentage of yield loss by studying two soybean cultivars Although root mass was not determined at either the Brown or AREC sites it was apparent that there was greater amount of roots occurring in the upper 45 cm of soil at the AREC. This would provide for a greater carrying capacity of J2 than would occur in the deeper and sandy soil at the Brown Farm. Other possible reasons for low number of nematodes extracted from Brown F arm include potential biological agents and the poor development of peanut in 2015.

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59 Figure 3 1. Season al fluctuation of Meloidogyne arenaria second stage juveniles in the soil profile of Georgia 06G at four depths in a Candler sand at the Brown Farm, Levy County, FL. 0 50 100 150 200 250 300 350 400 450 500 The number of Meloidogyne arenaria J2 extacted from 200 cm 3 of soil 0-30 cm 31-60 cm 61-90 cm 91-120 cm Sampling dates

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60 Figure 3 2. Seasonal fluctuation of Meloidogyne arenaria second stage juveniles in the soil profile of Georgia 06G at three depths in a Norfolk loamy sand soil at University of Georgia, Attapulgus Research and Education Center, Attapulgus, GA. 0 1,000 2,000 3,000 4,000 5,000 6,000 7,000 8,000 9,000 10,000 11,000 12,000 The number of Meloidogyne arenaria J2 extacted from 200 cm 3 of soil 0-15 cm 16-30 cm 31-45 cm

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61 Table 3 1 Significance of main and interactive effec ts of both fixed and random variables for the vertical distribution of Meloidogyne arenaria extracted from 200 cm 3 of soil collected from plots planted Georgia 06G at the Brown Farm (2014 2015). Effects Pr(>F) DAP 3.766e 15 *** Depth < 2.2e 16 *** DAPDepth 9.466e 11 *** Table 3 2. Significance of main and interactive effects of both fixed and random variables for the vertical distribution of Meloidogyne arenaria extracted from 200 cm 3 of soil collected from plots planted Georgia 06G at the Attapulgus Research and Education Center (2014 2015) Effects Pr(>F) DAP 9.893e 14 *** Depth 1.174e 08 *** DAPDepth 1.471e 10 *** Table 3 3. Number of Meloidogyne arenaria second stage juveniles extracted from 200 cm 3 of soil collected at four depths 0 30, 31 60, 61 90, and 91 120 cm deep from plots planted Georgia 06G at the Brown Farm (2014 2015). Date Depth 1 0 30 cm 31 60 cm 61 90 cm 91 120 cm 10 Apr 2014 13 Aa 39 Aabc 26 Aabc 8 Aab 19 May 2014 33 ABa 52 Babc 23 ABab 6 Aab 16 Jun 2014 37 Ba 28 ABab 12 ABab 3 Aab 17 Jul 2014 191 Ccd 90 Bbc 52 ABabcd 17 Aab 17 Aug 2014 306 Cd 436 Cd 165 Bd 47 Aab 08 Sep 2014 295 Cd 392 Cd 127 Bcd 58 Ab 05 Dec 2014 214 Bcd 162 Bc 58 Abcd 20 Aab 09 Jan 2015 16 Aa 27 Aab 19 Aab 8 Aab 20 Feb 2015 3 Aa 5 Aa 5 Aab 2 Aab 01 April 2015 2 Aa 2 Aa 2 Aab 2 Aab 28 May 2015 26 Aa 17 Aab 8 Aab 5 Aab 14 Jul 2015 4 Aa 2 Aa 2 Aa 1 Aa 14 Aug 2015 53 Bab 72 Babc 37 ABabc 11 Aab 14 Sep 2015 156 Cbcd 80 BCbc 35 ABabc 14 Aab 10 Oct 2015 59 Babc 36 ABab 17 ABab 9 Aab 1 Data are means of five replicates. Means within a row followed by a n upper case letter are not different according to F tests ( P different according to F tests ( P

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62 Table 3 4. Number of Meloidogyne arenaria second stage juveniles extracted from 200 cm 3 of soil collected at four depths 0 15, 16 30, and 31 45 cm deep from plots planted Georgia 06G at the Attapulgus Research and Education Center (2014 2015) Date Depth 1 0 15 cm 16 30 cm 31 45 cm 30 Jul 2014 151 Aa 185 Aab 185 Aab 22 Aug 2014 1,260 Aabc 1,049 Aabc 1,089 Aabc 22 Sep 2014 8,615 Bde 6,056 Bd 2,555 Abcd 30 Oct 2014 9,865 Be 6,488 Ad 5,136 Ad 11 Dec 2014 2,430 Abc 2,274 Acd 3,231 Acd 13 Feb 2015 174 Aa 213 Aabc 408 Aabc 02 Apr 2015 699 ABab 1,667 Babc 605 Aabc 19 May 2015 93 Aa 149 Aab 227Aab 03 Jul 2015 46 Aa 25 Aa 45 Aa 02 Aug 2015 29 Aa 67 Aab 17 Aa 01 Sep 2015 4,141 Bbc 2,371 Bbc 1,173 Aabc 04 Oct 2015 3,759 Bcd 6,265 Cd 1,753 Abcd 1 Data are means of six replicates. Means within a row followed by a n upper case letter are not different according to F tests ( P different according to F tests ( P

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63 Figure 3 3. Seasonal fluctuation of Meloidogyne arenaria second stage juveniles in the soil profile of Tifguard at four depths in a Candler sand at the Brown Farm, Levy County, FL and at three depths in a Norfolk loamy sand soil at University of Georgia, A ttapulgus Research and Education Center, Attapulgus, GA. 0 10 20 30 40 50 60 70 The number of Meloidogyne arenaria extracted from 200 cm 3 of soil 0-15 cm 16-30 cm 31-45 cm 0 5 10 15 20 The number of Meloidogyne arenaria extracted from 200 cm 3 of soil 0-30 cm 31-60 cm 61-90 cm 91-120 cm Brown Farm Attapulgus Research and Education Center

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64 Table 3 5. Significance of main and interactive effects of both fixed and random variables for the vertical distribution of Meloidogyne arenaria extracted from 200 cm 3 of soil collected from plots planted with Tifguard at the Brown Farm 2015 Effects Pr ( > F) DAP 4.488e 05 *** Depth < 2.2e 16 *** DAPDepth 0.002946 ** Table 3 6. Significance of main and interactive effects of both fixed and random variables for the vertical distribution of Meloidogyne arenaria extracted from 200 cm 3 of soil collected from plots planted with Tifguard at the Attapulgus Research and Educat ion Center 2015 Effects Pr ( > F) DAP 0.004385 ** Depth 0.671477 DAPDepth 0.035162 Table 3 7. Number of Meloidogyne arenaria second stage juveniles extracted from 200 cm 3 of soil collected at four depths 0 30, 31 60, 61 90, and 91 120 cm from plots planted with Tifguard at the Brown Farm 2015 Date Depth 1 0 30 cm 31 60 cm 61 90 cm 91 120 cm 01 April 2015 2 Aa 2 Aab 2 Aabc 2 Aab 28 May 2015 16 Bb 14 Bc 6 Ac 5 Ab 14 Jul 2015 3 Ca 1 Bca 1 Aba 0 Aa 14 Aug 2015 5 Ca 2 Ba 1 Bab 0 Aa 14 Sep 2015 16 Db 10 Cbc 5 Bbc 2 Aab 10 Oct 2015 1 Ba 1 Ba 1 Aba 0 Aa 1 Data are means of five replicates. Means within a row followed by a n upper case letter are not different according to F tests ( P different according to F tests ( P

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65 Table 3 8. Nu mber of Meloidogyne arenaria second stage juveniles extracted from 200 cm 3 of soil collected at four depths 0 15, 16 30, and 31 45 cm from plots planted with Tifguard at the Attapulgus Research and Education Center 2015 Date Depth 1 0 15 cm 16 30 cm 31 45 cm 19 May 2015 11 Aab 7 Aa 14 Aa 0 3 Jul 2015 21 Aab 30 Aa 60 Aa 0 2 Aug 2015 3 Aa 12 Aa 24 Aa 0 1 Sep 2015 25 Aab 27 Aa 33 Aa 0 4 Oct 2015 46 Bb 30 Aba 10 Aa 1 Data are means of six replicates. Means within a row followed by a n upper case letter are not different according to F tests ( P different according to F tests ( P before analysis.

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66 Table 3 9. Sand, silt, and clay distribution at different so il depths collected from Brown F arm, Levy County, FL and Uni versity of Georgia, Attapulgus R esearch and Education Center, Attapulgus, GA. Sites Depth (cm) Sand (%) Clay (%) Silt (%) Brown farm Candler sand 0 30 96 3 1 31 60 96.8 2.5 0.7 61 90 98 1.5 0.5 91 120 99 0.8 0.2 Attapulgus Research and Education Center Norfolk loamy sand 0 15 88.7 4.3 7 16 30 84 5 11 31 45 79 7 14

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67 CHAPTER 4 THE INFLUENCE OF TEMPERATURE ON THE SUSCEPTIBILITY OF CVS. TIFGUARD AND GEORGIA 06G PEANUT TO M eloidogyne arenaria Introduction Meloidogyne arenaria the peanut root knot nematode, is considered one of the most important soilborne pathogens affecting peanut in the southern USA. This species is prevalent in Alabama, Florida, Georgia, and Texas (Dickson, 2005; Ingram and Rodriguez Kabana, 1980; Starr and Morgan, 2002). The suppression of peanut yie ld s caused by this nematode can reach up to 50% or greater in heavily infested fields (Proite et al., 2008). Resistant cultivars potentially provide the most economic al and effective means of managing nematode disease s on agricultural crop s Up until 2001 there were no cultivated peanut with root knot nematode resistance, yet a number of Arachis spp. were observed to be highly resistant to the peanut root knot nematode (Baltensperger et al., 1986; Holbrook and Noe, 1990; Nelson et al., 1989). G enetic mater ial from a wild type peanut, A. cardenasii was introgre ssed into cultivated peanut resulting in the first root kn ot nematode resistant cultivar COAN (Simpson and Starr, 2001). The second peanut cultivar NemaTam resistant to M. arenaria was released in 200 2. This cultivar had the same resistant level as COAN, but had greater yield potential ( Simpson et al. 2003 ). The resistance gene in COAN was reported to have three different functions on M. arenaria (Bendezu and Starr, 2003). The first effect was to redu ce the number of the second stage juveniles (J2) of M. arenaria that penetrated roots; second, the resistance gene prevented most J2 from developing further, and third, the resistance gene delayed development of J2. No hypersensitive reaction was observed in the early stage of M. arenaria infection in COAN roots, which is considered as a common defense

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68 mechanism produced by resistant plants to disease pathogens (Bendezu and Starr, 2003). Neither of these two cultivars was considered for planting in Florid a because of the prevalence and severity of tomato spotted wilt virus (TSWV) and fungal diseases in the southeastern peanut growing reg ion ( Holbrook et al., 2008 ). In 2008, a runner type peanut cultivar ( Arachis hypogaea L. subsp. hypogaea var. hypogaea) Tifguard was released by the USDA ARS and the Georgia Agricultural Experiment Station (Holbrook and Shokes, 2002), a peanut cultivar with good field resistance to TSWV (Wells et al., TSWV and M. arenaria (Holbrook et al., 2008). Temperature is an important factor influencing the expression of resistance gen es in plants to parasitic nematodes. For example, it was observed that susceptible alfalfa plants attracted more juveniles of M. hapla than resistant plants, but the magnitude of this difference decreased when temperature increased (Griffin et al., 1971; 1 977). In addition, half of the resistant alfalfa plants grown at 32 had galled root s whereas no galled roots were observed at 28 Similarly, the number of mature M. incognita and M hapla in resistant and susceptible bean increased as the soil temper ature increased from 25 to 30 (Irizarry et al., 1971). Increased temperature also affected nematode resistance in soybean and tomato cultivars (Dropkin, 19 69 b ; Holtzmann, 1965). Greater penetration and development of M. incognita was observed in resistan t tomato roots at 30 and 34.5 than at 20 and 25 In other studies, on tomato, decreased hypersensitive necrotic response to nematode infection was

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69 reported at temperatures near or above 30 (Paulson and Webster, 1972). The Mi 1 gene in tomato confers resistance to root knot nematodes; however, this gene becomes nonfunctional when the temperature is at or above 28 ( Williamson, 1998 ). Changes in the response of root knot nematode resistance by high temperature has been reported under both greenhouse and field conditions (Philis and Vakis 1977; Tzortzakakis and Gowen 1996; Noling 2000). It appears that heat stress affects the functioning of resistant genes in host plants (A mmiraju et al., 2003). On the other hand, it is well known that root knot nematodes are dependent on temperature for vital physiological activities. Temperature affects root knot nematode migration, development, and the ts (Tyler, 1933). The optimal temperature for root knot nematode development was reported as 28 above which development was reduced. No development occurred at 36.5 (Tyler, 1933). The number of the second stage juveniles (J2) of M. arenaria in differ ent peanut genotypes at 2 and 7 days after inoculation (DAI) was compared by Bendezu and Starr (2003). Higher number of J2 penetrated the roots of the susceptible peanut Florunner than the resistant COAN. Since the release of the root knot nematode resista nt peanut cultivar there has been no study about how temperature affects the number and development of M. arenaria in peanut with the resistant gene. The objective of this study was (i) to determine the number of M. arenaria that penetrated roots of resist ant and susceptible peanut cultivars; and ( ii) to determine the effect of temperature on M. arenaria development in resistant and susceptible peanut cultivars.

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70 Materials and Methods Nematode O rigin Meloidogyne arenaria was isolated from infected peanut roots from an infested farm located in Levy County, FL and maintained on the susceptible tomato cultivar ( Solanum esculentum Mill. cv. Agriset 334) in the greenhouse. Polyacrylamide gel electrophoresis was used as an aid in identification of the species b ased on malate dehydrogenase and esterase phenotype s (Esbenshade and Triantaphyllou, 1985 ). Eggs were extracted from in fected tomato roots with 0.25% NaC IO (Hussey and Barker, 1973) method as modified by Boneti and Ferraz (1981). A modified Baermann funne l method was used for egg hatching ( Rodriguez and Pope, 1981 ). Second stage juveniles (J2) collected after a period of 48 hours were used as inoculum. Penetration and D evelopment of Meloidogyne arenaria P eanut seed of Tifguard and Georgia 06G were provided by the peanut breeder Corley Holbrook, USDA ARS, Tifton, GA T omato cv. Agriset 334 was used as a susceptible plant to ensure inoculum viability. Peanut seeds were surface treated by soaking in 0.6% Na Cl O for 1 min followed by rinsing in sterile distilled water (Bendezu and Starr, 2003). They were placed in a petri dish with a piece of moistened paper towel for germination at 28 for 5 days. Seedlings were transplanted into 250 ml plastic cups filled with autoclaved sand and grown at 28 for 7 days. H ole s were punched in to the cup bottom s to provide drainage. All seedlings were inoculated with 2,000 freshly hatched J2 of M. arenaria and then all seedlings were transferred to an environmental growth chambers at either 28, 31, or 34 with a 12 h photoperiod. The seedlings were hand watered daily.

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7 1 Data C ollection Three plants were removed at 5 day intervals following inoculation (DAI), washed in tap water, drie d with a paper towel, and weigh ed. Roots were fixed, stained and cleared for microscopic examination ( Byrd et al., 1983 ). The number and images of nematodes in the roots were recorded based on their developmental stages as follows: (i) second stage juvenile (J2); (ii) J2 with swollen body (late J2); (iii) the third and fourth stag e juvenile; ( iv) female; (vi) egg laying female; and (vii) male R ed food coloring (20%) was used to stain egg masses once they were observed on roots ( Thies et al., 2002 ). The experiment was repeated twice In the experiment 1, harvest ceased at 30 DAI, w hereas in the experiment 2, harvest was extended to 40 DAI in order to determine if egg masses of M. arenaria would form on Tifguard roots. Root knot nematode egg massed were collected (three egg masses per repetition per cultivar ) from both peanut cultivars once they were observed at 40 DAI. Statistical A nalysis The effects of temperature, host genotype, and DAI on the number of different developmental stages of M. arenaria per gram of root tissue were subjected to analysis of var iance using R programming software ( R 3.1.2 with R studio ). The means were was not included in the statistical analyses. Values were transformed to before analys is. The number of eggs per egg mass was subjected to analysis by Resistance Gene Marker A nalysis of Tifguard In the first experiment a young leaflet from any Tifguard plant with root knot nematode infection was collected and shipped to the National Environmentally Sound

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72 Production Agriculture Laboratory (NESPAL), Tifton, GA to confirm whether the infected plant contained the root knot nematode resistant gene. Any plant without the resistance gene was not included to the statistical analy ses. In the experiment 2 all the Tifguard seed used had a small portion of seed coat removed for testing for the resistance gene Rma by the NESPAL laboratory ( Nagy et al., 2009 ). Only seed that were positive for the resistant gene were used in experiment 2 Results Penetration of Meloidogyne arenaria in the Resistant and Susceptible Peanut C ultivars The root systems from Georgia 06G were generally 1 to 1.5 fold greater in size than that from Tifguard ( P (Table 4 1). Because of this, the number of ea ch developmental stage of M. arenaria per gram of roots was determined in the two peanut cultivars. In both experiments tomato was heavily infected assuring inoculum viability (Figure 4 1 6) The development of J2 was consistently faster in tomato than that which occurred in peanut roots regardless of cultivars Galls and egg masses were readily apparent on tomato roots at 15 DAI, whereas gall s and egg masses first became apparent on Georgia 06G roots at 20 DAI (Figure 4 1 6) Both peanut genotype s and temperature affected the number of nematode s in roots ( P (Table 4 1) In both experiment s, relative to the number of J2 found in roots at 5 DAI, the numbers in Tifguard and Georgia 06G decreased over the 30 to 40 period at all three temperatur es (Figure 4 7 ; 4 8 ). There was an increase in numbers of J2 infecting roots in both experiment s in response to temperatures above 28 In experiment one, a t day 5 the initial penetration of M. arenaria in Georgia 06G was greater at 31 than at 28 or 34 ( P experiment two the re was

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73 an increase in the number of M. arenaria found in Georgia 06G roots between 28 and 31 or 34 but there was no difference in numbers between 31 and 34 ( P (Figure 4 9) The number of nematode s in Tifguard was greater at 31 but no differences occurred between 31 and 34 in either experiment In experiment 1, there were higher numbers of M. arenaria J2 in resistant Tifguard roots at 34 than in susceptible Georgia 06G, whereas a higher number of nematode s was observed in susceptible Georgia 06G roots at 28 ( P in nematode number found in the susceptible and resistant peanut roots at 31 There was a difference of penetration between susceptible and resi stant cultivars at 28 and 34 ( P Development of Meloidogyne arenaria in the Resistant and Susceptible Peanut C ultivars In both experiment s females and males of M. arenaria were observed in Tifgu ard roots only at 34 (Figure 4 11 ). There was no development of M. arenaria in Tifguard roots at 28 or 31 ( Figure 4 1 2; 4 4 5) Although females were found in Tifguard at 34 no egg laying females were observed in experiment one over the 30 day period (Figure 4 3) whereas on average of three females with small egg masses were found in experiment two at 40 DAI (Figure 4 6; Figure 4 11 ). In the susceptible peanut cultivar, e gg laying females were first observed at 25 DAI at 28 and 20 days after inoculation at 31 in both experiment s (Figure 4 1 2; 4 4 5) However, there was a delay in the development of nematode at 34 compared with that at 31 in both experiments (Figure 4 2 3; 4 5 6) L owest number of egg laying females was found i n Geoorgia 06G roots 30 DAI at 34 whereas the highest number was found at 31 The numbers were similar to that at 28 in the first

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74 experiment (Figure 4 10) In experiment two h ighest n umber of egg laying females was observed at 31 at 40 DAI ( P ) (Figure 4 10) Necrotic lesions were formed in Tifguard roots in response to M. arenaria infection at all sampling dates ( Figure 4 12 ) Cell necrosis was found in the tissue surrounding the swollen J2, which apparently stop ped furth er development of the nematode Reproduction of Meloido gyne arenaria in Different Peanut G enotypes Numbers of e gg s per egg mass from Tifguard roots were different from those from Georgia 06G at 40 DAI ( P Th e average number of eggs per egg mass isolated from Tifguard was 13 whereas from Georgia 06G it was 244 Discussion Effect of P eanut G enotype s and T emperature s on Meloidogyne arenaria There was a strong effect of peanut genotype on the response to M. arenaria infection. Tifguard was reported to be highly resistant to M. arenaria infection (Holbrook et al., 2008 ) Alt hough nematode development occurre d readily in Georgia 06G there was only a small rate of development at 34 in Tifguard In Georgia 06 the first egg laying female s were observed at 2 0 or 25 DAI whereas J2 were not able to develop after penetration of Tifguard (except at 34 ). Temperature is considered to be an important factor on egg hatching, nematode migration, root invasion and development in host roots ( Tyler, 1933 ) At 31 more J2 entered Tifguard and G eorgia 06G roots compared with that at 28 and 34 In addition, final number s of M. arenaria infecting Georgia 06G also was influenced by higher temperature. In this study, t he temperature of 31 was the optimal for J2 to penetrate and parasit ize the susceptible peanut. This result was

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75 report where 28 was reported as the optimal temperature for nematode penetration and infection. Moreover, only 6 out of 26 root system s were reported to be infected by M. arenaria at 33 to 35 (Tyler, 1933) wh ereas all Georgia 06G and tomato plants were galled and had egg masses at 34 in this study. The discrepancy may indicate that root knot nematode s in Florida have adapted to high soil temperature s The F unction of Resistance G ene Based on the data from this studies, the number of egg laying females and eggs per egg mass were much less in the resistant cultivar Tifguard t han that in the susceptible cultivar Georgia 06G This confirms results reported by G arcia et al., 1996 about the funct ions of Mae and Mag genes found in the wild type Arachis spp which were resistant to M arenaria These two genes were discovered to be linked to Rma which is the dominant root knot nematode resistant gene in cultivated peanut (Nagy et al., 2010) Mae was reported to suppress egg production of M. arenaria whereas Mag is reporte d to inhibit formation of galls Hypersensitive R eaction The presence of host necrosis near the site where J2 were observed in the early stages of the host parasite interaction was contradictory to the observation by Choi et al (1999), and Bendezu and Starr (2003). This hypersensitive reaction (HR) mechanism has been well documented in the case of the Mi resistance gene in tomato to root knot nematode infection ( Paulson and W ebster 1 972 ; Williamson, 1999 ). After recognition of the nematode by the plant bearing a single r esistance gene, host defenses respond and are activated, leading to necrotized cell s surrounding the nematode. A resistant plant provide s the most effective and environmentally safe means to manage root knot nematodes, thus understanding the mechanisms involved in

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76 Meloidogyne virulence is critical for the sustainable management of these pathogens. In summary, this research demonstrates that soil temperature is not likely to be a factor in preventing the resistant gene in Tifguard from functioning to protect this peanut from root knot disease induced by M. arenaria.

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77 Table 4 1. Analysis of variance of root weight, number of Meloidogyne arenaria per gram root system of two peanut cultivars Experiment Experiment

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78 Figure 4 1 The effect of temperature on the number of different developmental stages of Meloidogyne arenaria in roots of two peanut cultivars and tomato recorded at 5 day intervals over a 30 day period at 28 (Experiment 1). 0 25 50 75 100 125 150 175 200 5d 10d 15d 20d 25d 30d 5d 10d 15d 20d 25d 30d 5d 10d 15d 20d 25d 30d Tifguard Georgia-06G Tomato Mean number of nematodes from each developmental stage per root system Male Egg-laying female Female J3/J4 Late-J2 J2

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79 Figure 4 2 The effect of temperature on the number of different developmental stages of Meloidogyne arenaria in roots of two peanut cultivars and tomato recorded at 5 day intervals over a 30 day period at 31 (Experiment 1). 0 25 50 75 100 125 150 175 200 5d 10d 15d 20d 25d 30d 5d 10d 15d 20d 25d 30d 5d 10d 15d 20d 25d 30d Tifguard Georgia-06G Tomato Mean number of nematodes from each developmental stage per root system Male Egg-laying female Female J3/J4 Late-J2 J2

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80 Figure 4 3 The effec t of temperature on the number of different developmental stages of Meloidogyne arenaria in roots of two peanut cultivars and tomato recorded at 5 day intervals over a 30 day period at 34 (Experiment 1). 0 25 50 75 100 125 150 175 200 5d 10d 15d 20d 25d 30d 5d 10d 15d 20d 25d 30d 5d 10d 15d 20d 25d 30d Tifguard Georgia-06G Tomato Mean number of nematodes from each developmental stage per root system Male Egg-laying female Female J3/J4 Late-J2 J2

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81 Figure 4 4 The effect of temperature on the number of different developmental stages of Meloidogyne arenaria in roots of two peanut cultivars and tomato recorded at 5 day intervals over a 40 day period at 28 (Experiment 2). 0 50 100 150 200 250 300 350 400 450 500 5d 10d 15d 20d 25d 30d 35d 40d 5d 10d 15d 20d 25d 30d 35d 40d 5d 10d 15d 20d 25d 30d 35d 40d Tifguard Georgia-06G Tomato Mean number of nematodes from each developmental stage per root system Male Egg-laying female Female J3/J4 Late-J2 J2

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82 Figure 4 5 The effect of temperatur e on the number of different developmental stages of Meloidogyne arenaria in roots of two peanut cultivars and tomato recorded at 5 day intervals over a 40 day period at 31 (Experiment 2). 0 50 100 150 200 250 300 350 400 450 500 5d 10d 15d 20d 25d 30d 35d 40d 5d 10d 15d 20d 25d 30d 35d 40d 5d 10d 15d 20d 25d 30d 35d 40d Tifguard Georgia-06G Tomato Mean number of nematodes from each developemental stage per root system Male Egg-laying female Female J3/J4 Late-J2 J2

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83 Figure 4 6 The effect of temperature on the number of different developmental stages of Meloidogyne arenaria in roots of two peanut cultivars and tomato recorded at 5 day intervals over a 40 day period at 34 (Experiment 2). 0 50 100 150 200 250 300 350 400 450 500 5d 10d 15d 20d 25d 30d 35d 40d 5d 10d 15d 20d 25d 30d 35d 40d 5d 10d 15d 20d 25d 30d 35d 40d Tifguard Georgia-06G Tomato Mean number of nematode from each developemental stages per root system Male Egg-laying female Female J3/J4 Late-J2 J2

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84 Figure 4 7 The mean number of Meloidogyne arenaria of all development stages in roots of Tifguard and Georgia 06G recorded at 5 day intervals over a 30 day period in experiment 1. 0 20 40 60 80 100 120 5D 10D 15D 20D 25D 30D Tifguard Mean number of nematodes per gram of roots 28 31 34 0 20 40 60 80 100 120 5D 10D 15D 20D 25D 30D Georgia-06G Mean number of nematodes per gram of root

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85 Figure 4 8 The mean number of Meloidogyne arenaria of all development stages in roots of Tifguard and Georgia 06G recorded at 5 day intervals over a 40 day period in experiment 2. 0 20 40 60 80 100 120 140 160 180 5D 10D 15D 20D 25D 30D 35D 40D Tifguard Mean number of nematode per gram of roots 28 31 34 0 20 40 60 80 100 120 140 160 180 200 220 240 260 5D 10D 15D 20D 25D 30D 25D 40D Georgia-06G Mean number of nematode per gram of roots

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86 Figure 4 9 The number of second stage juveniles (J2) per gram of root system of the resistant cultivar Tifguard and susceptible cultivar Georgia 06G 5 days after inoculation in two experiments Different letters over bars from the same experiment indicate significa nt differences at 0.05 level 0 20 40 60 80 100 120 28 31 34 5 days The number of J2 per gram of roots Tifguard Georgia-06G A B B Experiment 0 50 100 150 200 250 300 28 31 34 The number of J2 per gram of roots Tifguard Georgia-06G b a Experiment

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87 Figure 4 10. The number of egg laying females per gram of root system of the resistant cultivar Tifguard and susceptible cultivar Georgia 06G 30 or 40 days after inoculation in two experiments. Different letters over bars from the same experiment indicate significant differences at the 0.05 level 0 20 40 60 80 100 120 140 28 31 34 30 days The number of egg laying female per gram of roots Tifguard Georgia-06G B A B A A A 0 10 20 30 40 50 60 28 31 34 40 days The number of egg laying females per gram of roots Tifguard Georgia-06G Experiment a bc a c b b Experiment

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88 Figure 4 11 Different developmental stages of Meloidogyne arenaria in peanut and tomato roots at 34 Photos courtesy by author. Tifguard Georgia 06G Tomato 5 DAI 10 DAI 15 DAI 20 DAI 25 DAI 30 DAI 35 DAI 40 DAI

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89 Figure 4 12 Arrows point to necrotic lesions formed around the root knot nematode infection sites in Tifguard roots at 5 and 40 days after inoculation (DAI) Photos courtesy by author. 5 DAI 40 DAI

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90 CHAPTER 5 COMPARISON OF YIELD OF BREEDER TIFGUARD, ISOGENIC TIFGUARD AND GEORGIA 06G TREATED AND NONTREATED WITH 1,3 DICHLOROPROPENE Introduction Yield drag, an 'old idea' in plant breeding, is used in comparison of genetically engineered crop with nongenetically engineered crops ( Opling er, 1999 ). Yield drag refers to a negative effect on yield of crop plants that have a specific gene or a specific trait ( Benbrook, 1999 ). A 'drag' on yield has been reported in breeding soybeans selected for higher protein levels for decades. Though with c ontinued breeding efforts, this drag has been overcome to a certain extent by the development of high yielding and high protein lines, it remains a constant challenge (Hain and Lee, 2010). Genetically modified crops are unique because of an add itional gene being placed into one of the chromosome. Yield drag can occur if the transgene inserts in the coding region of a native gene and interrupts coding of gene products which might be associated with yield. On the other hand, genetically modified p lants are also being manipulated to produce a new protein in large quantities. The new protein could be made at the cost of the proteins that are normally produced because the pool of protein building blocks (amino acids) is limited. This could result in a shortage of other proteins and cause yield drag (Hain and Lee, 2010). Although not transgenic, Tifguard was generated by interspecific crossing and insertion of gene(s) coding for root knot nematode resistance. It is not known whether the gene insertion( s) cause yield drag in Tifguard. The yield produced by Tifguard was compared to other peanut cultivars planted in Florida from 2010 to 2013 ( Tillman et al., 2013 ). On average, the yield of Tifguard was ranked at 11th among 15 peanut cultivars.

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91 The objectiv e of this experiment is to compare yield from breeder Tifguard, isogenic Tifguard and Georgia 06G treated and nontreated with 1,3 dichloropropene (1,3 D). Materials and Methods F ield D esign and T reatments The trial was located at the University of Florida Plant Science Researc h and Education Unit, Citra, FL. An arbitrarily selected preplant soil sample from each plot was collected to determine the initial population density of nematodes. A 2.5 cm diameter cone shaped sampling tube was used to collect soil samples according to a zig zag pattern in each plot (Barker and Campbell, 1981). Six soil cores from each plot were mixed thoroughly and 200 cm 3 processed to extract second stage juveniles of M. arenaria 1, 3 D (Dow AgroSciences Inc., Indianapolis, IN) was applied at rates of 150 L/ ha 10 days preplant by a Yetter C oulter Rig The chemical was injected 20.3 cm deep usi ng a 63.5 cm diameter coulter and trailing chisel shank on the tractor mounted applicator. Ten days after chemical application, Tifguard breeder, isogenic Tifguard, and Georgia 06G peanut cultivars were planted. The planting and harvesting methods including preplant fertilizer and herbicid e application normally followed the IFAS Peanut Production G uide ( Stephone 1994 ). The field design was arranged in randomized complete block with two factors, chemical treatment and three different peanut cultivars (Tifguard breeder+1,3 D; Tifguard breeder; isogenic Tifguard+1,3 D; isogenic Tifguard; Geofgia 06G+1 ,3 D; Geofgia 06G ) The treatments were replicated four times, with each plot measuring 9.14 m long and 0.9 m row spacing. A preplant commercial fertilizer mix including minor elements (3 9 18) was applied broadcast ( 560 kg/ha ) and incorporated into soil before seeding. Gypsum was

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92 applied at 2 242 kg/ha at early peg initiation. Irrigation was applied via a pivot overhead traveling system on an as needed basis. Prowl was applied preplant to suppress early weed emergence. An over the top application of paraquat + 2, 4 D and Select were applied to suppress broad weeds and grasses, respectively. During the growing season, fungicide was applied every 14 to 20 days Peanut Harvest and Data C ollection The peanuts were dug and inverted 130 days post harvest P eanut growth were assessed at harvest based on a subjective rating scale from 1 to 10, where 1 refers the very poor growth and 10 refers the best growth in terms of height, width and general appearance of the plants ( Turner and Backman, 1991 ). Six groups of peanut roots were arbi trarily selected from each treatment at harvest to determine root knot nematode infection. The assessment index for galls and egg mass was based on the following scale: 0= no gall or egg mass, 1 = 1 2, 2 = 3 10, 3 = 11 30, 4 = 31 root system (Taylor and Sasser, 1978). After field digging for 3 days the plants were combined, the pods collected in cottage bags, and placed in peanut w agons for drying to 10 % moisture. Each sample was weighted and reco rded Statistical A nalysis The SAS proprietary statistical package was used to subject data to analysis of variance (ANOVA) and mean differences were separated by Duncan's multiple range test. Results Peanut Growth and R oot knot Nematode I nfection Georgia 06G had lower plant growth ratings than Tifgua rd or isogenic Tifguard (P ). There were no difference s in peanut growth between Georgia 06G treated and

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93 nontreated with 1,3 D nor was there an effect of 1,3 D on the peanut growth or yield of Tifguard and isogenic Tifguard (Table 5 1) A relatively small amount of galling was observed on roots from Georgia 06G and isogenic Tifguard without 1,3 D treatment. Both of them are susceptible to Meloidogyne arenaira There was relatively little infection observed on Georgia 06G with out 1,3 D whereas more than 30 galls or egg masses were found on isogenic Tifguard without 1,3 D treatment (Table 5 1) Y ield C omparison Georgia 06G had the lower yield than Tifgua However, t here was no difference of yield between Georgia 06G treated or nontreated with 1,3 The 1,3 D had no effect on yield produced by Tifgua rd treated and nontreated with 1,3 D ( P Similarity, there was no difference of yield between isogenic Tifguard treated and nontreated with 1,3 D ( P (Table 5 1) Soil Type, Texture, A nalysis The soil was a mixture of fine sandy Arredondo and span type (sand 95%, silt 3%, clay 2%; organic matter 1.5 %; pH 6.5). Moisture content of the soil at field capacity averages 13.5% to 30 cm deep and bulk density of the planting beds average 1.4 g/cm 3 Drainage was excellent with very little underlying clay. Discussion Yield is a key factor for profitability expectations and results. Yield drag is often used in comparing crop with genetic engineering to crops without genetic engineering or crop with application of herbicide or without herbicide ( Oplinger, 1999 ). Tifguard is t he first peanut cultivar with resistance to two pathogens. The yield performance of Tifguard was conducted and compared with other peanut cultivar in 2010 2013. Based

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94 on the pod yield from different peanut varieties, Tifguard is a good peanut cultivar wit h high yield compared with other peanut cultivar s (Tillman et al., 2013). In this study, however, Tifguard had higher pod yield compared with Georgia 06G. The isogenic Tifguard is also a peanut cultivar susceptible to M. arenaria The untreated isogenic Ti fguard had the highest pod yield among the six treatments albeit the highest infection was also found on it. Based on the statistical analysis, there is no significance of estimated yield among Tifguard and isogenic Tifguard with or without treatment. It s eems that yield drag was not observed on Tifguard performance. Moreover, the chemical treatment was not effective management for root knot nematode since the treated Georgia 06G and untreated Georgia 06G had the same yield performance. This is single year trial and results from any single year may not be sufficient to determine the yield performance of peanut variety. The results from one year trial are simply a reflection of the growing season that occurred in this year The multi year results are better for evaluation of the yield performance of peanut varieties

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95 Table 5 1. Effect of different treatments on plant growth, galling induced by root knot nematodes and yield of peanut in a field trial at Plant Science Research and Education Unit, Citra, FL, Spring Summer 2015. Treatment Plant growth rating 1 Gall index 2 Yield ( kg / ha ) 1. G eorgia 06G without 1,3 D 6.5 a 0.8 a 3 064 a 2. Georgia 06G with 1,3 D 6.8 a 0 a 2 957 a 3. Tifguard without 1,3 D 8.5 b 0 a 4 462 b 4. Tifguard with 1,3 D 9.4 c 0 a 4 731 b 5. Isogenic Tifguard 3 without 1,3 D 9.8 c 3 b 4 785 b 6. Isogenic Tifguard 3 with 1,3 D 9.9 c 0 a 4 408 b Data are means of four replications. Means within a column followed by a common letter are not different according to range test ( P 0.05 ) 1 Scale 1 10, where 1= very poor plant growth, 10 = very good plant growth 2 Scale 1 4, where 0 = 0 galls on roots or pegs and pods, 1 = 1 10 galls on roots and < 10 on pegs and pods, 3 = 11 100 galls on roots and 10 50 on pegs and pods, 4 = > 100 galls on roots and > 50 on pegs and pods. 3 Root knot nematode susceptible Tifguard.

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96 LIST OF REFERENCES Allen, E. K. 1981. The Leguminosae, A source book of characteristics, uses, and nodulation. Madison, WI: University of Wisconsin Press. Ammiraju J. S. S., J. C. Veremis, X. Huang P. A. Roberts, and I. Kaloshian. 2003. The heat stable root knot nematode resistance gene Mi 9 from Lycopersicon peruvianum is localized on the short arm of chromosome 6. Theoretical and Applied Genetics 106:478 484. Anonymous 2015. Acreage. National Agricultural Statistics Service (NASS), Agricultural Statistics Board, United States Department of Agriculture (USDA). Bale J. S., J. C. van Lenteren, and F. Bigler. 2008. Biological control and sustainable food production. Philosophical Transactions of the Royal Society 363:761 776. Baltensperger, D. D., G. M. Prine, and R. A. Dunn. 1986. Root knot nematode resistance in Arachis gl abrata Peanut Science 13:78 80. Barker, K. R., and C. J. Nusbaum. 1971. Diagnostic and advisory programs. Pp. 257 280 in B. M. Zuckerman, W. F. Mai, and R. A. Rhode, eds. Plant parasitic nematodes.Vol. 1, Academic Press, New York. Barker, K. R., and T. H A. Olthof. 1976. Relationship between nematode population densities and crop responses. Annual Review of Phytopathology 14:327 353. Barker, K. R. 1989. Yield relationships and population dynamics of Meloidogyne spp. on flue cured tobacco. Supplement to J ournal of Nematology 2:597 603. Barker, K. R., and C. L. Campbell. 1981. Sampling nematode population. Pp. 451 473 in B. M. Zuckerman and R. A. Rohde, eds. Plant parasitic nematodes, Vol. Academic Press, New York. Barker, K. R., G. A. Pederson, and G. L. Windham. 1998. Plant and nematode interactions. ASA, CSSA, SSA Publishers, Madison, WI. Baunacke, W. 1922. Untersuchungen zur biologie und bekampfung des rubennematoden Heterodera schachtii Schmidt. Arb. Biol. Reichsanst. Land und Forstw. 11:185 288. Blair, G., and G. Boivin. 1988. Spatial pattern and sequential sampling plan for Meloidogyne hapla in muck grown carrots. Phytopathology 78:604 607. Blair, G. 1989. A note on the development of Meloidogyne hapla on lettuce in organic soil. Phytoprotecti on 70:133 135. Been, T. H., and C. H. Schomaker. 2006. Distribution patterns and sampling. Pp. 302 326 in R. N. Perry and M. Moens, eds. Plant nematology. Oxfordshire: CABI.

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113 BIOGRAPHICAL SKETCH Weimin Yuan was born in 1985 to Mr. Youjun Yuan and Mrs. Hua Li in Jiangsu Province, P.R. China. She grew up in City of Nantong in China where she also obtained her primary and secondary education in Nantong No.1 Middle School of Jiangsu Province. In 2004, she was accepted in the B. S. plant protection program in Nanjing Forestry University. For her master degree in Nanjing Forestry University, she majored in Microbiology under the supervision of Dr. Xiaoqin W u and studied the endophytic bacteria present in Bursaphelenchus xylophilus She worked as a research assistant on project dealing with identification and function of endophytic bacteria using molecular and culture based methods. She was awarded as an outstanding student and got scholarship three times during seven years in Nanjing Forestry University. After finishing, she moved to Shanghai and work ed as a professional clerk in two private companies. Realizing the need to get more professional training and knowledge, she went to pursue a doctoral program at the University of Florida where she studied root knot nematodes under the supervision of Dr. Donald W. Dickson. Her research project was focused on evaluation of resistance to Meloidogyne arenaria in the peanut cultivar Tifguard.