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Reproduction and Identification of Root-Knot Nematodes on Perennial Ornamental Plants in Florida


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REPRODUCTION AND IDENTIFICATION OF ROOT-KNOT NEMATODES ON PERENNIAL ORNAMENTAL PLANTS IN FLORIDA By ROI LEVIN A THESIS PRESENTED TO THE GRADUATE SCHOOL OF THE UNIVERSITY OF FLORIDA IN PARTIAL FULFILLMENT OF THE REQUIREMENTS FOR THE DEGREE OF MASTER OF SCIENCE UNIVERSITY OF FLORIDA 2005

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Copyright 2005 by Roi Levin

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iii ACKNOWLEDGMENTS I would like to thank my chair, Dr. W. T. Crow, and my committee members, Dr. J. A. Brito, Dr. R. K. Schoellhorn, and Dr. A. F. Wysocki, for their guidance and support of this work. I am honored to have worked under their supervision and commend them for their efforts and contributions to their respective fields. I would also like to thank my parents. Through my childhood and adult years, they have continuously encouraged me to pursue my interests and dreams, and, under their guidance, gave me the freedom to steer opportunities, curiosities, and decisions as I saw fit. Most of all, I would like to thank my fiance, Melissa A. Weichert. Over the past few years, she has supported, encouraged, and loved me, through good times and bad. I will always remember her dedication, patience, and sacrifice while I was working on this study. I would not be the person I am today without our relationship and love.

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iv TABLE OF CONTENTS page ACKNOWLEDGMENTS.................................................................................................iii LIST OF TABLES.............................................................................................................vi LIST OF FIGURES...........................................................................................................ix ABSTRACT.....................................................................................................................xi ii CHAPTERS 1 INTRODUCTION AND LITERATURE REVIEW....................................................1 Introduction.................................................................................................................. .1 Meloidogyne spp...........................................................................................................2 Relationship with Ornamentals...................................................................................14 Objectives...................................................................................................................30 2 REPRODUCTION OF FOUR MELOIDOGYNE SPP. ON SEVERAL SPECIES OF PERENNIAL ORNAMENTAL PLANTS...........................................................31 Introduction.................................................................................................................31 Materials and Methods...............................................................................................33 Results........................................................................................................................ .40 Discussion...................................................................................................................60 3 IDENTIFICATION OF ROOT-KNOT NEMATODES............................................67 Introduction.................................................................................................................67 Objectives...................................................................................................................69 Materials and Methods...............................................................................................70 Results........................................................................................................................ .95 Discussion.................................................................................................................103 APPENDIX A PICTURES OF MELOIDOGYNE SPP. ESTERASE AND MALATE DEHYDROGENASE ISOZYME PHENOTYPES UNVEILED THROUGH POLYACRYLAMIDE GEL ELECTROPHORESIS ON PHASTSYSTEM AND MINI-PORTEIN 3 CELL APPARATUSES............................................................107

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v B COLLECTIVE RECORD OF THE HOST STATUS OF ORNAMENTAL PLANTS TO MELOIDOGYNE INCOGNITA M. JAVANICA M. ARENARIA AND M. HAPLA ........................................................................................................123 REFERENCES................................................................................................................184 BIOGRAPHICAL SKETCH...........................................................................................197

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vi LIST OF TABLES Table page 2-1 Crop and source of liners used for growth room and greenhouse experiments.......34 2-2 Crops, experimental sites, liner planting dates, inoculation dates, and study lengths for all crops in the growth room and greenhouse Meloidogyne spp. studies carried out at the University of Florida during 2003 to 2005......................38 2-3 Treatments, root-gall indexes, root weights, number of eggs per plant, number of eggs per g of roots, and dry shoot weights from the first Liriope muscari cv. Evergreen Giant growth room trial..........................................................................41 2-4 Treatments, root-gall indexes, root weights, number of eggs per plant, number of eggs per g of roots, and dry shoot weights from the second Liriope muscari cv. Evergreen Giant growth room trial..........................................................................42 2-5 Treatments, root-gall indexes, root weights, number of eggs per plant, number of eggs per g of roots, and dry shoot weights from the Liriope muscari cv. Evergreen Giant greenhouse experiment.................................................................44 2-6 Treatments, root-gall indexes, root weights, number of eggs per plant, number of eggs per g of roots, and dry shoot weights from the first Pittosporum tobira cv. Variegata growth room trial.....................................................................................45 2-7 Treatments, root-gall indexes, root weights, number of eggs per plant, number of eggs per g of roots, and dry shoot weights from the second Pittosporum tobira cv. Variegata growth room trial...............................................................................46 2-8 Treatments, root-gall indexes, root weights, number of eggs per plant, number of eggs per g of roots, and dry shoot weights from the Pittosporum tobira cv. Variegata greenhouse experiment............................................................................47 2-9 Treatments, root-gall indexes, root weights, number of eggs per plant, number of eggs per g of roots, and dry shoot weights from the first Salvia leucantha growth room trial..................................................................................................................49

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vii 2-10 Treatments, root-gall indexes, root weights, number of eggs per plant, number of eggs per g of roots, and dry shoot weights from the second Salvia leucantha growth room trial......................................................................................................50 2-11 Treatments, root-gall indexes, root weights, number of eggs per plant, number of eggs per g of roots, and dry shoot weights from the purple-corolla Salvia leucantha greenhouse experiment............................................................................52 2-12 Treatments, root-gall indexes, root weights, number of eggs per plant, number of eggs per g of roots, and dry shoot weights from the white-corolla Salvia leucantha greenhouse experiment............................................................................53 2-13 Treatments, root-gall indexes, root weights, number of eggs per plant, number of eggs per g of roots, and dry shoot weights from the first Odontonema cuspidatum growth room trial..................................................................................55 2-14 Treatments, root-gall indexes, root weights, number of eggs per plant, number of eggs per g of roots, and dry shoot weights from the second Odontonema cuspidatum growth room trial..................................................................................56 2-15 Treatments, root-gall indexes, root weights, number of eggs per plant, number of eggs per g roots, and dry shoot weights from the Odontonema cuspidatum greenhouse experiment.............................................................................................57 2-16 Treatments, root-gall indexes, root weights, number of eggs per plant, number of eggs per g of roots, and dry shoot weights from the first Musa acuminata ssp. zebrina growth room trial.........................................................................................58 2-17 Treatments, root-gall indexes, root weights, number of eggs per plant, number of eggs per g of roots, and dry shoot weights from the second Musa acuminata ssp. zebrina growth room trial.........................................................................................59 2-18 Treatments, root-gall indexes, root weights, number of eggs per plant, number of eggs per g of roots, and dry shoot weights from the Musa acuminata ssp. zebrina greenhouse experiment.............................................................................................61 2-19 Treatments, root-gall indexes, root weights, number of eggs per plant, number of eggs per g of roots, and dry shoot weights from the first Codiaeum variegatum cv. Gold Dust trial....................................................................................................62 2-20 Treatments, root-gall indexes, root weights, number of eggs per plant, number of eggs per g of roots, and dry shoot weights from the second Codiaeum variegatum cv. Gold Dust growth room trial...........................................................63 2-21 Treatments, root-gall indexes, root weights, number of eggs per plant, number of eggs per g of roots, and dry shoot weights from the Codiaeum variegatum cv. Gold Dust greenhouse experiment...........................................................................64

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viii 3-1 Known Meloidogyne spp. malate dehydrogenase and esterase relative migrations that were used as references against those that were revealed electrophoretically from females collected from several counties in Florida..........91 3-2 Enzyme stain concoctions used in staining malate dehydrogenase and esterase following electrophoresis using the PhastSystem....................................................94 3-3 Enzyme stain concoctions used in staining malate dehydrogenase and esterase following electrophoresis using the Mini-Protean 3 Cell.........................................96 3-4 Plant species, family, county, relative migration, isozyme phenotype, number of samples, and Meloidogyne spp. identified from ornamental plants collected in Florida and processed using the PhastSystem and Mini-Protean 3 Cell..................97 B-1 Sources and citations of publications referred to in Appendix B...........................124 B-2 Collective record of the host status of ornamental plants to Meloidogyne incognita M. javanica M. arenaria and M. hapla............................................... 125

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ix LIST OF FIGURES Figure page 3-1 Malate dehydrogenase and esterase relative electrophoretic migrations and enzyme phenotypes of Meloidogyne javanica as reported by several authors.......71 3-2 Malate dehydrogenase and esterase relative electrophoretic migrations and enzyme phenotypes of Meloidogyne incognita as reported by several authors......73 3-3 Malate dehydrogenase and esterase relative electrophoretic migrations and enzyme phenotypes of Meloidogyne arenaria as reported by Esbenshade and Triantaphyllou (1985c).............................................................................................75 3-4 Malate dehydrogenase and esterase relative electrophoretic migrations and enzyme phenotypes of Meloidogyne arenaria as reported by several authors.......77 3-5 Malate dehydrogenase and esterase relative electrophoretic migrations and enzyme phenotypes of Meloidogyne hapla ..............................................................79 3-6 Malate dehydrogenase and esterase relative electrophoretic migrations and enzyme phenotypes of Meloidogyne spp.................................................................81 3-7 Malate dehydrogenase and esterase relative electrophoretic migrations and enzyme phenotypes of unidentified Meloidogyne spp.............................................85 3-8 Malate dehydrogenase (Mdh) and esterase (Est) isozyme phenotypes, revealed using the mini-protean 3 cell apparatus, of Meloidogyne partityla females extracted from Carya illinoensis from Jefferson County on 31 March 2004........101 3-9 Malate dehydrogenase (Mdh) and esterase (Est) isozyme phenotype, revealed using the mini-protean 3 cell apparatus, of a Meloidogyne querciana female extracted from Viburnum odoratissimum cv. Awabuki from Hillsborough County on 25 January 2005....................................................................................102 A-1 PhastSystem gels exhibiting smeared malate dehydrogenase isozyme phenotypes whose relative migration could not be accurately measured..............107 A-2 Malate dehydrogenase and esterase isozyme phenotypes, revealed using the PhastSystem apparatus, of Meloidogyne females extracted from Hibiscus rosasinensis cv. Pink Versicolor from Alachua County on 02 April 2003...................108

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x A-3 Malate dehydrogenase and esterase isozyme phenotypes, revealed using the PhastSystem apparatus, of Meloidogyne females extracted from Rosmarinus officinalis from Suwannee County on 30 April 2003............................................108 A-4 Malate dehydrogenase and esterase isozyme phenotypes, revealed using the PhastSystem apparatus, of Meloidogyne females extracted from Callistemon viminalis from Lee County on 01 May 2003.........................................................109 A-5 Malate dehydrogenase and esterase isozyme phenotypes, revealed using the PhastSystem apparatus, of Meloidogyne females extracted from Syagrus romanzoffiana from Lee County on 27 May 2003.................................................109 A-6 Malate dehydrogenase isozyme phenotypes, revealed using the mini-protean 3 cell apparatus, of Meloidogyne females extracted from Callistemon viminalis from Lee County on 27 May 2003.........................................................................110 A-7 Eterase isozyme phenotypes, revealed using the mini-protean 3 cell apparatus, of Meloidogyne females extracted from Callistemon viminalis from Lee County on 27 May 2003...........................................................................................................110 A-8 Malate dehydrogenase isozyme phenotypes, revealed using the mini-protean 3 cell apparatus, of Meloidogyne females extracted from Pittosporum tobira from Alachua County on 18 December 2003.................................................................111 A-9 Esterase isozyme phenotypes, revealed using the mini-protean 3 cell apparatus, of Meloidogyne females extracted from Pittosporum tobira from Alachua County on 18 December 2003................................................................................111 A-10 Malate dehydrogenase isozyme phenotypes, revealed using the mini-protean 3 cell apparatus, of Meloidogyne females extracted from Ruscus aculeatus from Volusia County on 19 December 2003..................................................................112 A-11 Esterase isozyme phenotypes, revealed using the mini-protean 3 cell apparatus, of Meloidogyne females extracted from Ruscus aculeatus from Volusia County on 19 December 2003............................................................................................112 A-12 Malate dehydrogenase isozyme phenotypes, revealed using the mini-protean 3 cell apparatus, of Meloidogyne females extracted from Buddleia davidii from Pinellas County on 30 September 2003.................................................................113 A-13 Esterase isozyme phenotypes, revealed using the mini-protean 3 cell apparatus, of Meloidogyne females extracted from Buddleia davidii from Pinellas County on 30 September 2003............................................................................................113 A-14 Malate dehydrogenase isozyme phenotypes, revealed using the mini-protean 3 cell apparatus, of Meloidogyne females extracted from Brassica rapa cv. Shogoin from Alachua County on 06 January 2004..............................................114

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xi A-15 Esterase isozyme phenotypes, revealed using the mini-protean 3 cell apparatus, of Meloidogyne females extracted from Brassica rapa cv. Shogoin from Alachua County on 06 January 2004.....................................................................114 A-16 Malate dehydrogenase isozyme phenotypes, revealed using the mini-protean 3 cell apparatus, of Meloidogyne females extracted from Pittosporum tobira cv. Variegata from Lake County on 11 February 2004...............................................115 A-17 Esterase isozyme phenotypes, revealed using the mini-protean 3 cell apparatus, of Meloidogyne females extracted from Pittosporum tobira cv. Variegata from Lake County on 11 February 2004.........................................................................115 A-18 Malate dehydrogenase isozyme phenotypes, revealed using the mini-protean 3 cell apparatus, of Meloidogyne females extracted from Ruscus aculeatus from Volusia County on 22 February 2004....................................................................116 A-19 Esterase isozyme phenotypes, revealed using the mini-protean 3 cell apparatus, of Meloidogyne females extracted from Ruscus aculeatus from Volusia County on 22 February 2004..............................................................................................116 A-20 Malate dehydrogenase isozyme phenotypes, revealed using the mini-protean 3 cell apparatus, of Meloidogyne females extracted from Liriope muscari cv. Evergreen Giant from Hillsborough County on 01 July 2004...............................117 A-21 Esterase isozyme phenotypes, revealed using the mini-protean 3 cell apparatus, of Meloidogyne females extracted from Liriope muscari cv. Evergreen Giant from Hillsborough County on 01 July 2004...........................................................117 A-22 Malate dehydrogenase isozyme phenotypes, revealed using the mini-protean 3 cell apparatus, of Meloidogyne females extracted from Ophiopogon japonicus from Orange County on 16 August 2004...............................................................118 A-23 Esterase isozyme phenotypes, revealed using the mini-protean 3 cell apparatus, of Meloidogyne females extracted from Ophiopogon japonicus from Orange County on 16 August 2004.....................................................................................118 A-24 Malate dehydrogenase isozyme phenotypes, revealed using the mini-protean 3 cell apparatus, of Meloidogyne females extracted from Jesticia carnia from Hillsborough County on 20 August 2004..............................................................119 A-25 Esterase isozyme phenotypes, revealed using the mini-protean 3 cell apparatus, of Meloidogyne females extracted from Jesticia carnia from Hillsborough County on 20 August 2004.....................................................................................119 A-26 Malate dehydrogenase isozyme phenotypes, revealed using the mini-protean 3 cell apparatus, of Meloidogyne females extracted from Solenostemon scutellarioides cv. Elfers from Hillsborough County on 15 September 2004.......120

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xii A-27 Esterase isozyme phenotypes, revealed using the mini-protean 3 cell apparatus, of Meloidogyne females extracted from Solenostemon scutellarioides cv. Elfers from Hillsborough County on 15 September 2004................................................120 A-28 Malate dehydrogenase and esterase isozyme phenotypes, revealed using the mini-protean 3 cell apparatus, of Meloidogyne females extracted from Brassica oleracea from Orange County on 11 January 2004...............................................121 A-29 Malate dehydrogenase and esterase isozyme phenotypes, revealed using the mini-protean 3 cell apparatus, of Meloidogyne females extracted from Beta vulgaris from Orange County on 11 January 2004................................................121 A-30 Malate dehydrogenase and esterase isozyme phenotypes, revealed using the mini-protean 3 cell apparatus, of Meloidogyne females extracted from Phaseolus vulgaris from Hillsborough County on 25 January 2005.......................................122

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xiii Abstract of Thesis Presented to the Graduate School of the University of Florida in Partial Fulfillment of the Requirements for the Degree of Master of Science REPRODUCTION AND IDENTIFICATION OF ROOT-KNOT NEMATODES ON PERENNIAL ORNAMENTAL PLANTS IN FLORIDA By Roi Levin May 2005 Chair: W. T. Crow Major Department: Entomology and Nematology Meloidogyne spp. (root-knot nematodes) are serious pathogens of perennial and woody ornamental plants. Meloidogyne spp. directly limit plant vigor and, by the mechanical action and physiological responses to their feeding, expose their hosts to an array of pathogenic fungi and bacteria. The evaluation of the host status of perennial ornamental plants to root-knot nematodes can identify root-knot nematode resistant plant material, which may be used to replace infected hosts in landscapes. Six perennial ornamental species were evaluated for their host status to M. incognita race 2, M. javanica M. arenaria race 1, and M. mayaguensis in separate growth room and greenhouse experiments. Data from these experiments indicate that Liriope muscari cv. Evergreen Giant is a good host to M. incognita race 2, M. javanica and M. mayaguensis and a poor host to M. arenaria race 1. In addition, a purple-corolla form of Salvia leucantha and Musa acuminata ssp. zebrina cv. Rowe red are good hosts to the Meloidogyne spp. evaluated. Pittosporum tobira cv. Variegata, Odontonema cuspidatum

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xiv and Codiaeum variegatum cv. Gold Dust are nonhosts to the Meloidogyne spp. isolates evaluated. Twenty ornamental plants were identified as hosts to several Meloidogyne spp. via the speciation of root-knot nematode females that were dissected directly from their hosts roots. These females were identified primarily by evaluation of esterase (Est) and malate dehydrogenase (Mdh) isozyme phenotypes, unveiled following polyacrylamide gel electrophoresis. Resolved isozyme phenotypes indicate that Rosmarinus officinalis (rosemary), Syagrus romanzoffiana (queen palm), P. tobira Brassica rapa (turnip) cv. Shogoin, Brassica oleracea (kale), Phaseolus vulgaris (bean), L. muscari cv. Evergreen Giant, and Ophiopogon japonicus (mondo grass) are hosts to M. incognita Hibiscus rosa-sinensis (hibiscus) cv. Pink Versicolor, B. rapa cv. Shogoin, Ruscus aculeatus (ruscus), Beta vulgaris (chard), and Viburnum odoratissimum (Viburnum) cv. Awabuki are hosts to M. javanica Ruscus aculeatus and P. vulgaris are hosts to M. arenaria Callistemon viminalis (bottle brush), S. romanzoffiana and Solenostemon scutellarioides (coleus) cv. Elfers are hosts to M. mayaguensis Carya illinoensis (pecan) is a host to M. partityla In addition, Meloidogyne spp. that could not be identified on the basis of their Est and Mdh isozyme phenotypes were isolated from the following ornamental plants: Buddleia davidii (butterfly bush), P. tobira cv. Variegata, L. muscari cv. Evergreen Giant, and O. japonicus and Justicia carnea (flamingo plant).

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1 CHAPTER 1 INTRODUCTION AND LITERATURE REVIEW Introduction Perennial ornamentals are an important component of the nursery and floriculture industry. According to Hodges and Haydu (2003), the gross wholesale value for U.S.grown floriculture and nursery crops in 2001 reached $13.3 billion, of which $1.6 billion was produced in Florida alone. Over 2,500 species in roughly 500 genera are included in this category, and are widely distributed in the United States, Canada, and Europe (LaMondia, 1995, 1997). Widespread dissemination of perennial ornamentals presents an important avenue for distribution of root-knot nematodes ( Meloidogyne spp.) and other plant-pathogenic organisms (LaMondia, 1995). Root-knot nematodes cause estimated crop losses of 5 to 10% in major crops, and are considered the most widespread and destructive of all plant-pathogenic nematodes (Haseeb et al., 1984; Stokes, 1977; Walker et al., 1994). The lack of information regarding perennial ornamental crop losses due to plant-parasitic nematodes is attributed to the demand for research pertaining to agronomic crops, the long time period required for crop loss assessment on horticultural crops, and the vast array and interchangeability of available cultivated plant material (Walker and Melin, 1998b). Information regarding resistance of ornamental plants to root-knot nematodes is needed by extension personnel, the landscape industry, plant producers, and gardeners (Walker et al., 1994; Walker and Melin, 1998a). Furthermore, knowledge of the susceptibility of cultivated perennials to root-knot may alleviate postinstallation damage associated with these pathogens by the avoidance of highly

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2 susceptible cultivars and increased development and use of resistant cultivars (GiblinDavis et al., 1992). Incorporation of resistance into the genomes of perennial cultivars would greatly benefit the perennial plant industry (Walker and Melin, 1998a), as rotation with resistant species has been shown to successfully control root-knot nematodes in infested nursery fields (LaMondia, 1995). High demand and extensive shipments impact the rate of inspection for root-knot nematodes. Therefore, root-knot nematode management programs that are based on sanitation, resistance, tolerance, rotation, and exclusion, should be applied and developed in nurseries and landscapes to alleviate product losses, and to reduce the spread of the pathogens into uninfested field-grown nurseries and landscapes (Benson and Barker, 1985; LaMondia, 1997; Walker, 1980; Walker and Melin, 1998b). Care should be taken with the cultivation of tolerant perennials in root-knot nematode infested sites since annuals planted adjacent to such plants may become infected with root-knot nematodes that thrive on tolerant perennials (McSorley and Dunn, 1990; Rohde, 1972). Meloidogyne spp. Historical Background The first account of root-knot nematodes was by Berkeley, who in 1855 observed galls on roots of greenhouse-grown cucumber plants in England (Hartman and Sasser, 1985). Cornu first coined the name Anguillula marioni Cornu for root-knot nematodes, after observing root galls on Onobrychis sp. (sainfoin). Subsequently, root-knot nematodes were classified in the genera Heterodera or Anguillula In 1884, Mller classified root-knot nematodes as H. radicicola and in 1887 the type species Meloidogyne exigua was described by Gldi. In 1932, Goodey reclassified root-knot nematodes as H.

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3 marioni (Mller) Goodey. Differences in host responses led Chitwood to redescribe Meloidogyne exigua Gldi, M. incognita (Kofoid and White, 1919) Chitwood, M. javanica (Treub, 1885) Chitwood, and M. arenaria (Neal, 1889) Chitwood, and to describe M. hapla Chitwood and M. incognita acrita Chitwood, as a new species and variety, respectively (Christie, 1959). In his description of M. hapla Chitwood cited Abelia grandiflora as a host (Bernard and Witte, 1987). Chitwood also gave a general description of the genus Meloidogyne and differentiated it from the genus Heterodera (Hirschmann, 1985). Chitwoods diagnoses of the redescribed and newly described species were based on examinations of morphological features and morphometrics from all life stages of the evaluated species (Eisenback, 1985). These features included the perineal pattern, stylet morphology, and distance from the base of the stylet knobs to the dorsal esophageal gland opening (DEGO) (Christie, 1959). To date, more than 80 Meloidogyne spp. have been described (Randig et al., 2002). The current taxonomic status for the Meloidogyne spp. reviewed in this study is: phylum Nemata, order Tylenchida, suborder Tylenchina, superfamily Tylenchoidea, family Heteroderidae, subfamily Meloidogyninae, species M. incognita M. javanica M. arenaria and M. mayaguensis Rammah and Hirschmann, 1988 (Andrssy, 1976; Rammah and Hirschmann, 1988; Thorne, 1961). Meloidogyne incognita M. javanica and M. arenaria have been frequently encountered in Florida for many years. Meloidogyne mayaguensis however, was not detected frequently in Florida until the use of electrophoresis for the identification of root-knot nematodes became commonplace. Meloidogyne mayaguensis was described as a result of the reevaluation of a population that was tentatively identified as M. arenaria at North Carolina State University. Differentiating biological

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4 and morphological characters that part this species from the four common Meloidogyne species include a somatic chromosome number of 2n = 44 to 45 and unique enzyme phenotypes revealed through electrophoresis (Rammah and Hirschmann, 1988). Although the M. mayaguensis type species exhibits a morphologically unique perineal pattern, investigations by Brito et al. (2004) revealed M. mayaguensis isolates having a perineal pattern similar to that of M. incognita in Florida, making this feature unreliable for the identification of M. mayaguensis Identification The accurate identification of root-knot nematodes to species and host races is essential for their control and is a prerequisite to meaningful research. Many Meloidogyne species are easily identified based on distinct morphological characters and restricted host ranges. Several species are difficult to identify due to their similarity to other species and poor taxonomic descriptions. The four most common root-knot nematode species, composing 98% of all worldwide populations, are M. incognita M. javanica M. arenaria and M. hapla (Hussey, 1985a). Other Meloidogyne species, such as M. mayaguensis become increasingly important due to their uncommon virulence and increasing occurrence. Difficulty in identifying root-knot nematodes may result from morphological variations within and between populations from a same species. Since the reevaluation of Meloidogyne spp. by Chitwood in 1949, female perineal patterns became the dominant diagnostic character of the four most common Meloidogyne species. The perineal pattern presents several benefits that render it a valuable diagnostic tool. Aside from minor variations, perineal patterns are constant within populations and their source (females) is abundant in infected host roots. Other diagnostic features used in taxonomic

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5 identification include female stylets, male heads and stylets, and second-stage juvenile (J2) heads and stylets. Morphological features of the perineal patterns of M. incognita M. javanica M. arenaria and M. mayaguensis are describes as follows: Meloidogyne incognita Striae are smooth, wavy, sometimes in a zigzag pattern. Lateral lines are absent. A squarish, high dorsal arch containing a distinct whorl around the tail terminus is the most conspicuous diagnostic character of this perineal pattern (Eisenback, 1985). Meloidogyne javanica Striae are smooth and somewhat wavy. The dorsal arch is often low and rounded but may be high and squarish, frequently possessing a whorl in the tail terminus area. Unique to this species are distinct lateral ridges that run across the pattern, fading away around the tail terminus (Eisenback, 1985). Meloidogyne arenaria Striae are smooth and slightly wavy, often extended laterally, forming wings on one or both lateral sides of the pattern. Distinctive lateral ridges are absent, but are marked by forked, irregular lateral fields. The dorsal arch is low and indented near the lateral fields, forming rounded shoulders (Eisenback, 1985). Meloidogyne mayaguensis Striae are fine, continuous, and widely spaced. Lateral lines are inconspicuous or a single lateral line may be present on one side of the pattern. Dorsal arch is rounded, with a circular, striae-free tail terminus (Rammah and Hirschmann, 1988). Thirty percent of three isolates from Florida possess perineal patterns that differ from the type species and depict perineal patterns typical of M. incognita (Brito et al., 2004). Inconsistencies in host-parasite relationships lead to the erection of speciesspecific races based on their infection of specific crops, namely Gossypium hirsutum

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6 (cotton) cv. Deltapine 61, Nicotiana tabacum (tobacco) cv. NC 95, Capsicum frutescens (pepper) cv. California Wonder, Citrullus vulgaris (watermelon) cv. Charleston Gray, Arachis hypogaea (peanut) cv. Florunner, and Lycopersicon esculentum (tomato) cv. Rutgers. Based on their susceptibility to the differential hosts, M. incognita and M. arenaria were assigned four and two races, respectively. This host differential system allows for the rotation of the differential hosts as a means of maintaining low nematode populations and thus reduces crop losses. However, the identification of root-knot nematode species solely on the basis of the differential host test is unreliable due to the possibility of mixed populations, and should be used in conjunction with morphological, morphometric, and biochemical evaluations to determine root-knot nematode species (Hartman and Sasser, 1985). Problems in the morphological identification of Meloidogyne species, such as intra-species merging of morphological characters, rarely-seen characters, and lack of apparent differences between species, has encouraged much interest in the utilization of biochemical techniques as a complementary, routine method for the identification of rootknot nematodes (Hansen and Buecher, 1970). Enzyme phenotypes, unveiled through staining of polyacrylamide gel slabs following electrophoresis, have become a reliable, less subjective approach to identification of root-knot nematodes (Hussey, 1985a). The first study on root-knot nematode protein profile stability and its utilization in the identification of root-knot nematodes was conducted by Dickson et al. (1970), who established the usefulness of disc-electrophoresis in identification of root-knot nematodes. This early work also verified that protein profiles were stable within nematode species collected from different parts of the world and that infect an array of

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7 hosts. Further work by Dickson et al. (1971) examined lactate dehydrogenase (LDH), malate dehydrogenase (MDH), -glycerophosphate dehydrogenase (GDH), glucose-6phosphate dehydrogenase (G-6-PDH), acid phosphatase (AcP), alkaline phosphatase (AlkP), and esterase (EST) enzyme profiles from Meloidogyne javanica M. arenaria and M. hapla females, as well as from three life stages of M. incognita Dickson et al. (1971) revealed that enzyme profiles for GDH, MDH, G-6-PDH, and EST differed among the four studied Meloidogyne species. Furthermore, G-6-PDH enzymatic profiles from all evaluated species and GDH enzymatic profiles from M. incognita M. hapla and M. arenaria were monomorphic, while enzymatic profiles from all the remaining species evaluated were polymorphic. Among the enzymes evaluated for the characterization of Meloidogyne species, MDH and EST enzyme profiles were most variable with respect to electrophoretic mobility and therefore are most useful for differentiation of the four species evaluated by this method. However, MDH enzyme profiles did not differ between M. javanica and M. incognita and EST isoenzymes were detected from M. incognita and M. hapla isolates. Enzymatic profiles from the three life stages of M. incognita resulted in variable band numbers and electrophoretic migration for MDH and EST. The use of disk-electrophoresis for the assessment of Meloidogyne species protein phenotypes by Dickson et al. (1971) required the analysis of several specimens of the same species for the elucidation of a single protein phenotype. This method, therefore, rendered genetic analysis at the intraand interspecific levels impossible (Dalmasso and Berge, 1978). A breakthrough in biochemical speciation of Meloidogyne species was the use acrylamide gels as thin slabs (0.7-mm-thick) to electrophoretically separate proteins from individual Meloidogyne females. First used by Dalmasso and Berge (1978), the

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8 thin-slab method for polyacrylamide gel electrophoresis (PAGE) has proved to be useful in the identification of root-knot nematodes. This method allowed the speciation of 20 to 25 individual specimens from the same or different populations on single gels. Analyzing 22,000 specimens of root-knot nematodes using microscale electrophoresis, Dalmasso and Berge (1978) found that, of the enzymes elucidated, EST were the most useful for the differentiation of the common Meloidogyne species, primarily due to their polymorphic nature (Hussey, 1985a). Morphological characters, particularly female perineal patterns, are the primary method for routine root-knot nematode identification. However, perineal patterns are variable, and may lead to misidentification of aberrant populations and uncommon species. Conversely, biochemical analyses, particularly esterase phenotypes of young, egg-laying females, in conjunction with morphological and morphometric examinations, allow for precise, accurate diagnoses, thereby alleviating the confusion associated with morphological characters (Hussey, 1985a). In addition to morphological and morphometric analysis and the use of PAGE for the identification of root-knot nematodes, the use of single eggs or J2 for species identification via restriction fragment length polymorphism (RFLP) (Fargette et al, 1996), random amplified polymorphic deoxyribose nucleic acid (DNA) (RAPD) (Blok et al., 1997b), ribosomal DNA amplification (Blok et al., 1997a), and mitochondrial DNA (mtDNA) amplification (Blok et al., 2002), has been reported.

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9 Biology and Ecology Root-knot nematodes occur throughout the world and infect most major plant crops, and are responsible for substantial yield losses and reduced crop quality (Sasser and Carter, 1985). In Florida, root-knot nematodes are the most damaging plant-parasitic nematodes on agricultural crops (McSorley and Dunn, 1989). The life cycle of root-knot nematode is largely indifferent with respect to individual species host-parasite relationships and physiological characteristics (Christie, 1959). Root-knot nematode eggs are protected within a gelatinous egg mass produced by the female (Maggenti, 1987). Inside the egg, a first-stage root-knot nematode juvenile (J1) molts once prior to hatching into a J2. While egg hatching is usually spontaneous and does not correlate with plant-root stimuli, root diffusates have been shown to stimulate hatching (Hussey, 1985b). Once hatched, the now J2 move though the soil in search of a suitable feeding site (Christie, 1959). Root penetration by the pathogen involves the mechanical disruption of host tissues. However, cellulose and pectin-dissolving enzymes may also aid in the penetration process (Hussey, 1987). Upon penetration, J2 move within the root in a vertical manner, and often migrate toward and away from the root surface. Although penetration may occur anywhere in the root system, J2 are often observed aggregating and penetrating behind the root cap, near the meristematic zone. Other penetration sites include cracks and lesions of mature roots and areas of secondary root formation (Lewis, 1987). Furthermore, the site of one nematodes penetration often becomes attractive for other J2, leading to multiple infections in confined areas (Hussey, 1985b). Once established within the plant tissue, root-knot nematodes become sedentary endoparasites, and halt further movement or migration (Christie, 1959). Second-stage juveniles are

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10 often observed in cortical tissues about the zone of differentiation, their bodies vertically parallel with the root axis and heads settled in the periphery of the vascular tissue (Hussey, 1985b). Upon establishment of a suitable feeding site, sexually-undifferentiated J2 begin to modify the hosts physiology by transforming healthy, undifferentiated cells into specialized feeding sites referred to as giant cells. Modified cells exhibit nuclear, nucleolar, and surface hypertrophy, an increase in cytoplasmic density, organelle hyperplasia, and disappearance of the central vacuoles. While the nematodes two subventral glands are involved in giant cell initiation, dorsal gland secretions maintain giant cell development (Lewis, 1987). The ability of J2 to invade roots differs with respect to the root-knot nematode species, but symptoms of root-knot nematode penetration are often depicted as root-tip enlargement and root-growth retardation. While gall development is not essential for root-knot nematode survival, the development and maintenance of giant cells is critical for root-knot nematode development. This hostparasite relationship requires the developing root-knot nematode to feed on five to six viable giant cells. The inability to elicit a giant cell response results in nematode death or, if early in the J2 stage, migration out of the root in search of a suitable feeding site (Hussey, 1985b). Late in the J2 stage, following feeding initiation and giant cell formation, an increase in J2 width is observed, and the genital primordia attain their sexual characteristics prior to the second molt. The third-stage juvenile (J3) of both sexes is depicted by the J2 cuticle surrounding the J3, the loss of the stylet and the median esophageal bulb valve, and the loss of the tail spike, which becomes rounded. The J3 stage passes in a few hours, at which time a third molt gives rise to the fourth-stage juvenile (J4). In this stage, which lasts longer than the third stage, the median esophageal

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11 bulb valve is reformed and the excretory pore opens. The female rectal glands, uterus and vagina, and male vas deferens differentiate and enlarge, and the male undergoes metamorphosis, attaining an elongated, cylindrical shape. Following gonad differentiation in the fourth stage, a fourth and final molt reveals the adult nematodes enclosed in the three previous juvenile cuticles. At this stage the stylet reappears in both sexes, the perineal pattern is observed in females, and sperm production is initiated in males prior to the disappearance of the previously-molted cuticles (Triantaphyllou and Hirschmann, 1960). While male production occurs in most root-knot nematode species, M. hapla which reproduces through facultative meiotic parthenogenesis, produce relatively more males than M. incognita M. javanica and M. arenaria which reproduce through obligate mitotic parthenogenesis (Triantaphyllou, 1985). Similarly, M. mayaguensis reproduces through mitotic parthenogenesis (Rammah and Hirschmann, 1988). The pathogens parthenogenic reproductive behavior is adaptive to its unstable environmental and physiological conditions, namely the halt in female motility, short generation time, limiting habitat, and male infrequency. Parthenogenesis allows for the establishment of varying phenotypes through polyploidy, thereby increasing adaptation to unfavorable environmental conditions, including, but not limited to, fluctuating thermal gradients, drought, and low oxygen concentrations. Furthermore, such adaptations may allow for rapid generation turnover through rapid maturity of developing juvenile stages (Maggenti, 1987). Regardless of their reproductive behavior, Meloidogyne spp. produce less eggs and more males in response to increasing populations within a root system (Lewis, 1987). Upon maturation, females produce a gelatinous matrix through their rectal glands, into which they deposit eggs. The gelatinous matrix provides eggs with

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12 physical protection (Maggenti, 1987) and acts as a barrier to temperature fluctuations and water evaporation from eggs (Van Gundy, 1985). Meloidogyne spp. viability and development is influenced by various environmental and physical stresses, including temperature, soil texture, moisture, aeration, osmotic potential, and host suitability. Temperature has the greatest influence on egg development and hatching, growth, reproduction, and survival. Optimal temperatures for egg development of M. incognita M. javanica and M. arenaria are 10 to 15 C, and approximately 9 C for M. hapla Conversely, optimal temperatures for growth and development of juvenile and adult stages of M. incognita M. javanica and M. arenaria are 25 to 30 C. Although temperature extremes may inhibit reproduction, individual populations within certain species acclimatize to local temperature regimes. In general, relative cold tolerance is exhibited by M. hapla M. incognita M. arenaria and M. javanica in decreasing order, with M. hapla surviving subzero temperatures and M. javanica not surviving temperatures below 10 C. While Meloidogyne eggs do not survive in soils beyond one year, varying egg mass colors, from white to brown, have been attributed to egg dormancy and overwintering. Soil texture, moisture, and aeration compose a dynamic, complex environment, and influence Meloidogyne activity, reproduction, and pathogenicity. Continuously changing, soil texture is a solid-phase component of the soil environment. Vertically and horizontally shifting soil particles are interconnected by the liquid and gas phases of this dynamic system. Optimal temperature regimes and moisture levels of 40 to 60% of field capacity present the most advantageous conditions for nematode activity and metabolism. Moisture levels that are lower or higher than optimal reduce nematode activity due to drying soils and limiting oxygen concentrations,

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13 respectively (Van Gundy, 1985). It is widely accepted that Meloidogyne spp. activity is optimal in sandy loam soils and reproduction is greatest in fine sand (Wallace, 1969), primarily due to the low water holding capacity of such soils (Benson and Barker, 1985). However, Whitehead (1969) reported that eastern African population viability of M. incognita and M. javanica exhibited no correlation with soil texture. In addition, OBannon and Reynolds (1961) reported on heavy M. incognita infestations in coarsetextured soils planted to cotton in Arizona. Root-knot nematode females, examined in roots from different hosts, were variable in size. Highly susceptible plants exhibited large, robust females while less susceptible plants supported smaller females (Pant et al., 1983). Furthermore, Niblack and Bernard (1985) observed that M. hapla densities were positively correlated with Cornus florida and Acer rubrum nursery tree age. Meloidogyne spp. behavior is altered under various soil solution osmotic potentials. In drying, well-fertilized soil, nematodes are subjected to high osmotic pressures as the osmotic potential increases. Wallace (1969) observed the highest and lowest reproduction of M. javanica on Lycopersicon peruvianum (tomato) cv. Tatura Dwarf when subjected to high and low nutrient levels, respectively. However, M. javanica infectivity decreased with increased electrical conductivity of the soil solution, and J2 migration was observed from high to low salt concentrations (Van Gundy, 1985). The interacting environmental and biological factors that influence Meloidogyne spp. development, reproduction, and pathogenicity are complex and difficult to evaluate. Such factors as soil homogeneity, moisture, and temperature, in addition to changing osmotic and water potentials, interact with nematode development, reproduction, and pathogenicity, as well as various metabolic and developmental aspects of the host plant.

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14 Interactions between nematodes and plants, further complicated by inoculum densities and developmental levels, lead to the inevitable intricacy of this plant-parasite and biological relationship. In evaluating the effect of nematode populations on host plants, factors influencing the experimental environment are minimized to the studied components by controlling variables as temperature, moisture, lighting, host nutrition, initial inoculum density, and initial host development, in hope to reveal a significant relationship between pathogen and host, with minimum interference by unstudied factors (Van Gudny, 1985). Relationships with Ornamentals One of the most serious groups of pathogens to limit agricultural productivity, root-knot nematodes have been reported to infect many plants throughout the world (Zarina and Abid, 1995), and are considered the most damaging group of plant-parasitic nematodes in Florida (McSorley and Dunn, 1989). The woody ornamental/floriculture industry is one of the U.S.s fastest growing agricultural segments (Bernard and Witte, 1987). As serious pathogens of numerous woody ornamentals (Barker and Benson, 1977), Meloidogyne spp. have the potential to damage many important nursery crops (Benson and Barker, 1985) and form disease complexes with certain soil-borne fungal pathogens, thus increasing their hosts susceptibility to such pathogens (Nigh, 1972; Santamour and Riedel, 1993; Walker and Melin, 1998b). Unlike annual plants, damage thresholds levels do not apply to perennials, since low Meloidogyne spp. populations have the potential to increase and cause severe damage over a period of several years post-planting (LaMondia, 1995).

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15 In the U.S., root-knot nematodes have been reported to cause damage and reduced yields on ornamental crops in Alabama (Heald, 1967), Arizona (Nigh, 1972; Tarjan, 1952), California (Santo and Lear, 1976; Viglierchio, 1979), Connecticut (LaMondia, 1995, 1996, 1997), Florida (Giblin-Davis et al., 1992; Lehman, 1984a, 1984b; Lehman and Barnard, 1982; McSorley and Dunn, 1989, 1990; McSorley and Marlatt, 1983; Stokes, 1977, 1982), Georgia (Heald, 1967; Motsinger et al., 1977; Walker et al., 1994; Walker and Melin, 1998a, 1998b), Hawaii (Sher, 1954), North Carolina (Barker et al., 1979; Barker and Benson, 1977; Benson and Barker, 1982; Haasis et al., 1961; Rickard and Dupree, 1978, Sasser et al., 1966), Oklahoma (Nemec and Morrison, 1972; Nemec and Struble, 1968), Tennessee (Bernard and Witte, 1987; Bernard et al., 1994; Niblack and Bernard, 1985), Virginia (Eisenback, 1987), and Washington, DC (Santamour, 1992; Santamour and Riedel, 1993, 1995). Internationally, Meloidogyne spp. have been reported to damage ornamental crops in Australia (Wallace, 1969), Belgium (Coolen and Hendrickx, 1972; Stoffelen et al., 2000), Egypt (Montasser, 1995), France (De Waele and Davide, 1998), India (Ahuja and Arora, 1980; Haseeb et al., 1984, 1985; Khanna et al., 1998; Mishra and Mishra, 1997; Mishra and Misra, 1993; Misra and Mishra, 1997; Misra et al., 2002; Pant et al., 1983; Singh et al., 2000; Singh and Gupta, 1993), Iraq (Singh and Majeed, 1991), Ivory Coast (Adiko, 1988), Korea (Cho et al., 1996), Nigeria (Caveness and Wilson, 1977), Pakistan (Zarina and Abid, 1995), Saudi Arabia (Ibrahim and AlYahya, 2002), Spain (Jaizme-Vega et al., 1997), Tamil Nadu (Rajendran et al., 1975), and Trinidad and Tobago (Bala and Hosein, 1996). Although commonly observed, the potential for damage to many perennial ornamental crops by root-knot nematodes is unrecognized (Bernard and Witte, 1987).

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16 Meloidogyne spp. have been attributed as causing decline to growth of ornamental plants and thus hinder production and lead to reduced returns (Stokes, 1977). Meloidogyne incognita has been attributed to such effects on the production of Anthurium andraeanum and other tropical ornamentals (Bala and Hosein, 1996), certain flower bulbs (Montasser, 1995), and several Ilex spp. (Heald, 1967). In addition, production losses in field-grown Rosa spp. have been attributed to M. hapla (Santo and Lear, 1976). Furthermore, Meloidogyne spp. have been reported to reduce Dianthus caryophyllus production worldwide by 10 to 20% (Cho et al., 1996) and cause qualitative and quantitative decline in Indian Gladiolus hortulanus production (Khanna et al., 1998). Furthermore, in the middle Tennessee nursery-growing region, Meloidogyne hapla occurs in approximately 25% of nursery blocks (Bernard et al., 1994). Goff (1936) was one of the first researchers to conduct an extensive survey of the susceptibility of ornamental plants to Meloidogyne spp. In his survey, Goff noted the varying degrees of susceptibility among the tested plant species. Root-knot infected plants often exhibit symptoms that include root galls and root rots, shoot yellowing and chlorosis, stunted growth, and other symptoms commonly associated with nutritional deficiencies (Bala and Hosein, 1996; Bird, 1974; Misra et al., 2002, Santo and Lear, 1976; Zarina and Abid, 1995), resulting in general decline (Nigh, 1972), poor yield, and wilt diseases (Rajendran et al., 1975). Furthermore, photosynthetic rate reduction has been observed in response to root-knot nematode infections. Often, the ratio of food resources provided by the host plant and the root-knot nematode density determines the degree of host response to infection (Bird, 1974). However, Walker and Melin (1998b) observed greater plant growth in the presence of low plant-parasitic nematode

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17 populations than in their absence. Susceptible hosts severely infected with root-knot nematodes often decline as a linear function of time (Barker and Benson, 1977), and such decline is accelerated under unfavorable climatic conditions (Haasis et al., 1961), leading to severe disease symptoms in late summer, fall, and during periods of severe dry periods (Lehman, 1984a). The degree of root galling is dependant upon the infected plant species and/or cultivar and the root-knot nematode species, race, population, or even isolate (Bird, 1992; LaMondia, 1995; Rohde, 1972). Certain ornamentals infected with rootknot nematodes exhibit unique symptoms. Such plants include Sansevieria cylindrica which developed leaf discoloration and tip necrosis 4 to 5 months post-infection with M. incognita (Mishra and Mishra, 1997), Philodendron selloum which exhibits a reduction in leaf size when infected with M. incognita (Mishra and Misra, 1993), and Juniperus horizontalis var. Plumosa and Thuja orientalis cv. Dwarf Greenspike, which exhibit thickened roots and slight galling post-infection with Meloidogyne spp. (Nemec and Morrison, 1972). Furthermore, Gladiolus hortulanus plants infected with M. incognita race 2 exhibited leaf drying, reduction in floral stalk height and girth, and reduced number of florets (Khanna et al, 1998). Some plants exhibit minute galls following infection with Meloidogyne spp. In such conditions, root-knot nematode females can be seen protruding from the infected roots. Other plants, such as Rheum spp., Begonia spp., and Thunbergia spp. produce large galls, measuring up to 0.6 m in the latter case (Bird, 1974). In testing the susceptibility of numerous herbaceous perennial ornamentals to M. hapla however, LaMondia (1995, 1996) did not observe egg mass production in the absence of cellular hypertrophy.

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18 Mixed populations of Meloidogyne spp. are often observed parasitizing perennial ornamentals. In Pakistan, mixed populations of M. javanica and M. incognita have been reported on Solanum nigrum Cucumis melo ssp. melo var. flexuosus and Rosa indica (Zarina and Abid, 1995). Pant et al. (1983) did not observe infection signs or symptoms on Matthiola spp., Tagetes spp., Gaillardia spp., Chrysanthemum spp., and Zinnia spp. after inoculation of these plants with M. incognita Likewise, no signs or symptoms were observed in Areca catechu nine months after inoculation with Meloidogyne arenaria race 1, M. incognita races 1 and 3, M. javanica and M. hapla (McSorley and Dunn, 1989). In testing for pathogenicity of M. incognita on flower bulbs, Montasser (1995) rendered Amarylis vittata Clivia miniata Crinum longifolium and Narcissus tazetta which belong to Amaryllidaceae, and Agapanthus umbellatus Hysacinthus orientalis Lilium longiflorum and Tulip suaveolens which belong to Liliaceae, highly resistant. Likewise, Rajendran et al. (1975) found Barleria prionitis free of M. incognita infections in a pathogenicity experiment. Osborne and Jenkins (1963) observed M. hapla juveniles and light galling in Forsythia intermedia but indicated that invading juveniles failed to mature. Similar observations were reported by Bernard and Witte (1987) for Ligustrum sinense and Nandina domestica in which giant cells failed to develop following infection with M. hapla In addition, N. domestica was a nonhost for M. arenaria (Benson and Barker, 1982). Several landscape ornamentals were tested for their susceptibility to M. arenaria M. incognita races 1, 2, and 3, and M. javanica Of these, Photinia fraseri was tolerant to all the M. incognita isolates as well as to M. javanica Furthermore, egg mass indices for all evaluated Meloidogyne spp. except for M. javanica were low on Dracaena marginata while Ficus benjamina was highly susceptible to all Meloidogyne spp. except

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19 M. hapla (McSorley and Dunn, 1990). Several herbaceous perennials were evaluated for susceptibility to Meloidogyne spp. by Walker and Melin (1998a). Fragaria ananassa cv. Pink Panda, Monarda didyma cv. Blue Stockings, Phlox paniculata cv. Eva Cullum, Franz Shubert, and Oakington Blue, and Polygonum affine cv. Dimity did not support M. arenaria and M. incognita populations six weeks after inoculation. Geranium psilostemon procurrens cv. Ann Folkard did not support populations of M. arenaria while G. cinereum cv. Laurence Flatman did not support populations of M. incognita Evaluating galling of several herbaceous perennials by Meloidogyne spp., Walker and Melin (1998a) found no or very few galls on Aethionema cordifolium Echinacea purpurea Moranda citriodora or Patrinia scabiosifolia Similarly, Santo and Lear (1976) found Rosa noisettiana cv. Manetti a poor host to M. hapla while Coolen and Hendrickx (1972) found R. canina cv. Succes and Heinsohns Rekord poor hosts for M. hapla In testing the susceptibility of several annual bedding plants to Meloidogyne spp., McSorley and Frederick (1994) found that M. incognita race 1 caused no galls on Ageratum houstonianum cv. Blue Mink, Tagetes patula cv. Dwarf Primrose, Vinca rosea cv. Little Bright Eye, and Salvia splendens cv. Bonfire, and very light galling on Verbena hybrida cv. Florist and Zinnia elegans cv. Scarlet. In the same experiments, Ageratum houstonianum cv. Blue Mink, Lobularia maritime cv. Rosie ODay, and T. patula cv. Dwarf Primrose exhibited no and little infection symptoms post inoculation with M. javanica and M. arenaria race 1, respectively. Dianthus chinensis cv. Baby Doll Mix was slightly infected by M. javanica and M. arenaria while V. rosea cv. Little Bright Eye and Z. elegans cv. Scarlet were minimally infected by M. javanica and M. arenaria respectively. In evaluating carnation cultivars for M. incognita resistance, Fawzy et al.

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20 (1991) and Cho et al. (1996) reported on the resistance of several cultivars of Dianthus caryophyllus to M. incognita Bernard and Witte (1987) reported that M. hapla failed to reproduce on several species of Prunus and Rhododendron although Haasis et al. (1961) reported that Rhododendoron spp. and Camellia spp. supported light Meloidogyne spp. populations in field experiments. Giblin-Davis et al. (1992) evaluated the susceptibility of several Ixora spp. cultivars to M. incognita race 1 and M. javanica Although all Ixora spp. were susceptible to the Meloidogyne spp. isolates evaluated, above-ground disease symptoms were not observed 20 weeks after inoculation. Giblin-Davis et al. (1992) hypothesized that either the Ixora spp. cultivars tested were tolerant to the inoculated Meloidogyne spp. isolates, or that 20 weeks was not enough time for above-ground disease symptoms manifestation. LaMondia (1995, 1996) conducted extensive studies that evaluated the susceptibility of an array of perennial herbaceous ornamentals to M. hapla Approximately 30% of tested perennials were resistant to the isolate evaluated. In India, Khan and Khan (1989) reported on the resistance of Verbena bipinnatifida to M. incognita and presented nematicidal properties associated with root exudates from this plant. Santamour and Riedel (1993) tested 23 landscape trees for root-knot nematode resistance, of which six were nonhosts to M. arenaria races 1 and 2, M. hapla M. incognita and M. javanica including Magnolia grandiflora and Gleditsia triacanthos which are widely used native American landscape trees. Niblack and Bernard (1985) and Lehman and Barnard (1982) reported on M. hapla and M. incognita infecting Cornus florida and Niblack and Bernard (1985) reported on M. hapla infecting Acer rubrum and Prunus persica Several gymnosperms were nonhosts to M. hapla (Bernard and Witte, 1987) and resistant to M. incognita (Nemec and Struble, 1968). Walker and Melin

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21 (1998b) reported Cupressocyparis leylandi Cedrus deodara and Taxodium distichum to be nonhosts to M. incognita Bernard and Witte (1987) found no M. hapla reproduction on Acer palmatum and A. saccharum In Washington, Santamour and Riedel (1995) reported that Cercis canadensis C. chinensis and C. yunnanensis seedlings were severely galled by M. arenaria races 1 and 2, M. incognita M. javanica and M. hapla but only supported populations of M. hapla Reporting on the susceptibility of Acer spp. to Meloidogyne spp., Santamour (1992) found several Acer spp. resistant or tolerant to M. arenaria races 1 and 2, M. hapla M. incognita M. javanica and M. querciana and reported a wide range in susceptibility, even among progeny of a single tree. Several authors documented variable root gall sizes and shapes on different susceptible ornamental crops. Bernard and Witte (1987) documented that M. hapla induced galls were minute and difficult to detect in Abelia grandiflora Photinia fraseri Spirae bumalda cv. Froebelii, and S. vanhouttei Galls were intercalary and spindle-shaped on thick, fleshy roots of Hydrangea paniculata and Viburnum carlesii and spherical and terminal on Cornus florida roots infected with M. hapla However, C. florida infected with M. incognita exhibited spindle-shaped, intercalary galls. In testing certain cultivars of Tagetes patula to M. incognita M. arenaria and M. hapla Motsinger et al. (1977) found that only one cultivar, Tangerine, was free of root galls or egg masses. They further suggested that certain T. patula cultivars act as trap crops rather than produce nematicidal agents from their roots. Similarly, pathogenicity studies conducted in India found only T. erecta cv. Crackerjack to be resistant to M. javanica as no galls or penetration were observed on this cultivar following inoculation

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22 with the pathogen (Singh and Gupta, 1993). In addition, Eisenback (1987) tested eleven populations of M. hapla of different cytological forms and from different geographic origins for their pathogenicity on T. erecta cv. Carnation. Although most M. hapla populations did not reproduce on this host, several populations varied in their pathogenicity on this host, causing varying degrees of root galling and root proliferation in response to infection. Perhaps no group of plants has been studied more for their susceptibility to rootknot nematodes as those in the genus Ilex Susceptibility studies involving Ilex spp. generate conflicting results that may be attributed to nematode variants and differing host susceptibilities. Symptoms often observed in root-knot infected Ilex spp. include poor growth and vigor, foliage yellowing and bronzing, dieback of branches, root-system distortion, and in severe infections, death (Haasis et al., 1961). Furthermore, Ilex spp. foliage chlorosis is often associated with root galling (Heald, 1967). Meloidogyne incognita is frequently observed parasitizing Ilex spp. (Bernard et al., 1994). Nemec and Struble (1968) documented the pathogenicity of M. incognita on I. cornuta cv. Burfordii, I. crenata cv. Hetz, and I. cassine var. angustifolia Haasis et al. (1961) found I. crenata to be severely damaged by M. incognita and M. hapla Similarly, Barker and Benson (1977) found I. crenata cv. Convexa, Helleri, and Rotundifolia to be susceptible to M. arenaria and Stokes (1982) documented I. crenata to be frequently infected by M. javanica and M. incognita in commercial nurseries. On the contrary, Barker et al. (1979) found I. cornuta cv. Burfordii and I. vomitoria cv. Nana to be resistant and tolerant to M. arenaria respectively, and I. cornuta cv. Rotunda a poor host to M. arenaria McSorley and Dunn (1990) found I. cornuta cv. Burfordii to be a nonhost to M. arenaria M.

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23 incognita races 1 and 3, and M. javanica Sasser et al. (1966) observed no galling of I. crenata cv. Convexa, Latifolia, and Helleri, or I. cornuta cv. Burfordii following inoculation with M. hapla but I. crenata cv. Convexa, Latifolia, and Helleri were highly susceptible to M. incognita M. javanica and M. arenaria Heald (1967) indicated that I. crenata cv. Helleri was susceptible to M. incognita M. javanica M. arenaria and M. hapla in greenhouse experiments, in which galled roots were observed and plant weights were reduced in response to root-knot nematode infections. However, in these experiments symptoms of M. hapla were less apparent than those caused by the other root-knot nematode species. Other holly cultivars that did not support M. hapla reproduction include I. attanuata cv. Foster No. 2, I. crenata cv. Hetzii, and I. cornuta aquifolium cv. Nellie R. Stevens (Bernard and Witte, 1987). Accounting for these susceptibility variations among the various Ilex spp. cultivars, Heald (1967) suggested that plant selection variations and differing isolates of M. hapla may account for such inconsistencies. In an attempt to correlate galling inconsistencies with differing nematode isolates, Bernard et al. (1994) compared galling induced by one isolate of M. incognita from Tennessee and two isolates of M. hapla from Tennessee and North Carolina on numerous Ilex spp. cultivars. While all Ilex spp. cultivars supported reproduction of the M. incognita isolate, the M. hapla isolates differed in their pathogenicity on the various Ilex spp. cultivars, inducing variable gall numbers and sizes. This variability was attributed to geographic and host-adaptive variability of the M. hapla isolates. In examining the effect of root-knot nematode pathogenicity on shoot growth, Nemec and Struble (1968) found no differences in shoot growth between infected and non-infected plants after eight weeks in a greenhouse.

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24 Host plant resistance (HPR) to Meloidogyne spp. may involve several mechanisms, including hypersensitivity reactions, delay in Meloidogyne spp. juvenile maturation, reduction in numbers of giant cells, and reduction in cortical hypertrophy (Nemec and Morrison, 1972). Botanically, however, HPR also infers to the plants ability to withstand, lessen, oppose, or overcome nematode infections, while an immune plant suffers no injury (Rohde, 1972). The increasing availability of plant germplasm and advances in plant science technology may allow for the development of HPR for plant-parasitic nematode control. Such advancements are crucial with the progressing withdrawal of chemical nematicides (Roberts, 1992). An ornamental plant used as a rootstock in a Meloidogyne spp. infested area should exhibit resistance to infection by these pathogens. For example, several Hibiscus rosa-sinensis cultivars with the potential for use as rootstocks based on their reaction to inoculations with M. javanica and M. incognita have been identified (McSorley and Marlatt, 1983). Codiaeum variegatum Codiaeum variegatum (L.) Blume, commonly referred to as croton, is an evergreen, glabrous shrub in the family Euphorbiaceae that grows 2 m tall. Its petioled leaves are spirally-arranged, alternate, simple and entire or lobed, ovate-lanceolate and are marked with yellow, white, or red variegation (Bailey, 1958; Gilman, 1999a). While flowers occur in racemes from leaf axils, they are inconspicuous (Watkins and Sheehan, 1977). Codiaeum variegatum thrives in USDA zones 9b to 11, where large specimens may be used as foundation plants, hedges, and borders, while smaller specimen may be used as annual bedding plants. Additionally, it is commonly used as an interior foliage plant throughout the United States. Codiaeum variegatum is commercially propagated

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25 mostly through tip cuttings, and to a lesser extent through air layering and seed propagation. Potting media used for croton production should be light, slightly acidic, and possess good drainage. In the landscape, crotons grow best in well-draining sandy soils incorporated with organic matter. Croton producers must balance fertilizing and light requirements to produce a saleable plant, as too much fertilizer for the amount of light produces a leggy, weak plant, while too little fertilizer produces a woody, stunted plant. While most croton cultivars require high light, many cultivars that tolerate less light, including C. variegatum cv. Gold Dust, are available, and the market for indoorthriving cultivars is increasing. Insect pests of crotons include Tetranychus spp. (spider mites), Pseudococcus longispinus (long tailed mealybug), Planococcus citri (citrus mealybug), Maconellicoccus hirsutus (pink hibiscus mealybug), Ferrisia virgata (striped mealybug), Polyphagotarsonemus latus (broad mite), and numerous scale species. While stem galls, rots, rusts, scabs, and blights are sometimes encountered on crotons, they are not serious problems (Gilman, 1999a; Henny et al., 1991; Reeves and Bell, 1988; Stamps and Osborne, 2003). Nematode parasites of croton include Paratylenchus spp. (Ibrahim and Al-Yahya, 2002) and Hoplolaimus spp. (Sher, 1954). Salvia leucantha Salvia leucantha Cav. is a 1-m-tall herbaceous perennial shrub, commonly known as Mexican sage, which belongs to the family Labiatae. This plant possesses cylindrical, somewhat tapering white branches that are covered with fine wool. Its leaves, silver-gray colored and arranged in a whorl, possess a short petiole and are 5 to 15 cm long, lanceolate-linear, crenate, and are pubescent above and tomentose beneath. The plant exhibits many flowers that occur on 15 to 25-cm-long racemes that appear in the summer.

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26 Flowers possess a funnel-shaped, violet to lavender calyx and a white or purple corolla, to 2 cm long (Bailey, 1958; Gilman and Marshall, 1999; Anonymous, 2000). Salvia leucantha thrives in USDA zones 7 to 11 and is used in mass plantings, borders, and edging. In zones 9 to 11, flowers persist longer than in zones 7 and 8. The plant grows well in full sun and prefers well-drained, rich, sandy soils. Mexican sage is commercially propagated through cuttings and divisions (Gilman and Marshall, 1999). Buhrer (1938) reported on Meloidogyne spp. parasitizing S. leucantha On the contrary, the plant was reported resistant to M. incognita and M. javanica in India (Krishnaprasad, 1979). Liriope muscari Liriope muscari Bailey, commonly known as lilyturf, is a grass-like herbaceous perennial in the family Ruscaceae (Gilman, 1999b; Judd, 2003). The plant possesses a short, thick rhizome by which it spreads. The stemless, dark green leaves are stiff, long, linear, with an acute tip, and do not exceed the floral scapes, which are 25 to 70 cm high and occur in spring and summer. Scapes possess 10 or more whorls of up to seven flowers, each 0.6 to 0.7 cm across (Bailey, 1958). The plant grows in tufts of various heights, depending on the cultivar. Liriope muscari is often used as a ground cover or for edging, and prefers fertile, well-drained soils. Commercially, this plant is propagated though divisions and seldom through seed (Watkins and Sheehan, 1977). It is also often propagated through tissue culture. Liriope muscari thrives in USDA zones 6 to 10 in shady areas, but tolerates full sun except in the hottest areas of the south, as severe leaf tip burn occurs in such conditions. Various cultivars of lilyturf are available, including variegated foliage ( L. muscari cv. Variegata) and white and purple-colored flower varieties. Liriope muscari cv. Evergreen Giant grows to 30-cm-high and spreads slower

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27 than other cultivars. Recently, a leaf and crown rot disease, caused by Phytophthora palmivora has taken its toll on L. muscari in landscapes and nurseries, to the extent that L. muscari cv. Evergreen Giant may lose its popularity in the landscape industry (Leahy and Davison, 1999). Pittosporum tobira Pittosporum tobira Ait., also known as Japanese pittosporum, is a 3 to 6-m-high flowering evergreen shrub to small tree in the family Pittosporaceae. Its whorled, thick, leathery leaves are 5 to 10 cm long and are glabrous, with revolute margins. Japanese pittosporum is winter-flowering, possessing 1-cm-long white, fragrant flowers in terminal umbels. Pittosporum tobira is often grown as a shrub for foundations, hedges, mass plantings, as a screen, in planter boxes, and may also be trained as a small tree (Bailey, 1958; Gilman, 1999c, 1999d; Stamps, 2002). Furthermore, salt tolerance makes it useful in seaside plantings, where it is widely used as a hedge and windbreaker (Rinallo and Bennici, 1989). Since it is damaged by temperatures below -6 C, P. tobira is restricted to USDA zones 8 to 10. It prefers fertile, slightly acidic soils, is fairly drought tolerant, and requires minimal fertilization for optimal growth. Pittosporum tobira establishes well in partial shade to full-sun conditions (Stamps, 2002). Commercially, this plant is propagated through semi-hardwood cuttings under mist and seldom by seed (Bailey, 1958; Watkins and Sheehan, 1977). Several cultivars that have either been produced or assigned to the species include P. tobira cv. Compacta, Variegata, and Wheelers Dwarf, of which P. tobira cv. Variegata possesses thinner leaves than the type species that are variegated with white, and is heavily cultivated in Florida and California, mainly for landscape use and for floral designs (Bailey, 1928; Stamps, 1987, 2002). Diseases that

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28 often occur on P. tobira include angular leaf spot, caused by Cercospora pittospori Rhizoctonia aerial blight, caused by Rhizoctonia ramicola or R. solani southern blight, caused by Sclerotium rolfsii root rots, caused by Pythium spp. and Rhyzoctonia spp., and dieback, caused by Agrobacterium spp., Diaporthe spp., Diplodia spp., Nectriella spp., homopsis spp., and Sphaeropsis spp. Other, less common diseases include Alternaria leaf spot, caused by Alternaria tenuissima mushroom root rot, caused by Armillariella tabescens corticium limb blight, caused by Corticium salmonicolor rough bark disease, which may be caused by a virus, and several others (Chase and Simone, 2001). Pittosporum tobira is also a host for the tomato spotted wilt virus (Gera et al., 2000). This plant often exhibits symptoms of magnesium, iron, manganese, and copper deficiencies, especially when cultivated in high-pH soils (Dehgan, 1998). Insect pests often encountered on P. tobira include Aphis gossypii (melon aphid) and Icerya purchasi (cottony cushion scale) (Hamon and Fasulo, 1998). Buhrer (1938) observed Meloidogyne spp. infected P. tobira plants, and M. arenaria was detected on a declining hedge of P. tobira in Florida (Bureau of Nematology, 1989). The ectoparasitic nematode Belonolaimus longicaudatus has been associated with P. tobira cv. Variegatum (syn. P. tobira cv. Variegata) decline (Rhoades, 1989). Odontonema cuspidatum Odontonema cuspidatum (Nees) Kuntze (syn. O. strictum (Nees) Kuntze), commonly referred to as firespike, is a herbaceous perennial in the family Acanthaceae, which is hardy in USDA zones 8B to 11. It is a glabrous, erect shrub, 1 to 2-m-high. that forms clumps that generate from root suckers. It possesses simple, opposite, acuminate, short-petioled leaves that are entire and undulate, 10 to 30 cm long. Its tubular flowers

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29 are bright red, pink, white, or lavender, up to 2.5 cm long, and occur in long racemes. Flowers occur year-round in tropical climates and in the fall in temperate climates. This erect, compact shrub is often used in mass plantings, as a background plant, or a hedge, and often attracts butterflies and hummingbirds. Following the first frost, it dies to the ground, but comes back in the spring. For best flower development and persistence, O. cuspidatum is planted in full sun in fertile, sandy soil. This plant is commercially propagated by cuttings and divisions. Aside from Pseudococcus spp. (mealy bugs), no pests or pathogens have been reported on this plant (Bailey, 1958; Francis, 2004; Gilman and Delvalle, 1999; Watkins and Sheehan, 1977). Musa acuminata ssp. zebrina Musa acuminata Colla. ssp. zebrina Van Houtte ex Planch cv. Rowe Red (syn. Musa sumatrana Baccari cv. Rowe Red) is a cultivated banana in the family Musaceae that is used as a decorative ornamental. Descriptions surrounding this species have been erroneous and confused due to the age of specimens at the time of description. Additionally, the description of wrongly-named specimen, as evident by differing floral and fruit characters among the described specimen documented, may have contributed to the taxonomic confusion (Cheesman, 1985). Recent evidence has led to the reclassification of this plant as M. acuminata ssp. zebrina The cultivar Rowe Red is not described in the literature, and may have been designated to the species by its cultivators. Musa acuminata ssp. zebrina is rhizomatous and possesses red-ornamented green pseudostems 2.5 m high, which are formed via sheathing leaf bases, and continuously sucker from pseudostem bases. Its leaves are spirally arranged, 1.8 m long and 0.5 m wide, rounded at the base, glaucous, purple beneath, and irregularly purple-patched

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30 above, with a 0.3-m-long petiole. The floral spikes droop slightly, and the rachis on which they appear is pubescent. The fruit is inedible, dry, cylindrical and curved, 7 cm long and 1.3 cm diam. (Baker, 1893; Cheesman, 1985; Griffiths, 1994; Ricker, 1937). Musa spp. thrive through USDA zone 9, as long as they are protected in northern regions of zone 9b. Plants in this genus prefer moist, fertile soils and ample fertilization. Musa acuminata spp. zebrina cv. Rowe Red may be propagated via sucker divisions, but is produced commercially via tissue culture (Dehgan, 1998). Most Musa spp. are susceptible to Black Sigatoka, caused by Mycosphaerella fijiensis Yellow Sigatoka, caused by Mycosphaerella musicola and Panama Disease, caused by Fusarium oxysporum f. sp. cubense Plant-parasitic nematode pathogens of Musa spp. include M. incognita M. javanica M. arenaria and other Meloidogyne spp., Radopholus simillis Pratylenchus goodeyi P. coffeae and Helicotylenchus multicinctus (Adiko, 1988; De Waele and Davide, 1998; Jaizme-Vega et al., 1997; Stoffelen et al., 2000). Objectives The objectives of this research were to: 1. Evaluate the host status of several woody and perennial ornamental plants to Meloidogyne incognita race 2, M. javanica M. arenaria race 1, and M. mayaguensis in separate experiments. 2. Differentiate malate dehydrogenase and esterase phenotypes of Meloidogyne spp. that were collected from infected ornamental plants in Florida using PAGE. 3. Evaluate the usefulness of PAGE in differentiating Meloidogyne spp. for routine extension diagnostic purposes.

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31 CHAPTER 2 REPRODUCTION OF FOUR MELOIDOGYNE SPP. ON SEVERAL SPECIES OF PERENNIAL ORNAMENTAL PLANTS Introduction Root-knot nematodes ( Meloidogyne spp.) are the most damaging group of plantparasitic nematodes to ornamental plants in Florida (McSorley and Dunn, 1989). As serious pathogens of many woody ornamental species, root-knot nematodes limit productivity by damaging numerous nursery crops directly and by forming disease complexes with certain soil-borne fungal pathogens (Barker and Benson, 1977; Benson and Barker, 1985; Nigh, 1972; Santamour and Riedel, 1993; Walker and Melin, 1998b; Zarina and Abid, 1995). Furthermore, since root-knot nematode populations often thrive and cause damage on perennial hosts for many months and years, damage threshold levels do not apply for such plants (LaMondia, 1995). Symptoms associated with root-knot nematode infection include root galls and root rots, shoot chlorosis, stunted growth, and other symptoms commonly associated with nutritional deficiencies (Bala and Hosein, 1996; Bird, 1974; Misra et al., 2002; Santo and Lear, 1976; Zarina and Abid, 1995). Such symptoms are often associated with general decline (Nigh, 1972), poor yield, and wilt diseases (Rajendran et al., 1975). Published work on the susceptibility of woody ornamentals to Meloidogyne spp. is limited. In Florida, several authors (Giblin-Davis et al., 1992; Lehman, 1984a, 1984b; Lehman and Barnard, 1982; McSorley and Dunn, 1989, 1990; McSorley and Marlatt,

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32 1983; Stokes, 1982) reported on the pathogenicity of a number of Meloidogyne spp. to several perennial ornamental plants. Other reports of root-knot infection of ornamentals from Alabama (Heald, 1967), Arizona (Nigh, 1972), California (Santo and Lear, 1976; Viglierchio, 1979), Connecticut (LaMondia, 1995, 1996, 1997), Georgia (Heald, 1967; Walker and Melin, 1998a, 1998b), New Jersey (Davis and Jenkins, 1960), North Carolina (Barker et al., 1979; Barker and Benson, 1977; Benson and Barker, 1982; Haasis et al., 1961), Oklahoma (Nemec and Morrison, 1972; Nemec and Struble, 1968), Tennessee (Bernard and Witte, 1987; Bernard et al., 1994; Niblack and Bernard, 1985), and Washington, DC (Santamour, 1992; Santamour and Riedel, 1993, 1995) have been published. Compared to research on the pathogenicity of Meloidogyne spp. on agronomic crops, and taking into account the vast array of perennial species cultivated, research on the susceptibility of woody and perennial ornamental plants to root-knot nematodes is minimal. The objectives of these studies were to evaluate the host status of several woody and perennial ornamental plants to Meloidogyne incognita race 2, M. javanica M. arenaria race 1, and M. mayaguensis in separate experiments. The experiments were carried out in a controlled environmental chamber (growth room) and a greenhouse at the University of Florida. The plant species evaluated were Liriope muscari (Lilyturf) cv. Evergreen Giant, Pittosporum tobira (Pittosporum) cv. Variegata, Odontonema cuspidatum (Firespike), Codiaeum variegatum (Croton) cv. Gold Dust, Musa acuminata ssp. zebrina (ornamental banana) cv. Rowe Red, and Salvia leucantha (Mexican Sage). While cultivars of Salvia leucantha do not exist, two forms, one with a purple corolla and another with a white corolla, were tested in the greenhouse experiment.

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33 Materials and Methods General Cultivation Practices Planting Experimental plants were planted as liners in 15.24-cm plastic (Model AZE0600, ITML Horticultural Products, Ontario, CA) and clay pots for growth room and greenhouse experiments, respectively, each containing 800 cm3 of planting media. Liners were obtained from commercial propagators (Table 2-1). Several liners were tested for the presence of plant-parasitic nematodes by the rapid centrifugal-flotation technique (Jenkins, 1964) prior to planting. Planting media Experiments were identical with respect to the soil mixture used for planting experimental plants and maintaining nematode inocula. The soil mixture was a 2:1 ratio of sand and potting mix (Jungle Growth Professional Growers Mix, Statham, GA), respectively. The medium was tested for the presence of plant-parasitic nematodes via the rapid centrifugal-flotation method prior to the initiation of each experiment. Treatments Five root-knot nematode species were used in the growth room and greenhouse experiments. The nematodes were: (i) M. incognita race 2, (ii) M. javanica (iii) M. arenaria race 1, (iv) M. mayaguensis and (v) non-inoculated control. Exceptions include the 5 June 2003-inoculated Liriope muscari cv. Evergreen Giant and Pittosporum tobira cv. Variegata trials, in which no non-inoculated control treatments were included. Meloidogyne spp. The Meloidogyne spp. utilized in these experiments were originally obtained from J. A. Brito, Florida Department of Agriculture and Consumer Services, Division of Plant Industry, Gainesville, FL. They originated from single egg

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34 Table 2-1. Crop and source of liners used for growth room and greenhouse experiments. Crop Source Liriope muscari cv. Evergreen Giant Agri-Starts III, Inc., Eustis, FL Pittosporum tobira cv. Variegata Jons Nursery, Inc., Eustis, FL Liner Source, Inc., Mount Dora, FL Salvia leucantha purple corolla Hatchett Creek Farms, LLC, Gainesville, FL Robrick Nursery, Inc., Hawthorne, FL Salvia leucantha white corolla Yoder Brothers, Inc., Lancaster, PA Odontonema cuspidatum Robrick Nursery, Inc., Hawthorne, FL Codiaeum variegatum cv. Gold Dust Parrish Nurseries, Inc., Parkland, FL Musa acuminata ssp. zebrina cv. Rowe Red Agri-Starts I, Inc., Apopka, FL

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35 masses, and their species designation was confirmed using perineal patterns and isozyme phenotypes, resolved on polyacrylamide gel slabs following electrophoresis. Meloidogyne spp. Extraction and Inoculation Meloidogyne spp. egg and second-stage juvenile (J2) inocula were extracted from Lycopersicon esculentum Mill. (tomato) cv. Rutgers by the sodium hypochlorite (NaOCl) procedure (Hussey and Barker, 1973), via the shaking of infected roots in 0.53% NaOCl (Regular Ultra Bleach, Publix Super Markets, Lakeland, FL) solution for 30 seconds, followed by the immediate rinsing of the suspension with 10 liters of water. Meloidogyne spp. eggs and J2 were inoculated onto test and tomato plants for experimental and inoculum-increase purposes on the same day that the eggs were extracted from infected tomato roots. Meloidogyne spp. inocula were transported to the growth room or greenhouse in sealed 500-ml Erlenmeyer flasks. An aquarium air pump equipped with a 1-ml, 22.8-cm-long, disposable glass Pasteur pipet that was inserted into the Erlenmeyer flask, was used to keep the root-knot nematode inocula thoroughly and evenly suspended. Five thousand eggs and J2 were pipetted into three equidistant holes approximately 3-cm-deep in the pre-moistened soil surrounding the base of each plant. Test tomato plants were included in every experiment as a control for inoculum viability. Immediately after inoculation, the holes were covered with the same soil mixture used throughout the experiment. The bench area used for inoculation was thoroughly disinfected with 6.0% regular ultra bleach, and all pipets were replaced between inoculations of the respective Meloidogyne spp. All pots were kept on inverted clay drainage saucers for the duration of the experiment to avoid Meloidogyne spp. contamination among pots.

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36 Fertilization and watering In all the experiments, a solution of 0.21 g N, 0.09 g P, and 0.17 g K (0.21 g 20-20-20 soluble fertilizer, Grace-Sierra Horticultural Products, Milpitas, CA) in 200 ml water was applied to each pot weekly and plants were watered as needed. Pesticides Tetranychus spp. (spider mites) were encountered parasitizing P. tobira in these experiments. Bifenthrin was sprayed at a rate of 0.13 ml a.i./liter when needed. In addition, all tomato plants and all greenhouse-grown crops in these experiments were treated with imidacloprid at a rate of 12.5 mg a.i. per pot, for Bemisia spp. (whitefly). Photoperiod and temperature Growth room light was provided by 400-W general lighting metal halide lamps (Osram Sylvania, Danvers, MA), directed toward the experimental plants, which were situated on 1-m-tall benches. Lights were suspended 1 m above the bench tops, and light intensity was recorded at 256.18 52.82 mol s-1 m-2 at a distance of 35 cm above the bench top using a photometer (LI-COR, Model LI-189, Lincoln, NE). Plants were on a 14-hr light cycle, from 7:00 PM to 9:00 AM. Air and media temperatures in the growth room were maintained using an air conditioner and were measured using a standard thermometer at 26 to 32 C and 24 to 26 C, respectively. Experimental Design Growth room and greenhouse experiments were arranged in a randomized complete block design, with six and three replications in the growth room and greenhouse, respectively. Each experiment was conducted twice in the growth room and once in the greenhouse. While growth room experiments were harvested approximately

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37 two months post-inoculation, greenhouse experiments were conducted over five months or longer, as outlined in Table 2-2. The S. leucantha growth room trials differed with respect to the plant flower-color forms used. Plants used in the first growth room trial possessed a mixture of purpleand white-colored corolla S. leucantha plants, while plants used in the second growth room trial were limited to the purple-corolla form. Experimental Plants Processing and Meloidogyne spp. Egg Extraction and Counting All experiments were processed by their respective replications. While entire root systems were processed in the growth room experiments, 10 g of each root system were evaluated for plants in greenhouse experiments due to the extensive root system of these plants. Egg numbers from greenhouse-experimental plants were calculated based on total root weight of respective plants. Experimental plants were moved, by replication, from the growth room or greenhouse to the nematode assay laboratory at the University of Florida. Each plant was completely processed separately prior to processing of the remaining plants in the respective replication. Each plant was removed from the plastic pot and the soil surrounding its roots was gently shaken into a container. The above-ground portion of each plant was then cut and placed into a pre-weighed 25.4-cm x 33.0-cm manila envelope (Sparco, Moorestown, NJ), which was placed in a 70 C oven. Drying times were determined by weighing envelopes every 24 hours until no further weight change was detected. The remaining root system was then immersed in water to remove any adhering soil. The soil-free root system was patted dry and fresh root system weights and gall ratings (Taylor and Sasser, 1978) were determined and recorded. Eggs were extracted using 0.53% NaOCl (Hussey and Barker, 1973) by a procedure modified by Boneti and Ferraz

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38 Table 2-2. Crops, experimental sites, liner planting dates, inoculation dates, and study lengths for all crops in the growth room and greenhouse Meloidogyne spp. studies carried out at the University of Florida during 2003 to 2005. Crop Experimental site Liner planting date Inoculation date Study length (days) Liriope muscari cv. Evergreen Giant Ca 22 May 2003 5 June 2003 65 Liriope muscari cv. Evergreen Giant C 26 August 2003 9 September 2003 63 Liriope muscari cv. Evergreen Giant G 19 February 2004 4 March 2004 354 Pittosporum tobira cv. Variegata C 22 May 2003 5 June 2003 70 Pittosporum tobira cv. Variegata C 26 August 2003 9 September 2003 67 Pittosporum tobira cv. Variegata G 19 February 2004 4 March 2004 336 Salvia leucantha purple and white corolla mix C 25 October 2003 8 November 2003 68 Salvia leucantha purple corolla C 15 January 2004 29 January 2004 70 Salvia leucantha purple corolla G 19 February 2004 4 March 2004 154 Salvia leucantha white corolla G 19 February 2004 4 March 2004 182 Odontonema cuspidatum C 25 October 2003 8 November 2003 81

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39Table 2-2. Continued Crop Experimental site Liner planting date Inoculation date Study length (days) Odontonema cuspidatum C 15 January 2004 29 January 2004 78 Odontonema cuspidatum G 19 February 2004 4 March 2004 263 Codiaeum variegatum cv. Gold Dust C 22 June 2004 6 July 2004 79 Codiaeum variegatum cv. Gold Dust C 8 September 2004 22 September 2004 96 Codiaeum variegatum cv. Gold Dust G 22 June 2004 6 July 2004 239 Musa acuminata ssp. zebrina cv. Rowe Red C 22 June 2004 6 July 2004 72 Musa acuminata ssp. zebrina cv. Rowe Red C 8 September 2004 22 September 2004 106 Musa acuminata ssp. zebrina cv. Rowe Red G 22 June 2004 6 July 2004 205 aC = Growth-room, G = Greenhouse

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40 (1981). One ml of solution was drawn out of each treatment and put into a 1-ml, 48division equine egg parasite counting slide (Advanced Equine Products, Issaquah, WA), which was counted in its entirety. Three 1-ml aliquots were counted per treatment, and the average of the three counts was used for statistical analysis. Statistical Analysis Data were subjected to analysis using Analysis of Variance (ANOVA). Mean comparisons among the treatments were performed with Duncans multiple range test using SAS software (SAS Institute, Cary, NC). Host Status Classification The host status of the tested perennials was determined based on the reproduction factor (Rf), which was calculated by dividing the final root-knot nematode density per plant (Pf) by the inoculated root-knot density of 5,000 eggs and J2 (Pi). A Rf 1.0 was designated as a good host, 1.0 Rf 0.1 a poor host, and Rf < 0.1 a nonhost (Sasser et al., 1984). Results Liriope muscari cv. Evergreen Giant Results for the two L. muscari cv. Evergreen Giant growth room trials were heterogeneous and are therefore presented separately (Tables 2-3 and 2-4). In the first trial, there were no differences ( P 0.05) among treatments for root-gall index, root weight, or dry shoot weight. Differences ( P 0.05) were observed among treatments for number of eggs per plant and number of eggs per g of roots, where fewer eggs were produced by M. arenaria than by the other three species. In the second trial, there were no differences ( P 0.05) among treatments for root-gall index, root weight, or shoot dry

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41Table 2-3. Treatments, root-gall indexes, root weights, number of eggs per plant, number of eggs per g of roots, and dry shoot weights from the first Liriope muscari cv. Evergreen Giant growth room trial. Treatment Root-gall index Root Weight (g) Mean number of eggs per plant Mean number of eggs per g roots Dry Shoot Weight (g) Control ----------M. incognita race 2 1.17 0.82a 35.95 19.71 31,920.00 44,748.39 ab 848.43 1,047.04 a 7.62 4.26 M. javanica 1.33 1.63 36.33 23.16 19,897.78 36,366.07 a 492.63 608.77 a 8.25 3.17 M. arenaria race 1 0.83 0.82 37.67 20.08 377.78 1,131.08 b 9.11 26.02 b 8.30 3.70 M. mayaguensis 1.00 0.00 37.73 14.28 13,822.22 12,905.42 a 373.04 320.68 a 7.47 4.23 a0 = no galls; 1 = 1 to 2 galls; 2 = 3 to 10 galls; 3 = 11 to 30 galls; 4 = 31 to 100 galls; 5 = more than 100 galls per root sy stem (Taylor and Sasser, 1978). bData are means of six replications. Means in columns followed by a common letter are not different ( P 0.05) according to Duncans multiple range test. Egg numbers were log-transformed prior to data analysis, but non-transformed numbers are shown.

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42Table 2-4. Treatments, root-gall indexes, root weights, number of eggs per plant, number of eggs per g of roots, and dry shoot weights from the second Liriope muscari cv. Evergreen Giant growth room trial. Treatment Root-gall index (0-5) Root Weight (g) Mean number of eggs per plant Mean number of eggs per g roots Dry Shoot Weight (g) Control 0.00 0.00a 39.18 14.76 0.00 0.00 cb 0.00 0.00 c 12.77 4.22 M. incognita race 2 0.00 0.00 40.00 21.73 231.11 418.14 b 5.36 8.27 b 10.20 3.51 M. javanica 0.33 1.03 37.95 16.83 684.44 829.44 a 20.93 33.18 a 9.73 5.94 M. arenaria race 1 0.00 0.00 38.05 22.35 8.89 27.54 c 0.26 0.81 c 10.63 6.35 M. mayaguensis 0.00 0.00 36.70 24.73 1,000.00 833.61 a 30.51 38.39 a 10.93 8.26 a0 = no galls; 1 = 1 to 2 galls; 2 = 3 to 10 galls; 3 = 11 to 30 galls; 4 = 31 to 100 galls; 5 = more than 100 galls per root sy stem (Taylor and Sasser, 1978). bData are means of six replications. Means in columns followed by a common letter are not different ( P 0.05) according to Duncans multiple range test. Egg numbers were log-transformed prior to data analysis, but non-transformed numbers are shown.

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43 weight. Differences ( P 0.05) were observed among treatments for number of eggs per plant and number of eggs per g of roots, where the number of eggs produced by M. arenaria was not different ( P 0.05) from the nematode control. Furthermore, M. javanica and M. mayaguensis produced more eggs than M. incognita which produced more eggs than M. arenaria In the greenhouse experiment (Table 2-5), differences ( P 0.05) were observed across all parameters. In this experiment, M. incognita and M. mayaguensis produced more galls than the other treatments, and the control treatment had a significantly higher root weight than all other treatments. Meloidogyne incognita M. javanica and M. mayaguensis produced more eggs per plant than M. arenaria and the control treatment, from which no eggs were recovered. However, M. incognita and M. mayaguensis produced more eggs per g of roots than M. javanica which produced more eggs per g of roots than M. arenaria Finally, the control and M. incognita treatments had higher dry shoot weights than the M. javanica and M. arenaria treatments, and the M. mayaguensis treatment dry shoot weights were not different ( P 0.05) than any of the other treatment. The data suggest that L. muscari cv. Evergreen Giant is a good host to M. incognita race 2 (Rf = 97.1), M. javanica (Rf = 16.6), and M. mayaguensis (Rf = 91.0), and a poor host to M. arenaria race 1 (Rf = 0.12). Pittosporum tobira cv. Variegata Results for the two P. tobira cv. Variegata growth room trials were heterogeneous and are therefore presented separately in tables 2-6 and 2-7. The results from the greenhouse experiment are presented in table 2-8. Root galls were not observed on any of the plants in the three studies. No differences ( P 0.05) were observed among treatments for root

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44Table 2-5. Treatments, root-gall indexes, root weights, number of eggs per plant, number of eggs per g of roots, and dry shoot weights from the Liriope muscari cv. Evergreen Giant greenhouse experiment. Treatment Root-gall index (0-5) Root Weight (g) Mean number of eggs per plant Mean number of eggs per g roots Dry Shoot Weight (g) Control 0.00 0.00 bab 141.50 55.65 a 0.00 0.00 b 0.00 0.00 c 30.80 3.83 a M. incognita race 2 5.00 0.00 a 104.33 15.10 b 485,507.38 184,451.74 a 4,646.22 1,555.40 a 29.33 10.05 a M. javanica 0.00 0.00 b 70.93 8.11 b 83,099.47 283,116.92 a 1,100.44 3,742.78 b 21.40 4.40 b M. arenaria race 1 0.00 0.00 b 81.70 34.01 b 597.33 2,069.22 b 6.22 1.55 c 19.70 4.13 b M. mayaguensis 5.00 0.00 a 82.33 37.12 b 454,887.02 412,521.08 a 5,467.56 3,903.75 a 25.20 17.97 ab a0 = no galls; 1 = 1 to 2 galls; 2 = 3 to 10 galls; 3 = 11 to 30 galls; 4 = 31 to 100 galls; 5 = more than 100 galls per root sy stem (Taylor and Sasser, 1978). bData are means of three replications. Means in columns followed by a common letter are not different ( P 0.05) according to Duncans multiple range test. Egg numbers were log-transformed prior to data analysis, but non-transformed numbers are shown.

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45Table 2-6. Treatments, root-gall indexes, root weights, number of eggs per plant, number of eggs per g of roots, and dry shoot weights from the first Pittosporum tobira cv. Variegata growth room trial. Treatment Root-gall index (0-5) Root Weight (g) Mean number of eggs per plant Mean number of eggs per g roots Dry Shoot Weight (g) Control ----------M. incognita race 2 0.00 0.00a 11.67 6.91 13.33 44.62 abb 1.40 5.07 ab 11.67 2.44 M. javanica 0.00 0.00 12.62 4.63 53.33 168.65 a 4.32 13.10 a 12.67 2.33 M. arenaria race 1 0.00 0.00 12.10 8.07 13.33 44.62 ab 0.79 2.51 ab 12.05 3.63 M. mayaguensis 0.00 0.00 12.23 7.68 0.00 0.00 b 0.00 0.00 b 11.83 3.69 a0 = no galls; 1 = 1 to 2 galls; 2 = 3 to 10 galls; 3 = 11 to 30 galls; 4 = 31 to 100 galls; 5 = more than 100 galls per root sy stem (Taylor and Sasser, 1978). bData are means of six replications. Means in columns followed by a common letter are not different ( P 0.05) according to Duncans multiple range test. Egg numbers were log-transformed prior to data analysis, but non-transformed numbers are shown.

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46Table 2-7. Treatments, root-gall indexes, root weights, number of eggs per plant, number of eggs per g of roots, and dry shoot weights from the second Pittosporum tobira cv. Variegata growth room trial. Treatment Root-gall index (0-5) Root Weight (g) Mean number of eggs per plant Mean number of eggs per g roots Dry Shoot Weight (g) Control 0.00 0.00a 11.57 3.78 0.00 0.00 0.00 0.00 11.42 1.29 M. incognita race 2 0.00 0.00 12.22 4.16 71.11 322.95 5.19 23.57 11.82 1.34 M. javanica 0.00 0.00 11.65 3.05 8.89 43.55 0.67 3.30 11.07 4.05 M. arenaria race 1 0.00 0.00 11.82 1.24 8.89 43.55 0.74 3.63 11.58 1.27 M. mayaguensis 0.00 0.00 10.08 2.99 31.11 78.50 3.56 9.28 10.37 1.14 Data are means of six replications. No differences ( P 0.05) were observed according to Duncans multiple range test. a0 = no galls; 1 = 1 to 2 galls; 2 = 3 to 10 galls; 3 = 11 to 30 galls; 4 = 31 to 100 galls; 5 = more than 100 galls per root sy stem (Taylor and Sasser, 1978).

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47Table 2-8. Treatments, root-gall indexes, root weights, number of eggs per plant, number of eggs per g of roots, and dry shoot weights from the Pittosporum tobira cv. Variegata greenhouse experiment. Treatment Root-gall index (0-5) Root Weight (g) Mean number of eggs per plant Mean number of eggs per g roots Dry Shoot Weight (g) Control 0.00 0.00a 24.83 14.61 0.00 0.00 0.00 0.00 45.10 19.08 M. incognita race 2 0.00 0.00 23.13 5.32 0.00 0.00 0.00 0.00 49.47 8.32 M. javanica 0.00 0.00 26.87 8.26 0.00 0.00 0.00 0.00 46.37 6.84 M. arenaria race 1 0.00 0.00 25.30 23.30 0.00 0.00 0.00 0.00 43.40 6.54 M. mayaguensis 0.00 0.00 13.80 0.87 0.00 0.00 0.00 0.00 35.03 9.47 Data are means of three replications. No differences ( P 0.05) were observed according to Duncans multiple range test. a0 = no galls; 1 = 1 to 2 galls; 2 = 3 to 10 galls; 3 = 11 to 30 galls; 4 = 31 to 100 galls; 5 = more than 100 galls per root sy stem (Taylor and Sasser, 1978).

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48 gall index, root weight, or dry shoot weight in either of the growth room trials or the greenhouse experiment. There were differences ( P 0.05) among treatments for number of eggs per plant and number of eggs per g of roots in the first growth room trial, where M. javanica produced more eggs than M. mayaguensis which produced no eggs. However, in the second growth room trial and the greenhouse experiment no differences ( P 0.05) were found. The data suggests that P. tobira cv. Variegata is a nonhost to the Meloidogyne spp. isolates evaluated (Rf < 0.1). Salvia leucantha The results from the first and second purple-corolla S. leucantha growth room trials are heterogeneous, and are presented in tables 2-9 and 2-10, respectively. Rootknot nematode galls were not detected on any treatment in the first growth room trial (Table 2-9). Furthermore, no differences ( P 0.05) were observed among treatments for root weight or dry shoot weight. Differences ( P 0.05) were observed among treatments for the number of eggs per plant and the number of eggs per g of roots. The number of eggs per plant was not different ( P 0.05) between M. javanica and M. mayaguensis However, these two species produced more eggs than M. arenaria and M. incognita The number of eggs per g of roots produced by M. javanica was greater than the other species. Results for the second growth room trial (Table 2-10) indicate differences ( P 0.05) among treatments for root-gall index, number of eggs per plant, and number of eggs per g of roots. There were no differences ( P 0.05) among treatments for root weight and dry shoot weight in this trial. Root galls were only observed in the M. javanica treatment. The root-gall index for M. javanica was greater than that for the other species, in which root galls were not detected. For the

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49Table 2-9. Treatments, root-gall indexes, root weights, number of eggs per plant, number of eggs per g of roots, and dry shoot weights from the first Salvia leucantha growth room trial. Treatment Root-gall index (0-5) Root Weight (g) Mean number of eggs per plant Mean number of eggs per g roots Dry Shoot Weight (g) Control 0.00 0.00a 20.15 7.09 0.00 0.00 bb 0.00 0.00 b 14.35 2.59 M. incognita race 2 0.00 0.00 22.42 13.45 0.00 0.00 b 0.00 0.00 b 13.72 2.62 M. javanica 0.00 0.00 20.03 12.97 5,906.67 19,410.10 a 463.25 1,527.17 a 13.35 3.09 M. arenaria race 1 0.00 0.00 21.68 9.22 0.00 0.00 b 0.00 0.00 b 12.50 5.02 M. mayaguensis 0.00 0.00 18.68 9.16 2,053.33 9,207.13 a 94.03 402.60 b 11.75 8.95 a0 = no galls; 1 = 1 to 2 galls; 2 = 3 to 10 galls; 3 = 11 to 30 galls; 4 = 31 to 100 galls; 5 = more than 100 galls per root sy stem (Taylor and Sasser, 1978). bData are means of six replications. Means in columns followed by a common letter are not different ( P 0.05) according to Duncans multiple range test. Egg numbers were log-transformed prior to data analysis, but non-transformed numbers are shown.

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50Table 2-10. Treatments, root-gall indexes, root weights, number of eggs per plant, number of eggs per g of roots, and dry shoot weights from the second Salvia leucantha growth room trial. Treatment Root-gall index (0-5) Root Weight (g) Mean number of eggs per plant Mean number of eggs per g roots Dry Shoot Weight (g) Control 0.00 0.00 bab 16.05 4.03 0.00 0.00 d 0.00 0.00 d 9.17 2.57 M. incognita race 2 0.00 0.00 b 17.23 6.35 5,915.56 11,225.64 bc 326.02 568.49 bc 9.52 1.87 M. javanica 1.00 1.26 a 18.42 7.73 113,048.89 164,013.68 a 5,836.49 7,267.12 a 9.10 3.05 M. arenaria race 1 0.00 0.00 b 17.18 8.07 4,537.78 12,798.02 c 239.18 661.38 c 9.35 2.94 M. mayaguensis 0.00 0.00 b 16.57 4.88 36,333.33 126,118.27 b 2,298.54 8,382.65 b 9.20 2.10 a0 = no galls; 1 = 1 to 2 galls; 2 = 3 to 10 galls; 3 = 11 to 30 galls; 4 = 31 to 100 galls; 5 = more than 100 galls per root sy stem (Taylor and Sasser, 1978). bData are means of six replications. Means in columns followed by a common letter are not different ( P 0.05) according to Duncans multiple range test. Egg numbers were log-transformed prior to data analysis, but non-transformed numbers are shown.

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51 number of eggs per plant and the number of eggs per g of roots, the number of eggs produced followed the following order, from high to low: M. javanica M. mayaguensis and M. arenaria In the purple-corolla S. leucantha greenhouse experiment (Table 2-11), there were differences ( P 0.05) among treatments for root-gall index, number of eggs per plant, and number of eggs per g of roots. No differences ( P 0.05) were observed among treatments for root weight or shoot dry weight. No root-knot nematode galls were observed on the control and M. incognita treatments, but were observed on the M. javanica, M. arenaria and M. mayaguensis treatments. All four root-knot nematodes reproduced, and the number of eggs per plant and number of eggs per g of roots were not different ( P 0.05) among species. The data from the two growth room trials and the purple-corolla S. leucantha greenhouse suggest that this plant is a good host to the four Meloidogyne spp. evaluated. In the white-corolla S. leucantha greenhouse experiment (Table 2-12), there were differences ( P 0.05) among treatments for number of eggs per plant and number of eggs per g of roots. In both categories, M. incognita produced more eggs than M. mayaguensis Egg production was not observed in M. arenaria and M. javanica inoculated plants. No differences ( P 0.05) were observed among treatments for the root-gall index, root weight, or dry shoot weight categories. The data from this experiment suggests that the white-corolla S. leucantha is a good host to Meloidogyne incognita and M. mayaguensis

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52Table 2-11. Treatments, root-gall indexes, root weights, number of eggs per plant, number of eggs per g of roots, and dry shoot weights from the purple-corolla Salvia leucantha greenhouse experiment. Treatment Root-gall index (0-5) Root Weight (g) Mean number of eggs per plant Mean number of eggs per g roots Dry Shoot Weight (g) Control 0.00 0.00 bab 27.43 20.13 0.00 0.00 b 0.00 0.00 b 26.20 2.31 M. incognita race 2 0.00 0.00 b 22.63 7.58 162,494.58 112,613.54 a 7,050.67 2,469.93 a 22.97 13.46 M. javanica 1.83 1.53 a 23.67 11.01 447,239.47 1,014,030.26 a 16,539.56 32,408.80 a 28.17 3.52 M. arenaria race 1 1.33 3.06 a 32.83 20.99 159,567.56 181,554.78 a 4,521.78 3,312.55 a 25.40 10.67 M. mayaguensis 0.67 1.15 ab 27.17 7.62 227,586.04 416,191.95 a 9,284.44 19,449.29 a 25.57 8.31 a0 = no galls; 1 = 1 to 2 galls; 2 = 3 to 10 galls; 3 = 11 to 30 galls; 4 = 31 to 100 galls; 5 = more than 100 galls per root sy stem (Taylor and Sasser, 1978). bData are means of three replications. Means in columns followed by a common letter are not different ( P 0.05) according to Duncans multiple range test. Egg numbers were log-transformed prior to data analysis, but non-transformed numbers are shown.

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53Table 2-12. Treatments, root-gall indexes, root weights, number of eggs per plant, number of eggs per g of roots, and dry shoot weights from the white-corolla Salvia leucantha greenhouse experiment. Treatment Root-gall index (0-5) Root Weight (g) Mean number of eggs per plant Mean number of eggs per g roots Dry Shoot Weight (g) Control 0.00 0.00a 39.87 12.23 0.00 0.00 cb 0.00 0.00 c 27.17 5.19 M. incognita race 2 0.00 0.00 39.10 1.78 898,208.53 1,159,608.89 a 22,911.11 29,747.46 a 27.57 6.71 M. javanica 0.00 0.00 36.10 23.95 0.00 0.00 c 0.00 0.00 c 26.33 2.83 M. arenaria race 1 0.00 0.00 37.07 14.53 0.00 0.00 c 0.00 0.00 c 29.67 5.95 M. mayaguensis 0.33 1.15 37.63 19.86 256,939.64 192,705.08 b 7,000.89 2,825.53 b 26.97 4.92 a0 = no galls; 1 = 1 to 2 galls; 2 = 3 to 10 galls; 3 = 11 to 30 galls; 4 = 31 to 100 galls; 5 = more than 100 galls per root sy stem (Taylor and Sasser, 1978). bData are means of three replications. Means in columns followed by a common letter are not different ( P 0.05) according to Duncans multiple range test. Egg numbers were log-transformed prior to data analysis, but non-transformed numbers are shown.

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54 Odontonema cuspidatum Results for the two O. cuspidatum growth room trials were heterogeneous and are therefore presented separately in tables 2-13 and 2-14. The results from the greenhouse experiment are presented in table 2-15. Root galls were not observed on any of the plants in the three studies. No differences ( P 0.05) were observed among treatments for rootgall index, root weight, number of eggs per plant, number of eggs per g of roots, or dry shoot weight in either of the growth room trials or the greenhouse experiment. The data suggest that O. cuspidatum is a nonhost to the four Meloidogyne spp. evaluated. Musa acuminata ssp. zebrina Results for the two M. acuminata ssp. zebrina growth room trials were heterogeneous and are therefore presented separately in tables 2-16 and 2-17. Root galls and reproduction were observed for all Meloidogyne species in the three experiments. Differences ( P 0.05) among treatments for root weight or dry shoot weight were not detected in the first growth room trial (Table 2-16). The root galls and number of eggs per plant were not different ( P 0.05) among the root-knot nematode species. However, M. arenaria produced more eggs per g of roots than the other root-knot nematode species. In the second growth room trial (Table 2-17), M. mayaguensis produced more galls than M. javanica which produced more galls than M. arenaria and M. incognita Plants inoculated with M. mayaguensis had higher root weights than the other treatments, and M. arenaria and M. mayaguensis produced more eggs per plant and eggs per g of roots than M. javanica and M. incognita The dry shoot weight of the control treatment was greater than that of the root-knot nematode treatments.

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55Table 2-13. Treatments, root-gall indexes, root weights, number of eggs per plant, number of eggs per g of roots, and dry shoot weights from the first Odontonema cuspidatum growth room trial. Treatment Root-gall index (0-5) Root Weight (g) Mean number of eggs per plant Mean number of eggs per g roots Dry Shoot Weight (g) Control 0.00 0.00a 33.85 18.01 0.00 0.00 0.00 0.00 20.07 4.42 M. incognita race 2 0.00 0.00 32.22 9.12 0.00 0.00 0.00 0.00 19.00 1.62 M. javanica 0.00 0.00 29.78 21.93 4.44 21.77 0.44 2.13 18.58 5.79 M. arenaria race 1 0.00 0.00 33.42 22.44 0.00 0.00 0.00 0.00 18.95 5.80 M. mayaguensis 0.00 0.00 26.37 16.32 4.44 21.77 0.22 1.08 15.53 9.36 Data are means of six replications. No differences ( P 0.05) were observed according to Duncans multiple range test. a0 = no galls; 1 = 1 to 2 galls; 2 = 3 to 10 galls; 3 = 11 to 30 galls; 4 = 31 to 100 galls; 5 = more than 100 galls per root sy stem (Taylor and Sasser, 1978).

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56Table 2-14. Treatments, root-gall indexes, root weights, number of eggs per plant, number of eggs per g of roots, and dry shoot weights from the second Odontonema cuspidatum growth room trial. Treatment Root-gall index (0-5) Root Weight (g) Mean number of eggs per plant Mean number of eggs per g roots Dry Shoot Weight (g) Control 0.00 0.00a 11.57 3.78 0.00 0.00 0.00 0.00 16.22 6.90 M. incognita race 2 0.00 0.00 12.22 4.16 31.11 152.41 2.19 10.73 17.85 4.25 M. javanica 0.00 0.00 11.65 3.05 0.00 0.00 0.00 0.00 14.53 15.36 M. arenaria race 1 0.00 0.00 11.82 1.24 0.00 0.00 0.00 0.00 18.17 3.50 M. mayaguensis 0.00 0.00 10.08 2.99 0.00 0.00 0.00 0.00 18.63 2.27 Data are means of six replications. No differences ( P 0.05) were observed according to Duncans multiple range test. a0 = no galls; 1 = 1 to 2 galls; 2 = 3 to 10 galls; 3 = 11 to 30 galls; 4 = 31 to 100 galls; 5 = more than 100 galls per root sy stem (Taylor and Sasser, 1978).

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57Table 2-15. Treatments, root-gall indexes, root weights, number of eggs per plant, number of eggs per g roots, and dry shoot weights from the Odontonema cuspidatum greenhouse experiment. Treatment Root-gall index (0-5) Root Weight (g) Mean number of eggs per plant Mean number of eggs per g roots Dry Shoot Weight (g) Control 0.00 0.00a 96.40 18.20 0.00 0.00 0.00 0.00 44.03 5.64 M. incognita race 2 0.00 0.00 89.37 6.50 8.89 30.79 0.89 3.08 43.80 12.87 M. javanica 0.00 0.00 89.53 26.26 0.00 0.00 0.00 0.00 44.77 6.63 M. arenaria race 1 0.00 0.00 106.26 1.24 0.00 0.00 0.00 0.00 49.47 16.30 M. mayaguensis 0.00 0.00 87.10 21.38 0.00 0.00 0.00 0.00 42.67 6.47 Data are means of three replications. No differences ( P 0.05) were observed according to Duncans multiple range test. a0 = no galls; 1 = 1 to 2 galls; 2 = 3 to 10 galls; 3 = 11 to 30 galls; 4 = 31 to 100 galls; 5 = more than 100 galls per root sy stem (Taylor and Sasser, 1978

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58Table 2-16. Treatments, root-gall indexes, root weights, number of eggs per plant, number of eggs per g of roots, and dry shoot weights from the first Musa acuminata ssp. zebrina growth room trial. Treatment Root-gall index (0-5) Root Weight (g) Mean number of eggs per plant Mean number of eggs per g roots Dry Shoot Weight (g) Control 0.00 0.00 bab 23.63 5.44 0.00 0.00 b 0.00 0.00 c 9.52 1.66 M. incognita race 2 4.33 2.07 a 26.55 6.74 189,306.67 114,226.82 a 7,039.58 2,385.42 ab 7.70 6.60 M. javanica 5.00 0.00 a 26.07 5.01 139,053.33 90,863.31 a 5,292.61 3,214.97 b 9.72 1.80 M. arenaria race 1 4.67 1.63 a 24.62 4.47 199,737.78 142,003.68 a 8,119.14 5,689.47 a 9.75 2.50 M. mayaguensis 5.00 0.00 a 26.20 6.42 190,946.67 124,960.19 a 7,477.72 5,774.92 ab 9.32 0.93 a0 = no galls; 1 = 1 to 2 galls; 2 = 3 to 10 galls; 3 = 11 to 30 galls; 4 = 31 to 100 galls; 5 = more than 100 galls per root sy stem (Taylor and Sasser, 1978). bData are means of six replications. Means in columns followed by a common letter are not different ( P 0.05) according to Duncans multiple range test. Egg numbers were log-transformed prior to data analysis, but non-transformed numbers are shown.

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59Table 2-17. Treatments, root-gall indexes, root weights, number of eggs per plant, number of eggs per g of roots, and dry shoot weights from the second Musa acuminata ssp. zebrina growth room trial. Treatment Root-gall index (0-5) Root Weight (g) Mean number of eggs per plant Mean number of eggs per g roots Dry Shoot Weight (g) Control 0.00 0.00 dab 11.87 3.29 b 0.00 0.00 d 0.00 0.00 c 9.25 1.55 a M. incognita race 2 2.17 1.97 c 13.45 6.83 ab 130,472.22 83,342.51 c 9,684.54 4,138.28 b 7.37 1.53 b M. javanica 3.17 0.82 b 14.47 1.79 ab 170,611.11 92,654.84 bc 11,707.26 5,311.64 ab 7.30 1.91 b M. arenaria race 1 2.00 2.19 c 12.25 4.02 b 180,138.89 95,931.79 ab 15,126.25 9,704.36 a 7.45 1.66 b M. mayaguensis 4.67 1.03 a 15.83 4.34 a 239,500.00 108,061.51 a 15,167.20 6,381.53 a 7.50 1.99 b a0 = no galls; 1 = 1 to 2 galls; 2 = 3 to 10 galls; 3 = 11 to 30 galls; 4 = 31 to 100 galls; 5 = more than 100 galls per root sy stem (Taylor and Sasser, 1978). bData are means of six replications. Means in columns followed by a common letter are not different ( P 0.05) according to Duncans multiple range test. Egg numbers were log-transformed prior to data analysis, but non-transformed numbers are shown.

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60 In the greenhouse experiment (Table 2-18), no differences ( P 0.05) among treatments were observed for root weight and dry shoot weight, and no differences ( P 0.05) among root-knot nematode treatments were observed for number of eggs per plant or number of eggs per g of roots. However, M. mayaguensis -inoculated plants had a higher root-gall index than the other root-knot nematode-infected plants. The data suggest that M. acuminata ssp. zebrina cv. Rowe Red is a good host to the four Meloidogyne spp. evaluated. Codieaum variegatum cv. Gold Dust Results for the two C. variegatum cv. Gold Dust growth room trials were heterogeneous and are presented in tables 2-19 and 2-20, respectively. The results for the greenhouse experiment are presented in table 2-21. Root galls were not observed on any of the plants in the three studies. No differences ( P 0.05) were observed among treatments for root-gall index, root weight, number of eggs per plant, number of eggs per g of roots, or dry shoot weight in either of the growth room trials or the greenhouse experiment. The data suggest that C. variegatum cv. Gold Dust is a nonhost to the Meloidogyne spp. evaluated. Discussion The assignment of host statues to perennial ornamentals is ambiguous. Unlike annual crops, root-knot nematode populations may thrive on perennial hosts for many months and years. Therefore, studies on the host status of root-knot nematodes on perennials that result in the classification an immune or a poor host may be disproved by conducting longer-term studies that allow ample time for pathogen reproduction. The plants evaluated in this study were homogeneous in their response to the four

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61Table 2-18. Treatments, root-gall indexes, root weights, number of eggs per plant, number of eggs per g of roots, and dry shoot weights from the Musa acuminata ssp. zebrina greenhouse experiment. Treatment Root-gall index (0-5) Root Weight (g) Mean number of eggs per plant Mean number of eggs per g roots Dry Shoot Weight (g) Control 0.00 0.00 cab 26.83 17.92 0.00 0.00 b 0.00 0.00 b 24.00 14.76 M. incognita race 2 2.33 1.15 b 32.30 44.06 277,331.67 389,211.65 a 8,344.33 9,516.67 a 19.77 13.63 M. javanica 4.00 3.64 ab 58.30 27.61 454,100.00 433,463.29 a 8,522.22 10,650.06 a 25.60 8.63 M. arenaria race 1 3.33 2.31 ab 29.10 3.02 318,673.89 470,556.48 a 10,794.44 15,023.29 a 20.23 5.28 M. mayaguensis 4.67 1.15 a 39.50 36.43 236,578.33 515,546.34 a 5,350.00 10,144.02 a 18.67 10.07 a0 = no galls; 1 = 1 to 2 galls; 2 = 3 to 10 galls; 3 = 11 to 30 galls; 4 = 31 to 100 galls; 5 = more than 100 galls per root sy stem (Taylor and Sasser, 1978). bData are means of three replications. Means in columns followed by a common letter are not different ( P 0.05) according to Duncans multiple range test. Egg numbers were log-transformed prior to data analysis, but non-transformed numbers are shown.

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62Table 2-19. Treatments, root-gall indexes, root weights, number of eggs per plant, number of eggs per g of roots, and dry shoot weights from the first Codiaeum variegatum cv. Gold Dust trial. Treatment Root-gall index (0-5) Root Weight (g) Mean number of eggs per plant Mean number of eggs per g roots Dry Shoot Weight (g) Control 0.00 0.00a 11.93 3.44 0.00 0.00 0.00 0.00 11.58 3.24 M. incognita race 2 0.20 0.82 13.32 11.05 37.40 97.86 2.78 7.17 11.46 10.18 M. javanica 0.17 0.82 11.32 3.49 8.83 43.55 0.93 4.54 10.78 4.97 M. arenaria race 1 0.17 0.82 14.52 5.59 4.50 21.77 0.41 2.03 13.40 3.68 M. mayaguensis 0.00 0.00 12.10 3.33 26.67 130.64 1.92 9.40 11.33 2.99 Data are means of six replications. No differences ( P 0.05) were observed according to Duncans multiple range test. a0 = no galls; 1 = 1 to 2 galls; 2 = 3 to 10 galls; 3 = 11 to 30 galls; 4 = 31 to 100 galls; 5 = more than 100 galls per root sy stem (Taylor and Sasser, 1978).

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63Table 2-20. Treatments, root-gall indexes, root weights, number of eggs per plant, number of eggs per g of roots, and dry shoot weights from the second Codiaeum variegatum cv. Gold Dust growth room trial. Treatment Root-gall index (0-5) Root Weight (g) Mean number of eggs per plant Mean number of eggs per g roots Dry Shoot Weight (g) Control 0.00 0.00a 9.87 2.47 0.00 0.00 0.00 0.00 9.22 2.36 M. incognita race 2 0.17 0.82 10.25 5.80 0.00 0.00 0.00 0.00 8.37 3.42 M. javanica 0.33 1.03 8.05 3.39 0.00 0.00 0.00 0.00 7.53 3.80 M. arenaria race 1 0.33 1.03 8.37 6.41 0.00 0.00 0.00 0.00 7.07 3.52 M. mayaguensis 0.00 0.00 8.82 2.06 0.00 0.00 0.00 0.00 9.05 5.67 Data are means of six replications. No differences ( P 0.05) were observed according to Duncans multiple range test. a0 = no galls; 1 = 1 to 2 galls; 2 = 3 to 10 galls; 3 = 11 to 30 galls; 4 = 31 to 100 galls; 5 = more than 100 galls per root sy stem (Taylor and Sasser, 1978).

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64Table 2-21. Treatments, root-gall indexes, root weights, number of eggs per plant, number of eggs per g of roots, and dry shoot weights from the Codiaeum variegatum cv. Gold Dust greenhouse experiment. Treatment Root-gall index (0-5) Root Weight (g) Mean number of eggs per plant Mean number of eggs per g roots Dry Shoot Weight (g) Control 0.00 0.00a 37.03 8.88 0.00 0.00 0.00 0.00 33.17 10.58 M. incognita race 2 0.00 0.00 35.80 21.20 0.00 0.00 0.00 0.00 36.93 2.90 M. javanica 0.00 0.00 44.00 49.36 0.00 0.00 0.00 0.00 34.27 15.01 M. arenaria race 1 0.00 0.00 39.47 17.76 0.00 0.00 0.00 0.00 35.53 19.60 M. mayaguensis 0.00 0.00 54.80 17.16 0.00 0.00 0.00 0.00 42.33 6.86 Data are means of three replications. No differences ( P 0.05) were observed according to Duncans multiple range test. a0 = no galls; 1 = 1 to 2 galls; 2 = 3 to 10 galls; 3 = 11 to 30 galls; 4 = 31 to 100 galls; 5 = more than 100 galls per root sy stem (Taylor and Sasser, 1978).

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65 Meloidogyne spp. evaluated, with the exception of L. muscari cv. Evergreen Giant, which was a good host to M. incognita race 2, M. javanica and M. mayaguensis, and a poor host to M. arenaria race 1. Liriope muscari was classified as a nonhost to M. arenaria race 1 in the first and second growth room experiments, yielding Rf values of 0.07 and 0.002, respectively. However, its host status to this pathogen changed to poor (Rf = 0.12) in the greenhouse experiment. It is unclear whether M. arenaria race 1 is a good host under longer term field or greenhouse studies. Pittosporum tobira cv. Variegata was a nonhost to all the Meloidogyne spp. evaluated. Liriope muscari cv. Variegata was previously reported susceptible to M. hapla (LaMondia, 1996). This is the first report of any Meloidogyne spp. on L. muscari cv. Evergreen Giant. Pittosporum tobira cv. Variegata was a nonhost to the Meloidogyne spp. evaluated in this study. This cultivar was found infected with Meloidogyne spp. in Lake County, FL (chapter 3). Meloidogyne incognita M. arenaria and Meloidogyne sp. were previously reported on P. tobira by Nigh (1972), Bureau of Nematology (1989), and Goodey et al. (1965), respectively. However, P. tobira cv. Variegata is frequently infected by Meloidogyne spp., as evident by root galls and egg masses that are observed on infected plant roots (Levin, R., personal observation). It is possible that P. tobira cv. Variegata is a nonhost to the Meloidogyne spp. isolates evaluated. It is unlikely that the reproductive period in these studies was limiting, since a small number of eggs (Rf 0.01), which probably remained from the initial inocula, was retrieved from plants in the two growth room experiments, and no eggs were isolated from plants in the greenhouse experiment. Differences in host response and results that differ from field observations in this study may be attributed to the Meloidogyne spp. isolates evaluated. Additional work,

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66 involving the isolation, culture, and identification of root-knot nematodes from infected P. tobira cv. Variegata plants, is required to unveil the host status of this cultivar to Meloidogyne spp. The purple-corolla form of S. leucantha and M. sumatrana ssp. zebrina cv. Rowe Red were good hosts to the Meloidogyne spp. evaluated in these experiments. The whitecorolla form of S. leucantha was a good host to the M. incognita and M. mayaguensis isolates evaluated, and a nonhost to the M. javanica and M. arenaria isolates evaluated. Meloidogyne sp. was previously reported on Salvia leucantha (Goodey and Franklin, 1956, Goodey et al., 1965). Although Meloidogyne arenaria and M. incognita have been reported on numerous Musa acuminata cultivars (Goodey et al., 1965), the host status of M. acuminata ssp. zebrina cv. Rowe Red to the Meloidogyne spp. evaluated is reported here for the first time. Odontonema cuspidatum and Codiaeum variegatum cv. Gold Dust were nonhosts to the Meloidogyne spp. evaluated in these experiments. Odontonema cuspidatum and C. variegatum were previously reported as hosts to Radopholus similis and Hoplolaimus sp., respectively (Goodey et al., 1965). Since many C. variegatum cultivars are frequently encountered infected by Meloidogyne spp. (Levin, R., personal observation), the host status of C. variegatum cv. Gold Dust to the Meloidogyne spp. evaluated and to additional root-knot nematode species and races needs to be verified and evaluated. In addition, the usefulness of C. variegatum cv. Gold Dust as a root-knot nematode resistant rootstock for many Meloidogyne spp.-susceptible C. variegatum cultivars needs to be investigated.

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67 CHAPTER 3 IDENTIFICATION OF ROOT-KNOT NEMATODES Introduction The usefulness of Meloidogyne spp. isozymes, resolved by electrophoresis, as a tool for the identification of root-knot nematode species, has increased dramatically over the last 40 years. Most Meloidogyne spp. show species-specific esterase (Est) phenotypes. Some species, including M. exigua and M. naasi show nonspecific Est phenotypes. Therefore, the elucidation of a second enzyme phenotype, malate dehydrogenase (Mdh), is necessary for differentiation of such species. Dickson et al. (1971) proved that Est and Mdh phenotypes, resolved following disk electrophoresis, provide a reliable means for speciating Meloidogyne incognita M. javanica M. arenaria and M. hapla Disk electrophoresis analysis of Meloidogyne females by Dickson et al. (1970, 1971) and Hussey et al. (1972) utilized several nematodes of the same species per isozyme phenotype elucidated, rendering genetic analysis at the intraand interspecific levels impossible (Dalmasso and Berge, 1978). The use of polyacrylamide gel electrophoresis (PAGE) by Dalmasso and Berge (1978) provided the means to unveil single Meloidogyne female isozyme phenotypes following electrophoresis in a 0.7-mm thick slab gel. Later works (Carneiro et al., 1996, 1998, 2000; Dalmasso and Berge, 1978; Esbenshade and Triantaphyllou, 1985a, 1985b, 1985c, 1987; Fargette, 1987a, 1987b; Fargette and Braaksma, 1990; Pais and Abrantes, 1989; Starr et al, 1996; Yongfang et al., 1998) utilized innovations of thin-slab gel electrophoresis for the

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68 identification of single Meloidogyne females. Unfortunately, standard methodology for such techniques is not universal, and published works utilize various gel-forming apparatuses and methodologies, an array of enzyme stain concoctions, and varying electrophoresis run times. In addition, many authors fail to accurately describe their methods and results in detail. Esbenshade and Triantaphyllou (1985b) presented numerous Mdh and Est phenotypes and their associated relative electrophoretic migration (Rm) values (distance of protein in question relative to the migration distance of the bromophenol-blue dye). Each band was designated a number, representing its migration relative to other bands, and the numbers were grouped into categories represented by letters. Combinations of letters and numbers elucidate particular phenotypic patterns. The letter-number system, referred to in most works that describe Meloidogyne spp. Mdh and Est phenotypes, along with a M. javanica or M. hapla standard on the same gel, is useful for routine identification of several Meloidogyne species. However, isozyme phenotypes are limited in their usefulness when compared on different gels. Enzyme phenotypes may vary with environmental conditions, nematode life stage, and different populations or isolates (Caswell-Chen et al., 1993). Furthermore, since variability is inevitable between electrophoretic runs due to human error and unavoidable environmental differences at the time of gel and enzyme stain preparations, slight variations in isozyme phenotypes may lead to improper speciation. Therefore, until procedural materials and methods, evaluation methods, and the accurate reporting of Rm values are standardized, information including gel formulations and sizes, staining methods, electrophoresis run times, and precisely-measured Rm must be included in works that elucidate known or

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69 unknown Meloidogyne spp. isozyme phenotypes (Esbenshade and Triantaphyllou, 1985a, 1985b, 1985c; Evans, 1971). Isozyme phenotypes, Rm values, and methodological information have been described accurately (Dalmasso and Berge, 1978; Esbenshade and Triantaphyllou, 1985a, 1985b, 1985c; Fargette, 1987a, 1987b; Hussey et al., 1972; Yongfang et al., 1998), and are presented for M. javanica (Figure 3-1), M. incognita (Figure 3-2), M. arenaria (Figure 3-3 and 3-4), M. hapla (Figure 3-5), various Meloidogyne spp. (Figure 3-6), and for several unidentified Meloidogyne spp. isolates (Figure 3-7). Yongfang et al. (1998) and Cetintas et al. (2003) elucidated Mdh and Est isozyme patterns using a PhastSystem apparatus (Pharmacia Biotech AB, Uppsala, Sweden) and a Mini Protean 3 Cell apparatus, respectively, for routine identification of root-knot nematodes. Yongfang et al. (1998) determined that the PhastSystem is ideal for the rapid identification of root-knot nematode species. Furthermore, the authors reported that band pattern stability withstands differing root-knot nematode host species, host nutrition, sample origin and cultivation practices, and sample dosage. Although Mdh and Est patterns relative Rm were identical to previously-reported figures (Esbenshade and Triantaphyllou, 1985a, 1985b, 1985c), Yongfang et al. (1998) reported on a previously unobserved Est phenotype, referred to as J2 (Figure 3-1). A similar phenotype was reported for M. javanica from soybean in Brazil (Castro et al., 2003). Objectives The objectives of this study were to 1. Differentiate malate dehydrogenase and esterase phenotypes of Meloidogyne spp. that were collected from infected ornamental plants in Florida using PAGE.

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70 2. Evaluate the usefulness of PAGE in differentiating Meloidogyne spp. for routine extension diagnostic purposes. Materials and Methods Nematode Populations A total of 20 root samples of ornamental plants were collected from nurseries, botanical gardens, and residential plantings from nine counties in Florida. For root-knot nematode identification, young egg-laying females were dissected from naturally infected roots of each plant. Isozyme Analysis Root-knot nematode females, dissected from different galls of the root systems, were used for isozyme analyses following electrophoresis, using either PhastSystem or Mini-Protean 3 Cell. Meloidogyne spp. were identified by comparing specimen Mdh and Est phenotypes to those of previously published root-knot nematode species (Dalmasso and Berge, 1978; Esbenshade and Triantaphyllou, 1985c, 1990; Fargette, 1987a, 1987b; Hussey et al., 1972; Yongfang et al; 1998). The Mdh and Est Rm of known M. incognita M. javanica M. arenaria M. mayaguensis and M. partityla isolates are presented in table 3-1 (Dalmasso and Berge, 1978; Esbenshade and Triantaphyllou, 1985c; Fargette, 1987a, 1987b; Hussey et al., 1972; Yongfang et al., 1998). At least 26 females from each root sample were examined for species identification with the Mini-Protean 3 Cell, except for the Carya illinoensis sample, from which eight females were examined. However, either six or ten females from each root sample were examined for species identification with the PhastSystem. PhastSystem-run samples were processed on the

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71 Band Phenotype Designation Key Enzyme Band Rma () MDHb N1 23.0 MDH N3 23.0, 27.0, 30.0 MDH J4 19.0, 24.0, 30.0, 34.0 ESTc J3 46.0, 54.5, 58.9 EST J2 47.0, 59.0 EST J3b 30.0, 36.0, 38.0 EST P4 61.0, 81.0, 89.0 Rm () 0 40 80 20 60 100 MDH EST Sourcef Systemg N1 J3 1 1 N3 J3 1 1 N1 J2 2 2 J4 J3b 3 3 N/A d P4e 4 4

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Figure 3-1. Malate dehydrogenase and esterase relative electrophoretic migrations and enzyme phenotypes of Meloidogyne javanica as reported by several authors. aRelative electrophoretic migration. bMalate dehaydrogenase. cEsterase. dNo phenotype specified. eFargette (1987b) reported that esterase phenotype P4 is identical to esterase phenotype J3, with Rm differences attributed to electrophoresis apparatus variations (Esbenshade and Triantaphyllou, 1985c). f1 = Esbenshade and Triantaphyllou, 1985c; 2 = Yongfang et al., 1998; 3 = Fargette, 1987b. g1 = Pharmacia GE-2/4 apparatus, separating and stacking gels 7% and 4%, respectively; 2 = PhastSystem, 10-15% gel; 3 = Pharmacia apparatus, 7% gel; 4 = unspecified.

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73 Band Phenotype Designation Key Enzyme Band Rma () MDHb N1 23.0 MDH N3b 23.0, 28.3, 35. 0 ESTc I1 47.0 EST I1a 39.0 EST P1 71.0, 76.0 EST S1 43.8 ESTP765.0 71.0 Rm () 0 20 40 60 80 100 MDH EST Sourcef Systemg N3b I1 1 1 N1 S1 1 1 N1 S1 2 1 N/A d P1e 3 2 N 1 I1 1 1 N/A P7e3 2 N/A I1a 4 3

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Figure 3-2. Malate dehydrogenase and esterase relative electrophoretic migrations and enzyme phenotypes of Meloidogyne incognita as reported by several authors. aRelative electrophoretic migration. bMalate dehydrogenase. cEsterase. dNo phenotypes specified. eFargette (1987a, 1987b) reported that esterase phenotypes P1 and P7 are identical to esterase phenotypes I1 and S1, respectively, with Rm differences attributed to electrophoresis apparatus variations (Esbenshade and Triantaphyllou, 1985c). f1 = Esbenshade and Triantaphyllou, 1985c; 2 = Dalmasso and Berge, 1978; 3 = Fargette, 1987b; 4 = Hussey et al., 1972. g1 = Pharmacia GE-2/4 apparatus, separating and stacking gels 7% and 4%, respectively; 2 = Pharmacia apparatus, 7% gel; 3 = Polyanalyst apparatus, 7% gel.

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75 Band Phenotype Designation Key Enzyme Band Rma () MDHb N1 23.0 MDH N3 23.0, 27.0, 30.0 MDH N3b 23.0, 28.3, 35.0 ESTc A1 53.3 EST A2 53.3, 56.3 EST P3 50.7, 53.3, 56.3 EST S1-M1 43.8, 47.0 EST S2-M1 41.3, 43.8, 47.0 EST M3-F1 47.0, 50.0, 53.3, 56.8 Rm () 0 20 40 60 80 100 MDH EST Sourced Systeme N1 A3 1 1 N1 A1 1 1 N3 S1-M1 1 1 N3 A2 1 1 N1 S2-M1 1 1 N1 A2 1 1 N1 S1-M1 1 1 N3b A2 1 1 N1 M3-F1 1 1

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Figure 3-3. Malate dehydrogenase and esterase relative electrophoretic migrations and enzyme phenotypes of Meloidogyne arenaria as reported by Esbenshade and Triantaphyllou (1985c). aRelative electrophoretic migration. bMalate dehydrogenase. cEsterase. d1 = Esbenshade and Triantaphyllou, 1985c. e1 = Pharmacia GE-2/4 apparatus, separating and stacking gels 7% and 4%, respectively.

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77 Band Phenotype Designation Key Enzyme Band Rma () MDHb N1 23.0 MDH A5 19.0, 24.0, 30.0, 35.0, 40. 0 ESTc A1 54.0 EST A2a 43.0, 46.0 EST A3a 30.0, 36.0, 38.0 EST P2 65.0, 71.0 EST P5 79.0, 85.0 ESTP859.0 65.0 71.0 Rm () MDH EST Sourcef Systemg N/A d A2a 3 3 N/A P5e 4 4 N/A P2e 4 4 A5 A3a 2 2 N/A P8e 4 4 N1 A1 1 1 0 20 40 60 80 100

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Figure 3-4. Malate dehydrogenase and esterase relative electrophoretic migrations and enzyme phenotypes of Meloidogyne arenaria as reported by several authors. aRelative electrophoretic migration. bMalate dehydrogenase. cEsterase. dNo phenotypes specified. eFargette (1987a, 1987b) reported that esterase phenotypes P2, P5, and P8 are identical to esterase phenotypes S1-M1, A2, and S2-M1, respectively, with Rm differences attributed to electrophoresis apparatus variations (Esbenshade and Triantaphyllou, 198 5c). f1 = Yongfang et al., 1998; 2 = Dalmasso and Berge, 1978; 3 = Hussey et al., 1972; 4 = Fargette, 1987b. g1 = PhastSystem; 2 = Pharmacia GE-2/4 apparatus, separating and stacking gels 7% and 4%, respectively; 3 = Polyanalyst apparatus, 7% gel; 4 = Pharmacia apparatus, 7% gel.

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79 Rm () Band Phenotype Designation Key Enzyme Band Rma () MDHb H1 37.0 MDH H1a 50.0 ESTc H1 50.0 EST H1a 33.0 EST A1 53.3 0 MDH EST Sourcee Systemf H1 A1 1 1 H1a H1a 3 3 H1 H1 1,2 1 2 N/A d H1 1 1 20 40 60 80 100

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Figure 3-5. Malate dehydrogenase and esterase relative electrophoretic migrations and enzyme phenotypes of Meloidogyne hapla aRelative electrophoretic migration. bMalate dehydrogenase. cEsterase. dNo phenotypes specified. e1 = Esbenshade and Triantaphyllou, 1985c; 2 = Yongfang et al., 1998; 3 = Delmasso and Berge, 1978. f1 = Pharmacia GE-2/4 apparatus, separating and stacking gels 7% and 4%, respectively; 2 = PhastSystem; 3 = Pharmacia GE2/4 apparatus, separating and stacking gels 7% and 4%, respectively.

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81 Band Phenotype Designation Key Enzyme Band Rma () MDHb H1 37.0 MDH N1 23.0 MDH N1a 30.0 ESTc VS1 38.0, 43.8 EST VS1-S1a 38.0, 45.0 EST S2-M1 41.3, 43.8, 47.0 EST M3a 47.0, 52.5, 55.7 EST S1 43.8 EST P6 51.0, 56.0, 65.0, 71.0 EST P7 65.0, 71.0 EST P8 59.0, 65.0, 71.0 Rm () 0 20 40 60 80 100 MDH EST Sourcef Systemg N1a S1 1 1 N/A d P7e 2 2 N1 M3a 1 1 N1a VS1-S1 1 1 H1 VS1-S1a 1 1 N/A P6e2 2 N1a VS1 1 1 N1 S2-M1 1 1 N/A P8e 2 2 M carolinensis M chitwoodi M chitwoodi M cruciani M enterolobii M enterolobii M graminicola M hispanica M hispanica

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Figure 3-6. Malate dehydrogenase and esterase relative electrophoretic migrations and enzyme phenotypes of Meloidogyne spp. aRelative electrophoretic migration. bMalate dehydrogenase. cEsterase. dNo phenotypes specified. eFargette (1987b) reported that esterase phenotypes P6, P7, and P8 are identical to esterase phenotypes VS1-S1, S1, and S2M1, respectively, with Rm differences attributed to electrophoresis apparatus variations (Esbenshade and Triantaphyllou, 1985c) f1 = Esbenshade and Triantaphyllou, 1985c; 2 = Fargette, 1987b. g1 = Pharmacia GE-2/4 apparatus, separating and stacking gels 7% and 4%, respectively; 2 = Polyanalyst apparatus, 7% gel.

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83 Band Phenotype Designation Key Enzyme Band Rma () MDHb N1 23.0 MDH H1 37.0 MDH N1a 30.0 MDH N1b 35.0 MDH N3a 23.0, 25.5, 28.3 ESTc A1 53.3 EST M1 47.0, 50.0 EST VF1 65.8 EST VF1a 46.0 EST VS1 38.0, 43.8 EST S1 43.8 EST P7 75.0, 71.0 EST F1 56.8 Rm () 0 20 40 60 80 100 MDH EST Sourcef Systemg H1 M1 1 1 N1b VF1a 2 2 N1a VF1 1 1 N1a VS1 1 1 N1 A1 1 1 N1a S1 1 1 N/A d P7e 3 3 N3a F1 1 1 M microcephala M microtyla M nassi M naasi M oryzae M platani M platani M querciana

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Figure 3-6. Continued aRelative electrophoretic migration. bMalate dehydrogenase. cEsterase. dNo phenotypes specified. eFargette (1987b) claims that esterase phenotype P7 is identical to esterase phenotype S1, with Rm differences attributed to electrophoresis apparatus variations (Esbenshade and Triantaphyllou, 1985c). f1 = Esbenshade and Triantaphyllou, 1985c; 2 = Delmasso and Berge, 1978; 3 = Fargette, 1987b. g1 = Pharmacia GE-2/4 apparatus, separating and stacking gels 7% and 4%, respectively; 2 = Pharmacia GE-2/4 apparatus, separating and stacking gels 7% and 4%, respectively; 3 = Polyanalyst apparatus, 7% gel.

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85 Band Phenotype Designation Key Enzyme Band Rma () MDHb N1 23.0 MDH N3 23.0, 27.0, 30.0 MDH N3c 23.0, 30.0, 32.5 ESTc F1 56.8 EST P3 86.0 EST VS1 38.0, 43.8 EST VS1-S1 36.0, 43.8 EST VS1-M2 40.0, 53.3, 56.3 Rm () 0 20 40 60 80 100 MDH EST Sourcef Systemg N1 F1 1 1 N3c VS1-S1 1 1 N/A d P3e 2 2 N3c VS1-M2 1 1 N3 VS1 1 1 N1 VS1 1 1

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Figure 3-7. Malate dehydrogenase and esterase relative electrophoretic migrations and enzyme phenotypes of unidentified Meloidogyne spp. aRelative electrophoretic migration. bMalate dehydrogenase. cEsterase. dNo phenotypes specified. eFargette (1987b) claims that esterase phenotype P3 is identical to esterase phenotype F1, with Rm differences attributed to electrophoresis apparatus variations (Esbenshade and Triantaphyllou, 1985c). f1 = Esbenshade and Triantaphyllou, 1985c; 2 = Fargette, 1987b. g1 = Pharmacia GE-2/4 apparatus, separating and stacking gels 7% and 4%, respectively; 2 = Polyanalyst apparatus, 7% gel.

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87 Band Phenotype Designation Key Enzyme Band Rma () MDHb N1 23.0 MDH N3 23.0, 27.0, 30.0 MDH N3c 23.0, 30.0, 32.5 ESTc F1 56.8 EST P3 86.0 EST VS1 38.0, 43.8 EST VS1-S1 36.0, 43.8 ESTVS1-M240.0 53.3 56.3 Rm () 0 20 40 60 80 100 MDH EST Sourcef Systemg N1 F1 1 1 N3c VS1-S1 1 1 N/A d P3e 2 2 N3c VS1-M2 1 1 N3 VS1 1 1 N1 VS1 1 1

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Figure 3-7. Continued aRelative electrophoretic migration. bMalate dehydrogenase. cEsterase. dNo phenotypes specified. eFargette (1987b) claims that esterase phenotype P3 is identical to esterase phenotype F1, with Rm differences attributed to electrophoresis apparatus variations (Esbenshade and Triantaphyllou, 1985c). f1 = Esbenshade and Triantaphyllou, 1985c; 2 = Fargette, 1987b. g1 = Pharmacia GE-2/4 apparatus, separating and stacking gels 7% and 4%, respectively; 2 = Polyanalyst apparatus, 7% gel.

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89 Rm () MDH EST Sourced Systeme N1 VS1-M2 1 1 N5 A2 1 1 N1a S1-M1 1 1 N1 M3 1 1 N1a S2-M1 1 1 Band Phenotype Designation Key Enzyme Band Rma () MDHb N1 23.0 MDH N1a 30.0 MDH N5 23.0, 25.5, 28.3, 30.0, 32. 5 ESTc VS1-M240.0, 53.3, 56.3 EST S1-M1 43.8, 47.0 EST S2-M1 41.3, 43.8, 47.0 EST M3 47.0, 50.7, 55.7 ESTA253.3 56.3 0 20 40 60 80 100

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Figure 3-7. Continued aRelative electrophoretic migration. bMalate dehydrogenase. cEsterase. d1 = Esbenshade and Triantaphyllou, 1985c. e1 = Pharmacia GE-2/4 apparatus, separating and stacking gels 7% and 4%, respectively.

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91 Table 3-1. Known Meloidogyne spp. malate dehydrogenase and esterase relative migrations that were used as references against those that were revealed electrophoretically from females collected from several counties in Florida. Meloidogyne spp. Relative migration M. incognita MDHa 21.4 ESTb 42.9 M. javanica MDH 22.7 EST 41.8, 49.0, 54.5 M. arenaria MDH 18.0, 20.0, 22.0 EST 38.0, 40.0 M. mayaguensis MDH 29.0, 32.0, 35.0 EST 31.0, 41.0 M. partityla MDH 31.0 EST 38.0, 49.0, 51.0 aMalate dehydrogenase bEsterase

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92 same day of female extraction, while females extracted for processing in the MiniProtean 3 Cell apparatus were stored at -4 C for up to six months without apparent effects on band resolution. One M. javanica female was included in every PhastSystemrun gel while two females were included in every Mini-Protean 3 Cell-run gel as controls. The PhastSystem apparatus was set according to the PhastSystem user manual (Pharmacia Biotech AB, 1995) as follows: first step 400 volt (V) limit, 10.0 current (mA) limit, 2.5 power (W) limit, bed temperature 15 C, duration 10.0 volthours (Vh); second step 400 V limit, 10.0 mA limit, 1.5 W limit, bed temperature 15 C, duration 2.0 Vh; third step 400 V limit, 10.0 mA limit, 1.5 W limit, bed temperature 15 C, duration 100.0 Vh. Meloidogyne females were dissected from infected roots and placed in a BPI dish containing 0.85% sodium chloride (NaCl) at room temperature. Similarly, two M. javanica females were dissected from infected tomato cv. Rutgers roots and placed in 0.85% NaCl in a separate BPI dish. A piece of Parafilm (American National Can, Chicago, IL) was pressed against the wells of an eight or 12-well sample applicator (Pharmacia Biotech AB, Uppsala, Sweden) until the well impressions were clearly visible on the Parafilm. Five microliters of extraction buffer (20% w/v sucrose, 2% v/v Triton X-100, 0.01% w/v bromophenol blue) were pipetted into each well and one Meloidogyne female was removed from the NaCl solution and placed in each well, for a total of 8or 12-one Meloidogyne spp. samples, including the two M. javanica samples, depending on the sample applicator used (eight or 12 wells). A separate sterile plastic toothpick (Armonds Manufacturing Company, Bogart, GA) was then used to homogenate each Meloidogyne female. An 8or 12-point applicator (Pharmacia Biotech AB, Uppsala, Sweden), depending on the sample applicator used, was pressed on the wells to upload

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93 the samples and immediately placed in the appropriate slot. The system was activated and stopped when the light emitting diode (LED) display read 100 Vh. Gels were removed from the apparatus and immediately washed with dionized water for one minute. Gels were then incubated in the dark in freshly prepared Mdh stain for 15 minutes in a 37 C Isotemp incubator (Fisher Scientific, Hampton, NH), rinsed with dionized water for one minute, and then placed into freshly prepared Est stain for 30 to 45 minutes in the incubator at 37 C in the dark (Table 3-2). Gels were rinsed again with dionized water for one minute and then incubated in 50 ml fixative solution (10% v/v glycerol, 10% v/v acetic acid) for at least 24 hrs at room temperature. Sample preparation and enzyme separation and development using Mini-Protean 3 Cell was carried out as follows: Meloidogyne females were dissected from infected roots and separately placed in 0.6-ml eppendorf tubes (Fisher Scientific, Hampton, NH) containing 10 l sample buffer (BioRad, Hercules, CA). The tubes, each containing one female, were kept on ice while Meloidogyne females were extracted, and then placed at 4 C until electrophoresis was conducted. Females were macerated and 10 l of the supernatants were loaded into the appropriate wells of the polyacrylamide gel [4% stacking gel (pH 6.8), 8% separating gel (pH 8.8) with Tris-glycine buffer (BioRad, Hercules, CA)]. Two-ten l of Meloidogyne javanica homogenates were loaded onto each of two separate wells of the same gel as controls. Electrophoresis was conducted in a 0.75-mm-thick, 8% running and 4% stacking gel, in a Mini-Protean 3 Cell (BioRad, Hercules, CA). Electrophoresis was performed in 5 C at 80 V for the first 15 minutes, and then increased to 200 V until the bromophenol blue tracking dye migrated 53 cm from the bottom of the wells, approximately 33

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94 Table 3-2. Enzyme stain concoctions used in staining malate dehydrogenase and esterase following electrophoresis using the PhastSystem. Malate Dehydrogenase Stain Esterase Stain Stock Solution The following were mixed and stored in the dark at room temperature: 50.0 ml 1.0 M Tris-HCl, pH 8.0 50.0 ml 0.3 M DL-Malic Acid, pH 7.5 370.0 ml dionized water Stain Solution The following were mixed in the dark with a magnetic stirrer for five minutes immediately prior to gel staining: 47.0 ml stock solution 0.007 g -Nicotinamide Adenine Dinucleotide ( -NAD) 0.5 ml 0.1 M Nitroblue Tetrazolium (NBT) 0.5 ml 0.1 M Phenazine Methosulfate (PMS) Stock Solutions The following were mixed, adjusted to pH 7.4, and stored in the dark at room temperature: Sodium Phosphate Buffer 9.28 g Na2HPO4, Monobasic, anhydrous 3.21 g Na2HPO4, Dibasic, anhydrous 0.3 g EDTA 400.0 ml dionized water Approximately 20.0 ml 4 M NaOH (for pH adjustment) The following were stored at -4 C: 50.0 ml 1 g -Naphthyl Acetate in 50.0 ml acetone 0.015 g Fast Blue RR Salt Stain Solution The following were mixed in the dark for five minutes immediately prior to gel staining: 25.0 ml Sodium Phosphate Buffer 0.5 ml Naphthyl acetate in acetone 0.015 g Fast Blue RR Salt

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95 minutes. Malate dehydrogenase and Est stain concoctions were freshly prepared (Table 3-3). Gels were incubated in the dark in either Mdh stain for approximately 15 minutes or Est stain for approximately 30 minutes or until bands appeared, or stained for both enzymes at the same time intervals, in a 37 C Isotemp incubator. After incubation, gels were gently washed in deionized water and then placed in 50 ml freshly prepared fixative solution (20% v/v ethanol, 10% v/v glycerol) (BioRad, Hercules, CA) for at least 24 hours. Gels were preserved between two cellophane sheets in 14-cm x 14-cm gel drying frames (Sigma-Aldrich, St. Louise, MO). Results A total of 20 root samples were infected with root-knot nematodes. Six species of root-knot nematodes were identified, primarily by isozyme (Est and Mdh) analysis. The major root-knot nematode species identified and their percentage were M. incognita M. javanica M. mayaguensis M. arenaria M. partityla M. querciana and Meloidogyne spp. (Table 3-4). Meloidoygne querciana was identified from only one female. Morphological and morphometric analysis shold be conducted to confirm this species identification. Mixed populations were found on 40% of the samples. While Rm were measured for Mini-Protean 3 Cell-run samples, such measurements could not be obtained from PhastSystem-run samples since the bromophenol blue tracking dye did not persist on the gels following incubation in the fixative solution. Meloidogyne incognita was found infecting the following: Rosmarinus officinalis (rosemary) in Lamiaceae, Syagrus romanzoffiana (queen palm) in Arecaceae, Pittosporum tobira in Pittosporaceae (Mdh Rm 0.19, Est Rm 0.38), Brassica rapa (turnip) cv. Shogoin (Mdh Rm 0.21, Est Rm 0.43) and Brassica oleracea (kale) (Mdh Rm 0.21, Est Rm 0.43) in Brassicaceae, Phaseolus

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96 Table 3-3. Enzyme stain concoctions used in staining malate dehydrogenase and esterase following electrophoresis using the Mini-Protean 3 Cell. Malate Dehydrogenase Stain Esterase Stain Stock Solution The following were mixed and stored in the dark at 5 C: 1000.0 ml 0.05 M TRIS-HCl, pH 8.6 Stain Solution The following were mixed in the dark with a magnetic stirrer for five minutes immediately prior to gel staining: 50.0 ml 0.05 M TRIS-HCl, pH 8.6 0.013 g -Nicotinamide Adenine Dinucleotide ( -NAD) 0.01 g Nitroblue Tetrazolium (NBT) 0.003 g Phenazine Methosulfate (PMS) Stock Solutions The following were mixed, adjusted to pH 6.0, and stored at room temperature: 0.05 M Potassium Phosphate Buffer, pH 6.0 720.0 ml of 50.0 ml Potassium Phospahte monobasic in 1.0 liter dionized water, adjusted to pH 6.0 180.0 ml of 25.0 ml Potassium Phosphate dibasic in 500 ml dionized water, adjusted to pH 6.0 Stain Solution The following were mixed in the dark for five minutes and filtered immediately prior to gel staining: 50.0 ml 0.05 M Potassium Phosphate buffer, pH 6.0 0.05 g Fast Blue RR Salt The following were mixed together and added to the stain solution immediately prior to gel staining: 2.5 ml acetone 2.5 ml dionized water 0.05 g -Naphthyl Acetate

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97Table 3-4. Plant species, family, county, esterase isozyme phenotype, number of samples, and Meloidogyne spp. identified from ornamental plants collected in Florida and processed using the PhastSystem and Mini-Protean 3 Cell. Host species Family County Esterase isozyme phenotype Number of samples Meloidogyne spp. Hibiscus rosa-sinensisa Malvaceae Alachua J3 1 M. javanica Rosmarinus officinalisa Lamiaceae Suwannee J1 1 M. incognita Callistemon viminalisa Myrtaceae Lee VS1-S1 1 M. mayaguensis Syagrus romanzoffianaa Arecaceae Lee VS1-S1 I1 1 M. mayaguensis M. incognita Callistemon viminalisb Myrtaceae Lee VS1-S1 1 M. mayaguensis Pittosporum tobirab Pittosporaceae Alachua I1 1 M. incognita Ruscus aculeatusb Ruscaceae Volusia A2 1 M. arenaria Buddleia davidiib Loganiaceae Pinellas N/A 1 Meloidogyne spp.

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98Table 3-4. Continued Host species Family County Esterase isozyme phenotype Number of samples Meloidogyne spp. Brassica rapab Brassicaceae Alachua J3 I1 1 M. javanica M. incognita Pittosporum tobirab Pittosporaceae Lake N/A 1 Meloidogyne spp. Ruscus aculeatusb Ruscaceae Volusia J3 1 M. javanica Carya illinoesisb Juglandaceae Jefferson Ci3 1 M. partityla Liriope muscarib Ruscaceae Hillsborough I1 N/A 1 M. incognita Meloidogyne spp. Ophiopogon japonicus Ruscaceae Orange I1 N/A 1 M. incognita Meloidogyne spp. Justicia carneab Acanthaceae Hillsborough N/A 1 Meloidogyne spp. Solenostemon scutellarioidesb Lamiaceae Hillsborough VS1-S1 1 M. mayaguensis

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99Table 3-4. Continued Host species Family County Esterase isozyme phenotype Number of samples Meloidogyne spp. Brassica oleraceab Brassicaceae Orange I1 1 M. incognita Beta vulgarisb ChenopodiaceaeOrange J3 1 M. javanica Viburnum odoratissimumb Caprifoliaceae Hillsborough J3 F1 1 M. javanica M. quercianac Phaseolus vulgarisb Fabaceae Hillsborough I1 A2 1 M. incognita M. arenaria aProcessed with PhastSystem bProcessed with Mini-Protean 3 Cell cBased on MDH and EST phenotypes from one female

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100 vulgaris (bean) (Mdh Rm 0.16, 0.20, 0.22, Est Rm 0.38) in Fabaceae, and Liriope muscari (lilyturf) cv. Evergreen Giant (Mdh Rm 0.24, Est Rm 0.27) and Ophiopogon japonicus (mondo grass) (Mdh Rm 0.22, Est Rm 0.42) in Ruscaceae. Meloidogyne javanica was found infecting the following: Hibiscus rosa-sinensis (hibiscus) cv. Pink Versicolor in Malvaceae, B. rapa cv. Shogoin (Mdh Rm 0.21, Est Rm 0.42, 0.50, 0.54) in Brassicaceae, Ruscus aculeatus (ruscus) (Mdh Rm 0.20, Est Rm 0.40, 0.48, 0.52) in Ruscaceae, Beta vulgaris (chard) (Mdh Rm 0.23, Est Rm 0.42, 0.49, 0.55) in Chenopodiaceae, and Viburnum odoratissimum (Viburnum) cv. Awabuki (Mdh Rm 0.17, 0.20, 0.23, Est Rm 0.37, 0.44, 0.48) in Caprifoliaceae. Meloidogyne arenaria was found infecting R. aculeatus (Mdh Rm 0.18, 0.20, 0.22, Est Rm 0.38, 0.40) and P. vulgaris (Mdh Rm 0.17, 0.23, 0.27, Est Rm 0.44, 0.46). Meloidogyne mayaguensis was found infecting Callistemon viminalis (bottle brush) (Mdh Rm 0.29, 0.32, 0.35, Est Rm 0.31, 0.41) in Myrtaceae, S. romanzoffiana in Arecaceae, and Solenostemon scutellarioides (coleus) cv. Elfers (Mdh Rm 0.30, 0.33, 0.37, Est Rm 0.32, 0.39) in Lamiaceae. Meloidogyne partityla was found infecting Carya illinoensis (pecan) (Mdh Rm 0.31, Est Rm 0.38, 0.49, 0.51) in Juglandaceae. The phenotype designation Ci3, named after the hosts species and number of bands, was assigned to the Est isozyme phenotype resolved from the Meloidogyne females that were isolated from C. illinoensis (Figure 3-8). Meloidogyne querciana was found infecting V. odoratissimum cv. Awabuki (Mdh Rm 0.20, 0.24, 0.26, Est Rm 0.47) in Caprifoliaceae (Figure 3-9). Finally, several Meloidogyne spp. whose Mdh and Est patterns could not be discerned were found infecting Buddleia davidii (butterfly bush) (Mdh Rm 0.19; 0.21; 0.19, 0.23, Est Rm 0.46; 0.43, 0.46; 0.25, 0.43, 0.46; 0.43, 0.46, 0.49; 0.25, 0.43, 0.46, 0.49) in Loganiaceae, P.

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101 Rm Mdh N1 N1a N1a N1a N1a N1 Rm Est J3 Ci3 Ci3 Ci3 Ci3 Figure 3-8. Malate dehydrogenase (Mdh) and esterase (Est) isozyme phenotypes, revealed using the mini-protean 3 cell apparatus, of Meloidogyne partityla females extracted from Carya illinoensis from Jefferson County on 31 March 2004.

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102 Rm Mdh N3 N3a Est J3 F1 Figure 3-9. Malate dehydrogenase (Mdh) and esterase (Est) isozyme phenotype, revealed using the mini-protean 3 cell apparatus, of a Meloidogyne querciana female extracted from Viburnum odoratissimum cv. Awabuki from Hillsborough County on 25 January 2005.

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103 tobira cv. Variegata (Mdh Rm 0.23, Est Rm 0.47, 0.50; 0.21, 0.28; 0.28, 0.47, 0.50; 0.48, 0.51, 0.54, 0.56; 0.47) in Pittosporaceae, L. muscari cv. Evergreen Giant (Mdh Rm 0.24, Est Rm 0.27, 0.48) and O. japonicus (Est Rm 0.28, 0.42, 0.45) in Ruscaceae, and Justicia carnea (flamingo plant) (Mdh Rm 0.29, 0.33, 0.37; 0.30; 0.29, 0.33, Est Rm 0.29, 0.37; 0.49, 0.52) in Acanthaceae (Table 3-4). Discussion Malate dehydrogenase and Est isozyme phenotypes, unveiled following PAGE, present a method for the speciation of Meloidogyne spp. that is less subjective than perineal patterns and other such morphologicallyand morphometrically-based identification techniques. In most cases, Est isozyme phenotypes are species-specific. Only four Meloidogyne spp. gels that clearly revealed Mdh and Est phenotypes were successfully run using the PhastSystem apparatus. Gels run after 17 May 2003 exhibited smeared Mdh bands whose Rm measurements could not be accurately measured. Altering many parameters used in gel processing, extraction buffer, and electrophoretic apparatus settings on the PhastSystem and using freshly prepared reagent solutions whose lot numbers were confirmed as useful, many M. javanica females were run after 17 May 2003 in an attempt to resolve the problem surrounding the smeared Mdh bands, but none revealed clear Mdh phenotypes (Figure A-1). In addition, the bromophenol blue migrating line did not persist on PhastSystem-run gels after incubation in the fixative solution, making measurement of Rm impossible. Thus, it was decided to use the MiniProtean 3 Cell to carry out PAGE. Based on this work, it is concluded that, if isozymes are resolved well, the PhastSystem apparatus is useful for routine identification of M. incognita M. javanica and M. arenaria Mdh and Est isozyme phenotypes, based on their

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104 migration in respect to M. javanica Mdh and Est phenotypes only. However, since Rm cannot be measured, identification of uncommon Meloidogyne spp. on the PhastSystem apparatus is misleading and may result in incorrect speciation. Mini-Protean 3 Cell-run gels in this work clearly reveal Mdh and Est isozyme phenotypes after staining for these enzymes. Gels were stained separately for Mdh or Est, or the enzymes were stained on the same gel. The clearly observed bromophenol blue dye band and the relatively large gel size (60.5-mm x 91.0-mm) allowed for accurate Rm measurements. Based on this work, it is concluded that the Mini-Protean 3 Cell apparatus is useful for the speciation of Meloidogyne spp., based on unknown sample Rm measurements and isozyme phenotype position, which can be compared with Rm measurements and isozyme phenotypes of known Meloidogyne spp. Meloidogyne spp. Mdh and Est isozyme phenotypes presented in this study were referenced against perineal pattern-confirmed M. incognita M. javanica M. arenaria and M. mayaguensis which were run electrophoretically using the same methodology as the samples in questions (Table 3-1). In addition, cultured M. partityla females, provided by A. P. Nyczepir, United States Department of Agriculture, ARS, Southeastern Fruit and Tree Nut Research Laboratory, Byron, GA, on Carya illinoensis roots, were run electrophoretically using the same methodology as all other samples, and their Mdh and Est phenotypes and Rm measurements (Table 3-1) were used as a references against isozyme phenotypes in question. Since human error and environmental conditions vary slightly in the preparation, staining, and measuring of each gel, Rm values of Meloidogyne spp. presented in table 3-4 do not consistently conform to those presented in table 3-1. However, relative distances within isozyme phenotypes and band migrations

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105 relative to that of M. javanica controls do conform to the referenced Meloidogyne spp. presented in table 3-1, or are labeled as unidentified isozyme patterns. Meloidogyne incognita and Meloidogyne spp. were identified from P. tobira and P. tobira cv. Variegata, respectively. Meloidogyne incognita M. arenaria and Meloidogyne spp. were previously reported on P. tobira (Bureau of Nematology, 1989; Goodey and Franklin, 1956; Goodey et al., 1965; Nigh, 1972). In addition, Meloidogyne spp. were isolated from P. tobira cv. Variegata in this study. However, growth room and greenhouse studies conducted at the University of Florida (chapter 2) suggest that P. tobira cv. Variegata is a nonhost to M. incognita race 2, M. javanica M. arenaria race 1, and M. mayaguensis It is possible that P. tobira cv. Variegata is a nonhost to the Meloidogyne spp. isolates tested in these studies. Further investigation on the susceptibility of P. tobira cultivars to Meloidogyne spp. is required to alleviate this conflict. In addition, M. incognita and Meloidogyne spp. were isolated from O. japonicus Meloidogyne incognita acrita was previously reported on this plant (Goodey et al., 1965). Meloidogyne javanica and M. arenaria were isolated from R. aculeatus Meloidogyne spp. were also isolated from J. carnea This is the first report of any Meloidogyne spp. infecting these plants. Meloidogyne mayaguensis was identified from C. viminalis and S. romanzoffiana roots. This is the first report of M. mayaguensis infecting these plants, as well as the first report of M. mayaguensis in Lee and Hillsborough Counties. Meloidogyne partityla was found infecting C. illinoensis in Madison County in Florida. This is the first report of this root-knot nematode species in Florida.

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106 Uncommon Meloidogyne spp. may exhibit isozymes that may not be stable between or within populations, or possess phenotypic bands that stain at various intensities depending on the quantity of enzyme present. Therefore, until universal methodologies are accepted for the electrophoretic identification of Meloidogyne spp. via Mdh and Est phenotypes revealed following electrophoresis, the identification of uncommon Meloidogyne spp. must not involve enzyme phenotype comparisons among works that utilize different methodologies. Isozyme band phenotypes and Rm measurements must only be compared and referenced to M. javanica or M. hapla controls that are run concurrently with the unknown samples, and to isozyme phenotypes revealed through identical methodologies as those used to unveil the unknown samples. All isozyme phenotypes published must include well-described gel formulation and enzyme stain concoction methodologies, accurately-measured gel sizes, and Rm measurements of isozyme phenotypes, revealed on gels processed through identical methodologies, to be compared regardless of gel size.

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107 APPENDIX A PICTURES OF MELOIDOGYNE SPP. ESTERASE AND MALATE DEHYDROGENASE ISOZYME PHENOTYPES UNVEILED THROUGH POLYACRYLAMIDE GEL ELECTROPHORESIS ON PHASTSYSTEM AND MINIPORTEIN 3 CELL APPARATUSES Figure A-1. PhastSystem gels exhibiting smeared malate dehydrogenase isozyme phenotypes whose relative migration could not be accurately measured. Malate dehydrogenase Esterase

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108 PhastSystem-Run Gels Figure A-2. Malate dehydrogenase and esterase isozyme phenotypes, revealed using the PhastSystem apparatus, of Meloidogyne females extracted from Hibiscus rosasinensis cv. Pink Versicolor from Alachua County on 02 April 2003. Figure A-3. Malate dehydrogenase and esterase isozyme phenotypes, revealed using the PhastSystem apparatus, of Meloidogyne females extracted from Rosmarinus officinalis from Suwannee County on 30 April 2003. Malate dehydrogenase Esterase

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109 Figure A-4. Malate dehydrogenase and esterase isozyme phenotypes, revealed using the PhastSystem apparatus, of Meloidogyne females extracted from Callistemon viminalis from Lee County on 01 May 2003. Figure A-5. Malate dehydrogenase and esterase isozyme phenotypes, revealed using the PhastSystem apparatus, of Meloidogyne females extracted from Syagrus romanzoffiana from Lee County on 27 May 2003.

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110 Mini-Protean 3 Cell-Run Gels Figure A-6. Malate dehydrogenase isozyme phenotypes, revealed using the miniprotean 3 cell apparatus, of Meloidogyne females extracted from Callistemon viminalis from Lee County on 27 May 2003. Figure A-7. Eterase isozyme phenotypes, revealed using the mini-protean 3 cell apparatus, of Meloidogyne females extracted from Callistemon viminalis from Lee County on 27 May 2003.

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111 Figure A-8. Malate dehydrogenase isozyme phenotypes, revealed using the miniprotean 3 cell apparatus, of Meloidogyne females extracted from Pittosporum tobira from Alachua County on 18 December 2003. Figure A-9. Esterase isozyme phenotypes, revealed using the mini-protean 3 cell apparatus, of Meloidogyne females extracted from Pittosporum tobira from Alachua County on 18 December 2003.

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112 Figure A-10. Malate dehydrogenase isozyme phenotypes, revealed using the mini-protean 3 cell apparatus, of Meloidogyne females extracted from Ruscus aculeatus from Volusia County on 19 December 2003. Figure A-11. Esterase isozyme phenotypes, revealed using the mini-protean 3 cell apparatus, of Meloidogyne females extracted from Ruscus aculeatus from Volusia County on 19 December 2003.

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113 Figure A-12. Malate dehydrogenase isozyme phenotypes, revealed using the mini-protean 3 cell apparatus, of Meloidogyne females extracted from Buddleia davidii from Pinellas County on 30 September 2003. Figure A-13. Esterase isozyme phenotypes, revealed using the mini-protean 3 cell apparatus, of Meloidogyne females extracted from Buddleia davidii from Pinellas County on 30 September 2003.

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114 Figure A-14. Malate dehydrogenase isozyme phenotypes, revealed using the mini-protean 3 cell apparatus, of Meloidogyne females extracted from Brassica rapa cv. Shogoin from Alachua County on 06 January 2004. Figure A-15. Esterase isozyme phenotypes, revealed using the mini-protean 3 cell apparatus, of Meloidogyne females extracted from Brassica rapa cv. Shogoin from Alachua County on 06 January 2004.

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115 Figure A-16. Malate dehydrogenase isozyme phenotypes, revealed using the mini-protean 3 cell apparatus, of Meloidogyne females extracted from Pittosporum tobira cv. Variegata from Lake County on 11 February 2004. Figure A-17. Esterase isozyme phenotypes, revealed using the mini-protean 3 cell apparatus, of Meloidogyne females extracted from Pittosporum tobira cv. Variegata from Lake County on 11 February 2004.

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116 Figure A-18. Malate dehydrogenase isozyme phenotypes, revealed using the mini-protean 3 cell apparatus, of Meloidogyne females extracted from Ruscus aculeatus from Volusia County on 22 February 2004. Figure A-19. Esterase isozyme phenotypes, revealed using the mini-protean 3 cell apparatus, of Meloidogyne females extracted from Ruscus aculeatus from Volusia County on 22 February 2004.

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117 Figure A-20. Malate dehydrogenase isozyme phenotypes, revealed using the mini-protean 3 cell apparatus, of Meloidogyne females extracted from Liriope muscari cv. Evergreen Giant from Hillsborough County on 01 July 2004. Figure A-21. Esterase isozyme phenotypes, revealed using the mini-protean 3 cell apparatus, of Meloidogyne females extracted from Liriope muscari cv. Evergreen Giant from Hillsborough County on 01 July 2004.

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118 Figure A-22. Malate dehydrogenase isozyme phenotypes, revealed using the mini-protean 3 cell apparatus, of Meloidogyne females extracted from Ophiopogon japonicus from Orange County on 16 August 2004. Figure A-23. Esterase isozyme phenotypes, revealed using the mini-protean 3 cell apparatus, of Meloidogyne females extracted from Ophiopogon japonicus from Orange County on 16 August 2004.

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119 Figure A-24. Malate dehydrogenase isozyme phenotypes, revealed using the mini-protean 3 cell apparatus, of Meloidogyne females extracted from Jesticia carnea from Hillsborough County on 20 August 2004. Figure A-25. Esterase isozyme phenotypes, revealed using the mini-protean 3 cell apparatus, of Meloidogyne females extracted from Jesticia carnea from Hillsborough County on 20 August 2004.

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120 Figure A-26. Malate dehydrogenase isozyme phenotypes, revealed using the mini-protean 3 cell apparatus, of Meloidogyne females extracted from Solenostemon scutellarioides cv. Elfers from Hillsborough County on 15 September 2004. Figure A-27. Esterase isozyme phenotypes, revealed using the mini-protean 3 cell apparatus, of Meloidogyne females extracted from Solenostemon scutellarioides cv. Elfers from Hillsborough County on 15 September 2004.

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121 Figure A-28. Malate dehydrogenase and esterase isozyme phenotypes, revealed using the mini-protean 3 cell apparatus, of Meloidogyne females extracted from Brassica oleracea from Orange County on 11 January 2004. Figure A-29. Malate dehydrogenase and esterase isozyme phenotypes, revealed using the mini-protean 3 cell apparatus, of Meloidogyne females extracted from Beta vulgaris from Orange County on 11 January 2004. Malate dehydrogenase Esterase

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122 Figure A-30. Malate dehydrogenase and esterase isozyme phenotypes, revealed using the mini-protean 3 cell apparatus, of Meloidogyne females extracted from Phaseolus vulgaris from Hillsborough County on 25 January 2005.

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123 APPENDIX B COLLECTIVE RECORD OF THE HOST STATUS OF ORNAMENTAL PLANTS TO MELOIDOGYNE INCOGNITA M. JAVANICA M. ARENARIA AND M. HAPLA The following list (Table B-2) is a collective record of the host status of ornamental plants to Meloidogyne incognita M. javanica M. arenaria and M. hapla as reported by several authors and gathered from this study. The host statuses of listed plants are reported herein as they are in the original works from which they are reported. Scientific names were verified and misspelled names were corrected as listed by the International Plant Names Index (IPNI) (www.inpi.org). Referenced publications, referred to as Source in table B-2, are cited in table B-1.

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124 Table B-1. Sources and citations of publications referred to in Appendix B. Source Citation SourceCitation 1 Ahuja and Arora, 1980 29 McSorley and Marlatt, 1983 2 Bala and Hosein, 1996 30 McSorley, 1994 3 Barker and Benson, 1977 31 Mishra and Mishra, 1997 4 Barker et al., 1979 32 Mishra and Misra, 1993 5 Benson and Barker, 1982 33 Misra and Mishra, 1997 6 Bernard and Witte, 1987 34 Misra et al., 2002 7 Bernard et al., 1994 35 Montasser, 1995 8 Bureau of Nematology, 1989 36 Moreno et al., 1992 9 Caveness and Wilson, 1977 37 Motsinger et al., 1977 10 Cho et al., 1996 38 Nemec and Morrison, 1972 11 Coolen and Hendrickx, 1972 39 Nemec and Struble, 1968 12 Davis and Jenkins, 1960 40 Niblack and Bernard, 1985 13 Eisenback, 1987 41 Nigh, 1972 14 Giblin-Davis et al., 1992 42 Pant et al., 1983 15 Haasis et al., 1961 43 Rajendran et al., 1975 16 Haseeb and Pandey, 1987 44 Santamour and Riedel, 1993 17 Heald, 1967 45 Santamour and Riedel, 1995 18 Khan and Khan, 1989 46 Santamour, 1992 19 Khanna et al., 1998 47 Santo and Lear, 1976 20 Kirby, 1978 48 Sasser et al., 1966 21 LaMondia, 1995 49 Singh and Gupta, 1993 22 LaMondia, 1996 50 Singh and Majeed, 1991 23 LaMondia, 1997 51 Tarjan, 1952 24 Lehman, 1984b 52 Viglierchio, 1979 25 McSorley and Dunn, 1989 53 Walker and Melin, 1998b 26 McSorley and Dunn, 1990 54 Walker et al., 1994 27 McSorley and Frederick, 1994 55 Walker, 1980 28 McSorley and Frederick, 2001

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125Table B-2. Collective record of the host status of ornamental plants to Meloidogyne incognita M. javanica M. arenaria and M. hapla. M. incognita race M. javanica M. arenaria race M. hapla Plant species Cultivar Common Name 12 3 4? 12 ? Source Abelia grandiflora --Glossy abelia Sa 6 Acanthus spinosissimus --Bears breeches S 21 Acer campestre --Hedge maple SS SS 46 Acer davidii --Maple SS SS S 46 Acer francheti --Maple RR S R 46 Acer grandidentatum --Bigtooth maple SR S R 46 Acer grosseri --Grossers maple SS SS S 46 Acer macrophyllum --Big leaf maple SS SS S 46 Acer mono --Painted maple SS SS 46 Acer negundo --Boxelder SS SS S 46 Acer palmatum --Japanese maple R 6 Acer palmatum --Japanese maple SS SS S 46

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126 Table B-2. Continued M. incognita race M. javanica M. arenaria race M. hapla Plant species Cultivar Common Name 1 2 3 4? 12? Source Acer pentaphyllum --Maple RR S R 46 Acer platanoides --Norway maple SS SS R 46 Acer pseudoplatanus --Sycamore maple RS SS S 46 Acer rubrum --Red maple S 40 Acer rubrum --Red maple SR SS R 46 Acer saccharinum --Silver maple SS SS R 46 Acer saccharum --Sugar maple R 6 Acer saccharum --Sugar maple SR SS R 46 Acer tartaricum --Tartarian maple SS SS S 46 Acer truncatum --Shantung maple SS SS S 46 Acer velutinum --Maple SS SS R 46 Achillea Anthea Yarrow S S 53 Achillea Coronation Gold Yarrow R 21

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127 Table B-2. Continued M. incognita race M. javanica M. arenaria race M. hapla Plant species Cultivar Common Name 1 2 3 4? 12? Source Aconitum carmichaelii Arendsii Monkshood S 21 Aconitum lycoctonum ssp. ranunculifolium --Monkshood S 23 Acorus calamus --Calamus S 16 Acroclinium roseum --Paper flower R 1 Acroclinium roseum --Paper flower S 42 Adenophora confusa --Ladybells S 22 Aesculus flava --Yellow buckeye SS SS 44 Aethionema cordifolium --Stone cress R R 53 Agapanthus umbellatus --Agapanthus R 35 Ageratum conyzoides --Tropical whiteweed S 50 Ageratum houstonianum --Ageratum S 1 Ageratum houstonianum --Ageratum S 50

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128 Table B-2. Continued M. incognita race M. javanica M. arenaria race M. hapla Plant species Cultivar Common Name 1 2 3 4? 12? Source Ageratum houstonianum Blue Blazer Ageratum R 54 Ageratum houstonianum Blue Danube Ageratum R 54 Ageratum houstonianum Blue Mink Ageratum R R R 27 Ageratum houstonianum Hawaii White Ageratum R 54 Ageratum houstonianum Royal Delft Ageratum R 54 Aglaonema commutatum Treubii Philippine evergreen S 33 Ailanthus altissima --Tree of heaven RR RR R 44 Ajuga reptans --Common bugle S S S S S 26 Ajuga reptans Bronze Beauty Bugleweed S 23 Ajuga reptans Burgundy Glow Bugleweed S 21 Alcea rosea --Hollyhock S 50 Alcea rosea Chater's Doubles Hollyhock SL 21 Alchemilla mollis Improved Form Lady's mantle SL 21

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129 Table B-2. Continued M. incognita race M. javanica M. arenaria race M. hapla Plant species Cultivar Common Name 1 2 3 4? 12? Source Aloe perryi --Perry's aloe SS 16 Aloe vera --Aloe vera SS 16 Aloe vera --Aloe vera S 33 Aloe vera --Aloe vera S 2 Alpinia galanga --Greater galangal S 16 Amaryllis vittata --Amaryllis R 35 Amberboa moschata --Sweet sultan S 1 Anchusa azurea Dropmore Alkanet S 22 Andrographis paniculata --Andrographis S 16 Anemone Queen Charlotte Windflower S 22 Anemone coronaria --Cut-leaf anemone S 35 Anemone hupehensis var. j aponica --Japanese thimbleweed S 23

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130 Table B-2. Continued M. incognita race M. javanica M. arenaria race M. hapla Plant species Cultivar Common Name 1 2 3 4? 12? Source Antirrhinum majus --Antirrhinum S 1 Antirrhinum majus --Snapdragon S 42 Antirrhinum majus --Snapdragon S 50 Antirrhinum majus First Ladies Snapdragon S S S 27 Antirrhinum majus Margaret Snapdragon SS SS 51 Aquilegia caerulea Blue Star Columbine S 21 Arabis caucasica Compinkie Rock cress SL 21 Arctotis stoechadifolia --African daisy S 1 Arctotis stoechadifolia --African daisy S 50 Arctotis venusta --Blue eyed African daisy S 50 Areca catechu --Betel palm R R R RR 25 Argyreia nervosa --Elephant creeper S 16 Artemisia schmidtiana Silver Mound Silver mound S 21

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131 Table B-2. Continued M. incognita race M. javanica M. arenaria race M. hapla Plant species Cultivar Common Name 1 2 3 4? 12? Source Asclepias curassavica --Blood flower SS 16 Asclepias tuberosa --Butterfly weed R 22 Asparagus densiflorus --Sprenger's asparagus fern S 33 Aspidistra elatior --Cast iron plant S 33 Aster novae-angliae Harrington's Pink New England aster R 21 Aster novae-angliae September Ruby New England aster R 21 Aster novi-belgii Mt. Everest New York aster R 23 Astilbe arendsii Peach Blossom Feather flower S 21 Astrantia major Rose Symphony Great masterwort S 22 Asystasia gangetica --Chinese violet S 43 Aucuba japonica Variegata Acuba S 4 Balsamita major --Costmary R 50 Barleria prionitis --Porcupine flower R 43

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132 Table B-2. Continued M. incognita race M. javanica M. arenaria race M. hapla Plant species Cultivar Common Name 1 2 3 4? 12? Source Barringtonia acutangula --Freshwater mangrove SS 16 Basella alba --Ceylon spinach S 33 Begonia Cocktail Gin Begonia S 54 Begonia Cocktail Vodka Begonia SL 54 Begonia Encore White BronzeBegonia S 54 Begonia Party Love Begonia S 54 Begonia Pizzazz Deep Rose Begonia S 54 Belamcanda chinensis --Blackberry lily R 21 Bellis perennis --Lawn daisy S 1 Bellis perennis --Lawn daisy S 42 Bellis perennis --Lawn daisy S 50 Betula nigra Heritage River birch S 44

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133 Table B-2. Continued M. incognita race M. javanica M. arenaria race M. hapla Plant species Cultivar Common Name 1 2 3 4? 12? Source Betula platyphylla var. j aponica Japanese white birch SS S S 44 Betula populifolia --Gray birch SS SS 44 Boltonia asteroides Pink Beauty White dolls daisy S 22 Borago officinalis --Common borage S S S S 36 Brassica oleracea var. acephala Kale S 1 Buxus harlandii --Japanese boxwood SL 6 Buxus microphylla --Littleleaf boxwood S S 56 Buxus microphylla Japonica Littleleaf boxwood S 5 Buxus sempervirens --Common boxwood SL 6 Calathea zebrina --Zebra plant S 33 Calendula officinalis --Pot marigold S 1 Calendula officinalis --Pot marigold S 42

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134 Table B-2. Continued M. incognita race M. javanica M. arenaria race M. hapla Plant species Cultivar Common Name 1 2 3 4? 12? Source Calendula officinalis --Pot marigold S 50 Calendula persica --Calendula S 50 Callistemon citrinus --Crimson bottlebrush S 16 Callistemon lanceolatus --Crimson bottlebrush SS 16 Callistephus chinensis --China aster S 1 Callistephus chinensis --China aster S 50 Camellia japonica --Common camellia SS 15 Camellia sasanqua --Sasanqua camellia SS 15 Camellia sasanqua --Sasanque camellia S 39 Campanula poscharskyana --Bellflower S 21 Canna indica --Indian shot R 35 Catharanthus roseus Blush Cooler Madagascar periwinkle S S 28

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135 Table B-2. Continued M. incognita race M. javanica M. arenaria race M. hapla Plant species Cultivar Common Name 1 2 3 4? 12? Source Catharanthus roseus Carpet Dawn Madagascar periwinkle SL 54 Catharanthus roseus Grape Cooler Madagascar periwinkle SL R 28 Catharanthus roseus Little Blanche Madagascar periwinkle SL 54 Catharanthus roseus Little Bright Eyes Madagascar periwinkle S S 28 Catharanthus roseus Little Delicata Madagascar periwinkle SL 54 Catharanthus roseus Little Mixed Colors Madagascar periwinkle S S 28 Catharanthus roseus Peppermint Cooler Madagascar periwinkle S S 28 Catharanthus roseus Polka Dot Madagascar periwinkle SL 54 Cedrus deodara --Deodar cedar R 39 Celastrus dependens --Magzsudhi S 16 Celosia argentea --Silver cock's comb S 42 Celosia argentea Accession no. 8 Silver cock's comb SS 9 Celosia argentea Century Mix Silver cock's comb S S S 27

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136 Table B-2. Continued M. incognita race M. javanica M. arenaria race M. hapla Plant species Cultivar Common Name 1 2 3 4? 12? Source Celosia cristata --Crested cocks comb S 50 Celosia plumosa Apricot Brandy Celosia S 54 Celosia plumosa Castle Scarlet Celosia S 54 Celosia plumosa Fireglow Celosia S 54 Celosia plumosa Kimona Cream Celosia S 54 Celtis occidentalis --Common hackberry SS SS R 44 Centaurea cyanus --Corn flower S 50 Centaurea cyanus --Cornflower S 1 Centranthus ruber Albus Jupiters beard S 22 Cercis Canadensis --Eastern redbud tree SS SS S 45 Cercis canadensis var. texensis --Texas redbud SS SS S 45 Cercis chinensis --Chinese redbud SS SS S 45

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137 Table B-2. Continued M. incognita race M. javanica M. arenaria race M. hapla Plant species Cultivar Common Name 1 2 3 4? 12? Source Cercis racemosa --Chain flowered redbud SS SS S 45 Cercis siliquastrum --Judas tree SS SS S 45 Cercis yunnanensis --Yunnan redbud SS SS S 45 Chamaecyparis pisifera --Sawara cypress R 39 Cheiranthus cheiri --Aegean wallflower S 1 Cheiranthus cheiri --Aegean wallflower S 50 Chelone obliqua --Red turtlehead R 21 Chlorophytum comosum --Spider plant S 33 Chrysanthemum carinatum --Tricolor daisy R 42 Chrysanthemum carinatum --Tricolor daisy R 50 Chrysanthemum coccineum Giant Hybrids Pyrethum daisy S 21

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138 Table B-2. Continued M. incognita race M. javanica M. arenaria race M. hapla Plant species Cultivar Common Name 1 2 3 4? 12 ? Source Chrysanthemum coronarium --Crowndaisy R 1 Cimicifuga acerina --Fairy candles S 21 Cimicifuga dahurica --Asian bugbane S 21 Cimicifuga simplex White Pearl Single-stem bugbane S 21 Cissus quadrangularis --Veld grape S 16 Clarkia unguiculata --Elegant fairyfan S 1 Clarkia unguiculata --Elegant fairyfan S 50 Clematis Hagley Hybrid Clematis S 21 Clivia miniata --Kaffir lily S 33 Clivia miniata --Kaffir lily R 35 Codiaeum variegatum Gold Dust Croton R R R Chapter 2 Cordyline fruticosa --Ti plant S 33

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139 Table B-2. Continued M. incognita race M. javanica M. arenaria race M. hapla Plant species Cultivar Common Name 1 2 3 4? 12 ? Source Coreopsis stillmanii --Stillmans tickseed R 1 Coreopsis tinctoria --Golden tickseed R 50 Coreopsis verticillata Moonbeam Tickseed S 21 Cornus florida --Eastern flowering dogwood S 6 Cornus florida --Eastern flowering dogwood S 40 Cosmos bipinnatus --Garden cosmos R 1 Cosmos bipinnatus --Garden cosmos R 50 Cosmos sulphunus --Sulphur cosmos SL 42 Costus speciosus --Wild ginger SS 16 Cotoneaster horizontalis --Rockspray cotoneaster SL 39 Crataeva nurvala --Three-leaf caper S 16 Crinum longifolium --Hardy swamplily R 35 Crossandra undulifolia Orange Firecracker flower S 43

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140 Table B-2. Continued M. incognita race M. javanica M. arenaria race M. hapla Plant species Cultivar Common Name 1 2 3 4? 12 ? Source Crossandra undulifolia Red Firecracker flower S 43 Croton sparsiflorus --Croton S 42 Cucumis melo ssp. melo Flexuosus Snakemelon SS 57 Dahlia pinnata --Pinnate dahlia S 42 Dahlia variabilis --Dahlia S 1 Dahlia variabilis --Dahlia R 35 Dahlia variabilis --Dahlia S 50 Delphinium ajacis --Larkspur S 1 Delphinium ajacis --Larkspur S 50 Delphinium grandiflorum Blue Mirror Siberian larkspur S 21 Dendranthema indicum --Chrysanthemum R 50

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141 Table B-2. Continued M. incognita race M. javanica M. arenaria race M. hapla Plant species Cultivar Common Name 1 2 3 4? 12 ? Source Dendranthema morifolium --Florists daisy R 1 Dianella caerulea --Cerulean flaxlily S 33 Dianthus barbatus --Sweet william R 1 Dianthus barbatus --Sweet william S 50 Dianthus barbatus Indian Carpet Sweet william R 21 Dianthus caryophyllus --Carnation S 1 Dianthus caryophyllus --Carnation R 50 Dianthus caryophyllus Antalia Carnation SL 10 Dianthus caryophyllus Astra Carnation S 10 Dianthus caryophyllus Beta Carnation S 10 Dianthus caryophyllus Carmit Carnation SL 10 Dianthus caryophyllus Castelaro Carnation SL 10

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142 Table B-2. Continued M. incognita race M. javanica M. arenaria race M. hapla Plant species Cultivar Common Name 1 2 3 4? 12 ? Source Dianthus caryophyllus Darling Carnation S 10 Dianthus caryophyllus Desio Carnation SL 10 Dianthus caryophyllus Echo Carnation SL 10 Dianthus caryophyllus Elegance Korea Carnation S 10 Dianthus caryophyllus Espana Carnation SL 10 Dianthus caryophyllus Galil Carnation S 10 Dianthus caryophyllus Imperial White Sim Carnation S 10 Dianthus caryophyllus Izu Pink Carnation SL 10 Dianthus caryophyllus Kappa Carnation R 10 Dianthus caryophyllus Lena Carnation S 10 Dianthus caryophyllus Mars Carnation S 10 Dianthus caryophyllus Mercury Carnation SL 10 Dianthus caryophyllus Rachel Carnation SL 10

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143 Table B-2. Continued M. incognita race M. javanica M. arenaria race M. hapla Plant species Cultivar Common Name 1 2 3 4? 12 ? Source Dianthus caryophyllus Rara Carnation SL 10 Dianthus caryophyllus Red Corso Carnation SL 10 Dianthus caryophyllus Red Lena Carnation S 10 Dianthus caryophyllus Roland Carnation S 10 Dianthus caryophyllus Rony Carnation SL 10 Dianthus caryophyllus Sarinah Carnation S 10 Dianthus caryophyllus Saturn Carnation SL 10 Dianthus caryophyllus Saturnus Carnation SL 10 Dianthus caryophyllus Scarlet Elegance Carnation S 10 Dianthus caryophyllus Shinkibo Carnation S 10 Dianthus caryophyllus Target Carnation SL 10 Dianthus caryophyllus Tasman Carnation S 10 Dianthus caryophyllus Virgo Carnation S 10

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144 Table B-2. Continued M. incognita race M. javanica M. arenaria race M. hapla Plant species Cultivar Common Name 1 2 3 4? 12 ? Source Dianthus caryophyllus White Royalitee Carnation SL 10 Dianthus caryophyllus Yellow Dusty Carnation S 10 Dianthus chinensis --Rainbow pink S 42 Dianthus chinensis --Rainbow pink R 50 Dianthus chinensis Baby Doll Mix Rainbow pink S SL S 30 Dianthus chinensis Princess Scarlet Rainbow pink S 54 Dicentra spectabilis Alba White bleeding heart S 21 Dieffenbachia amoena Tropic Snow Dumb cane S 33 Digitalis ambigua --Yellow foxglove SL 21 Digitalis lanata --Grecian foxglove SS 16 Digitalis purpurea --Common foxglove S 16 Digitalis purpurea Excelsior Common foxglove SL 21 Doronicum orientale Magnificum Oriental false leopardbane S 21

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145 Table B-2. Continued M. incognita race M. javanica M. arenaria race M. hapla Plant species Cultivar Common Name 1 2 3 4? 12 ? Source Dracaena marginata --Dragontree SL SL S SL R 26 Echinacea purpurea --Purple coneflower R R 53 Echinacea purpurea Leuchtstern Purple coneflower R 22 Echinops bannaticus Taplow Blue Globe thistle SL 22 Elaeagnus pungens --Elaeagnus R 39 Elytraria acaulis --Scalystem S 16 Epimedium versicolor Sulphureum Yellow barrenwort R 22 Eriobotrya japonica --Loquat SS 41 Eschscholtzia californica --California poppy S 50 Eschscholtzia californica --Californian poppy S 1 Euonymus alatus Compacta Winged spindletree R 6 Euonymus japonicus --Japanese spindletree S 41 Euphorbia thymifolia --Gulf sandmat S 16

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146 Table B-2. Continued M. incognita race M. javanica M. arenaria race M. hapla Plant species Cultivar Common Name 1 2 3 4? 12 ? Source Evolulus glomeratus Blue Daze Evolvulus S 8 Fagus grandifolia --American beech RR SS R 44 Ficus benjamina --Weeping fig S S S S R 26 Ficus benjamina --Weeping fig S 33 Ficus elastica Decora Indian rubber tree S 33 Filipendula rubra Venusta Magnifica Queen of the prairie SL 21 Flacourtia indica --Governors plum S 16 Fragaria ananassa Pink Panda Hybrid strawberry R R 53 Freesia --Freesia S 35 Gaillardia grandiflora Goblin Blanket flower R 21 Gaillardia pulchella Picta Blanket flower R 42 Gardenia jasminoides --Gardenia S S 12 Gardenia jasminoides --Gardenia S SS 24

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147 Table B-2. Continued M. incognita race M. javanica M. arenaria race M. hapla Plant species Cultivar Common Name 1 2 3 4? 12 ? Source Gardenia jasminoides Radicans Gardenia SL 5 Gentiana sp. Benichidori Gentian S 22 Geranium Ann Folkard Cranesbill S S 53 Geranium magnificum --Cranesbill S 22 Geranium oxonianum Thurstonianum Cranesbill S 22 Geranium cinereum Laurence Flatman Cranesbill S S 53 Geranium dalmaticum --Dalmatian cranesbill S 21 Geranium endressii Wargrave Pink Cranesbill S 23 Gerbera jamesonii Nain-Crimson Barberton daisy SL 54 Ginkgo biloba --Maidenhair tree SS SS S 44 Gladiolus Oscar Gladiola S 19 Gladiolus Pink Friendship Gladiola S 19 Gladiolus Snow Princess Gladiola S 19

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148 Table B-2. Continued M. incognita race M. javanica M. arenaria race M. hapla Plant species Cultivar Common Name 1 2 3 4? 12 ? Source Gladiolus Sylvia Gladiola S 19 Gladiolus Vinks Glory Gladiola S 19 Gladiolus White Friendship Gladiola S 19 Gladiolus gandavensis --Gladiola S 35 Gladiolus spp. --Gladiola S 33 Gleditsia triacanthos --Threespined honeyocust RR RR R 44 Gymnema sylvestre --Miracle fruit S 16 Gypsophila elegans Covent Garden Showy babys breath S 30 Hedera helix --English ivy SL 39 Hedychium coronarium --White gingerlily SL 35 Helenium autumnale Brilliant Common sneezeweed R 21 Helianthus annuus --Common sunflower S 50 Helichrysum bracteatum --Bracted strawflower S 50

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149 Table B-2. Continued M. incognita race M. javanica M. arenaria race M. hapla Plant species Cultivar Common Name 12 3 4? 12 ? Source Helictotrichon sempervirens --Helictotrichon S 22 Heliopsis helianthoides Karat Sunflower heliopsis S 21 Hemerocallis Bright Banner Daylily SL 22 Hemerocallis aurantiaca --Hardy daylily SL 35 Hemerocallis fulva Marion Vaughn Common daylily S 33 Heuchera cylindrica Green Ivory Roundleaf alum root S S 53 Heucherella alba Bridget Bloom Foamy bells S S 53 Hibiscus rosa-sinensis --Hibiscus SS 41 Hibiscus rosa-sinensis Anderson Crepe Hibiscus SLSL 29 Hibiscus rosa-sinensis Delight Hibiscus SLR 29 Hibiscus rosa-sinensis Fancy Lady Hibiscus SSL 29 Hibiscus rosa-sinensis Florida Sunset Hibiscus SLR 29

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150 Table B-2. Continued M. incognita race M. javanica M. arenaria race M. hapla Plant species Cultivar Common Name 1 2 3 4? 12 ? Source Hibiscus rosa-sinensis Go-Go Girls Hibiscus SLSL 29 Hibiscus rosa-sinensis Kona Hibiscus S 29 Hibiscus rosa-sinensis Minerva Hibiscus SSL 29 Hibiscus rosa-sinensis Old Gold Hibiscus SLSL 29 Hibiscus rosa-sinensis Painted Lady Hibiscus SSL 29 Hibiscus rosa-sinensis Painted Lady Hibiscus SS 29 Hibiscus rosa-sinensis Philipino Hibiscus SSL 29 Hibiscus rosa-sinensis President Hibiscus SLSL 29 Hibiscus rosa-sinensis President Hibiscus SLSL 29 Hibiscus rosa-sinensis Pride of Hankins Hibiscus SS 29 Hibiscus rosa-sinensis Rowena Wedding Hibiscus SSL 29 Hibiscus rosa-sinensis Versicolor Pink Hibiscus S Chapter 3

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151 Table B-2. Continued M. incognita race M. javanica M. arenaria race M. hapla Plant species Cultivar Common Name 1 2 3 4? 12 ? Source Hyacinthus orientalis --Garden hyacinth R 35 Hydrangea paniculata Grandiflora Old fashioned snowball S 6 Hygrophila auriculata --Hygrophila SS 16 Hypericum polyphyllum --Saint John's wort S 21 Iberis amara --Iberis S 42 Ilex Calina Holly S S 7 Ilex Little Red Holly S S 56 Ilex attenuata Foster No. 2 Attenuate holly R 6 Ilex attenuata Foster No. 2 Attenuate holly S S 7 Ilex meserveae Blue Boy Blue holly S S 7 Ilex meserveae Blue Girl Blue holly S S 7 Ilex cassine Lowei Dahoon holly S S 7

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152 Table B-2. Continued M. incognita race M. javanica M. arenaria race M. hapla Plant species Cultivar Common Name 12 3 4? 12 ? Source Ilex cassine var. angustifolia --Dahoon holly S 39 Ilex cornuta Burfordii Chinese holly R R R R R 26 Ilex cornuta Burfordii Chinese holly S 39 Ilex cornuta Burfordii Chinese holly SS RR 48 Ilex cornuta Burfordii Holly R 4 Ilex cornuta Carissa Chinese holly R SL 56 Ilex cornuta Needlepoint Chinese holly S S 7 Ilex cornuta Rotunda Chinese holly S 4 Ilex cornuta I. aquifolium Nellie R. Stevens Holly S S 7 Ilex cornuta I. aquifolium Nellie R. Stevens Holly R 6 Ilex cornuta I. pernyi Lydia Morris Holly S S 7

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153 Table B-2. Continued M. incognita race M. javanica M. arenaria race M. hapla Plant species Cultivar Common Name 1 2 3 4? 12 ? Source Ilex crenata Bennetts CompactaJapanese holly S S 7 Ilex crenata Compacta Japanese holly S 5 Ilex crenata Compacta Japanese holly S S 56 Ilex crenata Convexa Japanese holly S 3 Ilex crenata Convexa Japanese holly S S 7 Ilex crenata Convexa Japanese holly SS SR 48 Ilex crenata Green Luster Japanese holly S S 56 Ilex crenata Green Lustre Japanese holly S S 7 Ilex crenata Helleri Japanese holly S 3 Ilex crenata Helleri Japanese holly SS SS 17 Ilex crenata Helleri Japanese holly SS SR 48 Ilex crenata Helleri Japanese holly S S 56 Ilex crenata Hetz Japanese holly S 39

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154 Table B-2. Continued M. incognita race M. javanica M. arenaria race M. hapla Plant species Cultivar Common Name 1 2 3 4? 12 ? Source Ilex crenata Hetzii Japanese holly R 6 Ilex crenata Latifolia Japanese holly SS SR 48 Ilex crenata Rotundifolia Japanese holly S 3 Ilex crenata Rotundifolia Japanese holly S S 7 Ilex glabra Compacta Inkberry S S 7 Ilex glabra Nordic Inkberry S S 7 Ilex glabra Shamrock Inkberry R R 56 Ilex integra I. pernyi Elegance Holly S S 7 Ilex opaca Jersey Princess American holly S 7 Ilex vomitoria Nana Yaupon holly S 4 Ilex vomitoria Schelling's Dwarf Yaupon holly R R 56 Ilex vomitoria Stokes Dwarf Yaupon holly S S 7 Impatiens balsamina --Spotted snapweed S 1

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155 Table B-2. Continued M. incognita race M. javanica M. arenaria race M. hapla Plant species Cultivar Common Name 1 2 3 4? 12 ? Source Impatiens balsamina --Spotted snapweed S 42 Impatiens balsamina --Spotted snapweed S 50 Impatiens holstii Aztec Impatiens S 55 Impatiens holstii Chickasaw Impatiens S 55 Impatiens holstii Chippewa Impatiens S 55 Impatiens holstii Creek Impatiens S 55 Impatiens holstii Fuchsia Impatiens S 55 Impatiens holstii Futura Red Impatiens S 55 Impatiens holstii Futura White Impatiens S 55 Impatiens holstii Garden Blue Impatiens S 55 Impatiens holstii Hopi Impatiens S 55 Impatiens holstii Maya Impatiens S 55 Impatiens holstii Navajo Impatiens S 55

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156 Table B-2. Continued M. incognita race M. javanica M. arenaria race M. hapla Plant species Cultivar Common Name 12 3 4? 12 ? Source Impatiens holstii PI 354261 Impatiens S 55 Impatiens holstii PI 354264 Impatiens S 55 Impatiens holstii PI 354265 Impatiens S 55 Impatiens holstii Scarlet Impatiens S 55 Impatiens holstii Scarlet Baby Impatiens S 55 Impatiens holstii Series F1 (Pink) Impatiens S 55 Impatiens holstii Series F1 (Rose) Impatiens S 55 Impatiens holstii Series F1 (Salmon) Impatiens S 55 Impatiens holstii Shawnee Impatiens S 55 Impatiens holstii Twinkles Impatiens S 55 Iris germanica Afternoon Delight German iris S 21 Iris pumila Elfin Queen Dwarf iris SL 21 Iris sibirica Maranantha Siberian iris R 21

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157 Table B-2. Continued M. incognita race M. javanica M. arenaria race M. hapla Plant species Cultivar Common Name 1 2 3 4? 12 ? Source Iris tingitana --Morocco iris S 35 Ixora arborea --Jungleflame S 16 Ixora casei Super King Ixora R R 14 Ixora coccinea --Scarlet jungleflame S S 14 Ixora coccinea Maui Scarlet jungleflame S S 14 Ixora coccinea Nora Grant Scarlet jungleflame R R 14 Ixora coccinea Petite Red Scarlet jungleflame S S 14 Ixora coccinea Petite Yellow Scarlet jungleflame SL S 14 Ixora coccinea Singapore Scarlet jungleflame SL S 14 Ixora coccinea I. Chinensis Bonnie Lynn Ixora S S 14 Jasminum humile --Italian yellow jasmine S 16 Jasminum nudiflorum --Winter jasmine S 39

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158 Table B-2. Continued M. incognita race M. javanica M. arenaria race M. hapla Plant species Cultivar Common Name 1 2 3 4? 12 ? Source Juglans nigra --Black walnut RR RR R 44 Juniperus chinensis Hetzii Glauca Blue hetzi juniper R 6 Juniperus conferta Blue Pacific Blue pacific shore juniper R 6 Juniperus excelsa Stricta Greek juniper SL 5 Juniperus horizontalis --Creeping juniper R 5 Juniperus horizontalis Douglasii Creeping juniper SL 39 Juniperus horizontalis Plumosa Creeping juniper R R 6 Juniperus horizontalis Plumosa Creeping juniper R R 37 Juniperus horizontalis Plumosa Creeping juniper R 39 Justicia betonica --Squirrels tail S 43 Kalanchoe blossfeldiana --Madgascar widows thrill S 33 Kalanchoe fedtschenkoi --South American air plant S 34 Kochia trichophylla --Kochia R 50

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159 Table B-2. Continued M. incognita race M. javanica M. arenaria race M. hapla Plant species Cultivar Common Name 1 2 3 4? 12 ? Source Koelreuteria paniculata --Golden rain tree SS SS S 44 Lantana camara Miss Huff South American lantana R R 56 Lantana camara New Gold South American lantana SL R 56 Lathyrus latifolis --Perennial seet pea S 21 Lathyrus odoratus --Sweet pea S 1 Lathyrus odoratus --Sweet pea S 50 Lavandula angustifolia Munstead Dwarf English lavender S 21 Lavendula spica --Lavender S S S S 36 Leucanthemum superbum Exhibition Shasta daisy R 21 Leucanthemum superbum Polaris Shasta daisy S 21 Leucanthemum maximum Alaska Shasta daisy S 30

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160 Table B-2. Continued M. incognita race M. javanica M. arenaria race M. hapla Plant species Cultivar Common Name 1 2 3 4? 12 ? Source Liatris scariosa White Spires Devils bite R 21 Ligularia dentata Desdemona Strain Leopard plant S 21 Ligustrum sinense Variegatum Chinese privet S 6 Lilium longiflorum --Easter lily R 35 Linaria bipartita --Clovenlip toadflax R 1 Linaria bipartita --Clovenlip toadflax R 50 Linaria cymbalaria --Toadflax S S 53 Linum grandiflorum --Flowering flax R 1 Linum grandiflorum Cocineum Flowering flax R 50 Liquidambar styraciflua --Sweetgum SR RS R 44 Liriope muscari --Lilyturf S 33 Liriope muscari Evergreen Giant Liliturf S Chapter 3

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161 Table B-2. Continued M. incognita race M. javanica M. arenaria race M. hapla Plant species Cultivar Common Name 1 2 3 4? 12 ? Source Liriope muscari Evergreen Giant Liliturf S S SL Chapter 2 Liriope muscari Variegata Lilyturf R 22 Lithospermum diffusum Grace Ward Stoneseed R 22 Lobelia gerardi Vedrariensis Lobelia R 23 Lobelia cardinalis Complement ScarletCardinal flower S 21 Lobelia erinus --Edging lobelia S 1 Lobelia erinus --Edging lobelia S 50 Lobularia maritima --Sweet alyssum S 1 Lobularia maritima --Sweet alyssum S 50 Lobularia maritima Rosie O'Day Sweet alyssum R R 27 Loropetalum chinense --Razzle bush S 39 Lupinus hartwegii --Hartwegs bluebonnet S 1

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162 Table B-2. Continued M. incognita race M. javanica M. arenaria race M. hapla Plant species Cultivar Common Name 1 2 3 4? 12 ? Source Lupinus polyphyllus Russel Hybrids Bigleaf lupin S 21 Lysimachia clethroides --Gooseneck yellow loosestrife S 22 Lythrum sp. Morden's Pink Loosestrife S 21 Maclura pomifera --Osageorange RR RR R 44 Magnolia soulangiana Alexandrina Saucer magnolia R 6 Magnolia grandiflora --Southern magnolia RR RR R 44 Malva alcea Fastigiata Vervain mallow S 22 Malva moschata Alba Musk mallow SL 21 Marguerita sp. ----S 42 Matthiola incana --Tensweeks stock R 42 Matthiola incana --Tenweeks stock S 50 Melia azedarach --Chinaberrytree SS 41

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163 Table B-2. Continued M. incognita race M. javanica M. arenaria race M. hapla Plant species Cultivar Common Name 12 3 4? 12 ? Source Melissa officinalis --Common balm S 16 Mesembranthemum tricolor --Ice plant S 50 Mesembranthemum tricolor --Ice plant S 1 Metasequoia glyptostroboides --Dawn redwood R 6 Miscanthus sinensis Silberfeder Silver feather S 22 Moluccella laevis --Bells of Ireland R 1 Monarda citriodora --Lemon bee balm R R 53 Monarda didyma Cambridge Scarlet Scarlet bee balm R 21 Monarda didyma Cambridge Scarlet Scarlet bee balm R R 53 Morus alba --White mulberry SS 41 Murraya koenigii --Curry leaf tree S 33

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164 Table B-2. Continued M. incognita race M. javanica M. arenaria race M. hapla Plant species Cultivar Common Name 1 2 3 4? 12 ? Source Musa sumatrana ssp. zebrina Rowe Red Ornamental banana S S S Chapter 2 Myosotis alpestris Indigo Blue Forget me not R 22 Nandina domestica --Heavenly bamboo R 5 Nandina domestica --Heavenly bamboo S 6 Narcissus tazetta --Cream narcissus R 35 Nasturtium majus --Nasturtium S 50 Nepeta nervosa --Clear blue catmint S S 53 Nicotiana plumbaginifolia --Tex-mex tobacco S 16 Nierembergia hippomanica --Dwarf cupflower S S 53 Nigella damascena --Devil in the bush S 1 Nigella damascena --Devil in the bush S 50

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165 Table B-2. Continued M. incognita race M. javanica M. arenaria race M. hapla Plant species Cultivar Common Name 1 2 3 4? 12 ? Source Ocimum basilicum --Sweet basil S S S S 36 Ocimum canum --Hoary basil SS 16 Ocimum gratissimum --African basil SS 16 Ocimum kilimandscharicum --Hoary basil SS 16 Odontonema cuspidatum --Firespike R R R Chapter 2 Operculina turpethum --St. Thomas lidpot S 16 Ophiopogon japonicus --Mondo grass S Chapter 3 Origanum majorana --Sweet marjoram S R R R 36 Origanum onites --Pot marjoram S S S S 36 Origanum vulgare --Oregano S R S S 36 Osmanthus fortunei --Fortunes osmanthus S 39

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166 Table B-2. Continued M. incognita race M. javanica M. arenaria race M. hapla Plant species Cultivar Common Name 1 2 3 4? 12? Source Pachysandra procumbens --Montain spurge R 21 Pachysandra terminalis --Japanese spurge S 21 Paederia foetida --Stinkvine SS 16 Papaver dubium --Blindeyes S 50 Papaver orientale Carousel Oriental poppy R 21 Papaver orientale Oriental Red Perennial Poppy S 30 Papaver rhoeas --Corn poppy S 1 Papaver rhoeas --Corn poppy S 50 Passiflora edulis var. f lavicarpa --Yellow passion fruit RR R 20 Patrinia scabiosaefolia --Patrinia SL R 53 Penstemon Purple Passion Talus slope penstemon S S 53

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167 Table B-2. Continued M. incognita race M. javanica M. arenaria race M. hapla Plant species Cultivar Common Name 1 2 3 4? 12? Source Penstemon Ruby Talus slope penstemon S S 53 Penstemon Sour Grapes Talus slope penstemon S S 53 Penstemon digitalis Husker Red Talus slope penstemon S 22 Peperomia magnoliaefolia --Peperomia S 33 Perovskia atriplicifolia --Russian sage S 22 Petunia hybrida --Garden petunia S 1 Petunia hybrida --Garden petunia S 18 Petunia hybrida --Garden petunia S 50 Petunia hybrida Dwarf Bedding Garden petunia S S S 27 Petunia hybrida Fire Chief Garden petunia S SL S 30 Petunia violacea --Petunia SL 42 Philodendron laciniatum --Philodendron S 33

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168 Table B-2. Continued M. incognita race M. javanica M. arenaria race M. hapla Plant species Cultivar Common Name 1 2 3 4? 12? Source Philodendron selloum --Philodendron S 32 Philodendron selloum --Philodendron S 33 Phlox drummondii --Annual phlox R 1 Phlox drummondii --Annual phlox R 50 Phlox paniculata Eva Cullum Garden phlox R R 53 Phlox paniculata Fairest One Garden phlox R 21 Phlox paniculata Franz Shubert Garden phlox R R 53 Phlox paniculata Oakington Blue Garden phlox R R 53 Phlox stolonifera Bruce's White Creeping phlox R 21 Photinia fraseri --Copper tip photinia S 6 Photinia fraseri --Copper tip photinia SLSL S SL 26 Physostegia virginiana Summer Snow Obedient plant SL 22 Pinus ponderosa --Ponderosa pine RR R 52

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169 Table B-1. Continued M. incognita race M. javanica M. arenaria race M. hapla Plant species Cultivar Common Name 1 2 3 4? 12? Source Pinus strobus --Eastern white pine R 6 Pinus virginiana --Virginia pine R 6 Pisum sativum --Garden pea S 50 Pittosporum tobira --Pittosporum S Chapter 3 Pittosporum tobira --Pittosporum S 8 Pittosporum tobira --Pittosporum S 41 Pittosporum tobira Variegata Variegated pittosporum R R R Chapter 2 Polemonium reptans Firmament Greek valerian S 21 Polianthes tuberosa --Tuberose S 35 Polianthes tuberosa Double Tuberose S 33 Polianthes tuberosa Single Tuberose S 33 Polygonum affine Dimity Polygonum R R 53

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170 Table B-2. Continued M. incognita race M. javanica M. arenaria race M. hapla Plant species Cultivar Common Name 12 3 4? 12? Source Poncirus trifoliatus --Hardy orange S 39 Portulaca grandiflora --Rose moss R 50 Potentilla nepalensis Miss Wilmott Red potentilla S 21 Primula japonica Red Field Hybrids Primrose R 23 Primula polyantha Crescendo Mix Primrose R 21 Prunus cistena --Cherry R 6 Prunus yedoensis --Tokyo cherry R 6 Prunus avium --Sweet cherry SS SS S 44 Prunus cerasifera --Cherry plum SS SS S 44 Prunus cerasifera Atropurpurea Cherry plum R 6 Prunus glandulosa --Flowering almond R 6 Prunus laurocerasus var. zabeliana --Cherry laurel SL 39

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171 Table B-2. Continued M. incognita race M. javanica M. arenaria race M. hapla Plant species Cultivar Common Name 1 2 3 4? 12? Source Prunus mahaleb --Mahaleb cherry SS SS S 44 Prunus serrulata Kwanzan Japanese flowering cherry R 6 Pyrus calleryana --Callery pear RR RR R 44 Rhododendron Cannon's Double Azalea R 6 Rhododendron Girard's Rose Azalea R 6 Rhododendron catawbiense Boursalt Catawba rosebay R 6 Rhoeo discolor --Boat lily S 33 Robinia pseudoacacia --Black locust SS SS S 44 Rosa Dr. Huey Rose S 47 Rosa canina --Dog rose S 11 Rosa canina Brgs Stachellose Dog rose S 11 Rosa canina Heinsohns Rekord Dog rose S 11

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172 Table B-2. Continued M. incognita race M. javanica M. arenaria race M. hapla Plant species Cultivar Common Name 1 2 3 4? 12? Source Rosa canina Inermis Dog rose S 11 Rosa canina Pfnder Dog rose S 11 Rosa canina Pollmers Dog rose S 11 Rosa canina Schmids Ideal Dog rose S 11 Rosa canina Succes Dog rose S 11 Rosa dumetorum Laxa Corymb rose S 11 Rosa indica --Cyme rose SS 57 Rosa multiflora --Japanese rose S 11 Rosa multiflora --Japanese rose S 47 Rosa noisettiana Manetti Rose SL 47 Rosa odorata --Tea rose S 47 Rosa rubiginosa --Sweetbriar rose S 11 Rosa sp. --Rose SS 41

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173 Table B-2. Continued M. incognita race M. javanica M. arenaria race M. hapla Plant species Cultivar Common Name 1 2 3 4? 12? Source Rosemarinus officinalis --Rosemary S S S S 36 Rudbeckia fulgida Goldsturm Orange coneflower R 23 Rudbeckia laciniata Gold Drop Cutleaf coneflower R 21 Ruscus aculeatus --Butchers broom S S Chapter 3 Salvia azurea Grandiflora Azure blue sage S 21 Salvia haematodes --Sage S 21 Salvia jurisicii --Sage S 21 Salvia leucantha --Mexican bush sage S S S Chapter 2 Salvia nemorosa Miss Indigo Sage S S 53 Salvia officinalis --Common sage S S S S 36 Salvia splendens --Scarlet sage R 1 Salvia splendens Bonfire Scarlet sage R R SL 27

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174 Table B-2. Continued M. incognita race M. javanica M. arenaria race M. hapla Plant species Cultivar Common Name 1 2 3 4? 12? Source Salvia splendens Carabiniere Red Scarlet sage SL 54 Salvia splendens Flare Scarlet sage R R 28 Salvia splendens Hotline Red Scarlet sage SL 54 Salvia splendens Hotline White Scarlet sage SL 54 Salvia splendens Lady in Red Scarlet sage R R 28 Salvia splendens Oxford Blue Scarlet sage S S 28 Salvia splendens Rhea Scarlet sage R 54 Salvia splendens Sea Breeze Scarlet sage SL R 28 Salvia splendens Victoria Scarlet sage R SL 28 Salvia splendens Victoria Blue Scarlet sage R 54 Sanguisorba obtusa --Burnet S 22 Sansevieria cylindrica --African bowstring hemp S 31 Sansevieria cylindrica --African bowstring hemp S 33

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175 Table B-1. Continued M. incognita race M. javanica M. arenaria race M. hapla Plant species Cultivar Common Name 1 2 3 4? 12? Source Sansevieria trifasciata Laurentii Vipers bowstring hemp S 33 Sassafras albidum --Sassafras RR RR R 44 Satureja hortensis --Summer savory S S S S 36 Satureja montana --Winter savory S S S S 36 Scabiosa caucasica Fama Pincushion flower S 21 Scadoxus multiflorus ssp. katherinae --Blood lily S 33 Scindapsus aureus --Centipede tongavine S 33 Scoparia dulcis --Licorice weed S 16 Sidalcea Party Girl Checkermallow R 22 Solanum nigrum --Black nightshade SS 57 Solenostemon scutellarioides Rainbow Coleus S S S 27 Solidago sphacelata Golden Fleece Autumn goldenrod R 22

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176 Table B-2. Continued M. incognita race M. javanica M. arenaria race M. hapla Plant species Cultivar Common Name 12 3 4? 12? Source Sophora japonica --Japanese pagoda tree S S 44 Spiraea bumalda Froebelii Spirea S 6 Spiraea vanhouttei --Van Houttes spirea S 6 Stachys byzantina Lanatna Lamb's ear S 21 Stokesia laevis Blue Danube Stokes aster SL 21 Syagrus romanzoffiana --Queen palm S Chapter 3 Syngonium podophyllum Variegatum American evergreen S 33 Syringa persica --Persian lilac S 39 Tagetes erecta --African marigold R 1 Tagetes erecta --African marigold R 42 Tagetes erecta --African marigold R 50 Tagetes erecta Carnation African marigold S 13

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177 Table B-2. Continued M. incognita race M. javanica M. arenaria race M. hapla Plant species Cultivar Common Name 1 2 3 4? 12? Source Tagetes erecta Inca Gold African marigold R 54 Tagetes patula --French marigold R 1 Tagetes patula --French marigold R 50 Tagetes patula Dwarf Primose French marigold R R R 27 Tagetes patula Golden Gate French marigold R 54 Tagetes patula Goldie French marigold S SS 37 Tagetes patula Petite Gold French marigold S SS 37 Tagetes patula Petite Harmony French marigold S SS 37 Tagetes patula Tangerine French marigold R RR 37 Tagetes sp. Buprees First WhitesMarigold S 49 Tagetes sp. Chrysanthemum Charm Marigold S 49 Tagetes sp. Climax Marigold S 49

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178 Table B-2. Continued M. incognita race M. javanica M. arenaria race M. hapla Plant species Cultivar Common Name 12 3 4? 12? Source Tagetes sp. Crackerjack Marigold R 49 Tagetes sp. FM 584 Marigold S 49 Tagetes sp. Giant Double African (FM 560) Marigold S 49 Tagetes sp. Giant Double African (FM 561) Marigold S 49 Tagetes sp. Giant Double African (FM 562) Marigold S 49 Tagetes sp. Golden Age (FM 581) Marigold S 49 Tagetes sp. Local Marigold S 49 Tagetes sp. Marigold Cinnabar (FM 608) Marigold S 49 Tagetes sp. Red Brocade (FM 597) Marigold S 49 Tagetes sp. Spanish Brocade (FM 597) Marigold S 49

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179 Table B-2. Continued M. incognita race M. javanica M. arenaria race M. hapla Plant species Cultivar Common Name 1 2 3 4? 12? Source Tagetes sp. Spry Hybrid Marigold S 49 Tagetes sp. Spun Gold Marigold S 49 Tagetes sp. Suttons White (FM 370) Marigold S 49 Tamarix gallica --French tamarisk S 16 Tanacetum parthenium --Feverfew S 21 Thalictrum speciosissimum --Meadow rue S 21 Thuja occidentalis --Eastern arborvitae SL 39 Thuja occidentalis Globosa Eastern arborvitae R 6 Thuja occidentalis Pyramidalis Nigra Eastern arborvitae R 6 Thuja orientalis Berkmanns Oriental arborvitae SL 39 Thuja orientalis Dwarf Greenspike Oriental arborvitae R 38 Thuja orientalis Dwarf Greenspike Oriental arborvitae S 39

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180 Table B-2. Continued M. incognita race M. javanica M. arenaria race M. hapla Plant species Cultivar Common Name 1 2 3 4? 12? Source Thymus serpyllum Album Lemon thyme S 22 Tradescantia sp. J. C. Weguelin Spiderwort R 21 Trollius Lemon Queen Globe flower S 22 Trollius chinensis Golden Queen Globe flower S 23 Tropaeolum majus --Garden nasturtium S 1 Tsuga canadensis --Canadian hemlock R 6 Tulipa suaveolens --Tulip R 35 Ulmus parvifolia --Chinese elm SS SS R 44 Uraria picta --Indian gooseberry SS 16 Venidium fastuosum --Monarch of the veld S 1 Verbascum phoeniceum Benary's Hybrid Purple mullein SL 21 Verbena hybrida Deep Blue Verbena SL 54 Verbena hybrida Florist Verbena S S S 30

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181 Table B-2. Continued M. incognita race M. javanica M. arenaria race M. hapla Plant species Cultivar Common Name 12 3 4? 12? Source Verbena hybrida Showtime Blaze Verbena S 54 Verbena bipinnatifida --Dakota mock vervain R 18 Verbena hybrida --Verbena S 1 Verbena hybrida --Verbena S 50 Verbena officinalis --Common verbena S 42 Veronica spicata Icicle Spiked speedwell S 21 Viburnum carlesii --Viburnum S 6 Viburnum odoratissimum Awabuki Sweet arrowwood S Chapter 3 Vinca minor Bowles Variety Common periwinkle R 21 Vinca rosea Little Bright Eye Madagascar periwinkle R SL S 27 Viola cucullata Priceana Marsh blue violet S 21 Viola tricolor --Johnny jump up S 1

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182 Table B-2. Continued M. incognita race M. javanica M. arenaria race M. hapla Plant species Cultivar Common Name 1 2 3 4? 12? Source Viola tricolor --Johnny jump up S 50 Viola tricolor Jolly Joker Johnny jump up S 30 Viola wittrockiana Coronation Gold Pansy S 54 Viola wittrockiana Jolly Joker Pansy S 54 Viola wittrockiana Padparadja Pansy S 54 Vitex agnus-castus --Chaste tree S 39 Washingtonia filifera --California fan palm SS 41 Woodfordia fruticosa --Dhaytaki SS 16 Zantedeschia aethiopica --White calla S 35 Zelkova serrata --Calla lily SS RS R 44 Zinnia elegans --Common zinnia S 1 Zinnia elegans --Common zinnia S 50 Zinnia elegans Scarlet Common zinnia SL S SL 27

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183 Table B-2. Continued M. incognita race M. javanica M. arenaria race M. hapla Plant species Cultivar Common Name 1 2 3 4? 12? Source Zinnia linearis --Narrow leaved zinnia R 42 aHost Response Key: R = resistant S = susceptible SL = slightly susceptible

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190 Hirschmann, H. 1985. The genus Meloidogyne and morphological characters differentiating its species. Pp. 79-92 in J. N. Sasser and C. C. Carter, eds. An advanced treatise on Meloidogyne vol. 1. Biology and control. North Carolina State University Graphics, Raleigh. Hodges, A. W., and J. J. Haydu. 2003. Commodity outlook 2003: U.S. and Florida ornamental plant markets. Extension Data Information Source FE374. Food and Resource Economics Department, University of Florida, Gainesville, FL. Hussey, R. S. 1987. Secretions of esophageal glands of Tylenchida nematodes. Pp. 221228 in J. A. Veech and D. W. Dickson, eds. Vistas on nematology. Society of Nematologists, Hyattsville, Maryland. Hussey, R. S. 1985a. Biochemistry as a tool in identification and its probable usefulness in understanding the nature of parasitism. Pp. 127-133 in J. N. Sasser and C. C. Carter, eds. An advanced treatise on Meloidogyne vol. 1. Biology and control. North Carolina State University Graphics, Raleigh. Hussey, R. S. 1985b. Host-parasite relationships and associated physiological changes. Pp. 143-153 in J. N. Sasser and C. C. Carter, eds. An advanced treatise on Meloidogyne vol. 1. Biology and control. North Carolina State University Graphics, Raleigh. Hussey, R. S., and K. R. Barker. 1973. A comparison of methods of collecting inocula of Meloidogyne spp., including a new technique. Plant Disease Reporter 57:10251028. Hussey, R. S., J. N. Sasser, and D. Huisingh. 1972. Disk-electrophoretic studies of soluble proteins and enzymes of Meloidogyne incognita and M. arenaria Journal of Nematology 4:183-189. Ibrahim, A. A. M., and F. A. Al-Yahya. 2002. Phytoparasitic nematodes associated with ornamental plants in Riyadh region, central Saudi Arabia. Alexandria Journal of Agricultural Research 47:157-167. Jaizme-Vega, M. C., P. Tenoury, J. Pinochet, and M. Jaumot. 1997. Interactions between the root-knot nematode Meloidogyne incognita and Glomus mosseae in banana. Plant and Soil 196:27-35. Jenkins, W. R. 1964. A rapid centrifugal-flotation technique for separating nematodes from soil. Plant Disease Reporter 48:692. Judd, W. S. 2003. The genera of Ruscaceae in the southeastern United States. Harvard Papers in Botany 7:93-149.

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197 BIOGRAPHICAL SKETCH Roi Levin was born 30 March, 1979, in Tel-Aviv, Israel. At the age of 14, he had moved to Florida, where he graduated from Marjory Stoneman Douglas High School in Parkland, Florida, in 1997. He began his studies at the University of Florida, Gainesville, Florida, in 1997, and earned a Bachelor of Science degree in animal sciences, with an emphasis on animal biology, in 2002. He began studies for his Master of Science degree in entomology and nematology at the Univeristy of Florida, under W. T. Crow, in 2002. The title of his thesis is “Reproduction and identification of root-knot nematodes on perennial ornamental plants in Florida.”

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I certify that I have read this study and that in my opinion it conforms to acceptable standards of scholarly presentation and is fully adequate, in scope and quality, as a dissertation for the degree of Doctor of Philosophy. _________________________________ W. T. Crow, Chair Assistant Professor of Entomology and Nematology I certify that I have read this study and that in my opinion it conforms to acceptable standards of scholarly presentation and is fully adequate, in scope and quality, as a dissertation for the degree of Doctor of Philosophy. _________________________________ J. A. Brito Biological Scientist IV I certify that I have read this study and that in my opinion it conforms to acceptable standards of scholarly presentation and is fully adequate, in scope and quality, as a dissertation for the degree of Doctor of Philosophy. _________________________________ R. K. Schoellhorn Associate Professor of Horticultural Science I certify that I have read this study and that in my opinion it conforms to acceptable standards of scholarly presentation and is fully adequate, in scope and quality, as a dissertation for the degree of Doctor of Philosophy. _________________________________ A. F. Wysocki Associate Professor of Food and Resource Economics This dissertation was submitted to the Graduate Faculty of the College of Agricultural and Life Sciences and to the Graduate School and was accepted as partial fulfillment of the requirements for the degree of Doctor of Philosophy. May 2005 _________________________________ Dean, College of Agricultural and Life Sciences _________________________________ Dean, Graduate School


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REPRODUCTION AND IDENTIFICATION OF ROOT-KNOT NEMATODES ON
PERENNIAL ORNAMENTAL PLANTS IN FLORIDA















By

ROI LEVIN


A THESIS PRESENTED TO THE GRADUATE SCHOOL
OF THE UNIVERSITY OF FLORIDA IN PARTIAL FULFILLMENT
OF THE REQUIREMENTS FOR THE DEGREE OF
MASTER OF SCIENCE

UNIVERSITY OF FLORIDA


2005


































Copyright 2005

by

Roi Levin















ACKNOWLEDGMENTS

I would like to thank my chair, Dr. W. T. Crow, and my committee members, Dr.

J. A. Brito, Dr. R. K. Schoellhorn, and Dr. A. F. Wysocki, for their guidance and support

of this work. I am honored to have worked under their supervision and commend them

for their efforts and contributions to their respective fields.

I would also like to thank my parents. Through my childhood and adult years,

they have continuously encouraged me to pursue my interests and dreams, and, under

their guidance, gave me the freedom to steer opportunities, curiosities, and decisions as I

saw fit.

Most of all, I would like to thank my fiancee, Melissa A. Weichert. Over the past

few years, she has supported, encouraged, and loved me, through good times and bad. I

will always remember her dedication, patience, and sacrifice while I was working on this

study. I would not be the person I am today without our relationship and love.















TABLE OF CONTENTS

page

A C K N O W L E D G M E N T S ................................................................................................. iii

LIST OF TABLES .............. ........ ............ ................ vi

LIST OF FIGURES ......... ......................... ............ ........... ix

A B S T R A C T ....................................................................................................... x iii

CHAPTERS

1 INTRODUCTION AND LITERATURE REVIEW ..................................................1

In tro d u ctio n .................................................................................. 1
M eloidogyne spp. ...................................................................2
R relationship w ith O rnam entals ......... ............................................ .................... 14
O b j e c tiv e s ........................................................................................................3 0

2 REPRODUCTION OF FOUR MELOIDOGYNE SPP. ON SEVERAL SPECIES
OF PERENNIAL ORNAMENTAL PLANTS ................................... ...................31

Introduction ............. ........................................................................ 3 1
M materials and M methods ....................................................................... ..................33
R results ................. .....................................................................................40
D discussion ............. ......... ............................................................... ......60

3 IDENTIFICATION OF ROOT-KNOT NEMATODES ........................................67

In tro d u ctio n ........................................................................................................... 6 7
O b j e ctiv e s ...................................................................................6 9
M materials and M methods ....................................................................... ..................70
R e su lts .................95............................................
D iscu ssio n .................103............................................

APPENDIX

A PICTURES OF MELOIDOGYNE SPP. ESTERASE AND MALATE
DEHYDROGENASE ISOZYME PHENOTYPES UNVEILED THROUGH
POLYACRYLAMIDE GEL ELECTROPHORESIS ON PHASTSYSTEM AND
MINI-PORTEIN 3 CELL APPARATUSES ................................ ............... 107









B COLLECTIVE RECORD OF THE HOST STATUS OF ORNAMENTAL
PLANTS TO MELOIDOGYNE INCOGNITA, M. JA VANICA, M. ARENARIA,
AND M HAPLA .......... ............ .... ................ ........ .......... 123

REFERENCES .................................. .. .. ... ... .. .................. 84

BIOGRAPHICAL SKETCH ...... ........ .. ................. .... ....................... 197















LIST OF TABLES


Table pge


2-1 Crop and source of liners used for growth room and greenhouse experiments .......34

2-2 Crops, experimental sites, liner planting dates, inoculation dates, and study
lengths for all crops in the growth room and greenhouse Meloidogyne spp.
studies carried out at the University of Florida during 2003 to 2005 ..................38

2-3 Treatments, root-gall indexes, root weights, number of eggs per plant, number of
eggs per g of roots, and dry shoot weights from the first Liriope muscari cv.
Evergreen Giant growth room trial ............................................... ............... 41

2-4 Treatments, root-gall indexes, root weights, number of eggs per plant, number of
eggs per g of roots, and dry shoot weights from the second Liriope muscari cv.
Evergreen Giant growth room trial ............................................... ............... 42

2-5 Treatments, root-gall indexes, root weights, number of eggs per plant, number of
eggs per g of roots, and dry shoot weights from the Liriope muscari cv.
Evergreen Giant greenhouse experiment ...................................... ............... 44

2-6 Treatments, root-gall indexes, root weights, number of eggs per plant, number of
eggs per g of roots, and dry shoot weights from the first Pittosporum tobira cv.
V ariegata growth room trial .............................................................................. 45

2-7 Treatments, root-gall indexes, root weights, number of eggs per plant, number of
eggs per g of roots, and dry shoot weights from the second Pittosporum tobira
cv. V ariegata grow th room trial ........................................ .......................... 46

2-8 Treatments, root-gall indexes, root weights, number of eggs per plant, number of
eggs per g of roots, and dry shoot weights from the Pittosporum tobira cv.
V ariegata greenhouse experim ent .................................. ............... ............... 47

2-9 Treatments, root-gall indexes, root weights, number of eggs per plant, number of
eggs per g of roots, and dry shoot weights from the first Salvia Iet'//linhi growth
ro om trial ............................................................................ 4 9









2-10 Treatments, root-gall indexes, root weights, number of eggs per plant, number of
eggs per g of roots, and dry shoot weights from the second Salvia le i',nt i/,
grow th room trial............ ... .......................................... .............. ......... ....... 50

2-11 Treatments, root-gall indexes, root weights, number of eggs per plant, number of
eggs per g of roots, and dry shoot weights from the purple-corolla Salvia
leucantha greenhouse experim ent ........................................... ......... ... ............... 52

2-12 Treatments, root-gall indexes, root weights, number of eggs per plant, number of
eggs per g of roots, and dry shoot weights from the white-corolla Salvia
leucantha greenhouse experim ent ........................................... ......... ... ............... 53

2-13 Treatments, root-gall indexes, root weights, number of eggs per plant, number of
eggs per g of roots, and dry shoot weights from the first Odontonema
cuspidatum growth room trial ............................................................................ 55

2-14 Treatments, root-gall indexes, root weights, number of eggs per plant, number of
eggs per g of roots, and dry shoot weights from the second Odontonema
cuspidatum growth room trial ............................................................................ 56

2-15 Treatments, root-gall indexes, root weights, number of eggs per plant, number of
eggs per g roots, and dry shoot weights from the Odontonema cuspidatum
greenhouse experim ent.................................................. ............................... 57

2-16 Treatments, root-gall indexes, root weights, number of eggs per plant, number of
eggs per g of roots, and dry shoot weights from the first Musa acuminata ssp.
zebrina grow th room trial............................................... .............................. 58

2-17 Treatments, root-gall indexes, root weights, number of eggs per plant, number of
eggs per g of roots, and dry shoot weights from the second Musa acuminata ssp.
zebrina grow th room trial............................................... .............................. 59

2-18 Treatments, root-gall indexes, root weights, number of eggs per plant, number of
eggs per g of roots, and dry shoot weights from the Musa acuminata ssp. zebrina
green ou se ex p erim ent.................................................................. .....................6 1

2-19 Treatments, root-gall indexes, root weights, number of eggs per plant, number of
eggs per g of roots, and dry shoot weights from the first Codiaeum variegatum
cv. G old D ust trial ............................... ..... .. ....... ...... ............... 62

2-20 Treatments, root-gall indexes, root weights, number of eggs per plant, number of
eggs per g of roots, and dry shoot weights from the second Codiaeum
variegatum cv. Gold Dust growth room trial ................................. ............... 63

2-21 Treatments, root-gall indexes, root weights, number of eggs per plant, number of
eggs per g of roots, and dry shoot weights from the Codiaeum variegatum cv.
G old D ust greenhouse experim ent .................................. ...................................... 64









3-1 Known Meloidogyne spp. malate dehydrogenase and esterase relative migrations
that were used as references against those that were revealed
electrophoretically from females collected from several counties in Florida ..........91

3-2 Enzyme stain concoctions used in staining malate dehydrogenase and esterase
following electrophoresis using the PhastSystem ......................... ................. 94

3-3 Enzyme stain concoctions used in staining malate dehydrogenase and esterase
following electrophoresis using the Mini-Protean 3 Cell................................ ...96

3-4 Plant species, family, county, relative migration, isozyme phenotype, number of
samples, and Meloidogyne spp. identified from ornamental plants collected in
Florida and processed using the PhastSystem and Mini-Protean 3 Cell ..................97

B-l Sources and citations of publications referred to in Appendix B......................... 124

B-2 Collective record of the host status of ornamental plants to Meloidogyne
incognita, M. javanica, M. arenaria, and M hapla........................................ 125














LIST OF FIGURES


Figure page

3-1 Malate dehydrogenase and esterase relative electrophoretic migrations and
enzyme phenotypes of Meloidogynejavanica, as reported by several authors .......71

3-2 Malate dehydrogenase and esterase relative electrophoretic migrations and
enzyme phenotypes ofMeloidogyne incognita, as reported by several authors......73

3-3 Malate dehydrogenase and esterase relative electrophoretic migrations and
enzyme phenotypes of Meloidogyne arenaria, as reported by Esbenshade and
T riantaphyllou (1985c).................................................. ............................... 75

3-4 Malate dehydrogenase and esterase relative electrophoretic migrations and
enzyme phenotypes of Meloidogyne arenaria, as reported by several authors .......77

3-5 Malate dehydrogenase and esterase relative electrophoretic migrations and
enzyme phenotypes of Meloidogyne hapla.................... ...................79

3-6 Malate dehydrogenase and esterase relative electrophoretic migrations and
enzyme phenotypes of Meloidogyne spp. .......................... ............... 81

3-7 Malate dehydrogenase and esterase relative electrophoretic migrations and
enzyme phenotypes of unidentified Meloidogyne spp. ....................... ...........85

3-8 Malate dehydrogenase (Mdh) and esterase (Est) isozyme phenotypes, revealed
using the mini-protean 3 cell apparatus, ofMeloidogyne partityla females
extracted from Carya illinoensis from Jefferson County on 31 March 2004 ........101

3-9 Malate dehydrogenase (Mdh) and esterase (Est) isozyme phenotype, revealed
using the mini-protean 3 cell apparatus, of a Meloidogyne querciana female
extracted from Viburnum odoratissimum cv. Awabuki from Hillsborough
C county on 25 January 2005.......................... ............................... ............... 102

A-1 PhastSystem gels exhibiting smeared malate dehydrogenase isozyme
phenotypes whose relative migration could not be accurately measured ............107

A-2 Malate dehydrogenase and esterase isozyme phenotypes, revealed using the
PhastSystem apparatus, of Meloidogyne females extracted from Hibiscus rosa-
sinensis cv. Pink Versicolor from Alachua County on 02 April 2003 .................108









A-3 Malate dehydrogenase and esterase isozyme phenotypes, revealed using the
PhastSystem apparatus, of Meloidogyne females extracted from Rosmarinus
officinalis from Suwannee County on 30 April 2003 ........................................108

A-4 Malate dehydrogenase and esterase isozyme phenotypes, revealed using the
PhastSystem apparatus, of Meloidogyne females extracted from Callistemon
viminalis from Lee County on 01 M ay 2003 ......................... ............... ......109

A-5 Malate dehydrogenase and esterase isozyme phenotypes, revealed using the
PhastSystem apparatus, of Meloidogyne females extracted from Syagrus
romanzoffiana from Lee County on 27 May 2003...........................................109

A-6 Malate dehydrogenase isozyme phenotypes, revealed using the mini-protean 3
cell apparatus, of Meloidogyne females extracted from Callistemon viminalis
from Lee County on 27 May 2003 ............................... ...............110

A-7 Eterase isozyme phenotypes, revealed using the mini-protean 3 cell apparatus, of
Meloidogyne females extracted from Callistemon viminalis from Lee County on
27 M ay 2003 ........................................................... ... ... ......... 110

A-8 Malate dehydrogenase isozyme phenotypes, revealed using the mini-protean 3
cell apparatus, of Meloidogyne females extracted from Pittosporum tobira from
A lachua County on 18 D ecem ber 2003 .................................................................111

A-9 Esterase isozyme phenotypes, revealed using the mini-protean 3 cell apparatus,
of Meloidogyne females extracted from Pittosporum tobira from Alachua
C county on 18 D ecem ber 2003 ...................... .. .. ........................ .... ............... 111

A-10 Malate dehydrogenase isozyme phenotypes, revealed using the mini-protean 3
cell apparatus, of Meloidogyne females extracted from Ruscus aculeatus from
Volusia County on 19 December 2003 ............. .............. ............ ...... ............ 112

A-11 Esterase isozyme phenotypes, revealed using the mini-protean 3 cell apparatus,
of Meloidogyne females extracted from Ruscus aculeatus from Volusia County
on 19 December 2003 ........ .............. .... ......... .. .. ..... ............. ... 12

A-12 Malate dehydrogenase isozyme phenotypes, revealed using the mini-protean 3
cell apparatus, of Meloidogyne females extracted from Buddleia davidii from
Pinellas County on 30 September 2003 ............. ............. ........... ........ ........... 113

A-13 Esterase isozyme phenotypes, revealed using the mini-protean 3 cell apparatus,
of Meloidogyne females extracted from Buddleia davidii from Pinellas County
on 30 Septem ber 2003 ........... ................................................. ....... .... ... .. .... 113

A-14 Malate dehydrogenase isozyme phenotypes, revealed using the mini-protean 3
cell apparatus, of Meloidogyne females extracted from Brassica rapa cv.
Shogoin from Alachua County on 06 January 2004 ..........................................114









A-15 Esterase isozyme phenotypes, revealed using the mini-protean 3 cell apparatus,
of Meloidogyne females extracted from Brassica rapa cv. Shogoin from
Alachua County on 06 January 2004 ........................................ ............ 114

A-16 Malate dehydrogenase isozyme phenotypes, revealed using the mini-protean 3
cell apparatus, of Meloidogyne females extracted from Pittosporum tobira cv.
Variegata from Lake County on 11 February 2004 ...............................................115

A-17 Esterase isozyme phenotypes, revealed using the mini-protean 3 cell apparatus,
of Meloidogyne females extracted from Pittosporum tobira cv. Variegata from
Lake County on 11 February 2004........................................... ......... ............... 115

A-18 Malate dehydrogenase isozyme phenotypes, revealed using the mini-protean 3
cell apparatus, of Meloidogyne females extracted from Ruscus aculeatus from
Volusia County on 22 February 2004 ...................................... ......... ............... 116

A-19 Esterase isozyme phenotypes, revealed using the mini-protean 3 cell apparatus,
of Meloidogyne females extracted from Ruscus aculeatus from Volusia County
on 22 February 2004 ........... .. ...................... ........ ..... .. ...... ........ .. .. 116

A-20 Malate dehydrogenase isozyme phenotypes, revealed using the mini-protean 3
cell apparatus, of Meloidogyne females extracted from Liriope muscari cv.
Evergreen Giant from Hillsborough County on 01 July 2004 ............................117

A-21 Esterase isozyme phenotypes, revealed using the mini-protean 3 cell apparatus,
of Meloidogyne females extracted from Liriope muscari cv. Evergreen Giant
from Hillsborough County on 01 July 2004....................... ............... 117

A-22 Malate dehydrogenase isozyme phenotypes, revealed using the mini-protean 3
cell apparatus, of Meloidogyne females extracted from Ophiopogonjaponicus
from Orange County on 16 August 2004 .................................... .....................118

A-23 Esterase isozyme phenotypes, revealed using the mini-protean 3 cell apparatus,
of Meloidogyne females extracted from Ophiopogonjaponicus from Orange
C county on 16 A ugu st 2004 ........................... .................................................... 118

A-24 Malate dehydrogenase isozyme phenotypes, revealed using the mini-protean 3
cell apparatus, of Meloidogyne females extracted from Jesticia carnia from
Hillsborough County on 20 August 2004 ............. ........ ......................119

A-25 Esterase isozyme phenotypes, revealed using the mini-protean 3 cell apparatus,
of Meloidogyne females extracted from Jesticia carnia from Hillsborough
C county on 20 A ugust 2004...................... .... .......... .................... ............... 119

A-26 Malate dehydrogenase isozyme phenotypes, revealed using the mini-protean 3
cell apparatus, of Meloidogyne females extracted from Solenostemon
scutellarioides cv. Elfers from Hillsborough County on 15 September 2004 .......120









A-27 Esterase isozyme phenotypes, revealed using the mini-protean 3 cell apparatus,
of Meloidogyne females extracted from Solenostemon scutellarioides cv. Elfers
from Hillsborough County on 15 September 2004 ............................................. 120

A-28 Malate dehydrogenase and esterase isozyme phenotypes, revealed using the
mini-protean 3 cell apparatus, of Meloidogyne females extracted from Brassica
oleracea from Orange County on 11 January 2004 ............................................ 121

A-29 Malate dehydrogenase and esterase isozyme phenotypes, revealed using the
mini-protean 3 cell apparatus, of Meloidogyne females extracted from Beta
vulgaris from Orange County on 11 January 2004............. ........ ............ 121

A-30 Malate dehydrogenase and esterase isozyme phenotypes, revealed using the
mini-protean 3 cell apparatus, of Meloidogyne females extracted from Phaseolus
vulgaris from Hillsborough County on 25 January 2005.................................122















Abstract of Thesis Presented to the Graduate School
of the University of Florida in Partial Fulfillment of the
Requirements for the Degree of Master of Science


REPRODUCTION AND IDENTIFICATION OF ROOT-KNOT NEMATODES ON
PERENNIAL ORNAMENTAL PLANTS IN FLORIDA

By

Roi Levin

May 2005

Chair: W. T. Crow
Major Department: Entomology and Nematology

Meloidogyne spp. (root-knot nematodes) are serious pathogens of perennial and

woody ornamental plants. Meloidogyne spp. directly limit plant vigor and, by the

mechanical action and physiological responses to their feeding, expose their hosts to an

array of pathogenic fungi and bacteria. The evaluation of the host status of perennial

ornamental plants to root-knot nematodes can identify root-knot nematode resistant plant

material, which may be used to replace infected hosts in landscapes. Six perennial

ornamental species were evaluated for their host status to M. incognita race 2, M.

javanica, M. arenaria race 1, and M mayaguensis in separate growth room and

greenhouse experiments. Data from these experiments indicate that Liriope muscari cv.

Evergreen Giant is a good host to M. incognita race 2, M. javanica, and M mayaguensis,

and a poor host to M arenaria race 1. In addition, a purple-corolla form of Salvia

leucantha and Musa acuminata ssp. zebrina cv. Rowe red are good hosts to the

Meloidogyne spp. evaluated. Pittosporum tobira cv. Variegata, Odontonema cuspidatum,









and Codiaeum variegatum cv. Gold Dust are nonhosts to the Meloidogyne spp. isolates

evaluated.

Twenty ornamental plants were identified as hosts to several Meloidogyne spp.

via the speciation of root-knot nematode females that were dissected directly from their

hosts' roots. These females were identified primarily by evaluation of esterase (Est) and

malate dehydrogenase (Mdh) isozyme phenotypes, unveiled following polyacrylamide

gel electrophoresis. Resolved isozyme phenotypes indicate that Rosmarinus officinalis

(rosemary), Syagrus romanzoffiana (queen palm), P. tobira, Brassica rapa (turnip) cv.

Shogoin, Brassica oleracea (kale), Phaseolus vulgaris (bean), L. muscari cv. Evergreen

Giant, and Ophiopogonjaponicus mondoo grass) are hosts to M. incognita. Hibiscus

rosa-sinensis (hibiscus) cv. Pink Versicolor, B. rapa cv. Shogoin, Ruscus aculeatus

(ruscus), Beta vulgaris (chard), and Viburnum odoratissimum (Viburnum) cv. Awabuki

are hosts to M. javanica. Ruscus aculeatus and P. vulgaris are hosts to M. arenaria.

Callistemon viminalis (bottle brush), S. romanzoffiana, and Solenostemon scutellarioides

(coleus) cv. Elfers are hosts toM. mayaguensis. Carya illinoensis (pecan) is a host toM.

partityla. In addition, Meloidogyne spp. that could not be identified on the basis of their

Est and Mdh isozyme phenotypes were isolated from the following ornamental plants:

Buddleia davidii (butterfly bush), P. tobira cv. Variegata, L. muscari cv. Evergreen

Giant, and 0. japonicus, and Justicia carnea (flamingo plant).














CHAPTER 1
INTRODUCTION AND LITERATURE REVIEW

Introduction

Perennial ornamentals are an important component of the nursery and floriculture

industry. According to Hodges and Haydu (2003), the gross wholesale value for U.S.-

grown floriculture and nursery crops in 2001 reached $13.3 billion, of which $1.6 billion

was produced in Florida alone. Over 2,500 species in roughly 500 genera are included in

this category, and are widely distributed in the United States, Canada, and Europe

(LaMondia, 1995, 1997). Widespread dissemination of perennial ornamentals presents

an important avenue for distribution of root-knot nematodes (Meloidogyne spp.) and

other plant-pathogenic organisms (LaMondia, 1995). Root-knot nematodes cause

estimated crop losses of 5 to 10% in major crops, and are considered the most widespread

and destructive of all plant-pathogenic nematodes (Haseeb et al., 1984; Stokes, 1977;

Walker et al., 1994). The lack of information regarding perennial ornamental crop losses

due to plant-parasitic nematodes is attributed to the demand for research pertaining to

agronomic crops, the long time period required for crop loss assessment on horticultural

crops, and the vast array and interchangeability of available cultivated plant material

(Walker and Melin, 1998b). Information regarding resistance of ornamental plants to

root-knot nematodes is needed by extension personnel, the landscape industry, plant

producers, and gardeners (Walker et al., 1994; Walker and Melin, 1998a). Furthermore,

knowledge of the susceptibility of cultivated perennials to root-knot may alleviate post-

installation damage associated with these pathogens by the avoidance of highly






2


susceptible cultivars and increased development and use of resistant cultivars (Giblin-

Davis et al., 1992). Incorporation of resistance into the genomes of perennial cultivars

would greatly benefit the perennial plant industry (Walker and Melin, 1998a), as rotation

with resistant species has been shown to successfully control root-knot nematodes in

infested nursery fields (LaMondia, 1995). High demand and extensive shipments impact

the rate of inspection for root-knot nematodes. Therefore, root-knot nematode

management programs that are based on sanitation, resistance, tolerance, rotation, and

exclusion, should be applied and developed in nurseries and landscapes to alleviate

product losses, and to reduce the spread of the pathogens into uninfested field-grown

nurseries and landscapes (Benson and Barker, 1985; LaMondia, 1997; Walker, 1980;

Walker and Melin, 1998b). Care should be taken with the cultivation of tolerant

perennials in root-knot nematode infested sites since annuals planted adjacent to such

plants may become infected with root-knot nematodes that thrive on tolerant perennials

(McSorley and Dunn, 1990; Rohde, 1972).

Meloidogyne spp.

Historical Background

The first account of root-knot nematodes was by Berkeley, who in 1855 observed

galls on roots of greenhouse-grown cucumber plants in England (Hartman and Sasser,

1985). Cornu first coined the name Anguillula marioni Cornu for root-knot nematodes,

after observing root galls on Onobrychis sp. (sainfoin). Subsequently, root-knot

nematodes were classified in the genera Heterodera or Anguillula. In 1884, Miller

classified root-knot nematodes as H. radicicola and in 1887 the type species Meloidogyne

exigua was described by Goldi. In 1932, Goodey reclassified root-knot nematodes as H.









marioni (Muller) Goodey. Differences in host responses led Chitwood to redescribe

Meloidogyne exigua Goldi, M. incognita (Kofoid and White, 1919) Chitwood, M.

javanica (Treub, 1885) Chitwood, and M arenaria (Neal, 1889) Chitwood, and to

describe M. hapla Chitwood and M. incognita acrita Chitwood, as a new species and

variety, respectively (Christie, 1959). In his description ofM. hapla, Chitwood cited

Abelia x grandiflora as a host (Bernard and Witte, 1987). Chitwood also gave a general

description of the genus Meloidogyne and differentiated it from the genus Heterodera

(Hirschmann, 1985). Chitwood's diagnoses of the redescribed and newly described

species were based on examinations of morphological features and morphometrics from

all life stages of the evaluated species (Eisenback, 1985). These features included the

perineal pattern, stylet morphology, and distance from the base of the stylet knobs to the

dorsal esophageal gland opening (DEGO) (Christie, 1959). To date, more than 80

Meloidogyne spp. have been described (Randig et al., 2002). The current taxonomic

status for the Meloidogyne spp. reviewed in this study is: phylum Nemata, order

Tylenchida, suborder Tylenchina, superfamily Tylenchoidea, family Heteroderidae,

subfamily Meloidogyninae, species M incognita, M. javanica, M. arenaria, and M

mayaguensis Rammah and Hirschmann, 1988 (Andrassy, 1976; Rammah and

Hirschmann, 1988; Thorne, 1961). Meloidogyne incognita, M. javanica, and M arenaria

have been frequently encountered in Florida for many years. Meloidogyne mayaguensis,

however, was not detected frequently in Florida until the use of electrophoresis for the

identification of root-knot nematodes became commonplace. Meloidogyne mayaguensis

was described as a result of the reevaluation of a population that was tentatively

identified as M. arenaria at North Carolina State University. Differentiating biological









and morphological characters that part this species from the four common Meloidogyne

species include a somatic chromosome number of 2n = 44 to 45 and unique enzyme

phenotypes revealed through electrophoresis (Rammah and Hirschmann, 1988).

Although the M. mayaguensis type species exhibits a morphologically unique perineal

pattern, investigations by Brito et al. (2004) revealed M mayaguensis isolates having a

perineal pattern similar to that ofM incognita in Florida, making this feature unreliable

for the identification of M mayaguensis.

Identification

The accurate identification of root-knot nematodes to species and host races is

essential for their control and is a prerequisite to meaningful research. Many

Meloidogyne species are easily identified based on distinct morphological characters and

restricted host ranges. Several species are difficult to identify due to their similarity to

other species and poor taxonomic descriptions. The four most common root-knot

nematode species, composing 98% of all worldwide populations, are M incognita, M.

javanica, M. arenaria, and M hapla (Hussey, 1985a). Other Meloidogyne species, such

as M. mayaguensis, become increasingly important due to their uncommon virulence and

increasing occurrence. Difficulty in identifying root-knot nematodes may result from

morphological variations within and between populations from a same species. Since the

reevaluation of Meloidogyne spp. by Chitwood in 1949, female perineal patterns became

the dominant diagnostic character of the four most common Meloidogyne species. The

perineal pattern presents several benefits that render it a valuable diagnostic tool. Aside

from minor variations, perineal patterns are constant within populations and their source

(females) is abundant in infected host roots. Other diagnostic features used in taxonomic









identification include female stylets, male heads and stylets, and second-stage juvenile

(J2) heads and stylets. Morphological features of the perineal patterns ofM. incognita,

M. javanica, M arenaria, and M mayaguensis are describes as follows:

Meloidolyne incognita. Striae are smooth, wavy, sometimes in a zigzag pattern.

Lateral lines are absent. A squarish, high dorsal arch containing a distinct whorl around

the tail terminus is the most conspicuous diagnostic character of this perineal pattern

(Eisenback, 1985).

Meloidogyne javanica. Striae are smooth and somewhat wavy. The dorsal arch is

often low and rounded but may be high and squarish, frequently possessing a whorl in the

tail terminus area. Unique to this species are distinct lateral ridges that run across the

pattern, fading away around the tail terminus (Eisenback, 1985).

Meloidogyne arenaria. Striae are smooth and slightly wavy, often extended

laterally, forming wings on one or both lateral sides of the pattern. Distinctive lateral

ridges are absent, but are marked by forked, irregular lateral fields. The dorsal arch is

low and indented near the lateral fields, forming rounded shoulders (Eisenback, 1985).

Meloidogyne mayaguensis. Striae are fine, continuous, and widely spaced.

Lateral lines are inconspicuous or a single lateral line may be present on one side of the

pattern. Dorsal arch is rounded, with a circular, striae-free tail terminus (Rammah and

Hirschmann, 1988). Thirty percent of three isolates from Florida possess perineal

patterns that differ from the type species and depict perineal patterns typical ofM.

incognita (Brito et al., 2004).

Inconsistencies in host-parasite relationships lead to the erection of species-

specific races based on their infection of specific crops, namely Gossypium hirsutum









(cotton) cv. Deltapine 61, Nicotiana tabacum (tobacco) cv. NC 95, Capsicum frutescens

(pepper) cv. California Wonder, Citrullus vulgaris (watermelon) cv. Charleston Gray,

Arachis hypogaea (peanut) cv. Florunner, and Lycopersicon esculentum (tomato) cv.

Rutgers. Based on their susceptibility to the differential hosts, M. incognita and M

arenaria were assigned four and two races, respectively. This host differential system

allows for the rotation of the differential hosts as a means of maintaining low nematode

populations and thus reduces crop losses. However, the identification of root-knot

nematode species solely on the basis of the differential host test is unreliable due to the

possibility of mixed populations, and should be used in conjunction with morphological,

morphometric, and biochemical evaluations to determine root-knot nematode species

(Hartman and Sasser, 1985).

Problems in the morphological identification of Meloidogyne species, such as

intra-species merging of morphological characters, rarely-seen characters, and lack of

apparent differences between species, has encouraged much interest in the utilization of

biochemical techniques as a complementary, routine method for the identification of root-

knot nematodes (Hansen and Buecher, 1970). Enzyme phenotypes, unveiled through

staining of polyacrylamide gel slabs following electrophoresis, have become a reliable,

less subjective approach to identification of root-knot nematodes (Hussey, 1985a). The

first study on root-knot nematode protein profile stability and its utilization in the

identification of root-knot nematodes was conducted by Dickson et al. (1970), who

established the usefulness of disc-electrophoresis in identification of root-knot

nematodes. This early work also verified that protein profiles were stable within

nematode species collected from different parts of the world and that infect an array of









hosts. Further work by Dickson et al. (1971) examined lactate dehydrogenase (LDH),

malate dehydrogenase (MDH), a-glycerophosphate dehydrogenase (GDH), glucose-6-

phosphate dehydrogenase (G-6-PDH), acid phosphatase (AcP), alkaline phosphatase

(AlkP), and esterase (EST) enzyme profiles from Meloidogynejavanica, M. arenaria,

and M hapla females, as well as from three life stages of M incognita. Dickson et al.

(1971) revealed that enzyme profiles for GDH, MDH, G-6-PDH, and EST differed

among the four studied Meloidogyne species. Furthermore, G-6-PDH enzymatic profiles

from all evaluated species and GDH enzymatic profiles from M incognita, M. hapla, and

M. arenaria were monomorphic, while enzymatic profiles from all the remaining species

evaluated were polymorphic. Among the enzymes evaluated for the characterization of

Meloidogyne species, MDH and EST enzyme profiles were most variable with respect to

electrophoretic mobility and therefore are most useful for differentiation of the four

species evaluated by this method. However, MDH enzyme profiles did not differ

between M. javanica and M incognita and EST isoenzymes were detected from M

incognita and M hapla isolates. Enzymatic profiles from the three life stages of M

incognita resulted in variable band numbers and electrophoretic migration for MDH and

EST. The use of disk-electrophoresis for the assessment of Meloidogyne species protein

phenotypes by Dickson et al. (1971) required the analysis of several specimens of the

same species for the elucidation of a single protein phenotype. This method, therefore,

rendered genetic analysis at the intra- and interspecific levels impossible (Dalmasso and

Berge, 1978). A breakthrough in biochemical speciation of Meloidogyne species was the

use acrylamide gels as thin slabs (0.7-mm-thick) to electrophoretically separate proteins

from individual Meloidogyne females. First used by Dalmasso and Berge (1978), the









thin-slab method for polyacrylamide gel electrophoresis (PAGE) has proved to be useful

in the identification of root-knot nematodes. This method allowed the speciation of 20 to

25 individual specimens from the same or different populations on single gels.

Analyzing 22,000 specimens of root-knot nematodes using microscale electrophoresis,

Dalmasso and Berge (1978) found that, of the enzymes elucidated, EST were the most

useful for the differentiation of the common Meloidogyne species, primarily due to their

polymorphic nature (Hussey, 1985a).

Morphological characters, particularly female perineal patterns, are the primary

method for routine root-knot nematode identification. However, perineal patterns are

variable, and may lead to misidentification of aberrant populations and uncommon

species. Conversely, biochemical analyses, particularly esterase phenotypes of young,

egg-laying females, in conjunction with morphological and morphometric examinations,

allow for precise, accurate diagnoses, thereby alleviating the confusion associated with

morphological characters (Hussey, 1985a). In addition to morphological and

morphometric analysis and the use of PAGE for the identification of root-knot

nematodes, the use of single eggs or J2 for species identification via restriction fragment

length polymorphism (RFLP) (Fargette et al, 1996), random amplified polymorphic

deoxyribose nucleic acid (DNA) (RAPD) (Blok et al., 1997b), ribosomal DNA

amplification (Blok et al., 1997a), and mitochondrial DNA (mtDNA) amplification (Blok

et al., 2002), has been reported.









Biology and Ecology

Root-knot nematodes occur throughout the world and infect most major plant

crops, and are responsible for substantial yield losses and reduced crop quality (Sasser

and Carter, 1985). In Florida, root-knot nematodes are the most damaging plant-parasitic

nematodes on agricultural crops (McSorley and Dunn, 1989). The life cycle of root-knot

nematode is largely indifferent with respect to individual species' host-parasite

relationships and physiological characteristics (Christie, 1959). Root-knot nematode

eggs are protected within a gelatinous egg mass produced by the female (Maggenti,

1987). Inside the egg, a first-stage root-knot nematode juvenile (J1) molts once prior to

hatching into a J2. While egg hatching is usually spontaneous and does not correlate with

plant-root stimuli, root diffusates have been shown to stimulate hatching (Hussey,

1985b). Once hatched, the now J2 move though the soil in search of a suitable feeding

site (Christie, 1959). Root penetration by the pathogen involves the mechanical

disruption of host tissues. However, cellulose and pectin-dissolving enzymes may also

aid in the penetration process (Hussey, 1987). Upon penetration, J2 move within the root

in a vertical manner, and often migrate toward and away from the root surface. Although

penetration may occur anywhere in the root system, J2 are often observed aggregating

and penetrating behind the root cap, near the meristematic zone. Other penetration sites

include cracks and lesions of mature roots and areas of secondary root formation (Lewis,

1987). Furthermore, the site of one nematode's penetration often becomes attractive for

other J2, leading to multiple infections in confined areas (Hussey, 1985b). Once

established within the plant tissue, root-knot nematodes become sedentary endoparasites,

and halt further movement or migration (Christie, 1959). Second-stage juveniles are









often observed in cortical tissues about the zone of differentiation, their bodies vertically

parallel with the root axis and heads settled in the periphery of the vascular tissue

(Hussey, 1985b). Upon establishment of a suitable feeding site, sexually-undifferentiated

J2 begin to modify the host's physiology by transforming healthy, undifferentiated cells

into specialized feeding sites referred to as giant cells. Modified cells exhibit nuclear,

nucleolar, and surface hypertrophy, an increase in cytoplasmic density, organelle

hyperplasia, and disappearance of the central vacuoles. While the nematode's two

subventral glands are involved in giant cell initiation, dorsal gland secretions maintain

giant cell development (Lewis, 1987). The ability of J2 to invade roots differs with

respect to the root-knot nematode species, but symptoms of root-knot nematode

penetration are often depicted as root-tip enlargement and root-growth retardation. While

gall development is not essential for root-knot nematode survival, the development and

maintenance of giant cells is critical for root-knot nematode development. This host-

parasite relationship requires the developing root-knot nematode to feed on five to six

viable giant cells. The inability to elicit a giant cell response results in nematode death

or, if early in the J2 stage, migration out of the root in search of a suitable feeding site

(Hussey, 1985b). Late in the J2 stage, following feeding initiation and giant cell

formation, an increase in J2 width is observed, and the genital primordia attain their

sexual characteristics prior to the second molt. The third-stage juvenile (J3) of both sexes

is depicted by the J2 cuticle surrounding the J3, the loss of the stylet and the median

esophageal bulb valve, and the loss of the tail spike, which becomes rounded. The J3

stage passes in a few hours, at which time a third molt gives rise to the fourth-stage

juvenile (J4). In this stage, which lasts longer than the third stage, the median esophageal









bulb valve is reformed and the excretory pore opens. The female rectal glands, uterus

and vagina, and male vas deferens differentiate and enlarge, and the male undergoes

metamorphosis, attaining an elongated, cylindrical shape. Following gonad

differentiation in the fourth stage, a fourth and final molt reveals the adult nematodes

enclosed in the three previous juvenile cuticles. At this stage the stylet reappears in both

sexes, the perineal pattern is observed in females, and sperm production is initiated in

males prior to the disappearance of the previously-molted cuticles (Triantaphyllou and

Hirschmann, 1960). While male production occurs in most root-knot nematode species,

M. hapla, which reproduces through facultative meiotic parthenogenesis, produce

relatively more males than M incognita, M. javanica, and M arenaria, which reproduce

through obligate mitotic parthenogenesis (Triantaphyllou, 1985). Similarly, M.

mayaguensis reproduces through mitotic parthenogenesis (Rammah and Hirschmann,

1988). The pathogen's parthenogenic reproductive behavior is adaptive to its unstable

environmental and physiological conditions, namely the halt in female motility, short

generation time, limiting habitat, and male infrequency. Parthenogenesis allows for the

establishment of varying phenotypes through polyploidy, thereby increasing adaptation to

unfavorable environmental conditions, including, but not limited to, fluctuating thermal

gradients, drought, and low oxygen concentrations. Furthermore, such adaptations may

allow for rapid generation turnover through rapid maturity of developing juvenile stages

(Maggenti, 1987). Regardless of their reproductive behavior, Meloidogyne spp. produce

less eggs and more males in response to increasing populations within a root system

(Lewis, 1987). Upon maturation, females produce a gelatinous matrix through their

rectal glands, into which they deposit eggs. The gelatinous matrix provides eggs with









physical protection (Maggenti, 1987) and acts as a barrier to temperature fluctuations and

water evaporation from eggs (Van Gundy, 1985). Meloidogyne spp. viability and

development is influenced by various environmental and physical stresses, including

temperature, soil texture, moisture, aeration, osmotic potential, and host suitability.

Temperature has the greatest influence on egg development and hatching, growth,

reproduction, and survival. Optimal temperatures for egg development ofM. incognita,

M. javanica, and M arenaria are 10 to 15 C, and approximately 9 C forM. hapla.

Conversely, optimal temperatures for growth and development of juvenile and adult

stages of M incognita, M. javanica, and M arenaria are 25 to 30 OC. Although

temperature extremes may inhibit reproduction, individual populations within certain

species acclimatize to local temperature regimes. In general, relative cold tolerance is

exhibited by M. hapla, M. incognita, M. arenaria, and M. javanica, in decreasing order,

with M. hapla surviving subzero temperatures and M. javanica not surviving

temperatures below 10 C. While Meloidogyne eggs do not survive in soils beyond one

year, varying egg mass colors, from white to brown, have been attributed to egg

dormancy and overwintering. Soil texture, moisture, and aeration compose a dynamic,

complex environment, and influence Meloidogyne activity, reproduction, and

pathogenicity. Continuously changing, soil texture is a solid-phase component of the soil

environment. Vertically and horizontally shifting soil particles are interconnected by the

liquid and gas phases of this dynamic system. Optimal temperature regimes and moisture

levels of 40 to 60% of field capacity present the most advantageous conditions for

nematode activity and metabolism. Moisture levels that are lower or higher than optimal

reduce nematode activity due to drying soils and limiting oxygen concentrations,









respectively (Van Gundy, 1985). It is widely accepted that Meloidogyne spp. activity is

optimal in sandy loam soils and reproduction is greatest in fine sand (Wallace, 1969),

primarily due to the low water holding capacity of such soils (Benson and Barker, 1985).

However, Whitehead (1969) reported that eastern African population viability of M

incognita and M. javanica exhibited no correlation with soil texture. In addition,

O'Bannon and Reynolds (1961) reported on heavy M incognita infestations in coarse-

textured soils planted to cotton in Arizona. Root-knot nematode females, examined in

roots from different hosts, were variable in size. Highly susceptible plants exhibited

large, robust females while less susceptible plants supported smaller females (Pant et al.,

1983). Furthermore, Niblack and Bernard (1985) observed that M hapla densities were

positively correlated with Cornusflorida and Acer rubrum nursery tree age.

Meloidogyne spp. behavior is altered under various soil solution osmotic

potentials. In drying, well-fertilized soil, nematodes are subjected to high osmotic

pressures as the osmotic potential increases. Wallace (1969) observed the highest and

lowest reproduction ofM. javanica on Lycopersiconperuvianum (tomato) cv. Tatura

Dwarf when subjected to high and low nutrient levels, respectively. However, M.

javanica infectivity decreased with increased electrical conductivity of the soil solution,

and J2 migration was observed from high to low salt concentrations (Van Gundy, 1985).

The interacting environmental and biological factors that influence Meloidogyne

spp. development, reproduction, and pathogenicity are complex and difficult to evaluate.

Such factors as soil homogeneity, moisture, and temperature, in addition to changing

osmotic and water potentials, interact with nematode development, reproduction, and

pathogenicity, as well as various metabolic and developmental aspects of the host plant.









Interactions between nematodes and plants, further complicated by inoculum densities

and developmental levels, lead to the inevitable intricacy of this plant-parasite and

biological relationship. In evaluating the effect of nematode populations on host plants,

factors influencing the experimental environment are minimized to the studied

components by controlling variables as temperature, moisture, lighting, host nutrition,

initial inoculum density, and initial host development, in hope to reveal a significant

relationship between pathogen and host, with minimum interference by unstudied factors

(Van Gudny, 1985).

Relationships with Ornamentals

One of the most serious groups of pathogens to limit agricultural productivity,

root-knot nematodes have been reported to infect many plants throughout the world

(Zarina and Abid, 1995), and are considered the most damaging group of plant-parasitic

nematodes in Florida (McSorley and Dunn, 1989). The woody ornamental/floriculture

industry is one of the U.S.'s fastest growing agricultural segments (Bernard and Witte,

1987). As serious pathogens of numerous woody ornamentals (Barker and Benson,

1977), Meloidogyne spp. have the potential to damage many important nursery crops

(Benson and Barker, 1985) and form disease complexes with certain soil-borne fungal

pathogens, thus increasing their hosts' susceptibility to such pathogens (Nigh, 1972;

Santamour and Riedel, 1993; Walker and Melin, 1998b). Unlike annual plants, damage

thresholds levels do not apply to perennials, since low Meloidogyne spp. populations

have the potential to increase and cause severe damage over a period of several years

post-planting (LaMondia, 1995).









In the U.S., root-knot nematodes have been reported to cause damage and reduced

yields on ornamental crops in Alabama (Heald, 1967), Arizona (Nigh, 1972; Tarjan,

1952), California (Santo and Lear, 1976; Viglierchio, 1979), Connecticut (LaMondia,

1995, 1996, 1997), Florida (Giblin-Davis et al., 1992; Lehman, 1984a, 1984b; Lehman

and Barnard, 1982; McSorley and Dunn, 1989, 1990; McSorley and Marlatt, 1983;

Stokes, 1977, 1982), Georgia (Heald, 1967; Motsinger et al., 1977; Walker et al., 1994;

Walker and Melin, 1998a, 1998b), Hawaii (Sher, 1954), North Carolina (Barker et al.,

1979; Barker and Benson, 1977; Benson and Barker, 1982; Haasis et al., 1961; Rickard

and Dupree, 1978, Sasser et al., 1966), Oklahoma (Nemec and Morrison, 1972; Nemec

and Struble, 1968), Tennessee (Bernard and Witte, 1987; Bernard et al., 1994; Niblack

and Bernard, 1985), Virginia (Eisenback, 1987), and Washington, DC (Santamour, 1992;

Santamour and Riedel, 1993, 1995). Internationally, Meloidogyne spp. have been

reported to damage ornamental crops in Australia (Wallace, 1969), Belgium (Coolen and

Hendrickx, 1972; Stoffelen et al., 2000), Egypt (Montasser, 1995), France (De Waele and

Davide, 1998), India (Ahuja and Arora, 1980; Haseeb et al., 1984, 1985; Khanna et al.,

1998; Mishra and Mishra, 1997; Mishra and Misra, 1993; Misra and Mishra, 1997; Misra

et al., 2002; Pant et al., 1983; Singh et al., 2000; Singh and Gupta, 1993), Iraq (Singh and

Majeed, 1991), Ivory Coast (Adiko, 1988), Korea (Cho et al., 1996), Nigeria (Caveness

and Wilson, 1977), Pakistan (Zarina and Abid, 1995), Saudi Arabia (Ibrahim and Al-

Yahya, 2002), Spain (Jaizme-Vega et al., 1997), Tamil Nadu (Rajendran et al., 1975),

and Trinidad and Tobago (Bala and Hosein, 1996). Although commonly observed, the

potential for damage to many perennial ornamental crops by root-knot nematodes is

unrecognized (Bernard and Witte, 1987).









Meloidogyne spp. have been attributed as causing decline to growth of ornamental

plants and thus hinder production and lead to reduced returns (Stokes, 1977).

Meloidogyne incognita has been attributed to such effects on the production of Anthurium

andraeanum and other tropical ornamentals (Bala and Hosein, 1996), certain flower

bulbs (Montasser, 1995), and several Ilex spp. (Heald, 1967). In addition, production

losses in field-grown Rosa spp. have been attributed toM. hapla (Santo and Lear, 1976).

Furthermore, Meloidogyne spp. have been reported to reduce Dianthus caryophyllus

production worldwide by 10 to 20% (Cho et al., 1996) and cause qualitative and

quantitative decline in Indian Gladiolus x hortulanus production (Khanna et al., 1998).

Furthermore, in the middle Tennessee nursery-growing region, Meloidogyne hapla

occurs in approximately 25% of nursery blocks (Bernard et al., 1994).

Goff (1936) was one of the first researchers to conduct an extensive survey of the

susceptibility of ornamental plants to Meloidogyne spp. In his survey, Goff noted the

varying degrees of susceptibility among the tested plant species. Root-knot infected

plants often exhibit symptoms that include root galls and root rots, shoot yellowing and

chlorosis, stunted growth, and other symptoms commonly associated with nutritional

deficiencies (Bala and Hosein, 1996; Bird, 1974; Misra et al., 2002, Santo and Lear,

1976; Zarina and Abid, 1995), resulting in general decline (Nigh, 1972), poor yield, and

wilt diseases (Rajendran et al., 1975). Furthermore, photosynthetic rate reduction has

been observed in response to root-knot nematode infections. Often, the ratio of food

resources provided by the host plant and the root-knot nematode density determines the

degree of host response to infection (Bird, 1974). However, Walker and Melin (1998b)

observed greater plant growth in the presence of low plant-parasitic nematode









populations than in their absence. Susceptible hosts severely infected with root-knot

nematodes often decline as a linear function of time (Barker and Benson, 1977), and such

decline is accelerated under unfavorable climatic conditions (Haasis et al., 1961), leading

to severe disease symptoms in late summer, fall, and during periods of severe dry periods

(Lehman, 1984a). The degree of root galling is dependant upon the infected plant species

and/or cultivar and the root-knot nematode species, race, population, or even isolate

(Bird, 1992; LaMondia, 1995; Rohde, 1972). Certain ornamentals infected with root-

knot nematodes exhibit unique symptoms. Such plants include Sansevieria cylindrica,

which developed leaf discoloration and tip necrosis 4 to 5 months post-infection with M

incognita (Mishra and Mishra, 1997), Philodendron selloum, which exhibits a reduction

in leaf size when infected with M incognita (Mishra and Misra, 1993), and Juniperus

horizontalis var. Plumosa and Thuja orientalis cv. Dwarf Greenspike, which exhibit

thickened roots and slight galling post-infection with Meloidogyne spp. (Nemec and

Morrison, 1972). Furthermore, Gladiolus x hortulanus plants infected with M incognita

race 2 exhibited leaf drying, reduction in floral stalk height and girth, and reduced

number of florets (Khanna et al, 1998). Some plants exhibit minute galls following

infection with Meloidogyne spp. In such conditions, root-knot nematode females can be

seen protruding from the infected roots. Other plants, such as Rheum spp., Begonia spp.,

and Thunbergia spp. produce large galls, measuring up to 0.6 m in the latter case (Bird,

1974). In testing the susceptibility of numerous herbaceous perennial ornamentals to M.

hapla, however, LaMondia (1995, 1996) did not observe egg mass production in the

absence of cellular hypertrophy.









Mixed populations of Meloidogyne spp. are often observed parasitizing perennial

ornamentals. In Pakistan, mixed populations of M javanica and M incognita have been

reported on Solanum nigrum, Cucumis melo ssp. melo var. flexuosus, and Rosa indica

(Zarina and Abid, 1995). Pant et al. (1983) did not observe infection signs or symptoms

on Matthiola spp., Tagetes spp., Gaillardia spp., Ch iiyiiiunhenuin spp., and Zinnia spp.

after inoculation of these plants with M incognita. Likewise, no signs or symptoms were

observed in Areca catechu nine months after inoculation with Meloidogyne arenaria race

1, M. incognita races 1 and 3, M. javanica, and M hapla (McSorley and Dunn, 1989). In

testing for pathogenicity ofM incognita on flower bulbs, Montasser (1995) rendered

Amarylis vittata, Clivia miniata, Crinum longifolium, and Narcissus tazetta, which

belong to Amaryllidaceae, and Agapanthus umbellatus, H)y-%i l/1\t orientalis, Lilium

longiflorum, and Tulip suaveolens, which belong to Liliaceae, highly resistant. Likewise,

Rajendran et al. (1975) found Barleria prionitis free ofM incognita infections in a

pathogenicity experiment. Osborne and Jenkins (1963) observed M haplajuveniles and

light galling in Fori ythiii intermedia, but indicated that invading juveniles failed to

mature. Similar observations were reported by Bernard and Witte (1987) for Ligustrum

sinense and Nandina domestic, in which giant cells failed to develop following infection

with M hapla. In addition, N. domestic was a nonhost for M arenaria (Benson and

Barker, 1982). Several landscape ornamentals were tested for their susceptibility to M

arenaria, M. incognita races 1, 2, and 3, and M. javanica. Of these, Photiniafraseri was

tolerant to all the M. incognita isolates as well as to M. javanica. Furthermore, egg mass

indices for all evaluated Meloidogyne spp. except for M. javanica were low on Dracaena

marginata, while Ficus benjamin was highly susceptible to all Meloidogyne spp. except









M. hapla (McSorley and Dunn, 1990). Several herbaceous perennials were evaluated for

susceptibility to Meloidogyne spp. by Walker and Melin (1998a). Fragaria x aananssa

cv. Pink Panda, Monarda didyma cv. Blue Stockings, Phlox paniculata cv. Eva Cullum,

Franz Shubert, and Oakington Blue, and Polygonum affine cv. Dimity did not support M.

arenaria and M. incognita populations six weeks after inoculation. Geranium

psilostemon x procurrens cv. Ann Folkard did not support populations ofM. arenaria,

while G. cinereum cv. Laurence Flatman did not support populations ofM incognita.

Evaluating galling of several herbaceous perennials by Meloidogyne spp., Walker and

Melin (1998a) found no or very few galls on Aethionema cordifolium, Echinacea

purpurea, Moranda citriodora, or Patrinia scabiosifolia. Similarly, Santo and Lear

(1976) found Rosa noisettiana cv. Manetti a poor host to M. hapla, while Coolen and

Hendrickx (1972) found R. canina cv. Succes and Heinsohn's Rekord poor hosts for M.

hapla. In testing the susceptibility of several annual bedding plants to Meloidogyne spp.,

McSorley and Frederick (1994) found that M incognita race 1 caused no galls on

Ageratum houstonianum cv. Blue Mink, Tagetespatula cv. Dwarf Primrose, Vinca rosea

cv. Little Bright Eye, and Salvia splendens cv. Bonfire, and very light galling on Verbena

x hybrida cv. Florist and Zinnia elegans cv. Scarlet. In the same experiments, Ageratum

houstonianum cv. Blue Mink, Lobularia maritime cv. Rosie O'Day, and T. patula cv.

Dwarf Primrose exhibited no and little infection symptoms post inoculation with M

javanica and M. arenaria race 1, respectively. Dianthus chinensis cv. Baby Doll Mix

was slightly infected by M. javanica and M. arenaria, while V rosea cv. Little Bright

Eye and Z elegans cv. Scarlet were minimally infected by M. javanica and M arenaria,

respectively. In evaluating carnation cultivars for M. incognita resistance, Fawzy et al.









(1991) and Cho et al. (1996) reported on the resistance of several cultivars of Dianthus

caryophyllus to M incognita. Bernard and Witte (1987) reported that M hapla failed to

reproduce on several species of Prunus and Rhododendron, although Haasis et al. (1961)

reported that Rhododendoron spp. and Camellia spp. supported light Meloidogyne spp.

populations in field experiments. Giblin-Davis et al. (1992) evaluated the susceptibility

of several Ixora spp. cultivars to M incognita race 1 and M javanica. Although all Ixora

spp. were susceptible to the Meloidogyne spp. isolates evaluated, above-ground disease

symptoms were not observed 20 weeks after inoculation. Giblin-Davis et al. (1992)

hypothesized that either the Ixora spp. cultivars tested were tolerant to the inoculated

Meloidogyne spp. isolates, or that 20 weeks was not enough time for above-ground

disease symptoms manifestation. LaMondia (1995, 1996) conducted extensive studies

that evaluated the susceptibility of an array of perennial herbaceous ornamentals to M

hapla. Approximately 30% of tested perennials were resistant to the isolate evaluated. In

India, Khan and Khan (1989) reported on the resistance of Verbena bipinnatifida to M

incognita and presented nematicidal properties associated with root exudates from this

plant. Santamour and Riedel (1993) tested 23 landscape trees for root-knot nematode

resistance, of which six were nonhosts toM. arenaria races 1 and 2, M. hapla, M.

incognita, and M. javanica, including Magnolia grandiflora and Gleditsia triacanthos,

which are widely used native American landscape trees. Niblack and Bernard (1985) and

Lehman and Barnard (1982) reported on M hapla and M incognita infecting Cornus

florida, and Niblack and Bernard (1985) reported on M hapla infecting Acer rubrum and

Prunuspersica. Several gymnosperms were nonhosts to M hapla (Bernard and Witte,

1987) and resistant to M. incognita (Nemec and Struble, 1968). Walker and Melin









(1998b) reported x Cupressocyparis leylandi, Cedrus deodara, and Taxodium distichum

to be nonhosts toM. incognita. Bernard and Witte (1987) found no M hapla

reproduction on Acerpalmatum and A. saccharum. In Washington, Santamour and

Riedel (1995) reported that Cercis canadensis, C. chinensis, and C. yunnanensis

seedlings were severely galled by M. arenaria races 1 and 2, M. incognita, M javanica,

and M. hapla, but only supported populations ofM. hapla. Reporting on the

susceptibility of Acer spp. to Meloidogyne spp., Santamour (1992) found several Acer

spp. resistant or tolerant toM. arenaria races 1 and 2, M. hapla, M. incognita, M.

javanica, and M. querciana, and reported a wide range in susceptibility, even among

progeny of a single tree.

Several authors documented variable root gall sizes and shapes on different

susceptible ornamental crops. Bernard and Witte (1987) documented that M. hapla-

induced galls were minute and difficult to detect in Abelia x grandiflora, Photinia x

fraseri, Spirae x bumalda cv. Froebelii, and S. x vanhouttei. Galls were intercalary and

spindle-shaped on thick, fleshy roots of Hydrangea paniculata and Viburnum carlesii,

and spherical and terminal on Cornusflorida roots infected with M. hapla. However, C.

florida infected with M. incognita exhibited spindle-shaped, intercalary galls.

In testing certain cultivars of Tagetespatula to M incognita, M. arenaria, and M

hapla, Motsinger et al. (1977) found that only one cultivar, Tangerine, was free of root

galls or egg masses. They further suggested that certain T. patula cultivars act as trap

crops rather than produce nematicidal agents from their roots. Similarly, pathogenicity

studies conducted in India found only T. erecta cv. Crackerjack to be resistant to M

javanica as no galls or penetration were observed on this cultivar following inoculation









with the pathogen (Singh and Gupta, 1993). In addition, Eisenback (1987) tested eleven

populations ofM. hapla of different cytological forms and from different geographic

origins for their pathogenicity on T. erecta cv. Carnation. Although most M hapla

populations did not reproduce on this host, several populations varied in their

pathogenicity on this host, causing varying degrees of root galling and root proliferation

in response to infection.

Perhaps no group of plants has been studied more for their susceptibility to root-

knot nematodes as those in the genus Ilex. Susceptibility studies involving Ilex spp.

generate conflicting results that may be attributed to nematode variants and differing host

susceptibilities. Symptoms often observed in root-knot infected Ilex spp. include poor

growth and vigor, foliage yellowing and bronzing, dieback of branches, root-system

distortion, and in severe infections, death (Haasis et al., 1961). Furthermore, Ilex spp.

foliage chlorosis is often associated with root galling (Heald, 1967). Meloidogyne

incognita is frequently observed parasitizing Ilex spp. (Bernard et al., 1994). Nemec and

Struble (1968) documented the pathogenicity ofM. incognita on I. cornuta cv. Burfordii,

L crenata cv. Hetz, and I. cassine var. angustifolia. Haasis et al. (1961) found I. crenata

to be severely damaged by M. incognita and M. hapla. Similarly, Barker and Benson

(1977) found I. crenata cv. Convexa, Helleri, and Rotundifolia to be susceptible to M.

arenaria, and Stokes (1982) documented I. crenata to be frequently infected by M.

javanica andM. incognita in commercial nurseries. On the contrary, Barker et al. (1979)

found I. cornuta cv. Burfordii and I. vomitoria cv. Nana to be resistant and tolerant to M.

arenaria, respectively, and I. cornuta cv. Rotunda a poor host toM. arenaria. McSorley

and Dunn (1990) found I. cornuta cv. Burfordii to be a nonhost toM. arenaria, M.









incognita races 1 and 3, andM.javanica. Sasser et al. (1966) observed no galling of I.

crenata cv. Convexa, Latifolia, and Helleri, or cornuta cv. Burfordii following

inoculation with M hapla, but I. crenata cv. Convexa, Latifolia, and Helleri were highly

susceptible to M. incognita, M. javanica, and M arenaria. Heald (1967) indicated that I.

crenata cv. Helleri was susceptible to M. incognita, M. javanica, M. arenaria, and M.

hapla in greenhouse experiments, in which galled roots were observed and plant weights

were reduced in response to root-knot nematode infections. However, in these

experiments symptoms ofM. hapla were less apparent than those caused by the other

root-knot nematode species. Other holly cultivars that did not support M hapla

reproduction include I. x attanuata cv. Foster No. 2, crenata cv. Hetzii, and I. cornuta

x aquifolium cv. Nellie R. Stevens (Bernard and Witte, 1987). Accounting for these

susceptibility variations among the various Ilex spp. cultivars, Heald (1967) suggested

that plant selection variations and differing isolates ofM. hapla may account for such

inconsistencies. In an attempt to correlate galling inconsistencies with differing

nematode isolates, Bernard et al. (1994) compared galling induced by one isolate of M

incognita from Tennessee and two isolates ofM. hapla from Tennessee and North

Carolina on numerous Ilex spp. cultivars. While all Ilex spp. cultivars supported

reproduction of the M incognita isolate, the M hapla isolates differed in their

pathogenicity on the various Ilex spp. cultivars, inducing variable gall numbers and sizes.

This variability was attributed to geographic and host-adaptive variability of the M hapla

isolates. In examining the effect of root-knot nematode pathogenicity on shoot growth,

Nemec and Struble (1968) found no differences in shoot growth between infected and

non-infected plants after eight weeks in a greenhouse.









Host plant resistance (HPR) to Meloidogyne spp. may involve several

mechanisms, including hypersensitivity reactions, delay in Meloidogyne spp. juvenile

maturation, reduction in numbers of giant cells, and reduction in cortical hypertrophy

(Nemec and Morrison, 1972). Botanically, however, HPR also infers to the plant's

ability to withstand, lessen, oppose, or overcome nematode infections, while an immune

plant suffers no injury (Rohde, 1972). The increasing availability of plant germplasm

and advances in plant science technology may allow for the development of HPR for

plant-parasitic nematode control. Such advancements are crucial with the progressing

withdrawal of chemical nematicides (Roberts, 1992). An ornamental plant used as a

rootstock in a Meloidogyne spp. infested area should exhibit resistance to infection by

these pathogens. For example, several Hibiscus rosa-sinensis cultivars with the potential

for use as rootstocks based on their reaction to inoculations with M. javanica and M.

incognita have been identified (McSorley and Marlatt, 1983).

Codiaeum variegatum

Codiaeum variegatum (L.) Blume, commonly referred to as croton, is an

evergreen, glabrous shrub in the family Euphorbiaceae that grows > 2 m tall. Its petioled

leaves are spirally-arranged, alternate, simple and entire or lobed, ovate-lanceolate and

are marked with yellow, white, or red variegation (Bailey, 1958; Gilman, 1999a). While

flowers occur in racemes from leaf axils, they are inconspicuous (Watkins and Sheehan,

1977). Codiaeum variegatum thrives in USDA zones 9b to 11, where large specimens

may be used as foundation plants, hedges, and borders, while smaller specimen may be

used as annual bedding plants. Additionally, it is commonly used as an interior foliage

plant throughout the United States. Codiaeum variegatum is commercially propagated









mostly through tip cuttings, and to a lesser extent through air layering and seed

propagation. Potting media used for croton production should be light, slightly acidic,

and possess good drainage. In the landscape, crotons grow best in well-draining sandy

soils incorporated with organic matter. Croton producers must balance fertilizing and

light requirements to produce a saleable plant, as too much fertilizer for the amount of

light produces a leggy, weak plant, while too little fertilizer produces a woody, stunted

plant. While most croton cultivars require high light, many cultivars that tolerate less

light, including C. variegatum cv. Gold Dust, are available, and the market for indoor-

thriving cultivars is increasing. Insect pests of crotons include Tetranychus spp. (spider

mites), Pseudococcus longispinus (long tailed mealybug), Planococcus citri (citrus

mealybug), Maconellicoccus hirsutus (pink hibiscus mealybug), Ferrisia virgata (striped

mealybug), Polyphagotarsonemus latus (broad mite), and numerous scale species. While

stem galls, rots, rusts, scabs, and blights are sometimes encountered on crotons, they are

not serious problems (Gilman, 1999a; Henny et al., 1991; Reeves and Bell, 1988; Stamps

and Osborne, 2003). Nematode parasites of croton include Paratylenchus spp. (Ibrahim

and Al-Yahya, 2002) and Hoplolaimus spp. (Sher, 1954).

Salvia lIiint ai t

Salvia lemn/ti,/ Cav. is a 1-m-tall herbaceous perennial shrub, commonly known

as Mexican sage, which belongs to the family Labiatae. This plant possesses cylindrical,

somewhat tapering white branches that are covered with fine wool. Its leaves, silver-gray

colored and arranged in a whorl, possess a short petiole and are 5 to 15 cm long,

lanceolate-linear, crenate, and are pubescent above and tomentose beneath. The plant

exhibits many flowers that occur on 15 to 25-cm-long racemes that appear in the summer.









Flowers possess a funnel-shaped, violet to lavender calyx and a white or purple corolla,

to 2 cm long (Bailey, 1958; Gilman and Marshall, 1999; Anonymous, 2000). Salvia

leucantha thrives in USDA zones 7 to 11 and is used in mass plantings, borders, and

edging. In zones 9 to 11, flowers persist longer than in zones 7 and 8. The plant grows

well in full sun and prefers well-drained, rich, sandy soils. Mexican sage is commercially

propagated through cuttings and divisions (Gilman and Marshall, 1999). Buhrer (1938)

reported on Meloidogyne spp. parasitizing S. leucantha. On the contrary, the plant was

reported resistant to M incognita and M. javanica in India (Krishnaprasad, 1979).

Liriope muscari

Liriope muscari Bailey, commonly known as lilyturf, is a grass-like herbaceous

perennial in the family Ruscaceae (Gilman, 1999b; Judd, 2003). The plant possesses a

short, thick rhizome by which it spreads. The stemless, dark green leaves are stiff, long,

linear, with an acute tip, and do not exceed the floral scapes, which are 25 to 70 cm high

and occur in spring and summer. Scapes possess 10 or more whorls of up to seven

flowers, each 0.6 to 0.7 cm across (Bailey, 1958). The plant grows in tufts of various

heights, depending on the cultivar. Liriope muscari is often used as a ground cover or for

edging, and prefers fertile, well-drained soils. Commercially, this plant is propagated

though divisions and seldom through seed (Watkins and Sheehan, 1977). It is also often

propagated through tissue culture. Liriope muscari thrives in USDA zones 6 to 10 in

shady areas, but tolerates full sun except in the hottest areas of the south, as severe leaf

tip burn occurs in such conditions. Various cultivars of lilyturf are available, including

variegated foliage (L. muscari cv. Variegata) and white and purple-colored flower

varieties. Liriope muscari cv. Evergreen Giant grows to 30-cm-high and spreads slower









than other cultivars. Recently, a leaf and crown rot disease, caused by Phytophthora

palmivora has taken its toll on L. muscari in landscapes and nurseries, to the extent that

L. muscari cv. Evergreen Giant may lose its popularity in the landscape industry (Leahy

and Davison, 1999).

Pittosporum tobira

Pittosporum tobira Ait., also known as Japanese pittosporum, is a 3 to 6-m-high

flowering evergreen shrub to small tree in the family Pittosporaceae. Its whorled, thick,

leathery leaves are 5 to 10 cm long and are glabrous, with revolute margins. Japanese

pittosporum is winter-flowering, possessing 1-cm-long white, fragrant flowers in terminal

umbels. Pittosporum tobira is often grown as a shrub for foundations, hedges, mass

plantings, as a screen, in planter boxes, and may also be trained as a small tree (Bailey,

1958; Gilman, 1999c, 1999d; Stamps, 2002). Furthermore, salt tolerance makes it useful

in seaside plantings, where it is widely used as a hedge and windbreaker (Rinallo and

Bennici, 1989). Since it is damaged by temperatures below -6 C, P. tobira is restricted

to USDA zones 8 to 10. It prefers fertile, slightly acidic soils, is fairly drought tolerant,

and requires minimal fertilization for optimal growth. Pittosporum tobira establishes

well in partial shade to full-sun conditions (Stamps, 2002). Commercially, this plant is

propagated through semi-hardwood cuttings under mist and seldom by seed (Bailey,

1958; Watkins and Sheehan, 1977). Several cultivars that have either been produced or

assigned to the species include P. tobira cv. Compacta, Variegata, and Wheeler's Dwarf,

of which P. tobira cv. Variegata possesses thinner leaves than the type species that are

variegated with white, and is heavily cultivated in Florida and California, mainly for

landscape use and for floral designs (Bailey, 1928; Stamps, 1987, 2002). Diseases that









often occur on P. tobira include angular leaf spot, caused by Cercosporapittospori,

Rhizoctonia aerial blight, caused by Rhizoctonia ramicola or R. solani, southern blight,

caused by Sclerotium rolfsii, root rots, caused by Pythium spp. and Rhyzoctonia spp., and

dieback, caused by Agrobacterium spp., Diaporthe spp., Diplodia spp., Nectriella spp.,

homopsis spp., and Sphaeropsis spp. Other, less common diseases include Alternaria leaf

spot, caused by Alternaria tenuissima, mushroom root rot, caused by Armillariella

tabescens, corticium limb blight, caused by Corticium salmonicolor, rough bark disease,

which may be caused by a virus, and several others (Chase and Simone, 2001).

Pittosporum tobira is also a host for the tomato spotted wilt virus (Gera et al., 2000).

This plant often exhibits symptoms of magnesium, iron, manganese, and copper

deficiencies, especially when cultivated in high-pH soils (Dehgan, 1998). Insect pests

often encountered on P. tobira include Aphis gossypii (melon aphid) and Iceryapurchasi

(cottony cushion scale) (Hamon and Fasulo, 1998). Buhrer (1938) observed

Meloidogyne spp. infected P. tobira plants, and M. arenaria was detected on a declining

hedge of P. tobira in Florida (Bureau of Nematology, 1989). The ectoparasitic nematode

Belonolaimus longicaudatus has been associated with P. tobira cv. Variegatum (syn. P.

tobira cv. Variegata) decline (Rhoades, 1989).

Odontonema cuspidatum

Odontonema cuspidatum (Nees) Kuntze (syn. O. strictum (Nees) Kuntze),

commonly referred to as firespike, is a herbaceous perennial in the family Acanthaceae,

which is hardy in USDA zones 8B to 11. It is a glabrous, erect shrub, 1 to 2-m-high. that

forms clumps that generate from root suckers. It possesses simple, opposite, acuminate,

short-petioled leaves that are entire and undulate, 10 to 30 cm long. Its tubular flowers









are bright red, pink, white, or lavender, up to 2.5 cm long, and occur in long racemes.

Flowers occur year-round in tropical climates and in the fall in temperate climates. This

erect, compact shrub is often used in mass plantings, as a background plant, or a hedge,

and often attracts butterflies and hummingbirds. Following the first frost, it dies to the

ground, but comes back in the spring. For best flower development and persistence, 0.

cuspidatum is planted in full sun in fertile, sandy soil. This plant is commercially

propagated by cuttings and divisions. Aside from Pseudococcus spp. (mealy bugs), no

pests or pathogens have been reported on this plant (Bailey, 1958; Francis, 2004; Gilman

and Delvalle, 1999; Watkins and Sheehan, 1977).

Musa acuminata ssp. zebrina

Musa acuminata Colla. ssp. zebrina Van Houtte ex Planch cv. Rowe Red (syn.

Musa sumatrana Baccari cv. Rowe Red) is a cultivated banana in the family Musaceae

that is used as a decorative ornamental. Descriptions surrounding this species have been

erroneous and confused due to the age of specimens at the time of description.

Additionally, the description of wrongly-named specimen, as evident by differing floral

and fruit characters among the described specimen documented, may have contributed to

the taxonomic confusion (Cheesman, 1985). Recent evidence has led to the

reclassification of this plant as M acuminata ssp. zebrina. The cultivar Rowe Red is not

described in the literature, and may have been designated to the species by its cultivators.

Musa acuminata ssp. zebrina is rhizomatous and possesses red-ornamented green

pseudostems > 2.5 m high, which are formed via sheathing leaf bases, and continuously

sucker from pseudostem bases. Its leaves are spirally arranged, < 1.8 m long and 0.5 m

wide, rounded at the base, glaucous, purple beneath, and irregularly purple-patched









above, with a 0.3-m-long petiole. The floral spikes droop slightly, and the rachis on

which they appear is pubescent. The fruit is inedible, dry, cylindrical and curved, < 7 cm

long and 1.3 cm diam. (Baker, 1893; Cheesman, 1985; Griffiths, 1994; Ricker, 1937).

Musa spp. thrive through USDA zone 9, as long as they are protected in northern regions

of zone 9b. Plants in this genus prefer moist, fertile soils and ample fertilization. Musa

acuminata spp. zebrina cv. Rowe Red may be propagated via sucker divisions, but is

produced commercially via tissue culture (Dehgan, 1998). Most Musa spp. are

susceptible to Black Sigatoka, caused by Mycosphaerellafijiensis, Yellow Sigatoka,

caused by Mycosphaerella musicola, and Panama Disease, caused by Fusarium

oxysporum f sp. cubense. Plant-parasitic nematode pathogens of Musa spp. include M.

incognita, M. javanica, M. arenaria, and other Meloidogyne spp., Radopholus simillis,

Pratylenchus goodeyi, P. coffeae, and Helicotylenchus multicinctus (Adiko, 1988; De

Waele and Davide, 1998; Jaizme-Vega et al., 1997; Stoffelen et al., 2000).

Objectives

The objectives of this research were to:

1. Evaluate the host status of several woody and perennial ornamental plants to

Meloidogyne incognita race 2, M. javanica, M. arenaria race 1, and M

mayaguensis, in separate experiments.

2. Differentiate malate dehydrogenase and esterase phenotypes of Meloidogyne

spp. that were collected from infected ornamental plants in Florida using

PAGE.

3. Evaluate the usefulness of PAGE in differentiating Meloidogyne spp. for

routine extension diagnostic purposes.















CHAPTER 2

REPRODUCTION OF FOURMELOIDOGYNE SPP. ON SEVERAL SPECIES OF
PERENNIAL ORNAMENTAL PLANTS


Introduction

Root-knot nematodes (Meloidogyne spp.) are the most damaging group of plant-

parasitic nematodes to ornamental plants in Florida (McSorley and Dunn, 1989). As

serious pathogens of many woody ornamental species, root-knot nematodes limit

productivity by damaging numerous nursery crops directly and by forming disease

complexes with certain soil-borne fungal pathogens (Barker and Benson, 1977; Benson

and Barker, 1985; Nigh, 1972; Santamour and Riedel, 1993; Walker and Melin, 1998b;

Zarina and Abid, 1995). Furthermore, since root-knot nematode populations often thrive

and cause damage on perennial hosts for many months and years, damage threshold

levels do not apply for such plants (LaMondia, 1995).

Symptoms associated with root-knot nematode infection include root galls and

root rots, shoot chlorosis, stunted growth, and other symptoms commonly associated with

nutritional deficiencies (Bala and Hosein, 1996; Bird, 1974; Misra et al., 2002; Santo and

Lear, 1976; Zarina and Abid, 1995). Such symptoms are often associated with general

decline (Nigh, 1972), poor yield, and wilt diseases (Rajendran et al., 1975).

Published work on the susceptibility of woody ornamentals to Meloidogyne spp.

is limited. In Florida, several authors (Giblin-Davis et al., 1992; Lehman, 1984a, 1984b;

Lehman and Barnard, 1982; McSorley and Dunn, 1989, 1990; McSorley and Marlatt,









1983; Stokes, 1982) reported on the pathogenicity of a number ofMeloidogyne spp. to

several perennial ornamental plants. Other reports of root-knot infection of ornamentals

from Alabama (Heald, 1967), Arizona (Nigh, 1972), California (Santo and Lear, 1976;

Viglierchio, 1979), Connecticut (LaMondia, 1995, 1996, 1997), Georgia (Heald, 1967;

Walker and Melin, 1998a, 1998b), New Jersey (Davis and Jenkins, 1960), North Carolina

(Barker et al., 1979; Barker and Benson, 1977; Benson and Barker, 1982; Haasis et al.,

1961), Oklahoma (Nemec and Morrison, 1972; Nemec and Struble, 1968), Tennessee

(Bernard and Witte, 1987; Bernard et al., 1994; Niblack and Bernard, 1985), and

Washington, DC (Santamour, 1992; Santamour and Riedel, 1993, 1995) have been

published. Compared to research on the pathogenicity of Meloidogyne spp. on

agronomic crops, and taking into account the vast array of perennial species cultivated,

research on the susceptibility of woody and perennial ornamental plants to root-knot

nematodes is minimal.

The objectives of these studies were to evaluate the host status of several woody

and perennial ornamental plants to Meloidogyne incognita race 2, M. javanica, M.

arenaria race 1, and M mayaguensis, in separate experiments. The experiments were

carried out in a controlled environmental chamber (growth room) and a greenhouse at the

University of Florida. The plant species evaluated were Liriope muscari (Lilyturf) cv.

Evergreen Giant, Pittosporum tobira (Pittosporum) cv. Variegata, Odontonema

cuspidatum (Firespike), Codiaeum variegatum (Croton) cv. Gold Dust, Musa acuminata

ssp. zebrina (ornamental banana) cv. Rowe Red, and Salvia leiinI/ tht (Mexican Sage).

While cultivars of Salvia leucantha do not exist, two forms, one with a purple corolla and

another with a white corolla, were tested in the greenhouse experiment.









Materials and Methods


General Cultivation Practices

Planting. Experimental plants were planted as liners in 15.24-cm plastic (Model

AZE0600, ITML Horticultural Products, Ontario, CA) and clay pots for growth room and

greenhouse experiments, respectively, each containing 800 cm3 of planting media. Liners

were obtained from commercial propagators (Table 2-1). Several liners were tested for

the presence of plant-parasitic nematodes by the rapid centrifugal-flotation technique

(Jenkins, 1964) prior to planting.

Planting media. Experiments were identical with respect to the soil mixture used

for planting experimental plants and maintaining nematode inocula. The soil mixture

was a 2:1 ratio of sand and potting mix (Jungle Growth Professional Growers Mix,

Statham, GA), respectively. The medium was tested for the presence of plant-parasitic

nematodes via the rapid centrifugal-flotation method prior to the initiation of each

experiment.

Treatments. Five root-knot nematode species were used in the growth room and

greenhouse experiments. The nematodes were: (i) M. incognita race 2, (ii) M. javanica,

(iii) M arenaria race 1, (iv) M mayaguensis, and (v) non-inoculated control. Exceptions

include the 5 June 2003-inoculated Liriope muscari cv. Evergreen Giant and Pittosporum

tobira cv. Variegata trials, in which no non-inoculated control treatments were included.

Meloidogyne spp. The Meloidogyne spp. utilized in these experiments were

originally obtained from J. A. Brito, Florida Department of Agriculture and Consumer

Services, Division of Plant Industry, Gainesville, FL. They originated from single egg









Table 2-1. Crop and source of liners used for growth room and greenhouse
experiments.


Crop


Source


Liriope muscari cv. Evergreen Giant

Pittosporum tobira cv. Variegata


purple corolla


Salvia lemin/tih white corolla

Odontonema cuspidatum

Codiaeum variegatum cv. Gold Dust

Musa acuminata ssp. zebrina cv. Rowe Red


Agri-Starts III, Inc., Eustis, FL

Jon's Nursery, Inc., Eustis, FL

Liner Source, Inc., Mount Dora, FL

Hatchett Creek Farms, LLC,
Gainesville, FL

Robrick Nursery, Inc., Hawthorne, FL

Yoder Brothers, Inc., Lancaster, PA

Robrick Nursery, Inc., Hawthorne, FL

Parrish Nurseries, Inc., Parkland, FL

Agri-Starts I, Inc., Apopka, FL


Salvia letI '/n//h,









masses, and their species designation was confirmed using perineal patterns and isozyme

phenotypes, resolved on polyacrylamide gel slabs following electrophoresis.

Meloidogyne spp. Extraction and Inoculation

Meloidogyne spp. egg and second-stage juvenile (J2) inocula were extracted from

Lycopersicon esculentum Mill. (tomato) cv. Rutgers by the sodium hypochlorite (NaOC1)

procedure (Hussey and Barker, 1973), via the shaking of infected roots in 0.53% NaOCI

(Regular Ultra Bleach, Publix Super Markets, Lakeland, FL) solution for 30 seconds,

followed by the immediate rinsing of the suspension with 10 liters of water.

Meloidogyne spp. eggs and J2 were inoculated onto test and tomato plants for

experimental and inoculum-increase purposes on the same day that the eggs were

extracted from infected tomato roots. Meloidogyne spp. inocula were transported to the

growth room or greenhouse in sealed 500-ml Erlenmeyer flasks. An aquarium air pump

equipped with a 1-ml, 22.8-cm-long, disposable glass Pasteur pipet that was inserted into

the Erlenmeyer flask, was used to keep the root-knot nematode inocula thoroughly and

evenly suspended. Five thousand eggs and J2 were pipetted into three equidistant holes

approximately 3-cm-deep in the pre-moistened soil surrounding the base of each plant.

Test tomato plants were included in every experiment as a control for inoculum viability.

Immediately after inoculation, the holes were covered with the same soil mixture used

throughout the experiment. The bench area used for inoculation was thoroughly

disinfected with 6.0% regular ultra bleach, and all pipets were replaced between

inoculations of the respective Meloidogyne spp. All pots were kept on inverted clay

drainage saucers for the duration of the experiment to avoid Meloidogyne spp.

contamination among pots.









Fertilization and watering. In all the experiments, a solution of 0.21 g N, 0.09 g

P, and 0.17 g K (0.21 g 20-20-20 soluble fertilizer, Grace-Sierra Horticultural Products,

Milpitas, CA) in 200 ml water was applied to each pot weekly and plants were watered as

needed.

Pesticides. Tetranychus spp. (spider mites) were encountered parasitizing P.

tobira in these experiments. Bifenthrin was sprayed at a rate of 0.13 ml a.i./liter when

needed. In addition, all tomato plants and all greenhouse-grown crops in these

experiments were treated with imidacloprid at a rate of 12.5 mg a.i. per pot, for Bemisia

spp. (whitefly).

Photoperiod and temperature. Growth room light was provided by 400-W general

lighting metal halide lamps (Osram Sylvania, Danvers, MA), directed toward the

experimental plants, which were situated on 1-m-tall benches. Lights were suspended 1

m above the bench tops, and light intensity was recorded at 256.18 + 52.82 [tmol-s-lm-2

at a distance of 35 cm above the bench top using a photometer (LI-COR, Model LI-189,

Lincoln, NE). Plants were on a 14-hr light cycle, from 7:00 PM to 9:00 AM. Air and

media temperatures in the growth room were maintained using an air conditioner and

were measured using a standard thermometer at 26 to 32 C and 24 to 26 C,

respectively.

Experimental Design

Growth room and greenhouse experiments were arranged in a randomized

complete block design, with six and three replications in the growth room and

greenhouse, respectively. Each experiment was conducted twice in the growth room and

once in the greenhouse. While growth room experiments were harvested approximately









two months post-inoculation, greenhouse experiments were conducted over five months

or longer, as outlined in Table 2-2. The S. leucantha growth room trials differed with

respect to the plant flower-color forms used. Plants used in the first growth room trial

possessed a mixture of purple- and white-colored corolla S. lei'mintl plants, while plants

used in the second growth room trial were limited to the purple-corolla form.

Experimental Plants Processing and Meloidogyne spp. Egg Extraction and Counting

All experiments were processed by their respective replications. While entire root

systems were processed in the growth room experiments, 10 g of each root system were

evaluated for plants in greenhouse experiments due to the extensive root system of these

plants. Egg numbers from greenhouse-experimental plants were calculated based on total

root weight of respective plants.

Experimental plants were moved, by replication, from the growth room or greenhouse to

the nematode assay laboratory at the University of Florida. Each plant was completely

processed separately prior to processing of the remaining plants in the respective

replication. Each plant was removed from the plastic pot and the soil surrounding its

roots was gently shaken into a container. The above-ground portion of each plant was

then cut and placed into a pre-weighed 25.4-cm x 33.0-cm manila envelope (Sparco,

Moorestown, NJ), which was placed in a 70 C oven. Drying times were determined by

weighing envelopes every 24 hours until no further weight change was detected. The

remaining root system was then immersed in water to remove any adhering soil. The

soil-free root system was patted dry and fresh root system weights and gall ratings

(Taylor and Sasser, 1978) were determined and recorded. Eggs were extracted using

0.53% NaOCl (Hussey and Barker, 1973) by a procedure modified by Boneti and Ferraz












Table 2-2. Crops, experimental sites, liner planting dates, inoculation dates, and study lengths for all crops in the growth room
and greenhouse Meloidogyne spp. studies carried out at the University of Florida during 2003 to 2005.


Crop

Liriope muscari cv. Evergreen Giant

Liriope muscari cv. Evergreen Giant

Liriope muscari cv. Evergreen Giant

Pittosporum tobira cv. Variegata

Pittosporum tobira cv. Variegata

Pittosporum tobira cv. Variegata

Salvia Alw'/uinh purple and white corolla
mix
Salvia kluiihtla purple corolla

Salvia kluiihtla purple corolla

Salvia kluiihtl, white corolla

Odontonema cuspidatum


Experimental
site

Ca

C

G

C

C

G

C

C

G

G

C


Liner planting date

22 May 2003

26 August 2003

19 February 2004

22 May 2003

26 August 2003

19 February 2004

25 October 2003

15 January 2004

19 February 2004

19 February 2004

25 October 2003


Inoculation date

5 June 2003

9 September 2003

4 March 2004

5 June 2003

9 September 2003

4 March 2004

8 November 2003

29 January 2004

4 March 2004

4 March 2004

8 November 2003


Study length
(days)
65

63

354

70

67

336

68

70

154

182

81












Table 2-2. Continued


Crop

Odontonema cuspidatum

Odontonema cuspidatum

Codiaeum variegatum cv. Gold Dust

Codiaeum variegatum cv. Gold Dust

Codiaeum variegatum cv. Gold Dust

Musa acuminata ssp. zebrina cv. Rowe Red

Musa acuminata ssp. zebrina cv. Rowe Red

Musa acuminata ssp. zebrina cv. Rowe Red

aC = Growth-room, G = Greenhouse


Experimental
site
C

G

C

C

G

C

C

G


Liner planting date

15 January 2004

19 February 2004

22 June 2004

8 September 2004

22 June 2004

22 June 2004

8 September 2004

22 June 2004


Inoculation date

29 January 2004

4 March 2004

6 July 2004

22 September 2004

6 July 2004

6 July 2004

22 September 2004

6 July 2004


Study length
(days)
78

263

79

96

239

72

106

205









(1981). One ml of solution was drawn out of each treatment and put into a 1-ml, 48-

division equine egg parasite counting slide (Advanced Equine Products, Issaquah, WA),

which was counted in its entirety. Three 1-ml aliquots were counted per treatment, and

the average of the three counts was used for statistical analysis.

Statistical Analysis

Data were subjected to analysis using Analysis of Variance (ANOVA). Mean

comparisons among the treatments were performed with Duncan's multiple range test

using SAS software (SAS Institute, Cary, NC).

Host Status Classification

The host status of the tested perennials was determined based on the reproduction

factor (Rf), which was calculated by dividing the final root-knot nematode density per

plant (Pf) by the inoculated root-knot density of 5,000 eggs and J2 (Pi). A Rf> 1.0 was

designated as a good host, 1.0 > Rf> 0.1 a poor host, and Rf < 0.1 a nonhost (Sasser et

al., 1984).

Results

Liriope muscari cv. Evergreen Giant

Results for the two L. muscari cv. Evergreen Giant growth room trials were

heterogeneous and are therefore presented separately (Tables 2-3 and 2-4). In the first

trial, there were no differences (P 0.05) among treatments for root-gall index, root

weight, or dry shoot weight. Differences (P 0.05) were observed among treatments for

number of eggs per plant and number of eggs per g of roots, where fewer eggs were

produced by M. arenaria than by the other three species. In the second trial, there were

no differences (P < 0.05) among treatments for root-gall index, root weight, or shoot dry












Table 2-3. Treatments, root-gall indexes, root weights, number of eggs per plant, number of eggs per g of roots, and dry shoot
weights from the first Liriope muscari cv. Evergreen Giant growth room trial.


Root-gall index


Root Weight Mean number of eggs Mean number of Dry Shoot
(g) per plant eggs per g roots Weight (g)


Control --- --- --- -
M. incognita race 2 1.17 0.82a 35.95 19.71 31,920.00 44,748.39 ab 848.43 1,047.04 a 7.62 4.26
M.javanica 1.33 1.63 36.33 23.16 19,897.78 36,366.07 a 492.63 608.77 a 8.25 3.17
M. arenaria race 1 0.83 0.82 37.67 20.08 377.78 1,131.08b 9.11 26.02 b 8.30 3.70
M. mayaguensis 1.00 + 0.00 37.73 14.28 13,822.22 12,905.42 a 373.04 320.68 a 7.47 4.23
a0 = no galls; 1 = 1 to 2 galls; 2 = 3 to 10 galls; 3 = 11 to 30 galls; 4 = 31 to 100 galls; 5 = more than 100 galls per root system
(Taylor and Sasser, 1978).
bData are means of six replications. Means in columns followed by a common letter are not different (P < 0.05) according to
Duncan's multiple range test. Egg numbers were log-transformed prior to data analysis, but non-transformed numbers are shown.


Treatment


1












Table 2-4. Treatments, root-gall indexes, root weights, number of eggs per plant, number of eggs per g of roots, and dry shoot
weights from the second Liriope muscari cv. Evergreen Giant growth room trial.


Root-gall index
(0-5)


Root Weight Mean number of eggs per Mean number of
(g) plant eggs per g roots


Control 0.00 0.00a 39.18 14.76 0.00 0.00 c 0.00 0.00 c 12.77 4.22
M. incognita race 2 0.00 0.00 40.00 21.73 231.11 418.14 b 5.36 8.27b 10.20 3.51
M.javanica 0.33 1.03 37.95 16.83 684.44 829.44 a 20.93 33.18 a 9.73 5.94
M. arenaria race 1 0.00 + 0.00 38.05 + 22.35 8.89 + 27.54 c 0.26 0.81 c 10.63 6.35
M. mayaguensis 0.00 + 0.00 36.70 + 24.73 1,000.00 + 833.61 a 30.51 + 38.39 a 10.93 8.26
a0 = no galls; 1 = 1 to 2 galls; 2 = 3 to 10 galls; 3 = 11 to 30 galls; 4 = 31 to 100 galls; 5 = more than 100 galls per root system
(Taylor and Sasser, 1978).
bData are means of six replications. Means in columns followed by a common letter are not different (P < 0.05) according to
Duncan's multiple range test. Egg numbers were log-transformed prior to data analysis, but non-transformed numbers are shown.


Treatment


Dry Shoot
Weight (g)


1









weight. Differences (P < 0.05) were observed among treatments for number of eggs per

plant and number of eggs per g of roots, where the number of eggs produced by M

arenaria was not different (P < 0.05) from the nematode control. Furthermore, M.

javanica and M. mayaguensis produced more eggs than M incognita, which produced

more eggs than M. arenaria.

In the greenhouse experiment (Table 2-5), differences (P 0.05) were observed across all

parameters. In this experiment, M. incognita and M mayaguensis produced more galls

than the other treatments, and the control treatment had a significantly higher root weight

than all other treatments. Meloidogyne incognita, M. javanica, and M. mayaguensis

produced more eggs per plant than M. arenaria and the control treatment, from which no

eggs were recovered. However, M. incognita and M. mayaguensis produced more eggs

per g of roots than M. javanica, which produced more eggs per g of roots than M.

arenaria. Finally, the control and M. incognita treatments had higher dry shoot weights

than the M. javanica and M. arenaria treatments, and the M. mayaguensis treatment dry

shoot weights were not different (P < 0.05) than any of the other treatment. The data

suggest that L. muscari cv. Evergreen Giant is a good host to M. incognita race 2 (Rf =

97.1), M. javanica (Rf = 16.6), and M. mayaguensis (Rf = 91.0), and a poor host to M

arenaria race 1 (Rf = 0.12).

Pittosporum tobira cv. Variegata

Results for the two P. tobira cv. Variegata growth room trials were heterogeneous and

are therefore presented separately in tables 2-6 and 2-7. The results from the greenhouse

experiment are presented in table 2-8. Root galls were not observed on any of the plants

in the three studies. No differences (P < 0.05) were observed among treatments for root-












Table 2-5. Treatments, root-gall indexes, root weights, number of eggs per plant, number of eggs per g of roots, and dry shoot
weights from the Liriope muscari cv. Evergreen Giant greenhouse experiment.


Root-gall index
(0-5)


Root Weight Mean number of eggs Mean number of Dry Shoot
(g) per plant eggs per g roots Weight (g)


Control 0.00 + 0.00 ba 141.50 55.65 a 0.00 + 0.00 b 0.00 + 0.00 c 30.80 3.83 a
M. incognita race 2 5.00 + 0.00 a 104.33 15.10 b 485,507.38 184,451.74 a 4,646.22 1,555.40 a 29.33 10.05 a
M.javanica 0.00 0.00 b 70.93 8.11 b 83,099.47 283,116.92 a 1,100.44 3,742.78 b 21.40 4.40b
M. arenaria race 1 0.00 + 0.00 b 81.70 34.01 b 597.33 2,069.22 b 6.22 1.55 c 19.70 4.13 b
M. mayaguensis 5.00 + 0.00 a 82.33 37.12 b 454,887.02 412,521.08 a 5,467.56 3,903.75 a 25.20 17.97 ab
a0 = no galls; 1 = 1 to 2 galls; 2 = 3 to 10 galls; 3 = 11 to 30 galls; 4 = 31 to 100 galls; 5 = more than 100 galls per root system
(Taylor and Sasser, 1978).
bData are means of three replications. Means in columns followed by a common letter are not different (P < 0.05) according to
Duncan's multiple range test. Egg numbers were log-transformed prior to data analysis, but non-transformed numbers are shown.


Treatment












Table 2-6. Treatments, root-gall indexes, root weights, number of eggs per plant, number of eggs per g of roots, and dry shoot
weights from the first Pittosporum tobira cv. Variegata growth room trial.


Root-gall index Root Weight Mean number of eggs Mean number of
(0-5) (g) per plant eggs per g roots


Dry Shoot Weight
(g)


Control --- --- --- -
M. incognita race 2 0.00 0.00a 11.67 6.91 13.33 44.62 abb 1.40 5.07 ab 11.67 2.44
M.javanica 0.00 + 0.00 12.62 4.63 53.33 168.65 a 4.32 + 13.10 a 12.67 + 2.33
M. arenaria race 1 0.00 + 0.00 12.10 + 8.07 13.33 44.62 ab 0.79 2.51 ab 12.05 + 3.63
M. mayaguensis 0.00 + 0.00 12.23 7.68 0.00 + 0.00 b 0.00 0.00 b 11.83 3.69
a0 = no galls; 1 = 1 to 2 galls; 2 = 3 to 10 galls; 3 = 11 to 30 galls; 4 = 31 to 100 galls; 5 = more than 100 galls per root system
(Taylor and Sasser, 1978).
bData are means of six replications. Means in columns followed by a common letter are not different (P < 0.05) according to
Duncan's multiple range test. Egg numbers were log-transformed prior to data analysis, but non-transformed numbers are shown.


Treatment


1












Table 2-7. Treatments, root-gall indexes, root weights, number of eggs per plant, number of eggs per g of roots, and dry shoot
weights from the second Pittosporum tobira cv. Variegata growth room trial.


Treatment Root-gall Root Weight Mean number of eggs Mean number of Dry Shoot
index (0-5) (g) per plant eggs per g roots Weight (g)
Control 0.00 0.00a 11.57 3.78 0.00 + 0.00 0.00 + 0.00 11.42 + 1.29
M. incognita race 2 0.00 + 0.00 12.22 4.16 71.11 322.95 5.19 23.57 11.82 1.34
M.javanica 0.00 + 0.00 11.65 3.05 8.89 43.55 0.67 3.30 11.07 4.05
M. arenaria race 1 0.00 + 0.00 11.82 + 1.24 8.89 43.55 0.74 3.63 11.58 1.27
M. mayaguensis 0.00 +0.00 10.08 2.99 31.11 78.50 3.56 9.28 10.37 1.14
Data are means of six replications. No differences (P < 0.05) were observed according to Duncan's multiple range test.
a0 = no galls; 1 = 1 to 2 galls; 2 = 3 to 10 galls; 3 = 11 to 30 galls; 4 = 31 to 100 galls; 5 = more than 100 galls per root system
(Taylor and Sasser, 1978).












Table 2-8. Treatments, root-gall indexes, root weights, number of eggs per plant, number of eggs per g of roots, and dry shoot
weights from the Pittosporum tobira cv. Variegata greenhouse experiment.


Treatment Root-gall Root Weight Mean number of eggs Mean number of Dry Shoot
index (0-5) (g) per plant eggs per g roots Weight (g)
Control 0.00 + 0.00a 24.83 14.61 0.00 + 0.00 0.00 + 0.00 45.10 + 19.08
M. incognita race 2 0.00 + 0.00 23.13 5.32 0.00 +0.00 0.00 0.00 49.47 8.32
M.javanica 0.00 + 0.00 26.87 8.26 0.00 + 0.00 0.00 + 0.00 46.37 6.84
M. arenaria race 1 0.00 + 0.00 25.30 + 23.30 0.00 + 0.00 0.00 + 0.00 43.40 + 6.54
M. mayaguensis 0.00 + 0.00 13.80 + 0.87 0.00 + 0.00 0.00 + 0.00 35.03 9.47
Data are means of three replications. No differences (P < 0.05) were observed according to Duncan's multiple range test.
a0 = no galls; 1 = 1 to 2 galls; 2 = 3 to 10 galls; 3 = 11 to 30 galls; 4 = 31 to 100 galls; 5 = more than 100 galls per root system
(Taylor and Sasser, 1978).









gall index, root weight, or dry shoot weight in either of the growth room trials or the

greenhouse experiment. There were differences (P < 0.05) among treatments for number

of eggs per plant and number of eggs per g of roots in the first growth room trial, where

M. javanica produced more eggs than M. mayaguensis, which produced no eggs.

However, in the second growth room trial and the greenhouse experiment no differences

(P < 0.05) were found. The data suggests that P. tobira cv. Variegata is a nonhost to the

Meloidogyne spp. isolates evaluated (Rf < 0.1).

Salvia lIhn, t(ihth

The results from the first and second purple-corolla S. leucantha growth room

trials are heterogeneous, and are presented in tables 2-9 and 2-10, respectively. Root-

knot nematode galls were not detected on any treatment in the first growth room trial

(Table 2-9). Furthermore, no differences (P < 0.05) were observed among treatments for

root weight or dry shoot weight. Differences (P < 0.05) were observed among treatments

for the number of eggs per plant and the number of eggs per g of roots. The number of

eggs per plant was not different (P < 0.05) between M. javanica and M. mayaguensis.

However, these two species produced more eggs than M.

arenaria and M. incognita. The number of eggs per g of roots produced by M.

javanica was greater than the other species. Results for the second growth room trial

(Table 2-10) indicate differences (P < 0.05) among treatments for root-gall index, number

of eggs per plant, and number of eggs per g of roots. There were no differences (P

0.05) among treatments for root weight and dry shoot weight in this trial. Root galls were

only observed in the M. javanica treatment. The root-gall index for M. javanica was

greater than that for the other species, in which root galls were not detected. For the












Table 2-9. Treatments, root-gall indexes, root weights, number of eggs per plant, number of eggs per g of roots, and dry shoot
weights from the first Salvia leucantha growth room trial.


Root-gall index
(0-5)


Root Weight Mean numb
(g) pl


er of eggs per Mean number of Dry Shoot
plant eggs per g roots Weight (g)


Control 0.00 +0.00a 20.15 7.09 0.00 0.00 b 0.00 0.00 b 14.35 2.59
M. incognita race 2 0.00 + 0.00 22.42 + 13.45 0.00 + 0.00 b 0.00 + 0.00 b 13.72 + 2.62
M.javanica 0.00 + 0.00 20.03 12.97 5,906.67 19,410.10 a 463.25 1,527.17 a 13.35 + 3.09
M. arenaria race 1 0.00 + 0.00 21.68 + 9.22 0.00 + 0.00 b 0.00 + 0.00 b 12.50 + 5.02
M. mayaguensis 0.00 + 0.00 18.68 9.16 2,053.33 9,207.13 a 94.03 402.60 b 11.75 8.95
a0 = no galls; 1 = 1 to 2 galls; 2 = 3 to 10 galls; 3 = 11 to 30 galls; 4 = 31 to 100 galls; 5 = more than 100 galls per root system
(Taylor and Sasser, 1978).
bData are means of six replications. Means in columns followed by a common letter are not different (P < 0.05) according to
Duncan's multiple range test. Egg numbers were log-transformed prior to data analysis, but non-transformed numbers are shown.


Treatment


1












Table 2-10. Treatments, root-gall indexes, root weights, number of eggs per plant, number of eggs per g of roots, and dry
shoot weights from the second Salvia /in'//t i/h growth room trial.


Treatment Root-gall index Root Weight Mean number of eggs Mean number of Dry Shoot
(0-5) (g) per plant eggs per g roots Weight (g)
Control 0.00 + 0.00 bab 16.05 4.03 0.00 + 0.00 d 0.00 + 0.00 d 9.17 2.57
M. incognita race 2 0.00 0.00 b 17.23 6.35 5,915.56 11,225.64bc 326.02 568.49 bc 9.52 1.87
M.javanica 1.00 + 1.26 a 18.42 7.73 113,048.89 164,013.68 a 5,836.49 7,267.12 a 9.10 3.05
M. arenaria race 1 0.00 + 0.00 b 17.18 8.07 4,537.78 12,798.02c 239.18 661.38 c 9.35 2.94
M. mayaguensis 0.00 + 0.00 b 16.57 4.88 36,333.33 126,118.27 b 2,298.54 8,382.65 b 9.20 2.10
a0 = no galls; 1 = 1 to 2 galls; 2 = 3 to 10 galls; 3 = 11 to 30 galls; 4 = 31 to 100 galls; 5 = more than 100 galls per root system
(Taylor and Sasser, 1978).
bData are means of six replications. Means in columns followed by a common letter are not different (P < 0.05) according to
Duncan's multiple range test. Egg numbers were log-transformed prior to data analysis, but non-transformed numbers are shown.









number of eggs per plant and the number of eggs per g of roots, the number of eggs

produced followed the following order, from high to low: M. javanica, M. mayaguensis,

and M arenaria.

In the purple-corolla S. leucantha greenhouse experiment (Table 2-11), there were

differences (P < 0.05) among treatments for root-gall index, number of eggs per plant,

and number of eggs per g of roots. No differences (P 0.05) were observed among

treatments for root weight or shoot dry weight. No root-knot nematode galls were

observed on the control and M incognita treatments, but were observed on the M

javanica, M arenaria, and M mayaguensis treatments. All four root-knot nematodes

reproduced, and the number of eggs per plant and number of eggs per g of roots were not

different (P < 0.05) among species. The data from the two growth room trials and the

purple-corolla S. /eI'/t d/n/t/ greenhouse suggest that this plant is a good host to the four

Meloidogyne spp. evaluated.

In the white-corolla S. ientthIuii/ut greenhouse experiment (Table 2-12), there were

differences (P < 0.05) among treatments for number of eggs per plant and number of eggs

per g of roots. In both categories, M. incognita produced more eggs than M

mayaguensis. Egg production was not observed in M. arenaria and M. javanica-

inoculated plants. No differences (P < 0.05) were observed among treatments for the

root-gall index, root weight, or dry shoot weight categories. The data from this

experiment suggests that the white-corolla S. leucantha is a good host to Meloidogyne

incognita and M. mayaguensis.












Table 2-11. Treatments, root-gall indexes, root weights, number of eggs per plant, number of eggs per g of roots, and dry
shoot weights from the purple-corolla Salvia leucantha greenhouse experiment.


Treatment Root-gall index Root Weight Mean number of eggs Mean number of Dry Shoot
(0-5) (g) per plant eggs per g roots Weight (g)
Control 0.00 + 0.00 ba 27.43 20.13 0.00 + 0.00 b 0.00 + 0.00 b 26.20 2.31
M. incognita race 2 0.00 + 0.00 b 22.63 7.58 162,494.58 112,613.54 a 7,050.67 2,469.93 a 22.97 + 13.46
M.javanica 1.83 1.53 a 23.67 11.01 447,239.47 1,014,030.26 a 16,539.56 32,408.80 a 28.17 3.52
M. arenaria race 1 1.33 + 3.06 a 32.83 20.99 159,567.56 181,554.78 a 4,521.78 3,312.55 a 25.40 + 10.67
M. mayaguensis 0.67 1.15 ab 27.17 7.62 227,586.04 416,191.95 a 9,284.44 19,449.29 a 25.57 8.31
a0 = no galls; 1 = 1 to 2 galls; 2 = 3 to 10 galls; 3 = 11 to 30 galls; 4 = 31 to 100 galls; 5 = more than 100 galls per root system
(Taylor and Sasser, 1978).
bData are means of three replications. Means in columns followed by a common letter are not different (P < 0.05) according to
Duncan's multiple range test. Egg numbers were log-transformed prior to data analysis, but non-transformed numbers are shown.












Table 2-12. Treatments, root-gall indexes, root weights, number of eggs per plant, number of eggs per g of roots, and dry
shoot weights from the white-corolla Salvia leani/tha greenhouse experiment.


Treatment Root-gall index Root Weight Mean number of eggs Mean number of Dry Shoot
(0-5) (g) per plant eggs per g roots Weight (g)
Control 0.00 + 0.00a 39.87 12.23 0.00 + 0.00 cb 0.00 + 0.00 c 27.17 + 5.19
M. incognita race 2 0.00 + 0.00 39.10 1.78 898,208.53 1,159,608.89 a 22,911.11 29,747.46 a 27.57 6.71
M.javanica 0.00 + 0.00 36.10 23.95 0.00 + 0.00 c 0.00 + 0.00 c 26.33 2.83
M. arenaria race 1 0.00 + 0.00 37.07 14.53 0.00 + 0.00 c 0.00 + 0.00 c 29.67 + 5.95
M. mayaguensis 0.33 + 1.15 37.63 19.86 256,939.64 192,705.08 b 7,000.89 + 2,825.53 b 26.97 + 4.92
a0 = no galls; 1 = 1 to 2 galls; 2 = 3 to 10 galls; 3 = 11 to 30 galls; 4 = 31 to 100 galls; 5 = more than 100 galls per root system
(Taylor and Sasser, 1978).
bData are means of three replications. Means in columns followed by a common letter are not different (P < 0.05) according to
Duncan's multiple range test. Egg numbers were log-transformed prior to data analysis, but non-transformed numbers are shown.









Odontonema cuspidatum

Results for the two 0. cuspidatum growth room trials were heterogeneous and are

therefore presented separately in tables 2-13 and 2-14. The results from the greenhouse

experiment are presented in table 2-15. Root galls were not observed on any of the plants

in the three studies. No differences (P < 0.05) were observed among treatments for root-

gall index, root weight, number of eggs per plant, number of eggs per g of roots, or dry

shoot weight in either of the growth room trials or the greenhouse experiment. The data

suggest that 0. cuspidatum is a nonhost to the four Meloidogyne spp. evaluated.

Musa acuminata ssp. zebrina

Results for the two M acuminata ssp. zebrina growth room trials were

heterogeneous and are therefore presented separately in tables 2-16 and 2-17. Root galls

and reproduction were observed for all Meloidogyne species in the three experiments.

Differences (P < 0.05) among treatments for root weight or dry shoot weight were not

detected in the first growth room trial (Table 2-16). The root galls and number of eggs

per plant were not different (P < 0.05) among the root-knot nematode species. However,

M. arenaria produced more eggs per g of roots than the other root-knot nematode

species. In the second growth room trial (Table 2-17), M. mayaguensis produced more

galls than M. javanica, which produced more galls than M arenaria and M. incognita.

Plants inoculated with M. mayaguensis had higher root weights than the other treatments,

and M arenaria and M. mayaguensis produced more eggs per plant and eggs per g of

roots than M. javanica and M. incognita. The dry shoot weight of the control treatment

was greater than that of the root-knot nematode treatments.












Table 2-13. Treatments, root-gall indexes, root weights, number of eggs per plant, number of eggs per g of roots, and dry
shoot weights from the first Odontonema cuspidatum growth room trial.


Treatment Root-gall Root Weight Mean number of eggs Mean number of Dry Shoot
index (0-5) (g) per plant eggs per g roots Weight (g)
Control 0.00 + 0.00a 33.85 + 18.01 0.00 + 0.00 0.00 + 0.00 20.07 4.42
M. incognita race 2 0.00 + 0.00 32.22 9.12 0.00 0.00 0.00 0.00 19.00 1.62
M.javanica 0.00 + 0.00 29.78 21.93 4.44 21.77 0.44 2.13 18.58 + 5.79
M. arenaria race 1 0.00 + 0.00 33.42 + 22.44 0.00 + 0.00 0.00 + 0.00 18.95 5.80
M. mayaguensis 0.00 + 0.00 26.37 16.32 4.44 + 21.77 0.22 1.08 15.53 + 9.36
Data are means of six replications. No differences (P < 0.05) were observed according to Duncan's multiple range test.
a0 = no galls; 1 = 1 to 2 galls; 2 = 3 to 10 galls; 3 = 11 to 30 galls; 4 = 31 to 100 galls; 5 = more than 100 galls per root system
(Taylor and Sasser, 1978).












Table 2-14. Treatments, root-gall indexes, root weights, number of eggs per plant, number of eggs per g of roots, and dry
shoot weights from the second Odontonema cuspidatum growth room trial.


Treatment Root-gall Root Weight Mean number of eggs Mean number of Dry Shoot
index (0-5) (g) per plant eggs per g roots Weight (g)
Control 0.00 0.00a 11.57 3.78 0.00 + 0.00 0.00 + 0.00 16.22 6.90
M. incognita race 2 0.00 + 0.00 12.22 4.16 31.11 + 152.41 2.19 10.73 17.85 4.25
M.javanica 0.00 + 0.00 11.65 3.05 0.00 + 0.00 0.00 + 0.00 14.53 15.36
M. arenaria race 1 0.00 + 0.00 11.82 1.24 0.00 + 0.00 0.00 0.00 18.17 3.50
M. mayaguensis 0.00 + 0.00 10.08 + 2.99 0.00 + 0.00 0.00 + 0.00 18.63 2.27
Data are means of six replications. No differences (P < 0.05) were observed according to Duncan's multiple range test.
a0 = no galls; 1 = 1 to 2 galls; 2 = 3 to 10 galls; 3 = 11 to 30 galls; 4 = 31 to 100 galls; 5 = more than 100 galls per root system
(Taylor and Sasser, 1978).












Table 2-15. Treatments, root-gall indexes, root weights, number of eggs per plant, number of eggs per g roots, and dry shoot
weights from the Odontonema cuspidatum greenhouse experiment.


Treatment Root-gall Root Weight Mean number of eggs Mean number of Dry Shoot
index (0-5) (g) per plant eggs per g roots Weight (g)
Control 0.00 + 0.00a 96.40 + 18.20 0.00 + 0.00 0.00 + 0.00 44.03 5.64
M. incognita race 2 0.00 + 0.00 89.37 + 6.50 8.89 + 30.79 0.89 + 3.08 43.80 + 12.87
M.javanica 0.00 + 0.00 89.53 + 26.26 0.00 + 0.00 0.00 + 0.00 44.77 + 6.63
M. arenaria race 1 0.00 + 0.00 106.26 1.24 0.00 0.00 0.00 + 0.00 49.47 16.30
M. mayaguensis 0.00 + 0.00 87.10 21.38 0.00 0.00 0.00 + 0.00 42.67 6.47
Data are means of three replications. No differences (P < 0.05) were observed according to Duncan's multiple range test.
a0 = no galls; 1 = 1 to 2 galls; 2 = 3 to 10 galls; 3 = 11 to 30 galls; 4 = 31 to 100 galls; 5 = more than 100 galls per root system
(Taylor and Sasser, 1978












Table 2-16. Treatments, root-gall indexes, root weights, number of eggs per plant, number of eggs per g of roots, and dry
shoot weights from the first Musa acuminata ssp. zebrina growth room trial.


Treatment Root-gall index Root Weight Mean number of eggs Mean number of Dry Shoot
(0-5) (g) per plant eggs per g roots Weight (g)
Control 0.00 + 0.00 ba 23.63 5.44 0.00 + 0.00 b 0.00 + 0.00 c 9.52 1.66
M. incognita race 2 4.33 2.07 a 26.55 6.74 189,306.67 114,226.82 a 7,039.58 2,385.42 ab 7.70 + 6.60
M.javanica 5.00 + 0.00 a 26.07 5.01 139,053.33 90,863.31 a 5,292.61 3,214.97 b 9.72 + 1.80
M. arenaria race 1 4.67 + 1.63 a 24.62 4.47 199,737.78 142,003.68 a 8,119.14 5,689.47 a 9.75 2.50
M. mayaguensis 5.00 + 0.00 a 26.20 6.42 190,946.67 124,960.19 a 7,477.72 5,774.92 ab 9.32 0.93
a0 = no galls; 1 = 1 to 2 galls; 2 = 3 to 10 galls; 3 = 11 to 30 galls; 4 = 31 to 100 galls; 5 = more than 100 galls per root system
(Taylor and Sasser, 1978).
bData are means of six replications. Means in columns followed by a common letter are not different (P < 0.05) according to
Duncan's multiple range test. Egg numbers were log-transformed prior to data analysis, but non-transformed numbers are shown.












Table 2-17. Treatments, root-gall indexes, root weights, number of eggs per plant, number of eggs per g of roots, and dry
shoot weights from the second Musa acuminata ssp. zebrina growth room trial.


Treatment Root-gall index Root Weight Mean number of eggs Mean number of eggs Dry Shoot
(0-5) (g) per plant per g roots Weight (g)
Control 0.00 + 0.00 dab 11.87 + 3.29 b 0.00 + 0.00 d 0.00 + 0.00 c 9.25 1.55 a
M. incognita race 2 2.17 1.97 c 13.45 6.83 ab 130,472.22 83,342.51 c 9,684.54 4,138.28 b 7.37 + 1.53 b
M.javanica 3.17 0.82 b 14.47 1.79 ab 170,611.11 92,654.84bc 11,707.26 5,311.64 ab 7.30 1.91 b
M. arenaria race 1 2.00 2.19 c 12.25 4.02 b 180,138.89 95,931.79ab 15,126.25 9,704.36 a 7.45 + 1.66 b
M. mayaguensis 4.67 + 1.03 a 15.83 + 4.34 a 239,500.00 108,061.51 a 15,167.20 6,381.53 a 7.50 1.99 b
a0 = no galls; 1 = 1 to 2 galls; 2 = 3 to 10 galls; 3 = 11 to 30 galls; 4 = 31 to 100 galls; 5 = more than 100 galls per root system
(Taylor and Sasser, 1978).
bData are means of six replications. Means in columns followed by a common letter are not different (P < 0.05) according to
Duncan's multiple range test. Egg numbers were log-transformed prior to data analysis, but non-transformed numbers are shown.









In the greenhouse experiment (Table 2-18), no differences (P < 0.05) among

treatments were observed for root weight and dry shoot weight, and no differences (P <

0.05) among root-knot nematode treatments were observed for number of eggs per plant

or number of eggs per g of roots. However, M. mayaguensis-inoculated plants had a

higher root-gall index than the other root-knot nematode-infected plants. The data

suggest that M acuminata ssp. zebrina cv. Rowe Red is a good host to the four

Meloidogyne spp. evaluated.

Codieaum variegatum cv. Gold Dust

Results for the two C. variegatum cv. Gold Dust growth room trials were

heterogeneous and are presented in tables 2-19 and 2-20, respectively. The results for the

greenhouse experiment are presented in table 2-21. Root galls were not observed on any

of the plants in the three studies. No differences (P < 0.05) were observed among

treatments for root-gall index, root weight, number of eggs per plant, number of eggs per

g of roots, or dry shoot weight in either of the growth room trials or the greenhouse

experiment. The data suggest that C. variegatum cv. Gold Dust is a nonhost to the

Meloidogyne spp. evaluated.

Discussion

The assignment of host statues to perennial ornamentals is ambiguous. Unlike

annual crops, root-knot nematode populations may thrive on perennial hosts for many

months and years. Therefore, studies on the host status of root-knot nematodes on

perennials that result in the classification an immune or a poor host may be disproved by

conducting longer-term studies that allow ample time for pathogen reproduction. The

plants evaluated in this study were homogeneous in their response to the four












Table 2-18. Treatments, root-gall indexes, root weights, number of eggs per plant, number of eggs per g of roots, and dry
shoot weights from the Musa acuminata ssp. zebrina greenhouse experiment.


Treatment Root-gall index Root Weight Mean number of eggs Mean number of eggs Dry Shoot
(0-5) (g) per plant per g roots Weight (g)
Control 0.00 + 0.00 cab 26.83 17.92 0.00 + 0.00 b 0.00 + 0.00 b 24.00 14.76
M. incognita race 2 2.33 + 1.15 b 32.30 44.06 277,331.67 389,211.65 a 8,344.33 9,516.67 a 19.77 13.63
M.javanica 4.00 + 3.64 ab 58.30 + 27.61 454,100.00 433,463.29 a 8,522.22 10,650.06 a 25.60 + 8.63
M. arenaria race 1 3.33 2.31 ab 29.10 3.02 318,673.89 470,556.48 a 10,794.44 15,023.29 a 20.23 + 5.28
M. mayaguensis 4.67 + 1.15 a 39.50 + 36.43 236,578.33 515,546.34 a 5,350.00 10,144.02 a 18.67 + 10.07
a0 = no galls; 1 = 1 to 2 galls; 2 = 3 to 10 galls; 3 = 11 to 30 galls; 4 = 31 to 100 galls; 5 = more than 100 galls per root system
(Taylor and Sasser, 1978).
bData are means of three replications. Means in columns followed by a common letter are not different (P < 0.05) according to
Duncan's multiple range test. Egg numbers were log-transformed prior to data analysis, but non-transformed numbers are shown.












Table 2-19. Treatments, root-gall indexes, root weights, number of eggs per plant, number of eggs per g of roots, and dry
shoot weights from the first Codiaeum variegatum cv. Gold Dust trial.


Treatment Root-gall Root Weight Mean number of eggs Mean number of Dry Shoot
index (0-5) (g) per plant eggs per g roots Weight (g)
Control 0.00 0.00a 11.93 3.44 0.00 + 0.00 0.00 + 0.00 11.58 3.24
M. incognita race 2 0.20 + 0.82 13.32 11.05 37.40 97.86 2.78 7.17 11.46 10.18
M.javanica 0.17 0.82 11.32 3.49 8.83 43.55 0.93 4.54 10.78 4.97
M. arenaria race 1 0.17 0.82 14.52 5.59 4.50 21.77 0.41 2.03 13.40 3.68
M. mayaguensis 0.00 +0.00 12.10 3.33 26.67 130.64 1.92 9.40 11.33 2.99
Data are means of six replications. No differences (P < 0.05) were observed according to Duncan's multiple range test.
a0 = no galls; 1 = 1 to 2 galls; 2 = 3 to 10 galls; 3 = 11 to 30 galls; 4 = 31 to 100 galls; 5 = more than 100 galls per root system
(Taylor and Sasser, 1978).












Table 2-20. Treatments, root-gall indexes, root weights, number of eggs per plant, number of eggs per g of roots, and dry
shoot weights from the second Codiaeum variegatum cv. Gold Dust growth room trial.


Treatment Root-gall Root Weight Mean number of eggs Mean number of Dry Shoot
index (0-5) (g) per plant eggs per g roots Weight (g)
Control 0.00 + 0.00a 9.87 + 2.47 0.00 + 0.00 0.00 + 0.00 9.22 2.36
M. incognita race 2 0.17 0.82 10.25 5.80 0.00 + 0.00 0.00 + 0.00 8.37 3.42
M.javanica 0.33 1.03 8.05 + 3.39 0.00 + 0.00 0.00 + 0.00 7.53 + 3.80
M. arenaria race 1 0.33 1.03 8.37 6.41 0.00 + 0.00 0.00 + 0.00 7.07 3.52
M. mayaguensis 0.00 + 0.00 8.82 + 2.06 0.00 + 0.00 0.00 + 0.00 9.05 5.67
Data are means of six replications. No differences (P < 0.05) were observed according to Duncan's multiple range test.
a0 = no galls; 1 = 1 to 2 galls; 2 = 3 to 10 galls; 3 = 11 to 30 galls; 4 = 31 to 100 galls; 5 = more than 100 galls per root system
(Taylor and Sasser, 1978).












Table 2-21. Treatments, root-gall indexes, root weights, number of eggs per plant, number of eggs per g of roots, and dry
shoot weights from the Codiaeum variegatum cv. Gold Dust greenhouse experiment.


Treatment Root-gall Root Weight Mean number of eggs Mean number of Dry Shoot
index (0-5) (g) per plant eggs per g roots Weight (g)
Control 0.00 0.00a 37.03 8.88 0.00 0.00 0.00 0.00 33.17+ 10.58
M. incognita race 2 0.00 + 0.00 35.80 + 21.20 0.00 + 0.00 0.00 + 0.00 36.93 + 2.90
M.javanica 0.00 + 0.00 44.00 49.36 0.00 + 0.00 0.00 + 0.00 34.27+ 15.01
M. arenaria race 1 0.00 + 0.00 39.47 17.76 0.00 + 0.00 0.00 + 0.00 35.53 + 19.60
M. mayaguensis 0.00 + 0.00 54.80 17.16 0.00 + 0.00 0.00 + 0.00 42.33 + 6.86
Data are means of three replications. No differences (P < 0.05) were observed according to Duncan's multiple range test.
a0 = no galls; 1 = 1 to 2 galls; 2 = 3 to 10 galls; 3 = 11 to 30 galls; 4 = 31 to 100 galls; 5 = more than 100 galls per root system
(Taylor and Sasser, 1978).









Meloidogyne spp. evaluated, with the exception ofL. muscari cv. Evergreen Giant, which

was a good host to M. incognita race 2, M. javanica, and M. mayaguensis, and a poor

host to M. arenaria race 1. Liriope muscari was classified as a nonhost to M arenaria

race 1 in the first and second growth room experiments, yielding Rf values of 0.07 and

0.002, respectively. However, its host status to this pathogen changed to poor (Rf = 0.12)

in the greenhouse experiment. It is unclear whether M arenaria race 1 is a good host

under longer term field or greenhouse studies. Pittosporum tobira cv. Variegata was a

nonhost to all the Meloidogyne spp. evaluated. Liriope muscari cv. Variegata was

previously reported susceptible to M. hapla (LaMondia, 1996). This is the first report of

any Meloidogyne spp. on L. muscari cv. Evergreen Giant.

Pittosporum tobira cv. Variegata was a nonhost to the Meloidogyne spp.

evaluated in this study. This cultivar was found infected with Meloidogyne spp. in Lake

County, FL (chapter 3). Meloidogyne incognita, M. arenaria, and Meloidogyne sp. were

previously reported on P. tobira by Nigh (1972), Bureau of Nematology (1989), and

Goodey et al. (1965), respectively. However, P. tobira cv. Variegata is frequently

infected by Meloidogyne spp., as evident by root galls and egg masses that are observed

on infected plant roots (Levin, R., personal observation). It is possible that P. tobira cv.

Variegata is a nonhost to the Meloidogyne spp. isolates evaluated. It is unlikely that the

reproductive period in these studies was limiting, since a small number of eggs (Rf <

0.01), which probably remained from the initial inocula, was retrieved from plants in the

two growth room experiments, and no eggs were isolated from plants in the greenhouse

experiment. Differences in host response and results that differ from field observations in

this study may be attributed to the Meloidogyne spp. isolates evaluated. Additional work,









involving the isolation, culture, and identification of root-knot nematodes from infected

P. tobira cv. Variegata plants, is required to unveil the host status of this cultivar to

Meloidogyne spp.

The purple-corolla form of S. lemt in/lh and M sumatrana ssp. zebrina cv. Rowe

Red were good hosts to the Meloidogyne spp. evaluated in these experiments. The white-

corolla form of S. leucantha was a good host to the M. incognita and M mayaguensis

isolates evaluated, and a nonhost to the M. javanica and M. arenaria isolates evaluated.

Meloidogyne sp. was previously reported on Salvia leucantha (Goodey and Franklin,

1956, Goodey et al., 1965). Although Meloidogyne arenaria andM incognita have been

reported on numerous Musa acuminata cultivars (Goodey et al., 1965), the host status of

M. acuminata ssp. zebrina cv. Rowe Red to the Meloidogyne spp. evaluated is reported

here for the first time.

Odontonema cuspidatum and Codiaeum variegatum cv. Gold Dust were nonhosts

to the Meloidogyne spp. evaluated in these experiments. Odontonema cuspidatum and C.

variegatum were previously reported as hosts to Radopholus similis and Hoplolaimus sp.,

respectively (Goodey et al., 1965). Since many C. variegatum cultivars are frequently

encountered infected by Meloidogyne spp. (Levin, R., personal observation), the host

status of C. variegatum cv. Gold Dust to the Meloidogyne spp. evaluated and to

additional root-knot nematode species and races needs to be verified and evaluated. In

addition, the usefulness of C. variegatum cv. Gold Dust as a root-knot nematode resistant

rootstock for many Meloidogyne spp.-susceptible C. variegatum cultivars needs to be

investigated.















CHAPTER 3
IDENTIFICATION OF ROOT-KNOT NEMATODES

Introduction

The usefulness of Meloidogyne spp. isozymes, resolved by electrophoresis, as a

tool for the identification of root-knot nematode species, has increased dramatically over

the last 40 years. Most Meloidogyne spp. show species-specific esterase (Est)

phenotypes. Some species, including M. exigua and M naasi, show nonspecific Est

phenotypes. Therefore, the elucidation of a second enzyme phenotype, malate

dehydrogenase (Mdh), is necessary for differentiation of such species. Dickson et al.

(1971) proved that Est and Mdh phenotypes, resolved following disk electrophoresis,

provide a reliable means for speciating Meloidogyne incognita, M. javanica, M. arenaria,

and M hapla. Disk electrophoresis analysis of Meloidogyne females by Dickson et al.

(1970, 1971) and Hussey et al. (1972) utilized several nematodes of the same species per

isozyme phenotype elucidated, rendering genetic analysis at the intra- and interspecific

levels impossible (Dalmasso and Berge, 1978). The use of polyacrylamide gel

electrophoresis (PAGE) by Dalmasso and Berge (1978) provided the means to unveil

single Meloidogyne female isozyme phenotypes following electrophoresis in a 0.7-mm

thick slab gel. Later works (Carneiro et al., 1996, 1998, 2000; Dalmasso and Berge,

1978; Esbenshade and Triantaphyllou, 1985a, 1985b, 1985c, 1987; Fargette, 1987a,

1987b; Fargette and Braaksma, 1990; Pais and Abrantes, 1989; Starr et al, 1996;

Yongfang et al., 1998) utilized innovations of thin-slab gel electrophoresis for the









identification of single Meloidogyne females. Unfortunately, standard methodology for

such techniques is not universal, and published works utilize various gel-forming

apparatuses and methodologies, an array of enzyme stain concoctions, and varying

electrophoresis run times. In addition, many authors fail to accurately describe their

methods and results in detail.

Esbenshade and Triantaphyllou (1985b) presented numerous Mdh and Est

phenotypes and their associated relative electrophoretic migration (Rm) values (distance

of protein in question relative to the migration distance of the bromophenol-blue dye).

Each band was designated a number, representing its migration relative to other bands,

and the numbers were grouped into categories represented by letters. Combinations of

letters and numbers elucidate particular phenotypic patterns. The letter-number system,

referred to in most works that describe Meloidogyne spp. Mdh and Est phenotypes, along

with aM. javanica or M hapla standard on the same gel, is useful for routine

identification of several Meloidogyne species. However, isozyme phenotypes are limited

in their usefulness when compared on different gels. Enzyme phenotypes may vary with

environmental conditions, nematode life stage, and different populations or isolates

(Caswell-Chen et al., 1993). Furthermore, since variability is inevitable between

electrophoretic runs due to human error and unavoidable environmental differences at the

time of gel and enzyme stain preparations, slight variations in isozyme phenotypes may

lead to improper speciation. Therefore, until procedural materials and methods,

evaluation methods, and the accurate reporting of Rm values are standardized,

information including gel formulations and sizes, staining methods, electrophoresis run

times, and precisely-measured Rm must be included in works that elucidate known or









unknown Meloidogyne spp. isozyme phenotypes (Esbenshade and Triantaphyllou, 1985a,

1985b, 1985c; Evans, 1971). Isozyme phenotypes, Rm values, and methodological

information have been described accurately (Dalmasso and Berge, 1978; Esbenshade and

Triantaphyllou, 1985a, 1985b, 1985c; Fargette, 1987a, 1987b; Hussey et al., 1972;

Yongfang et al., 1998), and are presented for M. javanica (Figure 3-1), M. incognita

(Figure 3-2), M. arenaria (Figure 3-3 and 3-4), M. hapla (Figure 3-5), various

Meloidogyne spp. (Figure 3-6), and for several unidentified Meloidogyne spp. isolates

(Figure 3-7).

Yongfang et al. (1998) and Cetintas et al. (2003) elucidated Mdh and Est isozyme

patterns using a PhastSystem apparatus (Pharmacia Biotech AB, Uppsala, Sweden) and a

Mini Protean 3 Cell apparatus, respectively, for routine identification of root-knot

nematodes. Yongfang et al. (1998) determined that the PhastSystem is ideal for the rapid

identification of root-knot nematode species. Furthermore, the authors reported that band

pattern stability withstands differing root-knot nematode host species, host nutrition,

sample origin and cultivation practices, and sample dosage. Although Mdh and Est

patterns relative Rm were identical to previously-reported figures (Esbenshade and

Triantaphyllou, 1985a, 1985b, 1985c), Yongfang et al. (1998) reported on a previously

unobserved Est phenotype, referred to as J2 (Figure 3-1). A similar phenotype was

reported for M. javanica from soybean in Brazil (Castro et al., 2003).

Objectives

The objectives of this study were to

1. Differentiate malate dehydrogenase and esterase phenotypes of Meloidogyne spp. that

were collected from infected ornamental plants in Florida using PAGE.









2. Evaluate the usefulness of PAGE in differentiating Meloidogyne spp. for routine

extension diagnostic purposes.

Materials and Methods

Nematode Populations

A total of 20 root samples of ornamental plants were collected from nurseries,

botanical gardens, and residential plantings from nine counties in Florida. For root-knot

nematode identification, young egg-laying females were dissected from naturally infected

roots of each plant.

Isozyme Analysis

Root-knot nematode females, dissected from different galls of the root systems,

were used for isozyme analyses following electrophoresis, using either PhastSystem or

Mini-Protean 3 Cell. Meloidogyne spp. were identified by comparing specimen Mdh and

Est phenotypes to those of previously published root-knot nematode species (Dalmasso

and Berge, 1978; Esbenshade and Triantaphyllou, 1985c, 1990; Fargette, 1987a, 1987b;

Hussey et al., 1972; Yongfang et al; 1998). The Mdh and Est Rm of knownM incognita,

M. javanica, M. arenaria, M. mayaguensis, and M partityla isolates are presented in

table 3-1 (Dalmasso and Berge, 1978; Esbenshade and Triantaphyllou, 1985c; Fargette,

1987a, 1987b; Hussey et al., 1972; Yongfang et al., 1998). At least 26 females from each

root sample were examined for species identification with the Mini-Protean 3 Cell,

except for the Carya illinoensis sample, from which eight females were examined.

However, either six or ten females from each root sample were examined for species

identification with the PhastSystem. PhastSystem-run samples were processed on the












Rm (x 100)

0 Band Phenotype Designation Key
Enzyme Band Rma (x 100)

MDHb N1 23.0
20- MDH N3 23.0, 27.0, 30.0

SMDH J4 19.0, 24.0, 30.0, 34.0
ESTc J3 46.0, 54.5, 58.9
40- EST J2 47.0, 59.0
EST J3b 30.0, 36.0, 38.0
EST P4 61.0, 81.0, 89.0
60




80



100
MDH N1 N3 N1 J4 N/Ad
EST J3 J3 J2 J3b P4e
Sourcef 1 1 2 3 4
Systemg 1 1 2 3 4
























Figure 3-1. Malate dehydrogenase and esterase relative electrophoretic migrations and enzyme phenotypes ofMeloidogyne
javanica, as reported by several authors.
aRelative electrophoretic migration.
bMalate dehaydrogenase.
CEsterase.
dNo phenotype specified.
eFargette (1987b) reported that esterase phenotype P4 is identical to esterase phenotype J3, with Rm differences attributed to
electrophoresis apparatus variations (Esbenshade and Triantaphyllou, 1985c).
1 = Esbenshade and Triantaphyllou, 1985c; 2 = Yongfang et al., 1998; 3 = Fargette, 1987b.
g1 = Pharmacia GE-2/4 apparatus, separating and stacking gels 7% and 4%, respectively; 2 = PhastSystem, 10-15% gel; 3 =
Pharmacia apparatus, 7% gel; 4 = unspecified.












Rm (x0) Band Phenotype Designation Key

0- Enzyme Band Rma (x 100)

MDHb N1 23.0
MDH N3b 23.0, 28.3, 35.0
20 ESTc I 47.0

SEST Ila 39.0
EST P1 71.0, 76.0
40- EST S1 43.8
SEST P7 65.0, 71.0


60




80



100

MDH N1 N3b N1 N1 N/Ad N/A N/A
EST II I S1 S1 Pie P7e Ila
Sourcef 1 1 1 2 3 3 4
Systemg 1 1 1 1 2 2 3
























Figure 3-2. Malate dehydrogenase and esterase relative electrophoretic migrations and enzyme phenotypes of Meloidogyne
incognita, as reported by several authors.
aRelative electrophoretic migration.
bMalate dehydrogenase.
CEsterase.
dNo phenotypes specified.
eFargette (1987a, 1987b) reported that esterase phenotypes P1 and P7 are identical to esterase phenotypes II and S1,
respectively, with Rm differences attributed to electrophoresis apparatus variations (Esbenshade and Triantaphyllou, 1985c).
fl = Esbenshade and Triantaphyllou, 1985c; 2 = Dalmasso and Berge, 1978; 3 = Fargette, 1987b; 4 = Hussey et al., 1972.
g1 = Pharmacia GE-2/4 apparatus, separating and stacking gels 7% and 4%, respectively; 2 = Pharmacia apparatus, 7% gel; 3
Polyanalyst apparatus, 7% gel.














Band Phenotvoe Designation Key


- -


- -
m m


m
- -
- -


Enzyme
MDHb
MDH
MDH
EST
EST
EST
EST


EST
EST


Band
N1
N3
N3b
Al
A2
P3
S1-MI
S2-M1


Rma (x100)
23.0
23.0, 27.0, 30.0
23.0, 28.3, 35.0
53.3
53.3, 56.3
50.7, 53.3, 56.3
43.8, 47.0
41.3, 43.8, 47.0


M3-F1 47.0, 50.0, 53.3, 56.8


N3 N1 N1
S1-M1 S2-M1 S1-M1
1 1 1
1 1 1
I I I
I I I


Rm
(x100)


MDH
EST
Sourced
System


N3b
A2
1
1
I
I


N1
M3-F1
1
1
I
I
























Figure 3-3. Malate dehydrogenase and esterase relative electrophoretic migrations and enzyme phenotypes of Meloidogyne
arenaria, as reported by Esbenshade and Triantaphyllou (1985c).
aRelative electrophoretic migration.
bMalate dehydrogenase.
CEsterase.
dl = Esbenshade and Triantaphyllou, 1985c.
el = Pharmacia GE-2/4 apparatus, separating and stacking gels 7% and 4%, respectively.













Rm Band Phenotype Designation Key
(x100)
(x 0) Enzyme Band Rma (x100)
0-
MDHb N1 23.0
MDH A5 19.0, 24.0, 30.0, 35.0, 40.0

20 ESTc Al 54.0
SEST A2a 43.0, 46.0
SEST A3a 30.0, 36.0, 38.0
SEST P2 65.0, 71.0
40
40 EST P5 79.0, 85.0

EST P8 59.0, 65.0, 71.0

60



80



100

MDH N1 A5 N/Ad N/A N/A N/A
EST Al A3a A2a P5e P2e P8e
Sourcef 1 2 3 4 4 4
Systemg 1 2 3 4 4 4
























Figure 3-4. Malate dehydrogenase and esterase relative electrophoretic migrations and enzyme phenotypes of Meloidogyne
arenaria, as reported by several authors.
aRelative electrophoretic migration.
bMalate dehydrogenase.
CEsterase.
dNo phenotypes specified.
eFargette (1987a, 1987b) reported that esterase phenotypes P2, P5, and P8 are identical to esterase phenotypes S1-M1, A2, and
S2-M1, respectively, with Rm differences attributed to electrophoresis apparatus variations (Esbenshade and Triantaphyllou, 1985c).
f = Yongfang et al., 1998; 2 = Dalmasso and Berge, 1978; 3 = Hussey et al., 1972; 4 = Fargette, 1987b.
gl = PhastSystem; 2 = Pharmacia GE-2/4 apparatus, separating and stacking gels 7% and 4%, respectively; 3 = Polyanalyst
apparatus, 7% gel; 4 = Pharmacia apparatus, 7% gel.













Rm
(x100)
0 Band Phenotype Designation Key

Enzyme Band Rma (x100)

20 MDHb H1 37.0
MDH Hla 50.0
ESTc H1 50.0
40 EST Hla 33.0
40
EST Al 53.3


60



80



100
MDH H1 H1 Hia N/Ad
EST H1 Al Hla H1
Source 1,2 1 3 1
Systemf 1,2 1 3 1
























Figure 3-5. Malate dehydrogenase and esterase relative electrophoretic migrations and enzyme phenotypes of Meloidogyne


hapla.
aRelative electrophoretic migration.
bMalate dehydrogenase.
CEsterase.
dNo phenotypes specified.
el = Esbenshade and Triantaphyllou, 1985c; 2 = Yongfang et al., 1998; 3 = Delmasso and Berge, 1978.
fl = Pharmacia GE-2/4 apparatus, separating and stacking gels 7% and 4%, respectively; 2 = PhastSystem; 3
2/4 apparatus, separating and stacking gels 7% and 4%, respectively.


Pharmacia GE-













Band PhenotvDe


- -


H1
VS1-Sla
1
1
I
I


Nla
S1
1
1


N/Ad
P7e
2
2


N1
M3a
1
1


Nla
VS1-S1
1
1
I
I


N/A
P6e
2
2


-
m


Nla
VS1
1
1
I
I


N1
S2-M1
1
1


Rm
(x100)

0


Band

H1
N1
Nla
VS1
VS1-Sla
S2-M1
M3a
S1
P6
P7
P8


Designation Key


Rma (x100)

37.0
23.0
30.0
38.0, 43.8
38.0, 45.0
41.3, 43.8, 47.0
47.0, 52.5, 55.7
43.8
51.0, 56.0, 65.0, 71.0
65.0, 71.0
59.0, 65.0, 71.0


N/A
P8e
2
2


Enzyme

SMDHb
MDH
MDH
ESTc
EST
EST
EST
EST
SEST
- EST
- EST


MDH
EST
Sourcef
Systemg


( !
























Figure 3-6. Malate dehydrogenase and esterase relative electrophoretic migrations and enzyme phenotypes of Meloidogyne
spp.
aRelative electrophoretic migration.
bMalate dehydrogenase.
CEsterase.
dNo phenotypes specified.
eFargette (1987b) reported that esterase phenotypes P6, P7, and P8 are identical to esterase phenotypes VS1-S1, S1, and S2-
Ml, respectively, with Rm differences attributed to electrophoresis apparatus variations (Esbenshade and Triantaphyllou, 1985c).
1 = Esbenshade and Triantaphyllou, 1985c; 2 = Fargette, 1987b.
g1 = Pharmacia GE-2/4 apparatus, separating and stacking gels 7% and 4%, respectively; 2 = Polyanalyst apparatus, 7% gel.














Band Phenotype Designation Key
Enzyme Band Rma (x100)
MDHb N1 23.0
MDH H1 37.0
MDH Nla 30.0


- -


- -


MDH
MDH
ESTc
EST
EST
EST
EST
EST
EST
EST


Nib
N3a
Al
Ml
VF1
VFla
VS1
S1
P7
Fl


35.0
23.0, 25.5, 28.3
53.3
47.0, 50.0
65.8
46.0
38.0, 43.8
43.8
75.0, 71.0
56.8


N1 H1 Nib Nla Nla Nla N/Ad N3a
Al M1 VFla VF1 VS1 S1 P7e Fl
1 1 2 1 1 1 3 1
1 1 2 1 1 1 3 1


Rm
(x100)
0


- -
I


MDH
EST
Sourcef
Systemg


B B
fi fi


g
~ ~

~3 X
























Figure 3-6. Continued
aRelative electrophoretic migration.
bMalate dehydrogenase.
CEsterase.
dNo phenotypes specified.
eFargette (1987b) claims that esterase phenotype P7 is identical to esterase phenotype Si, with Rm differences attributed to
electrophoresis apparatus variations (Esbenshade and Triantaphyllou, 1985c).
f = Esbenshade and Triantaphyllou, 1985c; 2 = Delmasso and Berge, 1978; 3 = Fargette, 1987b.
g1 = Pharmacia GE-2/4 apparatus, separating and stacking gels 7% and 4%, respectively; 2 = Pharmacia GE-2/4 apparatus,
separating and stacking gels 7% and 4%, respectively; 3 = Polyanalyst apparatus, 7% gel.















Band Phenotype Designation Key


Enzyme

MDHb
- MDH
- MDH
EST
EST

EST
EST
EST


Band

N1
N3
N3c
Fl
P3
VS1
VS1-S1
VS1-M2


Rma (x100)

23.0
23.0, 27.0, 30.0
23.0, 30.0, 32.5
56.8
86.0
38.0, 43.8
36.0, 43.8
40.0, 53.3, 56.3


N3c
VS1-M2
1
1


Rm
(x100)
0


MDH
EST
Sourcef
Systemg


N/Ad
P3e
2
2


N1
VS1
1
1


N3
VS1
1
1


N3c
VS1-S1
1
1
























Figure 3-7. Malate dehydrogenase and esterase relative electrophoretic migrations and enzyme phenotypes of unidentified
Meloidogyne spp.
aRelative electrophoretic migration.
bMalate dehydrogenase.
CEsterase.
dNo phenotypes specified.
eFargette (1987b) claims that esterase phenotype P3 is identical to esterase phenotype Fl, with Rm differences attributed to
electrophoresis apparatus variations (Esbenshade and Triantaphyllou, 1985c).
1 = Esbenshade and Triantaphyllou, 1985c; 2 = Fargette, 1987b.
g1 = Pharmacia GE-2/4 apparatus, separating and stacking gels 7% and 4%, respectively; 2 = Polyanalyst apparatus, 7% gel.