Nematode (Phylum Nematoda) community assemblages: A tool to implement environmentally-sound management strategies for root-knot nematodes (Meloidogyne spp.) in potato-based cropping systems Emil Engelbrecht Thesis submitted in fulfilment of the requirements for the award of the degree Master of Environmental Science at the North-West University (Potchefstroom Campus) May 2012 Supervisor: H. Fourie Co-supervisor: M. Daneel i ACKNOWLEDGEMENTS There are several people without whom this dissertation and the work it describes would not have been possible. I would like to thank all those people who have contributed towards the successful completion of this work. God our saviour who held his hand over us with every road trip we made. Without His guidance nothing I have done would have been possible. My sincere thanks and appreciation to Prof. Driekie Fourie, my supervisor and mentor during this project. Her patience, guidance, constructive criticism, sound advice and encouragement throughout the course of the study are greatly appreciated. It is Prof. who showed me how interesting and dynamic the field of nematology is. Dr. M. Daneel, co-supervisor, for her contribute and patience with all the questions and assistance. Dr. Suria Ellis, of the statistical consultation service, which provided assistance with statistical analyses. I am also grateful to Suria Bekker for her help with laboratory work. Drs. A. Malan, M, Marias and E. Van den Berg form the ARC-PPRI Pretoria for identification of species. Mr. G. Posthumus at The Western Free State Seed Potato Growers for funding this research and the potato producers at Christiana, Mrs. N. Coetzee, I. Easby, J. Easby, W. Easby, J. Greyling, C. Kriek, W. Ras, JF. Van der Merwe, and oom Lourens for their dedicated help with the sampling. Also to mr. JC. Olevano for calibrating the planting equipment used in the field trial. Mr. J. Fourie from Javeline seeds for supplying the Raphanus seeds that was used in the field trial. Finally, I express my appreciation and love to my parents and Jean-mar้ Truter, for their moral support and motivation during the whole project period, it is much appreciated. ii ABSTRACT Potato is the number one non-grain food commodity and fourth most important food crop worldwide. Plant-parasitic nematodes (PPN), particularly root-knot nematodes (RKN), are one of the economically most important constraints that adversely affect the quantity and quality of this crop. With the removal of many red-band Class 1 nematicides an increased need exists for producers to improve human, environmental and food safety. Accurate identification of PPN and non-parasitic (NPN) assemblages is, therefore, a prerequisite to investigate and verify alternative, environmentally-friendly management strategies to minimise damage to potato and other rotation crops. The objectives of the study were to assess the status of PPN and NPN in 31 potato fields during two consecutive growing seasons in the Christiana area (North-West Province) of South Africa (SA), with the emphasis on RKN. In addition, the effects of selected Brassicaceae species as cover and/green manure crops on RKN population levels as well as NPN assemblages was investigated in a field experiment. For both surveys, root and soil samples were obtained from various crops as well as grass cover crops from 31 fields of five producers. PPN and NPN were extracted and prominence values calculated, while the data were also subjected to parametric and non-parametric statistical analyses. RKN were identified, using the molecularly-based SCAR-PCR method, as the predominant PPN associated with potato, other crops and grasses included in potato-based cropping systems in this area. In terms of NPN, soil food web conditions for the majority of the 31 fields were classified as maturing since they were plotted within the “Stable Enriched” quadrant. In a field trial, conducted during the 2010/2011 cropping season, four Brassicaceae species were evaluated with regard to their host suitability and biofumigation effects in a field where high M. incognita population levels prevailed. Ethylenedibromide (EDBฎ) as well as untreated control treatments were also included in a RCBD with five replicates. Relatively high M. incognita egg and J2 numbers were maintained by all four cultivars, i.e. Doublet and Terranova, Calient้ and Nemat. The follow-up potato crop, planted seven weeks after the incorporation of the Brassicaceae amendments was subsequently parasitized severely in plots planted with all four Brassicaceae cultivars. According to NPN assemblages at termination of the trial, soil food web conditions in plots treated with EDBฎ as well as untreated control plots were classified as disturbed. In contrast, the status of soil food webs in the four Brassicaceae-amended treatments was classified as maturing. Results obtained during this study in terms of using NPN assemblages as a tool to classify soil health is the first in SA, particularly for annual crops such as potato. Furthermore, identification of RKN species in fields of these farmers will assist them in choosing cultivars of rotation crops and/or grass cover crops that has been identified as poor hosts of these parasites. Results emanating from this study are thus applicable in the agricultural sector and could add substantially in terms of our understanding and of both PPN and NPN and their effects in potato-based cropping systems. Key words: biofumigation, Brassicaceae, non-parasitic nematodes, potato, Raphanus, root-knot nematodes iii EKSERP As nie-graan voedselkommoditeit is aartappels w๊reldwyd eerste gelys en verteenwoordig dit die vierde belangrikste voedselgewas. Plantparasitiese aalwurms (PPA), veral knopwortelaalwurms (KWA), is een van die ekonomies belangrikste organismes wat die kwaliteit en opbrengs van aartappels nadelig be๏nvloed oral waar dit in die w๊reld verbou word. Met die onttrekking van al hoe meer Rooiband, Klas 1 aalwurmdoders vanaf w๊reldmarkte het ? toenemende behoefte laat ontstaan by produsente en industrie๋ om veiligheid ten opsigte van mense, die omgewing en voedselgewasse aan te spreek en te verbeter. Dus is akkurate identifisering van PPA asook nie-parasitiese aalwurm (NPA) groepe 'n voorvereiste om alternatiewe, omgewingsvriendelike bestuurstrategie๋ ten opsigte van hierdie organismes te ondersoek en te verifieer. Sodoende kan skade aan aartappels en ander opvolggewasse voorkom en/of tot die minimum beperk word. Die doelwitte van hierdie studie was om die status van PPA en NPA in 31 lande waarin aartappels elke ses tot sewe jaar in wisselbou met ander gewasse en grasdekgewasse geplant word, tydens twee opeenvolgende groeiseisoene in die Christiana gebied (Noord-Wes Provinsie) in Suid- Afrika (SA) te assesseer met die klem op KWA. Die effek van geselekteerde Brassicaceae spesies as dek- en groenbemestinggewasse op KWA bevolkingsvlakke en NPA groepe is ook in ? veldproef ge๋valueer. Aalwurmwortel- en grondmonsters van verskeie gewasse asook die grasdekgewasse wat as wisselbougewasse op die 31 lande verbou is, is geneem en ontleed vir die teenwoordigheid van beide PPN en NPA. Vervolgens is prominensiewaardes vir die teenwoordige aalwurmgenera/spesies/families bereken. Aalwurmdata is egter ook aan parametriese en nie-parametriese statistiese ontledings onderwerp. KWA is met behulp van die molekul๊r-gebaseerde SCAR-PCR metode ge๏dentifiseer as die dominante PPA groep wat teenwoordig was in hierdie lande waarop aartappels asook ander gewasse en grasse ingesluit word in die aartappelproduserende gebied waar hierdie studie uitgevoer was. In terme van NPA, is die status van die grondgesondheid in die meerderheid van die 31 lande geklassifiseer as ontwikkellend en stabiel, omdat laasgenoemde in die "Stabiele Verrykte" kwadrant van die vereenvoudigde voedselweb geklassifiseer was. Gedurende die 2010/2011 groeiseisoen is vier Brassicaceae spesies ge๋valueer met betrekking tot hul gasheerstatus en bio-berokings effek, in ? veldproef met ho๋ M. incognita bevolkingsvlakke. Etileendibromied (EDBฎ) sowel as ? onbehandelde kontrole is ook in die gerandomiseerde blokontwerp ingesluit met vyf herhalings vir elke behandeling. Relatiewe ho๋ M. incognita-eier en tweede jeugstadium (J2) bevolkingsvlakke was in wortels/knolle van al vier die Brassicaceae kultivars, nl. Doublet, Terranova, Calient้ en Nemat teenwoordig tydens die 50% blomperiode van hierdie gewasse. Wortels van die opvolgaartappelgewas (geplant sewe weke na die inwerking van die Brassicaceae kultivars se bogrondse dele) het baie ho๋ M. incognita bevolkingsvlakke getoon tydens knolinisiasie, terwyl eiers en J2 ook reeds in die jong ontwikkelde aartappelknolle teenwoordig was. Ten opsigte van NPA groeperings is gevind dat die status van die grond ten opsigte van gesondeheid in behandelings met EDBฎ sowel as die onbehandelde kontrole as versteur geklassifiseer was. In teenstelling daarmee, is die status van grondvoedselwebbe in die vier Brassicaceae- behandelde persele as ontwikkelend en stabiel geklassifiseer. Resultate wat tydens hierdie studie verkry is in terme van grondvoedselwebbe wat deur die teenwoordigheid van NPA groepe gereflekteer word, kan as 'n waardevolle instrument gebruik word om die gesondheid van gronde te klassifiseer en is ? eerste in SA, veral op eenjarige gewasse soos aartappels. Voorts sal die identifisering van KWA spesies in lande van hierdie produsente in die Christiana gebied hulle alreeds in staat stel om ingeligte keuses te maak ten opsigte van kultivars vir wisselbou- en/of grasdekgewasse wat ge๏dentifiseer is as swak gashere van hierdie parasiete tydens vorige studies. Resultate wat uit hierdie studie voortspruit is dus onmiddellik van praktiese toepassing in die landbousektor en kan aansienlik waarde toevoeg in terme van die begrip en kennis van beide PPA en NPA en die gevolge van hul teenwoordigheid in aartappelgebaseerde wisselboustelsels. Sleutelwoorde: aartappels, bio-beroking, Brassicaceae, knopwortelaalwurms, nie-parasitiese aalwurms, Raphanus iv TABLE OF CONTENTS ACKNOWLEDGEMENTS ...................................................................................................... i ABSTRACT ............................................................................................................................ ii EKSERP ……......................................................................................................................... iii CHAPTER 1 INTRODUCTION AND LITERATURE REVIEW ..................................................................... 1 1.1 Introduction ....................................................................................................................... 1 1.2 Potato (Solanum tuberosum) ............................................................................................ 2 1.2.1 Origin .................................................................................................................. 2 1.2.2 Classification ...................................................................................................... 2 1.2.3 Anatomy ............................................................................................................. 3 1.2.3.1 The potato plant ................................................................................... 3 1.2.3.2 The potato tuber .................................................................................. 3 1.3 Economic and social importance of potato ....................................................................... 4 1.3.1 Potato production in South Africa (SA) ............................................................... 5 1.3.2 Potato production in the Christiana area (North-West Province) ...................... 10 1.4 Plant-parasitic nematodes (PPN) associated with potato ............................................... 11 1.4.1 Root-knot nematodes (RKN; Meloidogyne spp.) .............................................. 12 1.4.1.1 Life cycle of root-knot nematodes (RKN) ........................................... 13 1.4.1.2 Interactions with other organisms ...................................................... 14 1.4.1.3 Damage and symptoms caused by root-knot nematodes (RKN)........ 14 1.4.2 Lesion nematodes (Pratylenchus spp.) ............................................................ 16 1.4.2.1 Life cycle of lesion nematodes ........................................................... 17 1.4.2.2 Interactions with other organisms ...................................................... 17 1.4.2.3 Damage and symptoms caused by lesion nematodes ...................... 17 1.4.3 The golden cyst nematode (GCN; Globodera rostochiensis) ........................... 18 1.4.3.1 Life cycle of the golden cyst nematode (GCN) ................................... 19 1.4.3.2 Interactions with other organisms ...................................................... 20 1.4.3.3 Damage and symptoms caused by the golden cyst nematode (GCN) ……….……………………………………………….. 20 1.4.4 Other plant-parasitic nematodes (PPN) associated with potato in South Africa (SA) ……………………………………………………………......... 21 1.5 Identification of plant-parasitic (PPN) and non-parasitic nematodes (NPN) ................... 22 v 1.6 Management practices to reduce plant-parasitic nematodes (PPN) .............................. 27 1.6.1 Chemical control ............................................................................................... 28 1.6.2 Alternative strategies to manage plant-parasitic nematodes (PPN) ................. 31 1.6.2.1 Host plant resistance .......................................................................... 31 1.6.2.2 Biological control …………………………............................................ 32 1.6.2.3 Quarantine ......................................................................................... 33 1.6.2.4 Crop and fallow rotations ................................................................... 34 1.6.2.5 Soil amendments and cover- and/or green manure crops ................. 37 1.7 Integrated nematode management and future strategies ............................................... 40 1.8 Objectives of this study ................................................................................................... 41 1.9 References ...................................................................................................................... 42 CHAPTER 2 IDENTIFICATION OF NEMATODE ASSEMBLAGES IN POTATO-BASED CROPPING SYSTEMS IN THE CHRISTIANA AREA (NORTH-WEST PROVINCE, SOUTH AFRICA) ……………………………………………………………………………….… 58 2.1 Introduction ...................................................................................................................... 58 2.2 Material and Methods ...................................................................................................... 59 2.2.1 Study area ......................................................................................................... 59 2.2.2 Nematode sampling (surveys) ………………….……………………………….... 67 2.2.3 Nematode extractions …………………………………………………….………... 68 2.2.3.1 Root samples (50g) ……………………………………………………... 68 2.2.3.2 Root samples (5g) ……………..………………………………………… 69 2.2.3.3 Soil samples (200g) ……………………….……………………….……. 70 2.2.4 Counting of nematodes ……………………………………………………………. 72 2.2.5 Species identification by means of morphological identification ………………. 73 2.2.5.1 Transfer of nematodes to anhydrous glycerine ………………………. 73 2.2.6 Rearing of root-knot nematode (RKN) species for molecular identification ….. 74 2.3 Data analysis ……………………………………………………………………………..……. 74 2.3.1 Calculation of prominence values ………………………………………………… 74 2.3.2 Parametric and non-parametric statistics ………………………...……………… 75 2.4 Results …………………………………………………………………………..……………… 77 2.4.1 Prominence values …….…….…………………………………………….………. 77 2.4.1.1 Plant-parasitic nematode (PPN) data …………………………………. 77 2.4.1.2 Non-parasitic nematode (NPN) data ……………………….………….. 86 2.4.2 Parametric and non-parametric statistics ………………………………………... 90 vi 2.4.2.1 Plant-parasitic nematode (PPN) data …………………………………. 90 2.4.2.2 Non-parasitic nematode (NPN) data …………………………………. 103 2.4.3 Faunal analyses for non-parasitic nematodes (NPN) …………………………. 109 2.5 Discussion ……………...…………………………………………………………………….. 115 2.6 Conclusions ………………………………………………………………………….…….…. 120 2.7 References ……………………………………………………………………………..…….. 121 CHAPTER 3 MOLECULAR IDENTIFICATION OF ROOT-KNOT NEMATODE SPECIES USING SEQUENCE CHARACTERIZED AMPLIFIED REGION BASED POLYMERASE CHAIN REACTION ASSAYS ………..…………………………………………………………. 126 3.1 Introduction …………………………………………………………………………………… 126 3.2 Material and methods ……………..………………………………………………………… 128 3.2.1 Deoxy-ribonucleic acid (DNA) extraction …………………………………….…. 128 3.2.2 Sequence characterized amplified region (SCAR) amplification …………..… 129 3.3 Results ………………………………………………………………………………………… 130 3.4 Discussion …………………………………………………………………………...……….. 131 3.5 Conclusions …………………..………………………………………………...……………. 132 3.6 References ………………………………………………………………...…………………. 134 CHAPTER 4 THE EFFECT OF COVER/GREEN MANURE CROPS ON PLANT-PARASITIC AND NON-PARASITIC NEMATODE ASSEMBLAGES IN A POTATO FIELD ……..…… 137 4.1 Introduction …………………………………………………………………………………… 137 4.2 Material and methods ……………………………………………………………………….. 138 4.2.1 Field trial ….………………………………………………………………………… 138 4.2.2 Nematode extractions …………………………………………………….………. 143 4.2.2.1 Root (50g and 5g) and tuber (20g) samples ……….…..……………. 143 4.2.2.2 Soil samples (200g) ……………………………………………….……144 4.2.3 Identification of Meloidogyne species from the Christiana trial site ….....….... 146 4.2.3.1 Morphological identification …………………………………………… 146 4.2.3.2 Molecular identification ………………………………………………… 146 4.2.4 Data analysis ……..………………….……………………………………………. 146 4.3 Results ………………………………………………………………………………………… 147 4.3.1 Identification of root-knot nematode (RKN) species ……………..………..….. 147 vii 4.3.2 Root and tuber data for Brassicaceae and potato crops ………..……………. 149 4.3.2.1 Roots and tubers (50g and 5g): Brassicaceae crops ………………. 149 4.3.2.2 Roots (50g and 5g) tubers (20g): Potato crop ………………………. 150 4.4 Soil data ………………………………………………………………………………………. 151 4.4.1 Plant-parasitic nematodes (PPN) …………………………………….…………. 151 4.4.2 Non-parasitic nematodes (NPN) ………………………………………………… 153 4.5 Discussion ……………………………………………………………………………………. 157 4.6 Conclusions …………………………………………………………………..……….……… 161 4.7 References …………………………………………………………………………………… 162 CHAPTER 5 GENERAL CONCLUSIONS AND FUTURE PROSPECTS ……........................................ 168 5.1 Conclusions ………………………………………………………………………..…………. 168 5.2 Future prospects …………………………………………………...………………………… 171 Appendix A ........................................................................................................................ 172 1 CHAPTER 1 INTRODUCTION AND LITERATURE REVIEW 1.1 Introduction With the progressive increase in the global human population, estimated to reach 9 billion in 2050 (FAO, 2009), it is of vital importance that agricultural and horticultural crops be produced and utilized optimally. Potato is one of the crops that are crucial in terms of food security, taking into account the rapid human population growth and increased hunger rates (IPC, 2011). In 2010, potato was the third largest staple food crop in South Africa (SA) in terms of production (2 071 930 Mt), with a gross value of ZAR 2.4 billion (FAO, 2012). However, seed and table potato crops are exposed to severe damage by various diseases and pests such as plant-parasitic nematodes (PPN) of which root-knot nematodes (RKN; Meloidogyne spp.) are economically the most important (Kleynhans, 1991; Kleynhans et al., 1996). PPN damage to potato is experienced on a wide scale despite the extensive use of synthetic nematicides on which more severe restrictions are being enforced continuously (Nyczepir & Thomas, 2009). This predicament is true for commercial as well as self-sustaining producers throughout the world, including SA where RKN pose the biggest problem to potato. Basic, as well as innovative, research to address the RKN problem in optimising sustainable production of this important staple food crop is quintessential for local producers, the industry and the economy. Since RKN has a very wide host range (Kleynhans et al., 1996; Moens, 2009), sustainable augmentation in population levels of RKN is also detrimental to production of other crops that are planted in rotation with potato. Management of the Western Free State Seed Potato Growers approached nematologists at the North-West University to assist producers in the Christiana potato- producing area in addressing the RKN problem they experience in their potato-based cropping systems. As a result, this study emanated with the introductory chapter focussing on the potato crop, its origin, classification, anatomy as well as economical and social importance. Furthermore, statistics on potato production in SA and in particular in the Christiana area are also referred to. A concise review of economically important PPN, with special reference to RKN, follows while a synopsis on the use of non- parasitic nematodes (NPN) as indicators of soil health is also provided. Methods used to identify both PPN and NPN are concurrently briefly referred to since accurate identification of genera, species and/or families present in crop fields, in this case potato, is crucial to ensure that management strategies are effective. Ultimately such strategies used to protect the crop against PPN are described briefly, with a final glance to the value of integrated management strategies that are, and can be, used in potato- 2 based cropping systems. Therefore, this study entailed i) surveys during two consecutive seasons to enable identification of both PPN and NPN in 31 fields of five potato producers in the Christiana area, and, ii) investigation of the effect of four Brassicaceae spp. cultivars crops on a RKN population (M. incognita) as well as assemblages of NPN occurring in a field of one of these producers. 1.2 Potato (Solanum tuberosum) 1.2.1 Origin Solanum tuberosum or the “Irish potato” is generally accepted to have originated in South America in the Peruvian and Bolivian Andes Mountains (Louw, 1982; Scurrah et al., 2005). It is not certain how this crop was introduced to SA but it has been suggested that potato tubers were brought from Holland to the Cape in the early 1800’s to be planted as food for seafarers (Louw, 1982; DAFF, 2003). 1.2.2 Classification The taxonomic classification of potato is as follows (GBIF, 2011): Kingdom: Plantae Phylum: Magnoliophyta Class: Magnoliopsida Order: Solanales Family: Solanaceae Genus: Solanum Species: S. tuberosum L. 3 1.2.3 Anatomy 1.2.3.1 The potato plant Potato is a herbaceous, annual plant that grows up to 100cm tall and generally produces several tubers below the ground during one growing season (FAO, 2008). Only the tubers of a potato plant are edible, while the leaves, sprouts and stems contain toxic components known as glycoalkaloids. These natural substances protect the plant against fungi and insects. Potato is vegetatively propagated, with a new plant growing from a tuber or piece of tuber which is referred to as the seed. The crop can be grown from sea level up to 4 700 meters above sea level (IPC, 2011) and shares the genus Solanum with at least a 1 000 other species, including tomato (S. lycopersicum) and eggplant (S. melongena). Solanum tuberosum is divided into two subspecies, namely andigena (adapted to short day conditions and mainly grown in the Andes) and tuberosum (cultivated around the world) (Hawkes, 1990; FAO, 2008). 1.2.3.2 The potato tuber As the potato plant grows, its compound leaves manufacture starch that is transferred to the ends of its underground stems (or stolons). These stolons thicken/expand to form a few or as many as 20 tubers underneath the soil surface (FAO, 2008), which represent genetic clones of the mother seed (IPC, 2011). Except for the tubers, potato plants also produce flowers and berries that contain between 100- 400 botanical seeds. These can be planted to produce new tubers, which will be genetically different from the mother plant (IPC, 2011). The number of tubers that actually reach maturity depends on a range of conditions such as available moisture, temperature and soil nutrients. Tubers may vary in shape and size and normally weigh up to 300g each (FAO, 2008). At the end of the growing season, the leaves and stems of potato plants naturally die down or are mechanically sprayed with herbicides to terminate their active growth stage. Newly formed tubers detach from the stolons and then serve as a nutrient store that allows the plant to survive the cold, later regrow and reproduce. Each tuber may contain 2-10 buds (or "eyes"), which are arranged in a spiral pattern around the tuber surface. These buds generate shoots that grow into new plants when conditions are favourable again for plant growth (FAO, 2008). 4 1.3 Economic and social importance of potato Potato forms an integral part of the global food system and is a major food crop in 57 of the more than 100 world countries where it is produced (Scurrah et al., 2005; IPC, 2011). It is the world’s number one non-grain food commodity and ranks fourth as the world’s most important food crop, after maize, wheat and rice. However, unlike the latter crops, potato is not a globally traded commodity and its prices are determined usually by local supply and demand (FAO, 2008). Presently, more than half of the global potato production is provided by developing countries (IPC, 2011). A recent survey by the FAO in more than 70 of the world's developing countries found that increases in potato prices are much lower than that for cereals. Potato is, therefore, highly recommended to support food security and can help low- income countries to alleviate food price increases (FAO, 2008). World potato production reached up to 321 million tonnes in 2010, with 2 million tonnes being harvested from 53 000ha that were planted with potato in SA (PSA, 2010). Potato is regarded as a universal food crop since it produces more food per unit of water applied during its growth stage than any other major crop and is up to seven times more efficient in using water than cereal crops (IPC, 2011). Except for its general use as a vegetable, potato is used for a variety of purposes. Less than 50% of potato grown worldwide is consumed fresh. The rest are processed as food products and ingredients, feed to cattle, pigs and chickens; processed into starch for industry as well as re-used as seed tubers for growing the next season's crop. However, global consumption of potato as a food commodity is shifting from fresh potato to value-added, processed food products. One of the main items in the latter category is frozen potato, which includes most of the french fries served in restaurants and fast-food chains worldwide (IPC, 2011). Since the majority of the world’s undernourished people live in developing countries (FAO, 2010), potato could fill a substantial demand for food and nutrition. Potato is an excellent, low fat source of carbohydrates, containing only one-fourth the calories that bread contains. When boiled, potato contains more protein and nearly twice the calcium than maize (IPC, 2011). Potato is also a good source of the B vitamins, while their skins are an excellent source of vitamin C. Potato also offers advantages as a subsistence crop because of its high yields and favourable response to intensive gardening techniques. Use of potato as a major food source has grown considerably because of the ease with which it can be manipulated genetically, its versatility in agronomic systems and the expanding number of uses for potato as food and a raw industrial material (Kiple & Ornelas, 2000). 5 1.3.1 Potato production in South Africa (SA) More than half of the global potato production currently occurs in Africa, Asia and Europe with SA ranking 28th. In Africa, SA currently ranks fourth in terms of potato production with Malawi being first, Egypt second and Algeria third (PSA, 2010). Although potato production only accounts for 53% of vegetable, 12% of horticulture and 3% of the total agricultural crops being produced in SA (PSA, 2010), the local potato industry has grown and established itself as one of the most important food providers in the country (DAFF, 2003). Local potato production has grown substantially during the past 15 years, namely from 1.2 million tonnes in 1990 to a record of 2 million tonnes in 2010 (PSA, 2010). During the same period the area on which the crop has been cultivated, however, actually declined from 66 000ha to 53 000ha, while the average yield and fresh produce have increased steadily (Figure 1.1) (PSA, 2010). Reasons for this could be due to the decrease in potato production under rain-fed conditions together with the increase in production under irrigation (Figure 1.2). The use of higher-yielding cultivars, for example Mondial, and better production practices also contributed to this scenario (F. Niederwieser, pers comm., April 2012). Locally potato is generally grown under irrigation in 16 regions (Figure 1.3). The top three potato production regions are situated in the Limpopo (18%), Eastern Free Sate (17%) and Western Cape (Sandveld region; 14%) provinces. There are an estimated 654 commercial producers that currently produce potato in SA (PSA, 2010). Yields of approximately 33 tonnes per ha (PSA, 2010) are generally realised and in some regions two potato crops can be produced per year (DAFF, 2011). Since potato is planted at different times of the year due to climatic differences in the various production areas, fresh potato is available for consumption throughout the year (DAFF, 2011). The most common and popular potato cultivar planted locally is Mondial, followed by Buffelspoort 1 (BP1) and Up-To-Date (UTD) (Figure 1.4) (PSA, 2010). Selection of these potato cultivars, however, depends on the specific production area and purpose (DAFF, 2011). In 2010, 68% of the total potato crop was utilized by local markets, while 17% were processed, 8% used as seed and 7% exported (PSA, 2010). In general SA boasts a sophisticated seed potato industry and an effective and growing processing sector (FAO, 2008). Currently the industry uses 380 000 tonnes of fresh potato, which is processed into french fries, frozen and chilled products and crisps (PSA, 2010). Annual local potato consumption is estimated at ฑ30kg per person (FAO, 2008). 6 Although the potato industry is strong, a wide range of pests and diseases as well as other constraints affect local production. Damage inflicted to potato planted locally for example accounts to 10% of tubers being degraded at fresh produce markets. Other factors that adversely affect the crop, resulting in the degrading of tubers are greening, mechanical damage, browning, PPN and others as shown in Figure 1.5 (PSA, 2010). Figure 1.1. Hectares and crop size for potato production in South Africa from 1990-2010 (PSA, 2010). 7 Figure 1.2. Hectares planted to potato in South Africa from 1987 – 2010, indicating rain-fed versus irrigation areas (PSA, 2010). 8 Figure 1.3. The areas where the sixteen potato production regions in South Africa are situated (Adapted from: https://research.cip.cgiar.org/confluence/display/GILBWEB/South+Africa by Thinus du Plessis, Germstyle, Namibia). N 0 100 200 Z 9 Figure 1.4. Potato cultivars planted in South Africa (PSA, 2010). Figure 1.5. Factors related to the degrading of potato tubers in South Africa (PSA, 2010). 10 1.3.2 Potato production in the Christiana area (North-West Province) Approximately 61 potato producers are actively growing potato on approximately 6 109ha of arable agricultural land that is situated in the region where this study was conducted on the border of the North-West and Free State provinces. Primary cultivars planted for table potato production on a total area of 2 226ha in this area are Mondial (66%), Up-To-Date (UTD) (22%) and BP1 (7%). For seed potato purposes, which is the main objective for producers in this region, cultivars Mondial (80%), BP1 (8%) and Fabula (6%) are planted on a total area of 3 883ha (PSA, 2010). In this area the average inspections during which tuber infection were recorded, as a result of RKN parasitism, has increased from an average of 4%, with symptoms visible during the 2001/2002 season to nearly 50% during the 2009/2010 season (Table 1.1) (J. van Vuuren, pers comm., May 2010). Table 1.1. The extent of root-knot nematode damage as recorded during tuber inspections (J. van Vuuren, pers comm., May 2010). Growing seasons Average RKN infection (%) for inspections done Number of inspections done on tubers Number of tuber inspections showing tuber infection Average inspections showing tuber infection (%) 2001/2002 0.018 736 30 4.08 2002/2003 0.043 372 45 12.1 2003/2004 0.065 407 101 24.82 2004/2005 0.019 371 42 11.32 2005/2006 0.049 513 110 21.44 2006/2007 0.018 489 68 13.91 2007/2008 0.019 458 79 17.25 2008/2009 0.02 528 100 18.94 2009/2010 0.093 697 347 49.78 11 1.4 Plant-parasitic nematodes (PPN) associated with potato Nematodes are pseudocoelomate, unsegmented worm-like animals that occur in almost every habitat and are either non-parasitic or parasitic. The latter parasitic nematodes infect and parasitize plants, animals and humans (Decraemer & Hunt, 2006). PPN are amongst the most important pest constraints that adversely influence the production of potato worldwide (Scurrah et al., 2005). Although occupying many different ecological niches, nematodes are small, essentially aquatic animals (Kleynhans et al., 1996; Hunt et al., 2005) that require at least a film of water to enable locomotion. Therefore, water content is a primary ecological factor that is important for them to locate and infect plants. Although many PPN species die in dry soils, others may survive in an anhydrobiotic state. Conversely, too much soil water may result in a lethal oxygen deficit for these organisms (Hunt et al., 2005). In turn, nematodes are parasitized or preyed upon by viruses, bacteria, fungi, insects, mites and other nematodes (Kleynhans et al., 1996). The life cycle of PPN generally comprises of an egg, four juvenile (three in some longidorids) and an adult stage(s) (Kleynhans et al., 1996; Decraemer & Hunt, 2006). Each juvenile moults once and the adult stage appears after the last moult. The second moult, which in Tylenchida occurs within the eggshell gives rise to the second-stage juvenile (J2), which represents the hatching and often infective stage (Kleynhans et al., 1996). All PPN species spend at least part of their life cycle in the soil. Their activities are determined by abiotic factors such as soil aeration, temperature, moisture, organic material and other soil properties as well as biotic factors that include availability and suitability of host plants, soil cultivation practices and the presence of pathogens (Kleynhans et al., 1996; Bridge & Starr, 2007). Most PPN individuals occur in the upper, cultivated soil layer within the root zones of their host plants (Kleynhans et al., 1996). Most crops are particularly vulnerable to attack by PPN during the seedling stage when the young root system is becoming established (Keetch & Milne, 1982). Feeding of PPN on plant tissue, their interaction with fungi and bacteria and/or their transmission of certain PPN-borne virus diseases can also significantly retard the growth, reduce the yield and/or adversely affect the quality of most crops (Keetch & Milne, 1982). Infection of crops by PPN is often associated with early senescence and proliferation of lateral roots (Scurrah et al., 2005) and this usually results in poor crop performance and sometimes crop failure. In fields these symptoms tend to appear in defined patches which often show abundant weed growth and enlarge with time (Kleynhans et al., 1996). Globally the potato cyst nematodes (PCN), Globodera pallida and G. rostochiensis (known as the Golden Cyst Nematode; 12 GCN) are generally regarded as the economically most important nematode parasites of potato. The latter PPN are followed by species of Meloidogyne, Ditylenchus, Pratylenchus, Nacobbus and Trichodorus (Winslow & Willis, 1972; Scurrah et al., 2005; Jones et al., 2011). In SA, 39 PPN species that belongs to 19 genera have been identified to infect potato (Keetch & Buckley, 1984; Kleynhans et al., 1996), with the most important nematode parasites of this crop being Meloidogyne spp. followed by Pratylenchus spp. (Kleynhans, 1978; Jones, et al., 2011). The golden cyst nematode, G. rostochiensis, has only been recorded in certain potato producing areas of SA such as Pretoria (Gauteng Province), surrounding Gauteng areas as well as in the Ceres and Sandveld areas (Western Cape Province) (Louw, 1982; Jones, et al., 2011) where they cause significant damage to the crop. Strict quarantine legislation, however, seems to limit the spread of this nematode species in local production areas. Unfortunately, the conditions that favour successful potato production are also favourable for multiplication and survival of PPN (Jones, 1970). For example, hatch and movement to roots occur most rapidly in sandy soils and this contributes to crops grown on such soils, like potato, being those most likely to suffer the heaviest damage by PPN (Trudgill et al., 1998). Key factors that influence the extent of PPN damage to potato tubers are the initial abundance of these parasites as well as the period that the crop is in the soil. If a high RKN population, for example, is present at planting or if infected tubers are planted, potato roots will be invaded by J2 of the particular PPN species present (Jones, et al., 2011). 1.4.1 Root-knot nematodes (RKN; Meloidogyne spp.) RKN are an economically important polyphagous group of highly adapted obligate endoparasitic organisms that attack potato roots and tubers in local plantings (Jones et al., 2011). These parasites are distributed worldwide and infect a wide range of plant species that are cultivated (Castagnone- Sereno, 2006; Moens et al., 2009). Meloidogyne species are present in most agricultural soils in southern Africa but especially prefer and cause damage to crops planted in sandy soils (Keetch & Buckley, 1984; Louw, 1982; Kleynhans et al., 1996). Six RKN species were reported to be associated with potato in SA (Kleynhans et al., 1996; Jones et al., 2011) with M. incognita and M. javanica being the most widespread and damaging and constituting 27% and 41%, respectively, of populations isolated from potato plantings (Coetzee, 1968). 13 Meloidogyne acronea, M. arenaria, M. chitwoodi and M. hapla have also been recorded to infect potato in SA, but are rare in commercial plantings and little is known about their distribution (Kleynhans et al., 1996; Jones et al., 2011). Collectively, Meloidogyne species are more damaging to crops such as potato than most other groups of PPN since individuals of this genus are: i) widely distributed throughout agricultural soils worldwide ii) complete several life cycles per growing season, resulting in high fecundity rates and iii) generally have very wide host ranges (Nyczepir & Thomas, 2009). Should these parasites not be managed effectively they can reach population densities that adversely affect vigour, yield (Nyczepir & Thomas, 2009) and quality of crops (Moens et al., 2009), such as potato. Due to the combination of factors outlined above and set against the potential financial losses that can occur as a result of RKN infection, growers invest heavily in control strategies to limit the impact of RKN. Currently the input cost for local potato production can amount to R100 000/ha, which usually includes a substantial amount for applying synthetic nematicides (CropLife, 2011; Jones et al., 2011). 1.4.1.1 Life cycle of root-knot nematodes (RKN) RKN typically feed and reproduce on modified, living plant cells within plant tissue such as roots and tubers in the case of potato where they induce specialized feeding cells that are referred to as giant cells (Scurrah et al., 2005). Mature females produce eggs in gelatinous masses composed of a glycoprotein matrix, which is excreted by rectal glands in the anal area. The gelatinous substance keeps the eggs together and protects them against environmental extremes and predation by other soil organisms (Moens et al., 2009). Depending upon environmental conditions, some J2 may enter diapause and remain in the egg during unfavourable conditions such as during dry seasons or during winter periods when low temperatures are experienced (De Guiran & Ritter, 1979; Hunt et al., 2005). In the latter cases, infective J2 may not hatch from eggs for several years after being laid (Hunt et al., 2005). Egg masses are usually deposited on the surface of galled potato roots and about 1cm below the surface of developing tubers, but they may also be embedded within the gall tissue. The egg mass initially has a soft, sticky and hyaline structure but becomes firmer and dark brown with age (Moens et al., 2009). Hatching J2 are mobile and can move relatively long distances of between 40-100cm, both horizontally and vertically within the soil profile when soil moisture levels are optimum. Mobility allows the J2 to find a suitable host (Eisenback & Hunt, 2009), where it will penetrate just behind the root tip or any place on 14 a tuber in the case of potato plants. After penetration, the invasive J2 starts to feed on plant cells that are usually located behind the root cap in the differentiated vascular tissue. It then migrates intercellularly through the cortex to the region of cell differentiation where the female becomes sessile and develops a permanent feeding site from which she withdraws nutrients (Karssen & Moens, 2006; Moens et al., 2009). In parthenogenetic RKN species, sex determinism depends strongly on environmental factors. Factors such as high population densities of the particular RKN species, nutritional deficiencies that occur in the host plant, presence of other plant pathogens, the level of resistance exhibited by the host plant as well as the concomitant limitation of food supply influence this phenomenon (Bird, 1971; Triantaphyllou, 1973; Castagnone-Sereno, 2006; Moens et al., 2009). When conditions are favourable, juveniles develop into females and into males when adverse conditions (as described above) prevail (Castagnone-Sereno, 2006; Moens et al., 2009). However, if such stress is imposed during nematode development, J2 that are developing as females can undergo sex reversal, producing intersexes or males (Triantaphyllou, 1973; Papadopoulou & Triantaphyllou, 1982). 1.4.1.2 Interactions with other organisms Secondary infection by other pathogens, i.e. fungi and/or bacteria often results in extensive decay of RKN-infected plant tissue (Moens et al., 2009). On potato, associations of RKN have been confirmed for bacterial wilt, Pseudomonas solanacearum and Erwinia spp. and fungi such as Verticillium spp., Fusarium spp. and Rhizoctonia solani (Brodie et al., 1993; Manzanilla-Lop้z et al., 2004). Resistance of potato to bacterial wilt is known to break down when the plant is simultaneously infected with M. incognita (Jatala et al., 1975; Jatala & Martin, 1977). 1.4.1.3 Damage and symptoms caused by root-knot nematodes (RKN) By disrupting the physiology of the host plant, RKN may not only reduce crop yield (quantitative damage), but in the case of potato also the product quality (qualitative damage) and are therefore of great economic and social importance (Karssen & Moens, 2006; Moens et al., 2009). RKN damage to potato is usually associated with light (high sand content) soils or peats (Winslow & Willis, 1972; Kleynhans et al., 1996). Loamy and sandy soils, usually rich in organic matter with good drainage and aeration, are most suitable for the cultivation of potato (Winslow & Willis, 1972; DAFF, 2011). Soil structure is also an important factor since pore size affects the ease with which nematodes can move 15 through the soil interstices. In general, sandy soils provide for optimal movement of RKN, while soils with high clay content or those with an excessively open texture inhibit movement of these parasites (Kleynhans et al., 1996; Hunt et al., 2005). An increase in the severity of RKN infections in local potato plantings as well as the resultant increased dependence on the use of chemical control by local producers, is due to favourable temperatures, soil structure and moisture status that prevail in potato- producing areas (Jones et al., 2011). Potato crops infected by RKN are, however, unlikely to exhibit above-ground symptoms (Scurrah et al., 2005). Typical galls or knots are usually only visible on the roots and/or tubers in fields where high RKN population levels occur (Martin, 1972; Jones et al., 2011). RKN-infected potato plants may, however, exhibit stunting, yellowing, early senescence and tend to wilt under moisture stress (Manzanilla-Lop้z et al., 2004; Scurrah et al., 2005). Below-ground symptoms could also result in the root system being reduced or abnormal growth of the roots, excessive branching of secondary roots and overall root galling may occur and be visible (Manzanilla-Lop้z et al., 2004; Scurrah et al., 2005). Galls are initiated and produced in response to growth regulators, proteins and glycoproteins introduced into the host from subventral oesophageal glands of the feeding J2 which causes hyperplasia and hypertrophy of cortical tissues surrounding the nematode and the giant cells (Jepson, 1987; Bridge & Starr, 2007). While both potato roots and tubers are generally infected by RKN J2, the first generation occurs mainly in the root system while J2 from the second generation usually infect the tubers (Santos, 2001). RKN- infected roots generally have galls or knots of various sizes and shapes depending on the nematode density and the species present (Manzanilla-Lop้z et al., 2004; Scurrah et al., 2005; Jones et al., 2011). It is, however, not recommended to identify RKN species using the size and shapes of galls (Eisenback et al., 1981; Manzanilla-Lop้z et al., 2004) that are present on potato roots/tubers. All RKN species that infect potato produce necrotic spots in the region between the tuber surface and the vascular ring. This is the result of tuber tissue reacting to the deposition of eggs and the gelatinous matrix (Scurrah et al., 2005). When a RKN infection is severe, tubers may rot in the soil before it is harvested (Martin, 1972) or even be deformed (Jones et al., 2011). The depth of J2 penetration into tubers varies but, depending on the tuber size, nematode females are usually found 1-2mm below the skin where it feeds on vascular tissue (Jatala, 1975; Jones et al., 2011). Swellings and lesion-like symptoms on the tuber surface as a result of RKN infection (Martin, 1972; Manzanilla-Lop้z et al., 2004) are more numerous in the “eye” part of the tubers (Figure 1.6), with lightly infected lesions more likely to be found in the largest tubers (Martin, 1972). At lifting, the lesions on the tubers itself may have a watery appearance and protrude above the surface of the tuber (Martin, 1972; Manzanilla-Lop้z et 16 al., 2004). It may, however, collapse later and resembles rough and crinkley scar tissue. Not all lesions are watery and blister-like in appearance and in many cases the presence of RKN results in small swellings only, which are often better detected by touch than by sight (Martin, 1972). Figure 1.6. Root-knot nematode infected potato tubers, with typical knot- like protuberances visible on the surface of the tubers (Suria Bekker, Unit of Environmental Sciences and Development, NWU, Potchefstroom Campus). 1.4.2 Lesion nematodes (Pratylenchus spp.) Pratylenchus species are distributed worldwide and are responsible for substantial yield loss in many agricultural and horticulture crops (Duncan & Moens, 2006). Individuals of this species were first found to infect potato in SA during 1953 in the Highveld region (Mpumalanga Province) (Koen, 1960). Seven lesion nematode spp. (Kleynhans et al., 1996) have been associated with potato in SA of which P. brachyurus is the most damaging (Koen, 1960; Jones et al., 2011). According to Van den Berg (1971), Koen (1960) reported P. brachyurus to be present in almost all potato-producing areas of SA. Martin (1972) also concluded that it is not uncommon to find lesion and RKN infecting the same tuber. Lesion nematodes can survive in the soil in the absence of host plants for a long time (Koen, 1960; Hunt et al., 2005). When the infected seed potato tuber decomposes in the soil, lesion nematodes are released and can subsequently parasitize a new host (Koen, 1965; Jones et al., 2011). Lesion nematodes can, therefore, be spread by the planting of infected tubers in soils not previously infested 17 with these parasites. Potato are usually not cultivated on the same field for two years in succession and these nematodes can thus parasitize other crops such as maize, wheat, oats and various grasses (i.e. teff) that are rotated with potato and in this way assist in maintaining populations of these nematodes in the soil until the next potato crop is planted (Koen, 1965). A wide range of weeds also serves as hosts for lesion nematodes in which they can be maintained in potato fields (Koen, 1965; Ntidi et al., 2012). 1.4.2.1 Life cycle of lesion nematodes Pratylenchus species are polycyclic, polyphagous, migratory root endoparasites (Mc Donald & Nicol, 2005). The life cycle of lesion nematodes generally resembles that of RKN in that it has four moults before the mature male or female stage are reached (Van den Berg, 1982). However, unlike RKN and PCN, lesion nematodes do not induce permanent feeding sites in their host plants. Instead they feed and reproduce while migrating between or through plant cells (Duncan & Moens, 2006). All developmental stages of Pratylenchus species are, however, vermiform, including the mature female (Manzanilla-Lop้z et al., 2004; Decraemer & Hunt, 2006), and are capable of invading roots and/or tubers of host plants such as potato (Van den Berg, 1982; Bridge & Starr, 2007). 1.4.2.2 Interactions with other organisms Potato roots/tubers infected with Pratylenchus species are, like RKN-infected roots/tubers, more susceptible to secondary invasion by other pathogenic soilborne organisms (Koen, 1960). Since lesion nematodes are migratory endoparasites, they are more prone to interact with other root pathogens and this way enhances the resultant root disease condition (Jones et al., 2011). These nematodes interact with a variety of other soilborne pathogens, most notably Fusarium and Phytophthora spp. They are also associated with the fungus Verticillium that can cause Verticillium-wilt in potato (Jones et al., 2011). Although no such interactions have been recorded locally for lesion nematode interaction with potato, it is likely that these parasites do still influence the incidence of various soil borne pathogens that occur in potato fields (Jones et al., 2011). 1.4.2.3 Damage and symptoms caused by lesion nematodes Pratylenchus species typically cause conspicuous, dark brown or black necrotic areas on the root or tuber surface of their host plants (Figure 1.7 A&B) (Van den Berg, 1982; Mc Donald & Nicol, 2005). As they migrate through the plant tissue they cause mechanical destruction of cells (Brodie et al., 1993). 18 Symptoms are also usually associated with lesions on the roots and/or tubers that can turn from dark brown to red (Jones et al., 2011). Large numbers of lesion nematodes can, however, damage the roots/tubers to such an extent that the growth of the host plant is adversely affected (Koen, 1967; Manzanilla-Lop้z et al., 2004). When the potato plant dies or the above-ground parts are removed, the roots decay and large numbers of lesion nematodes migrate into the soil to attack the tubers (Koen, 1967). Koen and Hogewind (1967) found that lesion nematode-infected tubers are usually firm at lifting but when stored soon wither, loose mass and become hard. Symptoms on tubers can also represent purple-brown lesions that usually occur on the bottom of the tuber at the end of the growing season (Jones et al., 2011). The migration of lesion nematodes from the dead and dying roots is mainly responsible for the increased population levels in the soil, as only a few nematodes generally leave the tubers once they invaded and infected it. Another result of the dying-off or removal of the aerial parts of the plant is that soil temperatures may rise by as much as 6ฐC, which may stimulate the activity and development of PPN, such as lesion and RKN nematodes (Koen, 1967). Figure 1.7. (A&B). Brown to black necrotic lesions on potato roots (A) and tubers (B) as a result of root- lesion nematode damage (A: http://jnkvv.nic.in/IPM%20Project/nematode.html) (B: H. Kawagoe & H. Nakasono, APS; http://keys.lucidcentral.org/keys/sweetpotato/key/Sweetpotato%20Diagnotes/Media/ Html /TheProblems/Nematodes/LesionNematode/Lesion%20nematode.htm). 1.4.3 The golden cyst nematode (GCN; Globodera rostochiensis) Cyst-forming or cyst nematodes, belonging to the family Heteroderidae, are one of the most specialized and successful PPN pests that infect and parasitize agricultural and horticultural crops (Turner & Evans, 1998). The GCN is believed to have evolved along with their principle host, potato, in the highlands of Peru and Bolivia (Scurrah et al., 2005). The GCN was reported for the first time in SA in A B 19 1971 where it infected potato roots on an irrigated farm North of Pretoria and subsequently on small holdings around Johannesburg and Bon Accord (Gauteng Province) (Knoetze et al., 2004). In 1999, it was again recorded in the Hankey (Eastern Cape Province), Randfontein (Gauteng Province), Mitchells Plain, Ceres and the Sandveld areas (Western Cape Province). The biological race of G. rostochiensis present in SA appears to be pathotype A (Jones et al., 2011). The GCN, like other cyst nematodes, usually have a very narrow host range and are able to infect potato and some other Solanaceae crops such as tomato, nightshade (S. nigrum), bittersweet (S. dulcamara) and tobacco (Nicotiana tabacum) (Turner & Evans, 1998; Subbotin et al., 2010). In heavily GCN-infested fields where no control measures are applied, yields can be less than the mass of the tubers planted (Mai, 1977; Jones et al., 2011). Except for directly suppressing yields, GCN can indirectly cause economic losses by interacting with other micro-organisms and this way cause even higher yield losses than those caused by the nematodes alone (Mai, 1977; Turner & Evans, 1998). The GCN are regarded as one of the most important pests of potato since they cause substantial reductions in yields of susceptible potato cultivars planted in heavily infested fields (Trudgill et al., 1998). The lack of inexpensive nematicidal treatments that result in an adequate level of control under such GCN-infested field conditions also contributes to the latter scenario. The relative ease with which GCN can be spread from one area/field to another, their persistence as viable cysts in soil for up to 30 years and their high reproductive capacity in the presence of a growing potato plant contributes to its importance as a major pest of potato worldwide (Winslow & Willis, 1972; Turner & Evans, 1998). 1.4.3.1 Life cycle of the golden cyst nematode (GCN) The life cycle of GCN, are well adapted towards the host plants they infect and they can survive in various environments (Turner & Evans, 1998; Nicol et al., 2011). Eggs inside cysts can remain viable in soil for long periods of time (Scurrah et al., 2005) i.e. it can survive for more than 20 years (Oostenbrink, 1966; Subbotin et al., 2010) in soils under severe temperature (-15ฐC) stress or in desiccated soils (Scurrah et al., 2005). The eggs contain the infective J2 and are stimulated to hatch as a result of potato root exudates (Scurrah et al., 2005; Nicol et al., 2011). Hatching of up to 80% of the eggs can be attained under favourable environmental conditions. J2 enters the root near the root tip, migrates intracellulary towards the vascular cylinder and induce a feeding cell or syncytial “transfer cell” (Manzanilla-Lop้z et al., 2004; Nicol et al., 2011). The J3 and J4 life stages subsequently develop, become sedentary and feed from the syncytia. The 4th moult gives rise to the mature, round and 20 swollen female, which protrudes from the root/tuber of potato. The males are slender and vermiform, leave the roots, mate and fertilize the females (Nicol et al., 2011). After mating, eggs develop within the female. When the female dies the cuticle forms a protective cyst, encapsulating between 200–500 eggs. The life cycle is then completed and may take up to three months to repeat. The small yellow or brown coloured cysts (pinhead size) falls off the roots/tubers, with J2 hatching from the eggs it contains as soon as root exudates of a suitable host plant are excreted (Nicol et al., 2011). GCN usually completes one generation during a potato growing season (Morris, 1971). 1.4.3.2 Interactions with other organisms GCN not only cause wounds in roots/tubers, but also provide entry sites for other micro-organisms as is the case with RKN and lesion nematodes. This is of particular importance to a wide range of fungi and bacteria that are important pathogens of potato (Turner & Evans, 1998; Scurrah et al., 2005). A number of PCN species have been found to interact with Fusarium wilt species causing wilt disease, for example G. tabacum on tobacco. Another interaction is that of G. pallida and the bacterium Ralstonia solanacearum on potato in which the nematode enhances damage caused by the associated wilt. The economic effect of these interactions varies but their effect can be important with high value crops such as potato. Therefore, the need for research aimed at host-parasite relationships is crucial in order for effective management strategies to be developed (Turner & Rowe, 2006). 1.4.3.3 Damage and symptoms caused by the golden cyst nematode (GCN) In potato plantings, GCN generally does not cause any specific above-ground symptoms nor any typical symptoms on the roots and/tubers (Kleynhans, 1978; Scurrah et al., 2005). However, at flowering of the crop round, yellow or cream cysts are clearly visible on roots (Figure 1.8) when the plant is carefully removed from the soil (Brown, 1969; Kleynhans, 1978; Scurrah et al., 2005). Root injury by GCN, causes stress and reduces the uptake of water and nutrients which in turn cause stunting, yellowing and other discolouration as well as wilting of the foliage (Trudgill et al., 1998; Scurrah et al., 2005). Growth of infected potato plants is retarded and fields heavily infested with GCN usually result in patches of plants showing symptoms such as yellowing of leaves and/or retarded plant growth, especially when low rainfall is experienced (Nicol et al., 2011). Root systems can also be reduced and become abnormally branched and brownish in colour. Where low GCN densities occur, tuber sizes are reduced whereas at higher densities both the number and size of tubers can be reduced 21 (Subbotin et al., 2010). Apart from yield reduction of the potato crop, the financial benefits are reduced by the costs of control measures and subsequent reduction of marketable produce (Nicol et al., 2011). Figure 1.8. Golden cyst nematode females visible on roots of potato (Xiaohong Wang, US Department of Agriculture; http://www.sciencephoto.com/media/139385/enlarge). 1.4.4 Other plant-parasitic nematodes (PPN) associated with potato in South Africa (SA) Though PPN other than Meloidogyne spp., Pratylenchus spp. and G. rostochiensis have been recorded to parasitize potato in SA (Keetch & Buckley, 1984; Kleynhans et al., 1996; Jones et al., 2011), they have not been associated with crop and/or quality losses. These include various species of Anguina, Aphelenchus, Aphelenchoides, Ceocenamus, Criconema, Helicotylenchus, Mesocriconema, Nanidorus, Paratrichodorus, Rotylenchus, Rotylenchulus, Tylenchus, Tylenchorhynchus, Xiphinema, as well as Ditylenchus africanus (peanut pod nematode) and Radopholus similis (Keetch & Buckley, 1984; Kleynhans et al., 1996). Internationally, however, the root tip feeders belonging to the Paratrichodorus, Trichodorus and Nanidorus genera are considered as important pests of potato (Scurrah et al., 2005; Bridge & Starr, 2007). They have the potential to cause severe damage to the root system of young developing potato plants, while they also have the ability to transmit viral diseases (Scurrah et al., 2005; Bridge & Starr, 2007) like the tobacco rattle virus that cause tuber spraing or corkey ringspot in potato (Louw, 1982; Manzanilla-L๓pez et al., 2004). This virus together with its vector N. minor does occur in SA and were reported from soil samples of seven of the nine provinces where potato has been planted in potato- 22 producing regions. Trichodorus spp., however, are all endemic to the natural veld and indigenous forests in SA and have not been associated with any agricultural and/or horticultural crop to date (M. Marias, pers comm., January 2011). Although Paratrichodorus species occurs widely in sandy soils planted to potato, their concomitant occurrence with tobacco rattle virus have not yet been reported in SA (M. Marias, pers comm., January 2011). 1.5 Identification of plant-parasitic (PPN) and non-parasitic nematodes (NPN) Accurate identification of nematodes is the cornerstone upon which all aspects of research, advisory work, implementation of quarantine legislation and selection of control strategies is based. At present classical taxonomy, using morphological characteristics to describe and identify nematode genera and species, is being replaced to a large extent by the use of more sophisticated molecular methods (Perry & Moens, 2006). In some cases, molecular and bar-coding techniques used to identify nematodes may even replace traditional morphological identification of nematodes (Perry & Moens, 2006). This is mainly due to a progressive decline in the numbers of expert taxonomists that were trained to use the traditional morphological approach to identify these parasites. However, from a worldwide perspective, both classical and molecular techniques are required at present to ensure the accurate identification of nematodes (Perry & Moens, 2006). However, to differentiate between PPN species of the same genus, for example some Meloidogyne spp., using morphological characteristics of their perineal patterns alone often prove to be inconclusive (Zijlstra et al., 2000; Hu et al., 2011). Reasons for this are that a morphological approach requires much skill and experience (Jepson, 1987) since considerable similarity exists between some species with regard to the formation of striae of the cuticle that represent the perineal pattern (Zijlstra et al., 2000; Hu et al., 2011). The occurrence of high intra-species variation also contributes to accurate identification of RKN being elusive (Hartman & Sasser, 1985; Zijlstra et al., 2000; Hu et al., 2011). DNA-based diagnostics, however, provide attractive solutions for reliable identification of PPN that are considered to be economically important, such as RKN (Ziljstra et al., 2000), and have been applied with success by a number of researchers (Powers et al., 1986; Piotte et al., 1992; Xue et al., 1992; Castagnone- Sereno et al., 1993; Fargette et al., 1996, Ziljstra et al., 2000; Fourie et al., 2001; Berry et al., 2008). Before research programmes or proper management strategies can be implemented, particularly where quarantine organisms are concerned, accurate and reliable identification of RKN are absolute prerequisites (Piotte et al., 1992; Zijlstra et al., 2000). Accurate identification of Meloidogyne spp. is 23 also important for proper selection of non-host crops for rotation purposes or for the use of a resistant cultivar, when/if available (Thomason & Caswell, 1987). DNA-based methodology has been used extensively during the past few decades for the identification of the economically most important RKN species (Hu et al., 2011). According to literature, M. incognita, M. javanica, M. hapla, M. arenaria, and M. chitwoodi are discriminated from each other mainly by using COII polymerase chain reaction (PCR) and restriction fragment length polymorphism (RFLP) (Powers & Harris, 1993). Meloidogyne incognita, M. javanica and M. arenaria are, however, distinguished from each other by means of PCR using sequence-characterized amplified region (SCAR) primers (Zijlstra et al., 2000; Meng et al., 2004). In addition, M. incognita or M. javanica could also be discriminated by using real-time PCR (Berry et al., 2008; Toyota et al., 2008). More recently, various molecular techniques have also been developed to identify an emerging RKN species, M. enterolobii, that is causing problems in a variety of crops and include analyses of mitochondrial DNA (mtDNA) (Blok et al., 2002; Brito et al., 2004; Xu et al., 2004; Tigano et al., 2005; Zhuo et al., 2010) and ribosomal DNA (rDNA) intergenic regions (IGS) (Blok et al., 1997; Adam et al., 2007), Research-to-date has demonstrated the effectiveness of identifying and differentiating Meloidogyne spp. using molecular techniques. This approach together with the use of morphological methods is fundamental for research programmes and proper management strategies to be implemented, since it could improve progress and crop management decisions (Powers & Harris, 1993; Zijlstra et al., 2000). On the other hand, morphological identification of nematodes (both PPN and NPN) to family and/or genus level generally poses no problems and is usually done as a standard method. Since nematodes feed on a wide variety of soil organisms their greatest apparent morphological diversity can be seen in the head and mouth structures, which are closely related to their feeding habits. According to Yeates et al. (1993) the following nematode feeding or trophic groups are recognised: 1. Herbivores or plant-feeders (PPN) - feeds on host plants using a stomatostylet (Tylenchida, Aphelenchida), onchiostylet (Triplonchida) or odontostylet (Dorylaimida). 2. Fungivores or hyphal feeders - NPN that penetrate fungal hyphae using a stomato- or odontostylet. In addition to obligate hyphal feeders, this group also includes the alternative life cycle of some invertebrate parasites (e.g. Deladenus spp.). 24 3. Bacterivores or bacterial feeders - NPN that feed on any prokaryote food source. These organisms (e.g. Rhabditis spp., Alaimus spp.) ingest their prey through a narrow or broad mouth (e.g. Diplogaster spp.), followed by an oesophagus with strong muscles. 4. Substrate feeders - ingestion of substrate material may be incidental to bacterial feeding, predation and unicellular, eukaryote feeding in many NPN. Mouths of these NPN ranges from short and broad too long and narrow and teeth may be present, suggesting a more predatory life style. The expression “non-selective deposit feeding” used for aquatic nematodes relates to substrate feeding NPN. 5. Predators or animal feeders - NPN that ingest invertebrates such as protozoa, nematodes and rotifers either as “ingesters” (e.g. Diplogaster spp., Mononchus spp., Nygolaimus spp.) or as “piercers” (e.g. Seinura spp., Labronema spp., Laimaphelenchus spp.) by sucking body fluids of their prey through a narrow stylet. 6. Feeders on eukaryotes - NPN that feed on diatoms or other algae, as well as fungal spores and yeast cells. Examples of this trophic group are Achromadora spp., Diplogaster spp. and Fictor spp. 7. Omnivores - NPN that feed on a wide range of foods occurring in more than one trophic level (Yeates et al., 2009) (particularly combining feeding types 2-6). These species are restricted to a few members of the Dorylaimida. Examples are Actinolaimus spp., Aporcelaimellus spp. and Kochinema spp. 8. Dispersal or infective stages of animal parasites - includes life stages of animal parasitic nematodes that occur in the soil, for example invertebrate- (e.g. Deladenus spp., Heterorhabditis spp.) or vertebrate parasites (e.g. Strongiloides spp.). When these stages feed and contribute to soil processes, they should be included in other appropriate categories. If they die in the soil they contribute to the nutrient pool. Families (e.g. Rhabditidae and Diplogasteridae) that use animals as phoretic (transport) hosts are not included in this group. The relationship between nematode community structure and various agricultural practices (e.g. Wang et al., 2004), however, generally refer to only five main trophic groups of NPN viz. bacterivores, fungivores, herbivores, omnivores and predators which are the groups that are mainly focused upon in research of this nature. 25 Figure 1.9 below represents a graphical illustration and explanation of the proposed faunal profile into which soil can be categorised according to the presence, abundance and diversity of NPN, soil nematodes (Ferris et al., 2001). Figure 1.9. Functional guilds of non-parasitic, soil nematodes characterized by their respective feeding habit (trophic group) and by life history characteristics as expressed along a colonizer–persister (cp) scale (after Bongers & Bongers, 1998) (Graphical illustration from Ferris et al., 2001). The Enrichment Trajectory (representing the Enrichment Index; EI) and the Structure Trajectory (representing the Structural Index; SI) are both based on the indicator importance of functional guilds of nematodes and are thus descriptors of food web conditions (Neher et al., 2004). This way quantification of the state of the soil food web through the EI (a measure of opportunistic bacterivore and fungivore nematodes), the Channel Index (CI; indicator of predominant decomposition pathways) and the SI (indicator of food web state affected by stressor disturbance) can be determined (Neher et al., 2004). Quadrant A Stress Enriched Quadrant B Stable Enriched Quadrant C Stable Depleted Quadrant D Stress Depleted 26 Functional guilds of nematodes are defined as a matrix of their feeding habits, their biological, ecological as well as life history characteristics which are all incorporated in the cp classification and ultimately graphically illustrated in the simplified food web (Neher et al., 2004). Thus, the Ba1 functional guild comprises cp1 bacterivores such as those in the Rhabditidae and Panagrolaimidae families; Ba2 those grouped in the Cephalobidae, Monhysteridae and Plectidae families and Ba3 those belonging to the Prismatolaimidae family (Ferris & Matute, 2003; Neher et al., 2004). NPN representing bacterial-feeding in cp1 and fungivores in cp2 are indicators of enrichment (e), while nematodes of all feeding habits classified as cp2 are considered basal (b) to both enrichment and structure trajectories. Ultimately, nematodes of all feeding habits in cp3-5 are indicators of structure (s). Functional guilds of NPN, soil nematodes are characterized by feeding habit (trophic group) and by life history characteristics (Bongers & Bongers, 1998). Indicator guilds of soil food web conditions (basal, structured, enriched) are then designated and weightings of the guilds along the structure and enrichment trajectories are provided, for determination of the Enrichment Index and Structure Index of the food web (Neher et al., 2004). Although the enrichment trajectory can de dissected further to determine flow down fungal and bacterial decomposition channels according to the Channel Index, soil food webs in fields in the Christiana area sampled during this study were only categorised according to the Enrichment and Structure Trajectories to indicate the basic status of these fields in terms of soil health. An explanation of terminology (Ferris et al., 2001) important to understand and interpret faunal graphs are given below: • Colonizer-persister (cp) scale: Assignment of taxa of soil and freshwater nematodes to a 1-5 linear scale according to their r and K characteristics. • cp1: Short generation time, small eggs, high fecundity, mainly bacterivores, feed continuously in enriched media, form dauer larvae as microbial blooms subside. • cp2: Longer generation time and lower fecundity than the cp-1 group, very tolerant of adverse conditions and may become cryptobiotic. Feed more deliberately and continue feeding as resources decline. Mainly, bacterivores and fungivores. • cp3: Longer generation time, greater sensitivity to adverse conditions. Fungivores, bacterivores and carnivores. 27 • cp4: Longer generation time, lower fecundity, greater sensitivity to disturbance. Besides the other trophic roles, smaller omnivore species. • cp5: Longest generation time, largest body sizes, lowest fecundity, greatest sensitivity to disturbance. Predominantly carnivores and omnivores. 1.6 Management practices to reduce plant-parasitic nematodes (PPN) Producers in intensive crop production areas worldwide usually have to choose the practice(s) that is best suited to reduce PPN populations that damage their crops. Such decisions are mainly influenced by crop history, characteristics of a particular crop as well as PPN species that are present in a given field (Nyczepir & Thomas, 2009). Traditional knowledge and agricultural practices are being recalled and reviewed on a frequent basis to assess its potential to address damage caused by PPN in current potato-based cropping systems (Manzanilla-L๓pez et al., 2004). The control of PPN, in this case RKN, is basically aimed at maintaining their population levels below economic threshold densities since total eradication of these parasites is virtually impossible (Keetch & Milne, 1982; Kleynhans et al., 1996). Therefore, management approaches currently used to manage PPN are more holistic, including a combination of tools based on producers’ needs rather than scientific ideology (Sikora et al., 2005). Crop protectors and scientists should also acquire knowledge on the ecological nature of plant diseases (Manzanilla-L๓pez et al., 2004) as well as other important pests that attack crops. While the main focus previously was on the eradication of PPN in the soil and crop tissue by means of synthetically-derived chemical substances, a wider and collective view emerged during the last decade which is aimed at applying sustainable nematode management strategies with some yield loss being accepted (Sikora et al., 2005). Increasingly important is the need to take into consideration the impact of the pest management strategies on biodiversity (both fauna and flora) and the ecological balance in soils (Moens et al., 2009). Even though there are differences in the various approaches to control PPN, the ultimate goals remain the same, namely to reduce PPN populations and enable sustainable crop production by increasing yields at cost-effective levels (Sikora et al., 2005). Although multiple nematode management strategies are available to reduce levels of RKN in potato- based cropping systems, only a few will be elaborated on. 28 1.6.1 Chemical control Chemical control is the most expensive but also the quickest and usually the only practical way to control PPN that infect high-value cash crops such as potato (Kleynhans, 1978). The main advantage of chemical control compared to other methods is that target PPN populations such as RKN, are usually reduced to very low levels within a few days after application of a nematicide. The producer can thus grow a crop during or soon after nematicide application (Keetch & Milne, 1982). At present the use of synthetically-derived nematicides is the predominant and most popular strategy applied by local potato producers to minimise damage to their produce (Jones et al., 2011). A wide range of nematicides, fumigants and non-fumigants are registered in SA for use on a range of agricultural and horticultural crops and in particular on potato (Nel et al., 2007; CropLife, 2011; Table 1.2). These nematicides are generally classified into two main groups, namely fumigant and non- fumigant (contact and systemic) products with a third group being added recently and which include the seed treatment nematicidal applications (Nyczepir & Thomas, 2009). Generally the most effective nematicidal treatment, however, is fumigation of soils prior to planting using various products such as methyl bromide or EDBฎ (ethylenedibromide), Teloneฎ, Telopicฎ or Herbifumeฎ (Sikora & Fernแndez, 2005; CropLife, 2011). Methyl bromide is, however, not available for commercial use in SA anymore (Nel et al., 2007; CropLife, 2011), with potato producers still being heavily reliant on the use of EDBฎ and other fumigants mentioned earlier. Another consideration is that chemical control of PPN on potato in SA is very expensive and is only a short term solution because populations of these parasites can increase to reach damaging thresholds within a couple of months after application. This could lead to more applications as well as higher dosage rates of the product being applied. Therefore, chemical control should only be used on a preventative rather than curative basis (Jones et al., 2011). Although the use of nematicides is at present still an important tool for controlling PPN, its dominance as a control strategy is progressively decreasing (Nyczepir & Thomas, 2009). Many of the nematicides that were previously available have been or are being removed from agricultural markets due to worldwide environmental and human health awareness issues. Therefore, effective control practices that employ alternatives to conventional chemical control should increasingly be investigated and implemented (Batchelor, 2002). 29 Also, due in part to the small market share of nematicides relative to herbicides, insecticides and other pesticides, there is little prospect for the development of novel chemistry in terms of nematicides in the near future (Moens et al., 2009). This increases the risk that PPN could adapt to the few chemicals that are still registered for use on crops such as potato. The latter could result in the selection of PPN populations developing resistance against a particular nematicide (Kleynhans et al., 1996) that is used continuously on potato crops. 30 Table 1.2. Synthetic nematicides currently registered on potato in South Africa to reduce population levels of plant-parasitic nematodes (PPN) (CropLife, 2011). Mode of action Active(s) F/T Concentration Product name Reg no. Company name Fumigant 1,3 Dichloropropene FU-O 1 110 g l-1 Telone IIฎ 5223 Dow AgroSciences SA (Pty) Ltd Fumigant 1,3 Dichloropropene GE 1 110 g l-1 DD92ฎ 4988 BASF South Africa (Pty) Ltd Systemic Aldicarb GR 150 g kg-1 Sanacarbฎ 4717 Bayer (Pty) Ltd Systemic Aldicarb GR 150 g kg-1 Temikฎ 590 Bayer (Pty) Ltd Contact Cadusafos EW 100 g l-1 Rugby 10 MEฎ 6368 FMC SA (Pty) Ltd Contact Cadusafos GR 100 g kg-1 Rugby 10 Gฎ 4110 FMC SA (Pty) Ltd Contact & systemic Ethoprophos GR 150 g kg-1 Mocap GRฎ 6985 Bayer (Pty) Ltd Systemic Fenamiphos CS 240 g l-1 Nemacur 240 CSฎ 7247 Bayer (Pty) Ltd Systemic Fenamiphos EC 400 g l-1 Fenamiphos 400 ECฎ 7167 Erintrade cc t/a RT Chemicals Systemic Fenamiphos EC 400 g l-1 Nemafenฎ 7310 Novon Crop Protection Systemic Fosthiazate EC 900g l-1 Nemathorin 900 ECฎ 5612 ISK Biosciences Systemic Fosthiazate GR 100 g kg-1 Nemathorin 100 GRฎ 5613 ISK Biosciences Fumigant Furfural EC 900 g l-1 Crop Guardฎ 6864 Illovo Sugar Ltd Fumigant Ethylenedibromide VB 1 800 g l-1 EDBฎ 4509 Cropchem Contact & systemic Oxamyl (root-knot nematodes) GR 100 g kg-1 Blockade Gฎ 7552 Klub M5 Contact & systemic Oxamyl (root-knot nematodes) GR 100 g kg-1 Oxagran 100 GRฎ 6406 Nialcor Contact & systemic Oxamyl (root-knot nematodes) GR 100 g kg-1 Vydate 100 GRฎ 3945 Du Pont de Nemours Int SA Contact & systemic Oxamyl SL 250 g l-1 Oxatak SLฎ 7589 Meridian Agrochemical Co.(Pty) Ltd Contact & systemic Oxamyl SL 250 g l-1 Oxadateฎ 7588 Tsunami Plant Protection (Pty) Ltd Contact & systemic Oxamyl (root-knot nematodes) SL 310 g l-1 Blockade SLฎ 7345 Klub M5 Contact & systemic Oxamyl (root-knot nematodes) SL 310 g l-1 Platoon SLฎ 7370 Villa Crop Protection (Pty) Ltd Contact & systemic Oxamyl (root-knot nematodes) SL 310 g l-1 Vydate SLฎ 5057 Du Pont de Nemours Int SA Contact & systemic Oxamyl (root-knot nematodes) WSC 310 g l-1 Vigour 310 SLฎ 8088 Agchem 31 1.6.2 Alternative strategies to manage plant-parasitic nematodes (PPN) 1.6.2.1 Host plant resistance Breeding of plants resistant to specific species of for example RKN or cyst nematodes is probably one of the most economical and effective methods of control since it affords the grower at least the following advantages, namely: • should the growth and yield of the resistant plant be comparable to those of its susceptible counterpart, its use to control nematodes will involve little additional expense (Keetch & Milne, 1982; Cook & Starr, 2005; Nyzepir & Thomas, 2009) and • soil and climatic conditions required for the cultivation of resistant plants are not as critical as those necessary for the efficient use of nematicides (Keetch & Milne, 1982). Host plant resistance is generally defined as the ability of the host to minimise penetration, development, reproduction and fecundity of PPN and ultimately result in minimal damage to the host plant (Bos & Parlevliet, 1995; Hussey & Jansen, 2002; Roberts, 2002). On the other hand, crops highly susceptible to attack by PPN, such as RKN, allow a rapid build-up of population levels of these parasites since they support optimal development and reproduction (Bos & Parlevliet, 1995; Roberts, 2002) of the parasites even when low initial population densities are present at planting (Keetch & Milne, 1982; Whitehead & Turner, 1998). Resistance/susceptibility on the one hand and tolerance/sensitivity on the other hand are defined as independent, relative qualities of a host plant’s reaction to nematode infection based on comparison between a susceptible and resistant genotype (Bos & Parlevliet, 1995). Various mechanisms exist in terms of host plant resistance, including non-preference (antixenosis), antibiosis and tolerance (Bos & Parlevliet, 1995). Antixenosis refers to the response of a pathogen to a host plant that lacks characteristics to serve as a host, resulting from negative reactions or total avoidance during the search for food, oviposition sites or shelter. Antibiosis, on the other hand, includes all adverse effects exerted by the host plant on the biology, i.e. survival, development and reproduction of the pathogen (Bos & Parlevliet, 1995). Host plants that suffer little damage, even when heavily infected with nematodes that would severely damage susceptible host plants are, on the other hand, classified as tolerant (Bos & Parlevliet, 1995). Furthermore, a host plant is hypersensitive when high levels of damage are suffered, even when relatively lightly infected with PPN (Bos & Parlevliet, 1995). 32 However, only a few plants have been classified as being immune to PPN (Keetch & Milne, 1982) which entails total exclusion of the pathogen (Bos & Parlevliet, 1995). Most resistant plant cultivars that are used in cropping systems are usually resistant to only one species of RKN (vertical/qualitative resistance) (Roberts, 2002). Such resistant hosts may also be susceptible to other PPN genera or even to other populations or biotypes of the same species (Roberts, 2002; Starr & Mercer, 2009). The lack of resistance to multiple nematode genera is one of the main limitations of nematode-resistant plants. A further complication may be the occurrence of biotypes within a given RKN species that are able to overcome the resistance of a plant (Roberts, 2002). Quantitative/horizontal resistance on the other hand refers to a host that is resistant against more than one species/race/population of a specific PPN (Roberts, 2002). In terms of RKN in particular, inclusion of host plant resistance in potato cropping sequences together with non-host crops is suggested to be one of the most economical ways to reduce population levels of these parasites (Scurrah et al., 2005). None of the potato cultivars available in SA are, however, resistant against lesion nematodes, PCN or RKN that occur in local potato-producing soils (Jones et al., 2011). In fact, no potato breeding material adapted to warmer climates of the world that confer resistance to thermophyilic RKN species (M. incognita and M. javanica) had been developed by the mid 2000’s (Scurrah et al., 2005). However, the dominant Rmc1 gene in S. bulbocastanum (wild potato) has been reported to confer resistance to M. chitwoodi, M. fallax and M. hapla, which represents cryophilic RKN species that prefers colder climates (Brown et al., 1996; Cook & Starr, 2006; Evans & Perry, 2009). On the other hand, potato cultivars resistant to PCN are available in Australia, Europe, New Zealand and the USA (Finlay et al., 1998; Cook & Starr, 2006) but not in SA (Jones et al., 2011). The only way to use available cultivars to address RKN problems in local potato plantings is to plant short growing cultivars that could be lifted before too many life cycles of RKN developed in their roots and tubers. This way population levels of these parasites in soils could be limited, while damage to tubers could also be reduced substantially or even prevented (Jones et al., 2011). 1.6.2.2 Biological control Biological control is defined as the management of plant diseases and pests by using living organisms or antagonists such as predators and parasites of organisms that kill or damage their hosts (Moens et 33 al., 2009). Furthermore, the use of microbes that indirectly influence the establishment, function and survival of pathogens and pests adversely is included in this definition (Moens et al., 2009). A wide range of bacterial and fungal pathogens and antagonists have been evaluated for use as biological agents to reduce population levels of economically important PPN, such as RKN, PCN and Pratylenchus spp., of potato. These include active antagonists such as Paecilomyces lilacinus, Bacillus firmus, B. chitinosporus, B. laterosporus, B. licheniformis, Pochonia chlamydospona, Pasteuria penetrans, Burkholderia cepacia, Rhizobium spp., Pseudomonas spp., Glomus spp., Fusarium spp. and Trichoderma spp. (Hallman et al., 2009). Studies with P. lilacinus showed that the fungus consistently infected eggs (80-90%) and occasionally females of M. incognita that parasitized potato roots in Peru. The same fungus also penetrated 70-90% of eggs inside G. pallida cysts (Jatala et al., 1979). Except for fungi and bacteria that act as biological antagonists of PPN, other natural-occurring predatory nematodes as well as protozoa that live in soil have also been documented to feed on PPN in local agricultural soils (Keetch & Milne, 1982; Sikora, 1992). It is, however, important that population levels of such micro-organisms are stimulated by implementing soil-friendly approaches to optimize the impact thereof. Such strategies include the incorporation of organic material, i.e. manures, compost and above-ground parts of cover crops, which is seldom used by local potato producers. However, although notable effects on PPN population levels have been obtained with the application of biological agents itself under experimental conditions, their use in local, annual cropping systems such as potato-based sequences at present is very rare (Jones et al., 2011). 1.6.2.3 Quarantine Quarantine aims at excluding or intercepting exotic PPN, that cause major yield and quality problems of crops in certain regions of the world, or isolating and eradicating them once they have been detected and/or become established (Keetch & Milne, 1982). Such strategies are considered a preventive and not a curative approach that are applied to prevent and/or stop the introduction and/or increased dissemination of economically important PPN into a country, local region or planting site (Nyczepir & Thomas, 2009). The prevention of an alien, PPN pest from becoming established in a country, area or field, however, is seldom achieved because of the efficient natural spread of these parasites, the large scale movement of infected plant and planting material, the ability of PPN to establish themselves rapidly on host plants and the difficulty in completely eradicating them from infected plants or infested 34 soil (Nyczepir & Thomas, 2009). Furthermore, the information on which the implementation of control measures is based is derived almost exclusively from the sampling of bulk consignments of soil or plant material (Nyczepir & Thomas, 2009). Since only a few small samples can, however, be examined the chances of detecting one or two live PPN in a consignment of potato for example are minimal (Keetch & Milne, 1982). The long-distance movement of plant material supported by the global markets for agricultural products has also supported the spread of important PPN around the world and between countries on the same continent (Sikora et al., 2005). Therefore, quarantine services cannot indefinitely prevent the introduction into and establishment of alien PPN species in an area. The best that can be done is to delay and monitor the latter scenario (Swart, 2011). Currently, the PPN classified as quarantine organisms for local purposes are G. rostochiensis, Ditylenchus destructor, M. chitwoodi and M. fallax (Swart, 2011). Control of GCN for example in SA is achieved mainly by quarantine measures since no potato plantings are permitted on GCN-infested fields (Jones et al., 2011). The nematode Ditylenchus africanus was first isolated in 1987 in SA from the pods of groundnuts and was initially identified as D. destructor (De Waele et al., 1989). The latter PPN is regarded as a serious pest of potato in parts of Europe and the Soviet Union (Hooper, 1973). Experiments, however, later showed that all local potato cultivars tested were identified as poor hosts to the South African population of this PPN species and that no damage was caused to the tubers (De Waele et al., 1991). The South African population was therefore, later considered to be a new species and described as D. africanus, which does not pose a risk to potato (Wendt et al., 1995). The RKN M. chitwoodi and M. fallax are also quarantine organisms and important pests of potato in Europe, North America, New Zealand and Australia where it causes serious economic losses due to direct damage to the plant and tuber (Nicol et al., 2011). M. chitwoodi was reported for the first time in SA on potato in the Maclear area (Eastern Cape Province). Since then it has been reported on potato in the Clocolan (Free State Province), Boston and Mooiriver areas (KwaZulu-Natal Province) (Jones et al., 2011). Although Fourie et al. (2001) reported the first record of M. fallax from groundnut in the Vaalharts area (Northern Cape Province), the distribution of this species as well as M. chitwoodi in potato production areas remain unknown. 1.6.2.4 Crop and fallow rotations The basic concept of nematode control using crop rotation is to only grow crops susceptible to PPN sporadically to reduce populations of these parasites and prevent them from damaging crops (Keetch & 35 Milne, 1982; Coyne et al., 2009). This way it may also prevent low PPN population densities from increasing to levels at which these parasites would cause economic losses to the potato and other rotation crops (Keetch & Milne, 1982; Coyne et al., 2009; Jones et al., 2011). Crops that are good hosts to RKN and should not be included in local rotation cycles with potato include Fabaceae crops (beans and peas), maize (Zea mays), Solanaceae crops (tomato, paprika, tobacco) and lucern (Medicago sativa) in particular (Jones et al., 2011). Inclusion of maize in potato- based systems could for example lead to high RKN populations since M. incognita and M. javanica are the predominant economically important nematode pests in maize-production areas where potato are also cultivated (Riekert, 1996; Riekert & Henshaw, 1998). Ngobeni et al. (2010), however, reported various local maize hybrids that are resistant to both M. incognita and M. javanica, which can be used by local potato producers to reduce RKN population in such areas. As part of crop rotation systems, cover crops that are non-hosts to PPN have already been used successfully in various crop rotation systems worldwide (Sikora et al., 2005; Coyne et al., 2009). A variety of cover crops/grasses are listed as poor hosts of the predominant M. incognita and M. javanica that occur in local potato-producing areas (Taylor & Martin, 1960; Fourie et al., 2007; Jones et al., 2011) and could be used in such rotation systems. These include grasses like Bloubuffel grass (Cenchrus ciliaris), Borseltjie grass (Anthephora pubescens), Crotalaria spp., Kleinbuffels grass (Panicum coloratum), mustards (Brassica spp.), Oulands grass (Eragrostis curvula cv. Ermelo), Rhodes grass (Chloris gayana), Smutsvinger grass (Digitaria eriantha) and Vetiver grass (Vetivera zizanioides). Van Biljon (2004) also reported that several Sorghum spp., wheat (Triticum spp.), oats (Avena sativa) and alfalfa (Medicago sativa) cultivars as well as a variety of grass species are resistant to M. incognita and M. javanica, while Fourie et al. (2006) listed the soybean (Glycine max) cultivar Egret as resistant to both M. incognita and M. javanica. The current situation in SA is that potato producers generally include the crop in a 3-4 year rotation system with maize and wheat (Sellschop & Du Toit, 1948; Black, 2008), while producers in the Christiana area only plant potato every 7-8 years. Wheat, maize and grasses are used as the main rotation crops and/or cover crops in these cropping systems while some fields are also often left fallow for a year. The biggest advantage of a crop rotation strategy compared to most other cultural control methods or the use of a nematicide is that it generally produces an economic return on investment (Nyczepir & Thomas, 2009). If sufficient markets are available to support profitable production of crops that are poor 36 or non-hosts of RKN, studies can be done to exploit host range limitations of many PPN species. Key factors in determining economic viability of such an approach are (Nyczepir & Thomas, 2009): • accurate identification of PPN species, RKN in particular in the case of potato in SA, and knowledge of the host range for these target species, • knowledge about crops that are non-hosts to these PPN species and that are agronomically suited to the geographic region, and • ensuring that populations of other pathogens or pests are not significantly enhanced by the rotation crop. However, the implementation of crop rotation is not as simple as a nematicide application because rotation crops often require specialized farm equipment and production practices (Halbrendt & LaMondia, 2004; Nyczepir & Thomas, 2009). Crop rotation per se also frequently results in less profit per unit of land, while use of varieties resistant to RKN in this case may carry yield penalties when compared with their susceptible counterparts (Nyczepir & Thomas, 2009). Moreover RKN and lesion nematodes have very wide host-ranges which is the main factor that limits the use of crop rotation as a management strategy. The latter limitation is aggravated by the occurrence of physiological races of Meloidogyne species since crops that are tolerant to some races may be susceptible to others (Keetch & Milne, 1982). Except for rotating crops, a period(s) of fallowing is usually included for a specific field(s) within cropping cycles (Nyczepir & Thomas, 2009). Keeping a land free of all vegetation for a specific time is known as clean fallowing. It can be done by frequently tilling or ploughing the soil or by eliminating all plant growth by means of herbicide application (Keetch & Milne, 1982). Some crop rotations incorporate an extended period of time where the land is left fallow before replanting of another agricultural crop (Nyczepir & Thomas, 2009). The general objective in using a fallow rotation is to reduce the population of PPN prior to replanting the site through starvation, desiccation and increase in soil temperature (Keetch & Milne, 1982; Nyczepir & Thomas, 2009). The apparent benefits of applying fallowing to crop fields to reduce PPN populations are clear. Nearly all PPN are suppressed simultaneously, with minimal expenses being associated with this strategy compared to planting and maintaining a crop (Nyczepir & Thomas, 2009). Many potato growers involved in intensive crop production systems do, however, not rely on fallowing as a primary tactic to suppress RKN. Lack of income from the fallow land, the expenses associated with regular weed control which is preferably every three weeks to 37 prevent RKN reproduction and potentially deleterious effects from soil erosion are all factors that contribute towards the limited acceptance of fallowing (Halbrendt & LaMondia, 2004; Sikora et al., 2005; Nyczepir & Thomas, 2009). However, essential to the success of any rotation cycle is effective weed control because the presence of a variety of weed species that grow in potato fields can result in the build-up of high RKN and other PPN populations despite the poor-host status of for example a resistant and/or non-host crop (McSorley et al., 1994; B้lair & Parent, 1996). Various weed species have been associated with economically important RKN as well as other PPN that occur in local agricultural soils (Keetch & Buckley, 1984; Ntidi et al., 2012). Occurrence of such weeds in potato fields may have serious effects for local producers in terms of RKN infection of the crop. 1.6.2.5 Soil amendments and cover- and/or green manure crops Soil amendments, such as livestock or poultry manure and organic compost are often applied to soils used for intensive annual crop production (Nyczepir & Thomas, 2009). Suppression of PPN due to application of such soil amendments has been recorded and is suggested to result from the enhanced activity of antagonists of PPN in response to elevated levels of organic matter decomposition (Widmer et al., 2002). The control of PPN is however, seldom the primary reason for such applications. Amendments are rather applied as low cost sources of plant nutrients, acceptable methods for the disposal of animal waste or other agricultural by-products or to improve soil properties (Nyczepir & Thomas, 2009). As organic waste production increases in the future and disposal options become more restricted, the addition of soil amendments to intensive cropping systems is, however, likely to increase (Zasada et al., 2008). Application of such amendments, however, poses problems for local commercial potato producers whose crop fields are big, usually in excess of 20ha. Huge amounts of manures or compost are needed in such cases to obtain the desired effect in terms of increasing the organic content of soils and concurrently the numbers of soilborne antagonists of PPN (Coyne et al., 2009). A strategy that promises potential to support/supplement the incorporation of organic substances is, however the use of green manure crops. A more effective and viable method to manage PPN is to use plant species of which the aerial parts can be cut off, slashed and ploughed into the soil as a green manure (Keetch & Milne, 1982). Cover and green manure crops that are intended to suppress PPN populations are increasingly used (Nyczepir & Thomas, 2009). In addition such crops prevent/reduce 38 soil erosion and increase soil quality. Crops that are poor or non-hosts for RKN and prevent the build- up of population levels of these parasites are the best to use for this purpose (Timper et al., 2006). Brassicaceae crops, also referred to as cruciferous crops, offer a potential alternative to the use of synthetically-derived nematicides in intensive cropping systems to reduce PPN populations (Akhtar & Alam, 1991; Sarwar et al., 1998; Bending & Lincoln, 2000; Zasada & Ferris, 2004). These crops produce glucosinolates (GL), that when hydrolysed, produce isothiocyanates that are biocidal to a range of soil organisms (Akhtar & Alam, 1991; Sarwar et al., 1998; Bending & Lincoln, 2000; Zasada & Ferris, 2004). The term biofumigation is used to indicate that volatile substances are produced through microbial degradation of organic amendments that result in significant toxic activity towards PPN and/or other soilborne pests or diseases (Bello, 1998). The efficacy of a range of Brassicaceae species is used to reduce population levels of PPN (Mojtahedi et al., 1991; Lazzeri et al., 2004), other soilborne pathogens (Ramirez-Villapudua & Munnecke, 1988; Subbarao et al., 1999), insects (Williams et al., 1993) and weeds (Brown & Morra, 1995) in agricultural systems. The modes-of-action of these plant species in terms of reducing PPN is suggested to be the production of glucosinolate-degradation products (i.e. isothiocyanates, thiocyanates and nitrogenous substances) that are toxic to nematodes (Lazzeri et al., 1993; Zasada et al., 2003), but beneficial towards stimulation of communities of microbial communities present in the amended soil that are antagonistic to nematodes (Cohen et al., 2005). Various members of the Brassicaceae family have been used as green manures to manage RKN in particular (Mojtahedi et al., 1991; Lazzeri et al., 2004). To achieve the maximum benefit from utilizing Brassicaceae species as cover and/or green manure crops for the control of soilborne pathogens, a Brassicaceae species with a high potential of GL production and also one that produces large quantities of biomass in the selected geographic location or environment should be selected (Kirkegaard & Sarwar, 1998; Morra & Kirkegaard, 2002; Zasada et al., 2003). Research by Zasada et al. (2003) demonstrated that amended materials of Brassicaceae species differed substantially in terms of their effect on PPN and other soilborne organisms. For example, population levels of the citrus nematode Tylenchulus semipenetrans was consistently suppressed by brassicaceous amendments (Zasada et al., 2003), while its effect on Fusarium spp. (Ramirez-Villapudua & Munnecke, 1988) and weeds varied (Brown & Morra, 1995). The latter authors suggested that for consistent and reliable suppression of pests and diseases in crop management systems amended with Brassicaceae species, it is crucial to investigate and understand all component mechanisms that have an impact against the target soilborne organisms present. 39 Another important factor to keep in mind when chemical biofumigant properties of Brassicaceae crops is exploited for use in a cropping system(s) is that such species should not be good hosts of the target PPN (Stirling & Stirling, 2003). Such species of Brassicaceae exists and according to Pattison et al. (2006) Raphanus sativus represents an example of such a species that generally had the most consistent resistance of the 43 Brassicaceae cultivars screened for host suitability against Meloidogyne species during their study. It also proved to be a robust performer under tropical conditions where it was less sensitive to hot conditions as well as attack by insect pests. Other authors, however, argue that the resistance of Brassicaceae cultivars to RKN appear to be more related to genotype than its GL content (Potter et al., 1998). However, other studies indicated that the suppression of M. javanica and M. arenaria after soil incorporation of Brassicaceae tissues is not related to the GL content, but that mechanisms other than biofumigation may have caused reduction in numbers of these parasites (McLeod & Steel, 1999). The latter results, however, contradict those by Kirkegaard and Sarwar (1998); Morra and Kirkegaard (2002); Zasada et al. (2003) and Curto et al. (2005). The latter authors evaluated and selected different Brassicaceae species, namely E. sativa, B. juncea and R. sativus as well as Capparaceae species for their high active GL content and biocidal activity against M. incognita J2. Accordingly all varieties screened could be used in pest management programmes based on their ability to reduce M. incognita infestations in the soil due to their high GL content and subsequent good nematicidal activity. However, although used widely, controversy still exists about the ability of Brassicaceae crops to reduce populations of PPN through biofumigation. The potential reduction of PPN populations in the soil following incorporation of the green manures may be offset by the reproduction of the PPN in the roots of Brassicaceae plants susceptible to these parasites during their growth period (McLeod et al., 2001). It will thus be necessary to define the best strategy to obtain significant PPN control by careful selection and optimal use of cover and green manure crops, such as Brassicaceae, with biofumigation properties in rotation systems (Curto et al., 2005). At present the use of such crops in local cropping systems, including potato, is limited. There is, however, a movement in this direction since producers realise that alternatives must be investigated and found due to the withdrawal of various effective, synthetically-derived nematicides that were traditionally used to address the RKN problem in local potato plantings. 40 1.7 Integrated nematode management and future strategies At present, substantial gaps exist in our knowledge about multiple interactions between the soil ecosystem, weeds and other pests (including PPN) and diseases (Nyczepir & Thomas, 2009). Development of nematode-integrated management programmes thus necessitates analysis of all biotic and abiotic factors, on target PPN populations (Sikora et al., 2005). Furthermore, cost-benefit ratios for grower acceptance should be calculated to estimate the maximum impact a particular management strategy can have in reducing PPN population levels in cropping systems (Sikora et al., 2005). Such integrated approaches are designed to attain high crop yields, while simultaneously reducing PPN as well as other pest, disease and weed problems. Ultimately, the reduction of erosion and improvement of soil fertility should be selection dependant on the main crop, in this case potato (Sikora et al., 2005). Scurrah et al. (2005) listed the use of RKN-resistant cultivars and rotation with non-host crops as the economical most important tools to reduce RKN numbers in potato-based cropping systems effectively. Although RKN problems in local potato producing areas are at present still mainly dealt with by means of chemical control (Jones et al., 2011), planting of RKN-free tubers, crop rotation as well as the inclusion of poor-host rotation crops (Sikora et al., 2005) are all strategies currently used by local potato producers to protect their produce. The wide host range of the predominant PPN parasite of potato, i.e. RKN (Jones et al., 2011), however, continue to pose challenges to producers in terms of aligning available management strategies optimally to address this problem. Local producers realise that the management of PPN in the future will not be dependent on only one of these strategies, but that logical and effective integration thereof will be crucial to enable them to farm sustainably. Most important is that such an integrated approach should be economically acceptable to producers (Sikora et al., 2005). However, an integrated approach should be dynamic and flexible and may require frequent adjustment in response to changes related to the production system (e.g. new cultivars that enter the market, economic factors, environmental conditions or availability of registered nematicides and/or biological products) (Nyczepir & Thomas, 2009). Fortunately, the challenges that the agricultural sector face these days provide opportunities to adapt recent technological, molecular as well as biological, advances together with novel emerging chemical and biological agents for the management of PPN such as RKN (Nyczepir & Thomas, 2009). Once the latter is attained, the primary goal of any integrated pest management system namely increased crop yield (quality and quantity) by using a combination of the management approaches (discussed earlier) to target PPN (Nyczepir & Thomas, 2009) can and will be realised. Therefore, research on alternative control strategies and 41 management practices and also the integration of the latter to minimize the impact of these parasites on local potato production is critical to assist the producers and the industry. The latter constitute the main aim of this study since prior to this research no work has been done on potato in SA to determine the efficacy of Brassicaceae cover crops and its use as biofumigation agents in reducing RKN populations (Berry & Wiseman, 2003; Berry & Rhodes, 2006; Kruger et al., 2011). The present study was thus initiated since producers in the Christiana area realized that they should incorporate strategies, other than chemical control, to address the RKN problems they experience in their potato plantings. 1.8 Objectives of this study • To assess the status of PPN and NPN in 31 potato fields of five producers in the Christiana area (North-West Province), with the emphasis on RKN. • To investigate the effects of selected Brassicaceae species as both cover and/green manure crops on both PPN and NPN population levels under natural occurring environmental conditions in a field experiment. 42 1.9 References Adam, M.A.M., Phillips, M.S. & Blok, V.C. 2007. 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Nematology 2: 847-853. 58 CHAPTER 2 IDENTIFICATION OF NEMATODE ASSEMBLAGES IN POTATO-BASED CROPPING SYSTEMS IN THE CHRISTIANA AREA (NORTH-WEST PROVINCE, SOUTH AFRICA) 2.1 Introduction Root and tuber crops, such as potato, are the most important food commodities produced in many subtropical and tropical countries (Scurrah et al., 2005). The importance of plant-parasitic nematodes (PPN) on such food commodities, particularly potato, are often overlooked because these small, soilborne organisms seldom cause specific above-ground symptoms (Manzanilla-L๓pez et al., 2004). The frequent lack of distinct symptoms caused by PPN often result in damage to aerial parts of crops which are confused with problems caused by other factors, i.e. soil structure and nutrition deficiencies that can negatively impact the growth and development of crops (Kleynhans et al., 1996; Manzanilla- L๓pez et al., 2004; Bridge & Starr, 2007). The latter scenario generally leads to the slow, invisible build- up and spread of potentially pathogenic PPN populations that ultimately damage crops (Manzanilla- L๓pez et al., 2004). Correct diagnosis of a malady is, therefore, the starting point for successful management of PPN associated with crops and should be followed by accurate identification of the genera, species and/or families by a trained nematologist. Ample literature exists about PPN that are associated with potato in various world countries (Scurrah et al., 2005; Bridge & Starr, 2007), including South Africa (SA) (Keetch & Buckley, 1984; Kleynhans et al., 1996; Jones et al., 2011). The root-knot nematode (RKN) species, M. incognita and M. javanica have been listed as the predominant PPN that cause downgrading of potato on local markets. Lesion (Pratylenchus spp.) and the golden cyst nematode (GCN; Globodera rostochiensis) also affect the crop adversely in some local production areas (Jones et al., 2011). However, little is known about the total nematode complex, which includes both PPN and non-parasitic nematodes (NPN) that are associated and adversely affect the crop in local potato-plantings areas. Besides the presence of damaging PPN, it is also crucial to obtain information about the NPN assemblages, representing beneficial nematodes that occur in local fields. Only then can nematode control strategies be planned and applied successfully to prevent eradication of the beneficial nematodes (Yeates et al., 2009). The positive role that NPN plays in agroecosystems has been studied extensively, particularly during the last two decades (Bongers, 1990; Bongers & Ferris, 1999; Ferris et 59 al., 2001). However, in SA efforts to address soil health has only been attended to during the past five years with local nematologists also starting to focus on the role of beneficial nematodes. The current approach by local potato farmers is that they do not produce potato crops that are susceptible to RKN more than once every three or four years on the same field (Sellschop & Du Toit, 1948; Black, 2008). However, to minimise problems caused by RKN as well as other diseases, potato producers in the Christiana area implement an eight year crop rotation cycle during which potato is planted only once (G. Posthumus, pers comm., March 2010). Despite the latter, damage to tubers harvested in this area has increased from 4% to nearly 50% with visible symptoms of RKN infection from the 2001/2002 to the 2009/2010 season (see Table 1.1; Chapter 1) (J. van Vuuren, pers comm., May 2010). To recommend and/or develop viable and effective crop rotation systems for producers in the Christiana area it is imperative to first identify and determine population levels of the nematode complexes present in fields where potato is included. Once the latter has been done, recommendations about the use of cover and rotation crops, also those that are currently planted by these producers, can be made. At present, grasses such as Eragrostis and Digitaria species are planted as cover crops, which are mainly utilized as horse or cattle feed rather than a RKN management strategy. In addition, rotation crops such as maize, onion, sunflower and wheat are also included in these cropping sequences. As mentioned in the introductory part of this dissertation, information about the host suitability of grasses and grain crops to economically important RKN species is available. However, recommendations for their inclusion can only be made once the RKN species that cause problems to potato plantings in fields of these producers have been identified (Taylor & Martin, 1960; Fourie et al., 2007; Jones et al., 2011). Therefore, the first part of this study consisted of the comprehensive sampling of 31 fields of five potato-producers in the Christiana area (North-West Province of SA). 2.2 Material and methods 2.2.1 Study area This study was conducted near Christiana (S 28ฐ09.312, E 024ฐ59.671), situated in the south eastern part of the North-West Province of SA (Figure 2.1), with an altitude of approximately 1 050-1 400m above sea level (Mucina & Rutherford, 2006). This area is located in the Savanna biome of the Kimberly Thornveld and is characterized by summer and autumn rainfall and dry winters (Mucina & 60 Rutherford, 2006). Annual rainfall generally ranges between 300-500mm with mean minimum and maximum temperatures between -3.9ฐC and 37.4ฐC, respectively (Mucina & Rutherford, 2006). The nematode survey encompassed the sampling of both soil and root samples from six fields of five producers each (Figure 2.2; Tables 2.1–2.5), except for one producer where seven fields were sampled. Figure 2.1. A map of South Africa indicating where the study area is located near Christiana (North-West Province of South Africa) (Adapted from: http://cybercapetown.com/Maps/SouthAfrica). N CHRISTIANA 61 Figure 2.2. A detailed aerial view of the 31 individual fields of five producers in the Christiana area from which nematode samples were collected during the 2010 and 2011 growing seasons (Dirk Cilliers, Unit of Environmental Sciences and Development, NWU, Potchefstroom Campus, compiled the map using Google Earth, 2012). 62 Table 2.1. Detail information of fields (location, size and soil characteristics) as well as production practices, rotation/cover crops planted and nematicides applied where nematode samples were collected during the 2010 and 2011 growing seasons for producer 1 (Mr. J. Easby). Field no GPS coordinate Field size (ha) Soil type Production practice Cover and/or rotation crop Crops/Grasses planted Potato planting before survey was conducted/cv Nematicides applied on previous potato crops % Sand % Silt % Organic C 2010 2011 1 (TG2) S 28ฐ03.607ด E 25ฐ05.149ด 88 93.9 3.2 0.19 - 2 x disc - 2 x plough - Eragrostis - Maize - Potato Eragrostis Eragrostis 2004/Mondial - EDBฎ a - Temikฎ 2 (TG3) S 28ฐ03.607ด E 25ฐ05.149ด 88 93.7 1.2 0.22 - 2 x disc - 2 x plough - Eragrostis - Maize - Potato Eragrostis Potato/Mondial 2004/Mondial - EDBฎ - Temikฎ 3 (CG3) S 28ฐ03.607ด E 25ฐ05.149’ 88 93.9 1.1 0.17 - 2 x disc - 2 x plough - Eragrostis - Maize - Potato Eragrostis Potato/Mondial 2004/Mondial - EDBฎ - Temikฎ 4 (TK4) S 28ฐ03.607ด E 25ฐ05.149ด 28 94 1.1 0.17 - 2 x disc - 2 x plough - Eragrostis - Maize - Potato Potato/Mondial Eragrostis 2004/Mondial - EDBฎ - Temikฎ 5 (T6T5) S 28ฐ03.607ด E 25ฐ05.149’ 36 94 1.1 0.17 - 2 x disc - 2 x plough - Eragrostis - Maize - Potato Potato/Mondial Maize 2003/Mondial - EDBฎ - Temikฎ 6 (T6T7) S 28ฐ03.607ด E 25ฐ05.149ด 36 93.9 1.1 0.19 - 2 x disc - 2 x plough - Eragrostis - Maize - Potato Eragrostis Potato/Sifra 2005/Mondial - EDBฎ - Temikฎ aEDBฎ = ethylenedibromide 63 Table 2.2. Detail information of fields (location, size and soil characteristics) as well as production practices, rotation/cover crops planted and nematicides applied where nematode samples were collected during the 2010 and 2011 growing seasons for producer 2 (Mr. JF. Van der Merwe). Field no GPS coordinates Field size (ha) Soil type Production practice Cover and/or rotation crop Crops/Grasses planted Potato planting before survey was conducted/cv Nematicides applied on previous potato crops % Sand % Silt % Organic C 2010 2011 7 (VD6) S 28.059230ฐ E 25.195532ฐ 40 94 1.1 0.16 - Cut - Plough - Digitaria - Maize - Onion - Potato Potato/Sifra Fallow 2004/Mondial - EDBฎ a - Mocapฎ 8 (LH10) S 28.075963ฐ E 25.171617ฐ 22 94.1 1.1 0.16 - Cut - Plough - Digitaria - Potato Potato/Fabula Digitaria 2000/Sparta - EDBฎ - Nemacurฎ 9 (LH13) S 28.065838ฐ E 25.180370ฐ 22 93.9 1.1 0.23 - Cut - Plough - Digitaria - Potato Potato/Mondial Digitaria Never - EDBฎ - Mocapฎ 10 (LH15) S 28.058582ฐ E 25.159005ฐ 22 96 1.1 0.15 - Cut - Plough - Digitaria - Onion - Potato Onion Fallow 2003/Mondial - EDBฎ - Nemacurฎ 11 (LH20) S 28.091382ฐ E 25.240698ฐ 22 93.8 3.3 0.21 - Cut - Plough - Maize - Potato - Sunflower - Wheat Fallow Potato/Sifra Never - EDBฎ - Mocapฎ 12 (VS6) S 28.003375ฐ E 25.208412ฐ 22 94 1.1 0.28 - Cut - Plough - Digitaria - Maize - Potato Fallow Digitaria 2007/Mondial - EDBฎ - Mocapฎ aEDBฎ = ethylenedibromide 64 Table 2.3. Detail information of fields (location, size and soil characteristics) as well as production practices, rotation/cover crops planted and nematicides applied where nematode samples were collected during the 2010 and 2011 growing seasons for producer 3 (Mr. J. Greyling). Field no GPS coordinates Field size (ha) Soil type Production practice Cover and/or rotation crop Crops/Grasses planted Potato planting before survey was conducted/cv Nematicides applied on previous potato crops % Sand % Silt % Organic C 2010 2011 13 (A1) S 28บ07.948ด E 25บ00.003ด 32 93.9 1.1 0.3 - Plough - Rip - Digitaria - Potato Digitaria Potato/Mondial 2005/Mondial - EDBฎ a - Mocapฎ - Vydateฎ 14 (A6) S 28บ08.949ด E 25บ03.277ด 32 91.8 1.2 0.31 - Plough - Rip - Digitaria - Maize - Potato Potato/Mondial Maize Never - EDBฎ - Mocapฎ - Nemacurฎ - Vydateฎ 15 (B1) S 28บ08.099ด E 25บ03.656ด 32 93.8 1.2 0.37 - Plough - Rip - Digitaria - Potato Digitaria Potato/Sifra 2005/Mondial - EDBฎ - Mocapฎ - Vydateฎ 16 (B2) S 28บ08.099ด E 25บ03.650ด 32 91.9 1.1 0.34 - Plough - Rip - Digitaria - Maize - Potato Digitaria Digitaria 2006/Mondial - EDBฎ - Mocapฎ - Vydateฎ 17 (SP2) S 28บ07.854ด E 25บ02.543ด 34 91.6 1.2 0.48 - Plough - Rip - Digitaria - Maize - Potato Digitaria Potato/Mondial 2005/Mondial - EDBฎ - Mocapฎ - Vydateฎ 18 (SP11) S 28บ05.540ด E 24บ58.590ด 30 93.9 1.1 0.23 - Plough - Rip - Digitaria - Maize - Panicum Potato Digitaria Potato/Mondial 2005/Mondial - EDBฎ - Mocapฎ - Vydateฎ 19 (S5) S 28บ07.943ด E 24บ59.998ด 20 91.9 1.1 0.36 - Plough - Rip - Digitaria - Eragrostis - Maize - Potato Digitaria Digitaria 2007/Mondial - EDBฎ - Mocapฎ - Vydateฎ aEDBฎ = ethylenedibromide 65 Table 2.4. Detail information of fields (location, size and soil characteristics) as well as production practices, rotation/cover crops planted and nematicides applied where nematode samples were collected during the 2010 and 2011 growing seasons for producer 4 (Mr. W. Easby). Field no GPS coordinates Field size (ha) Soil type Production practice Cover and/or rotation crop Crops/Grasses planted Potato planting before survey was conducted/cv. Nematicides applied on previous potato crops % Sand % Silt % Organic C 2010 2011 20 (P1) S 27.988130ฐ E 25.110988ฐ 20 91.8 1.2 0.36 - Plough - Rip - Maize - Potato - Onion Maize Fallow 2007/Mondial - EDBฎ a 21 (P2) S 27.995517ฐ E 25.123238ฐ 30 91.9 1.1 0.33 - Plough - Rip - Digitaria - Maize - Potato Digitaria Digitaria 2000/Mondial - Nemacurฎ 22 (VH1) S 27.998832ฐ E 25.112532ฐ 60 91.8 1.2 0.29 - Cut - Plough - Maize - Potato - Wheat Fallow Potato/Mondial 2005/Mondial - Nemacurฎ - Temikฎ 23 (VH2) S 28.00420ฐ E 25.121402ฐ 50 93.8 1.2 0.26 - Cut - Plough - Maize - Potato - Wheat Potato/Mondial Maize 2006/Mondial - Nemacurฎ - Temikฎ 24 (VH3) S 28.002718ฐ E 25.130312ฐ 50 93.7 1.2 0.27 - Cut - Plough - Maize - Potato - Wheat Fallow Potato/Mondial 2004/Mondial - Nemacurฎ - Temikฎ 25 (VH5) S 28.005967ฐ E 25.148875ฐ 50 94 1.1 0.23 - Plough - Rip - Maize - Potato - Wheat Fallow Fallow 2006/Mondial - EDBฎ aEDBฎ = ethylenedibromide 66 Table 2.5. Detail information of fields (location, size and soil characteristics) as well as production practices, rotation/cover crops planted and nematicides applied where nematode samples were collected during the 2010 and 2011 growing seasons for producer 5 (Mr. W. Ras). Field no GPS coordinates Field size (ha) Soil type Production practice Cover and/or rotation crop Crops/Grasses planted Potato planting before survey was conducted/cv Nematicides applied on previous potato crops % Sand % Silt % Organic C 2010 2011 26 (D4) S 28ฐ07.665ด E 25ฐ00.265ด 60 91.8 1.2 0.25 - Disc rip - Plough - Eragrostis - Potato Fallow Potato/Mondial 2002/Mondial - EDBฎ a - Vydateฎ 27 (GH1) S 28ฐ09.106ด E 25ฐ00.323ด 60 93.9 1.2 0.31 - Disc rip - Plough - Eragrostis - Potato Eragrostis Eragrostis 2009/Mondial - EDBฎ - Vydateฎ 28 (GH17) S 28ฐ09.563ด E 25ฐ02.731ด 60 93.8 1.2 0.29 - Disc rip - Plough - Eragrostis - Potato Fallow Potato/Mondial 2003/Mondial - EDBฎ - Vydateฎ 29 (UK1) S 28ฐ08.850ด E 24ฐ59.834ด 60 91.7 1.2 0.36 - Disc rip - Plough - Eragrostis - Potato Eragrostis Eragrostis 2008/Mondial - EDBฎ - Vydateฎ 30 (UK4) S 28ฐ09.312ด E 24ฐ59.671ด 60 91.7 1.2 0.38 - Disc rip - Plough - Eragrostis - Potato Fallow Potato/Mondial 2002/Mondial - EDBฎ - Vydateฎ 31 (UK5) S 28ฐ08.722ด E 24ฐ58.184ด 60 93.7 1.2 0.44 - Disc rip - Plough - Eragrostis - Potato Eragrostis Eragrostis 2006/Mondial - EDBฎ - Vydateฎ aEDBฎ = ethylenedibromide 67 2.2.2 Nematode sampling (surveys) Due to the nature of the crop rotation cycles practiced by potato producers in the Christiana area, not all the fields were planted with potato at the time when nematode samples were collected. Some fields were planted with maize, onion or grass cover crops (Figure 2.3A; Tables 2.1-2.5) while others were left fallow (Figure 2.3B; Tables 2.1-2.5). The time of nematode sampling coincided with flowering of the potato crop during both the 2010 and 2011 growing seasons. This was done to access population levels as well as changes of PPN and NPN and to advise growers on strategies to implement in order to alleviate the PPN problem in their potato crops. Soil samples of each field were also analysed to obtain information on the physical characteristics of the soil and are listed for each of the fields in Tables 2.1-2.5. Due to a technical error, pH measurements for the 31 fields were not analysed. Figure 2.3 (A & B). A field planted with a grass cover crop (A) and a field left fallow (B) in the Christiana area where nematode samples were collected during the 2010 and 2011 growing seasons. Nematode sampling was done in 31 fields of five potato producers in the Christiana area both during the 2010 and 2011 growing seasons. For the 2010 season, samples were obtained from 16 to 18 March and from 14 to 16 February during 2011. Soil and root samples were collected from fields planted with crops (i.e. potato, maize and onion) as well as those planted with grasses. Only soil samples were, however, collected from fallow fields. Field sizes ranged between 20-88ha and only fields planted with potato, maize or onion at the time of sampling were irrigated from the Vaalriver. Fields planted with grasses were rain fed. For each field, 40 soil and root (crops and grasses) or only soil samples (fallow fields) were randomly collected at a depth of approximately 30cm using garden spades. This procedure was followed since A B 68 the top 15-20cm of soil where the majority of the root systems of crops planted in this area occur within the soil profile is also where PPN generally occur (Hooper et al., 2005). Root and corresponding soil samples from each field were placed in plastic bags, marked and transported to the Nematology Laboratory of the North-West University in Potchefstroom (North-West Province) where it were stored at 6-8บC until nematode extraction was performed. The latter was usually within five days after nematode sampling. Before extraction, soil as well as roots of the 40 samples obtained from each field were mixed thoroughly and divided into 4 sub-samples, which were then each subjected to nematode extractions as described below. 2.2.3 Nematode extractions 2.2.3.1 Root samples (50g) The adapted NaOCl method was used for the extraction of RKN eggs and females from the 50g root sub-samples. Principle This particular method is based on the use of NaOCl that dissolves the gelatine layer that surrounds the egg masses that has been produced by RKN females on the surface/within root segments (Riekert, 1995). Procedure a) A 50g root sample was taken from each of the four sub-samples that were collected from each field and cut into 1cm pieces and mixed thoroughly. b) Each root sample was shaken for 4 minutes in 400ml of a 1% NaOCl solution to break down the gelatinous matrix surrounding the RKN eggs and releases them from the roots. c) This mixture containing the eggs and J2’s was decanted through a range of nested sieves, which from bottom to top consisted of a 10-, 25-, 45-, 63-, 75-, 250- and ?m-mesh sieves. 7he latter sieve ensXres less Flogging on the bottom ?m-mesh sieve during the extraction process. A vacuum pump (Edwards high vacuum pump, RPM1725/1426, 60/50Hz) was also connected to the 10-?m mesh sieve to apply suction and enhance the movement of the NaOCl-water solution through the range of sieves. d) Root fragments on the top 710?m-mesh sieve were washed thoroughly with tap water for about 1 minute. 69 e) Eggs and J2 collected on the 10?m-mesh sieve were collected with running tap water into a 100ml sample bottle for counting. 2.2.3.2 Root samples (5g) Nematodes were extracted from 5g root samples using the adapted sugar-flotation method (Hooper et al., 2005). • Sugar-flotation method Principal This method relies on the density, size and gravity of nematodes and is based on maceration of root and tuber material. Procedure a) A 5g sample was taken from the four sub-samples from each of the fields sampled. The roots were cut into 1cm pieces and mixed thoroughly. b) Each root sample was macerated in 250ml tap water at high speed in a domestic blender for 90 seconds to release nematodes from the root tissue. c) The water suspension containing the nematodes and root fragments were subsequently decanted on a ?m-mesh sieve that was nested on a ?m-mesh sieve. d 7he root pieFes on the ?m-mesh sieve were washed thoroughly with running tap water for 2 minutes, the debris discarded and the residue on the 25?m-mesh sieve collected and decanted into a 50ml centrifuge tube for each separate sub-sample. f) Two cubic cm of kaolin was added to each of the tubes containing the nematodes in suspension, stirred well and centrifuged at 4000rpm for 1 minute. g) The supernatant that contained mainly debris was decanted and a sucrose solution with a specific gravity of 1.15g/cm3 added to each tube. The mixture containing the nematodes was stirred well and again centrifuged at 1 800g for 1 minute. h 7he sXpernatant Fontaining the nematodes was deFanted onto a ?m-mesh sieve and rinsed well with tap water to remove the sucrose solution. The residue on the sieve that contains the nematodes was collected in a 100ml sample bottle for counting and identification purposes. 70 • The importance of kaolin (Coolen & D’Herde, 1972) Kaolin is a clay mineral with a specific gravity of 2.6 and consists of particles that are approximately 2-?m in si]e. AlthoXgh the densit\ of kaolin is greater than that of nematodes, kaolin particles are small and flat. Therefore, these particles sink to the bottom of the centrifuge tube at a slower pace than the nematodes. Kaolin thus spreads out to form a layer over the loose sediment that contains the nematodes and seals it off when the supernatant is decanted. The nematodes thus stays sealed-in under the kaolin layer. After addition of the sucrose solution, the kaolin layer must be broken to enable mixing of the sediment and nematodes with the sucrose solution. This way the nematodes are brought into suspension in the sugar solution and will stay in suspension during and after the centrifuging process. Kaolin also precipitates during the second centrifugation with the sugar solution and prevent re-mixing of the sedimented debris when the sugar solution, containing the nematodes, is decanted on the ?m-mesh sieve. Finally, a suspension of nematodes in clear water is obtained for counting and identification. 2.2.3.3 Soil samples (200g) Extraction of nematodes, both PPN and NPN, from 200g of each of the four soil sub-samples, was done using the elutriator method (Seinhorst, 1962). • Elutriator method Principle The method is based on the difference in size, shape and sedimentation rate between nematodes and soil particles. It also depends on the mobility of nematodes since the nematode-water suspension obtained after elutriation is placed on tissues through which the nematodes move to be collected in clear water for counting and identification purposes. The Elutriator set-up uses an upward stream of tap water that enables the nematodes and fine particles to float in the extraction column, whereas heavier particles concurrently settle in the lower part of the apparatus. The nematode suspension in the extraction column is decanted through a side-outlet and sieved to remove the fine particles. The nematodes and washings are then washed from the sieves and placed on tissues. Nematodes move through the tissues into the water, collected in counting bottles and transferred to counting dishes for counting and identification purposes. 71 Procedure a) Rubber stoppers were firmly secured to the bottom of the each elutriator and a 5L bucket was placed on the top shelf next to the pipe of each elutriator. b) Each elutriator was filled to the top outlet pipe with tap water and the pressure dials adjusted to 75ml/min. c) A large ?m-mesh sieve was placed on top of a funnel and the funnel then placed into an open 2L Erlenmeyer flask. d) A 200g soil sub-sample was placed in a 200ml plastic beaker. Soil particles and debris with a diameter of more than 1mm were removed by staying on top of a ?m-mesh sieve. All the soil was subsequently washed through the sieve into the Erlenmeyer flask. e) The sieve and funnel were inverted and cleaned with high pressure spray using tap water to remove all debris and dirt (these were now ready to use for the next sub-sample). f) The Erlenmeyer flask was filled to the brim with tap water. The lid was then wetted with water and placed on top of the flask. Holding the sample carefully and ensuring that the lid was closed, the sample was shaken well by turning the flask upside down 3 times. The flask and lid were then placed upside down on the holder at the top of the elutriator. g) While holding the lid with one hand, the rubber stopper was gently opened to release the sample containing the nematodes into the elutriator allowing it to flow through the elutriating column for 20 minutes. Throughout the process it was checked that the lid of the Erlenmeyer flask did not block. h) After 20 minutes the remaining contents from the flask was emptied into the bucket and the bucket was left for a further 10 minutes to collect the rest of the nematode-water suspension that was still inside the elutriator column at the top. The bucket was then placed on the bottom shelf at the end of the elutriator and the clip on the bottom pipe was removed. This way the rest of the nematode-water suspension was transferred from the elutriator into the bucket. i) The bucket with the nematode-water suspension was left standing for at least 1 hour for the nematodes to settle. j) The nematodes were then concentrated by passing the suspension in the bucket through a series of six stacked sieves. The bigger nematodes (Xiphinema, Longidorus) were retained on the two top sieves (100ตm-mesh), while the smaller nematodes (Pratylenchus, Meloidogyne, Helicotylenchus etc.) were retained on the four bottom sieves (50ตm-mesh). k) The contents of the bucket were slowly poured onto the top sieve (1st 100ตm-mesh). Using a hand- held bottle filled with tap water, the nematodes on this sieve were washed into a clean 200ml beaker. The sieve was then turned upside down and whatever remained on it was washed onto the next 72 sieve below. Then the top sieve was cleaned with high pressure water spray and was ready for use for the next sample. The contents of the second sieve were washed (as described above) onto the bottom sieve (2nd 100ตm-mesh) into the same 200ml beaker. The remaining nematodes and washings on the second sieve was washed onto the third, smaller 50ตm sieve below and the second sieve cleaned as describe above for the first sieve. The same procedure as for the first two sieves were followed and the nematodes and washings on the third sieve washed into another a clean 200ml beaker. These steps were repeated until all nematodes and washings on the four 50ตm-mesh sieves had been washed into the beaker. m) Both beakers containing the suspended nematodes were allowed to stand for at least 30 minutes. n) Brown and blue sieves (both 38ตm-mesh) were then lined with 3 ply tissue and the brown sieve placed on top. All the sieves were wetted with tap water. o) Liquid from the 1st beaker was poured onto the mesh of the brown sieve and the sieve was placed on top of a 150mm brown plastic saucer. p) Liquid from the 2nd beaker was then poured onto the mesh of the blue sieve and the blue sieve placed on top of a second 150mm brown saucer. Afterwards 50ml distilled water was added to the base of each saucer and both sieves were covered with another brown saucer. q) After being left for 48 hours, in order for the nematodes to migrate through the tissue and mesh of the sieves into the water, the sieves were lifted and the excess water drained into the bottom of the brown saucers. r) The nematode-water suspension from the saucer was poured into a clean, 100ml measuring cylinder. s) The nematode-water suspension in the measuring cylinder was left to stand for 90 minutes before counting and identification of the nematodes suspended was done. 2.2.4 Counting of nematodes PPN that were extracted and of which mature specimens were present were identified to species level using morphological characteristics. The same procedure was followed for NPN, only for the 2011 survey, although the latter were only identified to order or family level. RKN species, however, were identified using molecular techniques, namely the SCAR-PCR method (Chapter 3) (Zijlstra, 2000; Zijlstra et al., 2000). PPN and NPN were counted in a De Grisse counting dish using a stereo microscope at 100x magnification. RKN nematode eggs and J2 from the 5g root as well as NPN from the 200g soil samples 73 were counted the same way without prior fixing of the nematodes. PPN per 200g soil were also counted with concurrent identification to genus level but fixed afterwards to enable accurate morphological species identification as described below. 2.2.5 Species identification by means of morphological features For species identification, PPN were transferred to anhydrous glycerine and mounted on microscope slides according to the paraffin ring method (see paragraph 2.2.5.1). 2.2.5.1 Transfer of nematodes to anhydrous glycerine It is crucial to study the finer morphological structures of nematodes for identification at species level. Nematodes were, therefore, fixed in a heated formaldehyde-propionic-acid-water (FPG) solution (100ml of a 40% formalin solution, 10ml propionic acid and 890ml distilled water) [Verbal communication, Dr. Antoinette Swart, Taxonomist ARC-Plant Protection Research Institute (PPRI), National Nematode Collection, Pretoria, South Africa, 22 November 2010] in an incubator at 40บC for 72 hours and then transferred to a glycerin solution (De Grisse, 1965). The method is described in detail below. Procedure Nematodes to be fixed were transferred to distilled water in a Syracuse dish (5ml capacity). Most of the water in which the nematode was suspended was carefully removed after the suspension was allowed to stand for at least one hour and the nematodes settled to the bottom of the dish. An FPG solution containing 100ml of a 40% formalin solution, 10ml propionic acid, 890ml distilled water and just enough picric acid to stain the solution citrus yellow was prepared and heated in a water bath to between 60- 70บC. The hot fixative was added to the dish containing the nematodes and placed in a petri dish with a closed lid, which was then placed in a desiccator with an atmosphere of FPG solution. The latter was obtained by adding FPG solution to the bottom of the dessiccator. The desiccator was placed in an oven at 38-40ฐC for three days. After three days the lid of the petri dish was removed and half of the FPG solution drawn off from the nematode suspension. Solution 1 (200ml of a 95% ethanol solution, 10ml glycerine, 790ml distilled water) was then added to the small dish containing the nematodes. Without replacing the cover of the petri dish the nematodes were put back in the desiccator, with an atmosphere of a 95% ethanol solution, for 12 hours. After 12 hours half of Solution 1 was drawn off and replaced with Solution 2 (950ml of a 95% ethanol solution, 50ml glycerine). The petri dish was then partially covered (so that slow evaporation could take place) and placed in an incubator at 38-40บC for 74 two to three days or until all the ethanol had evaporated. The small glass dish with nematodes was removed from the incubator and placed in a desiccator (that contained silica at the bottom) for another two days after which they were ready to be mounted. • The paraffin-ring method A copper tube with a diameter of 1.5cm was heated, pressed in paraffin wax and touched on the surface of a glass slide. As a result, a paraffin ring was formed on the slide. A small drop of glycerine was placed in the middle of the wax ring. Nematodes were individually fished from the eyeglass using a needle, transferred to the middle of the ring and sealed off by slipping a cover glass over the paraffin ring. The slide was heated over an open/warm plate to melt the paraffin wax and this way sealing in the nematodes. Finally the outside edge of the cover glass was sealed off with colourless acrylic nail hardener (Cutex). The mounted nematode specimens were sent to the nematode taxonomists, Drs. A. Malan, M. Marias and E. Van den Berg (ARC-Plant Protection Research Institute, Pretoria) for identification to species level. 2.2.6 Rearing of root-knot nematode (RKN) species for molecular identification Seedlings of the susceptible tomato cultivar Rodade was grown in 4L pots containing soil samples from the 31 fields of the five producers to rear RKN populations for identification purposes. Soil were collected from each locality and placed in separate, 4L plastic pots in a greenhouse which had no temperature regulation. Susceptible tomato seedlings (cv. Rodade) were planted in each pot to attract RKN J2’s and establish RKN populations for each locality. Infected tomato plants with visible galling were removed approximately 56 days later for identification of Meloidogyne species using molecular- based methods as described in Chapter 3. 2.3 Data analysis 2.3.1 Calculation of prominence values Nematode data obtained as a result of both surveys conducted during 2010 and 2011, respectively, was subjected first to the calculation of prominence values (PV) using Excel (Microsoft Windows XP). PV takes into consideration the population density (PD) as well as frequency of occurrence (%) of the 75 relevant nematode family/genus/species. For this study, PV were calculated only for nematode genera and families. The following equation (De Waele & Jordaan, 1988) was used to calculate PV: PV = population density x frequency of occurrence /10 Population density of each nematode genus/family at each locality (producers’ fields) was calculated as follows: total number of the genus/family present in each field number of samples per field in which the genus/family occurred Frequency of occurrence of each nematode genus/family at each locality (producers’ fields) was calculated as follows: number of samples per field in which the genus/family occurred x 100 total number of sample obtained per field 2.3.2 Parametric and non-parametric statistics Data for PPN (2010 and 2011) and NPN (2011) was transformed to log10(x+1) to remove the inherent variability before analyses. Due to general non-uniformity of nematode data it was also subjected to the Kruskal-Wallis test for non-parametric data and the Mann-Whitney U, T-test and Anova tests for parametric data (Statistica for Windows Version 10). For these analyses the various cultivation practices, nematicides and rotation sequences applied to the 31 fields were grouped into categories and assigned values (Table 2.6) to enable identification of those parameters that had a significant effect on the population levels of the various nematode genera/family identified during the surveys. In addition, NPN data obtained during the 2011 season were subjected to a faunal analysis based on calculating Enrichment Indices (EI) and Structural Indices (SI) using the Faunalizer program originally described by Bongers and Ferris (1999) and improved by Ferris et al. (2001). Nematode families present in soil samples were categorised along a coloniser-persister (cp) continuum (Bongers, 1990). This system recognizes that taxa of monophyletic families are adapted similarly to specific environmental conditions and food sources through anatomical and physiological commonalities. These taxa are then assigned a 1-5 linear scale according to their r and K strategies. Frequency of occur ence 76 Subsequently, enrichment and structure indices were calculated according to Ferris et al. (2001), which are the descriptors of the food web condition at a particular site. These trajectories allow a graphic representation of the faunal profile in order to quantify the soil food web state through the enrichment index (a measure of opportunistic bacterivore and fungivore nematodes) and the structure index (indicator of food web state affected by stress or disturbance). Results obtained by subjecting nematode data to these statistical analyses indicated whether enrichment-opportunist bacterivore nematodes increased rapidly in response to low C/N plant materials or whether population levels of opportunist bacterivore nematodes increased/decreased. Trends in terms of population shifts of fungivore nematodes as well as predator nematodes are also recorded using this simplified food web analysis for NPN (Ferris et al., 2001). Categorisation of these genera and family into functional guilds are based upon designated feeding habits, namely ba = bacterivores, ca = carnivore, fu = fungivores, om = predator (omnivore). Suffix numbers for each guild represents cp values (Bongers, 1990) for the various nematode taxa as reflected in Table 2.23. Table 2.6. Categories assigned to various rotation sequences, cultivation practices and nematicides applied on 31 fields of five potato producers in the Christiana area (North-West Province). Value assigned Cultivation practices Nematicides Rotation sequences 1 Plough EDBฎ a only Grass and potato 2 Rip and Plough Other nematicides Grass, maize and potato 3 - EDBฎ and other nematicides (combined) Maize, wheat and potato 4 - - Maize and/or grass and potato aEDBฎ=ethylenedibromide 77 2.4 Results 2.4.1 Prominence values 2.4.1.1 Plant-parasitic nematode (PPN) data • Roots (50g and 5g) Table 2.7. Meloidogyne species data for fields of five producers in the Christiana area sampled during the 2011 growing season as represented by prominence values (PV), frequency of occurrence (FO; %) and population density (PD) per 50g roots of various crops and grasses. Field no PV PD FO 23 701 7009 100 5 276 2755 100 19 53 1050 25 14 12 172 50 15 7 102 50 7 5 52 100 29 4 82 25 4 4 76 25 3 1 28 25 12 0.3 6 25 Extraction of Meloidogyne spp. from 50g roots of crops and grasses sampled using the adapted NaOCl method (Riekert, 1995) was done only during the 2011 survey. This was due to unforeseen technical problems experienced with extraction apparatus of the Nematology Laboratory during the 2010 season. RKN eggs and J2 were extracted from crops and grass roots from 10 of the 31 fields sampled during 2011 and were identified as M. incognita (fields 3, 14, 19, 23 and 29) and M. javanica (fields 5 and 29) using molecular techniques (see Chapter 3). Mature RKN females from fields 4, 7, 12 and 15 could not be obtained for molecular or morphological identification purposes for the purpose of this study (see Chapter 3). PV for RKN ranged from 0.3 (field 12) to 701 (field 23), PD from 6 to 7009 and FO from 25% to 100% for the respective fields. Fields 23, 5 and 19 had the highest PV, PD and FO for RKN while field 12 had the lowest (Table 2.7). 78 Table 2.8. Data for endo- and semi-endoparasitic nematode genera/family in fields of the five producers from the Christiana area sampled during the 2010 and 2011 growing seasons as represented by prominence values (PV), frequency of occurrence (FO; %) and population density (PD) per 5g roots of various crops and grasses. 2010 Survey 2011 Survey Meloidogyne spp. Hoplolaimidae Meloidogyne spp. Pratylenchus spp. Ditylenchus spp. Field no PV PD FO Field no PV PD FO Field no PV PD FO Field no PV PD FO Field no PV PD FO 14 45 450 100 5 2 33 25 23 39 451 75 10 5 56 75 29 2 15 100 31 23 227 100 21 0.5 10 25 14 4 43 75 23 1 14 75 31 0.9 11 75 23 16 219 50 5 3 44 50 3 0.9 14 50 1 15 294 25 15 1 21 25 1 0.6 9 50 20 9 125 50 12 0.5 11 25 14 0.3 7 25 17 8 83 100 3 0.5 10 25 18 0.2 4 25 6 7 78 75 26 0.2 4 25 21 5 61 75 16 2 42 25 2 1 19 25 79 Meloidogyne spp. were the predominant endoparasitic genus identified from 5g roots of crops and grasses sampled during both the 2010 and 2011 growing seasons (Table 2.8). RKN J2 were present in 10 and seven of the 31 fields sampled during the 2010 and 2011 growing seasons, respectively, and were identified as M. incognita (fields 20 and 3) and M. javanica (fields 16 and 26) using molecular techniques (see Chapter 3). Mature RKN females from fields 1, 2, 6, 15, 17, 21 and 31 could not be obtained for molecular or morphological identification purposes for the purpose of this study (see Chapter 3). During 2010 Meloidogyne spp. were followed only by individuals of the Hoplolaimidae family in terms of predominance, while Pratylenchus and Ditylenchus genera ranked second and third in 2011 after Meloidogyne spp. (Table 2.8). PV for Meloidogyne spp. ranged from 1 to 45, PD from 19 to 450 and FO from 25% to 100%. During 2010 individuals of the Hoplolaimidae were present only in roots of crop plants from 2 of the 31 fields sampled and had low PV (0.5 to 2) and PD (10 to 33) with individuals occurring in only 25% of the samples collected. During 2011 PV for Meloidogyne spp. ranged from 0.2 to 39, PD from 4 to 451 and FO from 25% to 75% (Table 2.8). PV, PD and FO for Pratylenchus and Ditylenchus spp. were substantially lower with the highest being 5 and 2, 56 and 15 and 75% and 100%, respectively. 80 • Soil (200g) Table 2.9. Data for endoparasitic nematode genera/family in fields of the five producers in the Christiana area sampled during the 2010 and 2011 growing seasons as represented by prominence values (PV), frequency of occurrence (FO; %) and population density (PD) per 200g soil. 2010 Survey 2011 Survey Meloidogyne spp. Pratylenchus spp. Heteroderidae Meloidogyne spp. Pratylenchus spp. Field no PV PD FO Field no PV PD FO Field no PV PD FO Field no PV PD FO Field no PV PD FO 25 111 1112 100 25 34 340 100 31 3 32 100 23 344 3442 25 23 14 137 100 23 68 682 100 22 17 165 100 29 1 20 25 14 77 762 100 25 9 87 100 14 63 630 100 20 12 117 100 5 32 317 100 3 7 67 100 29 46 462 100 24 8 90 75 22 9 90 100 16 6 60 100 20 14 137 100 3 7 67 100 27 3 37 75 6 5 47 100 9 12 117 100 8 6 62 100 18 3 60 25 18 4 40 100 8 4 42 100 16 6 62 100 24 3 60 25 9 3 37 75 22 3 32 100 4 6 57 100 20 2 23 75 20 2 22 100 11 3 37 75 9 5 52 100 1 1 20 50 1 2 30 50 24 3 33 75 10 4 47 75 16 1 20 50 21 1 17 75 31 3 30 75 26 3 32 100 6 1 20 25 27 1 20 50 30 2 27 75 7 1 15 50 10 1 20 25 5 1 20 25 21 2 40 25 19 2 22 100 15 1 20 25 7 1 20 25 15 2 23 75 2 2 23 75 26 1 20 25 8 1 20 25 7 1 15 50 21 2 30 25 25 0.8 10 75 14 1 15 50 4 1 13 75 11 1 20 50 9 0.5 10 25 30 1 20 25 12 0.5 10 25 6 0.7 10 50 29 0.5 10 25 31 0.7 10 50 16 0.5 10 25 12 0.5 10 25 31 0.5 10 25 17 0.5 10 25 28 0.5 10 25 18 0.5 10 25 26 0.5 10 25 23 0.5 10 25 28 0.5 10 25 30 0.5 10 25 29 0.5 10 25 31 0.5 10 25 81 Nematode species identification could only be done for fields of which 5g root and their corresponding soil (200g) samples contained mature specimens of endoparasitic genera and/or families listed in Table 2.9. For the other fields only juvenile specimens were present and in some cases very low numbers of these genera and/or families occurred, which complicated complete identification of individuals to species level. Meloidogyne spp. were the predominant endoparasitic genus identified from 200g soil samples collected from the 31 fields during both the 2010 and 2011 surveys. RKN J2 were present in 19 and 18 of the 31 fields sampled during the 2010 and 2011 seasons, respectively (Table 2.9). During 2010 RKN species identified from these fields were M. incognita (fields 9, 11, 22 and 25) and M. javanica (field 28) using molecular techniques (see Chapter 3), while M. incognita was identified during 2011 only for field 18. Mature RKN females could not be obtained after in vivo rearing of RKN J2 present in soil samples from the fields 1, 4, 6, 7, 8, 10, 12, 15, 21, 24, 27, 30 and 31 in separate pots planted with the tomato cv. Rodade for 56 days. During 2010 Meloidogyne spp. were followed by individuals of Pratylenchus spp. and the Heteroderidae family and in 2011 only by individuals of Pratylenchus spp. in terms of its predominance (Table 2.9). PV for Meloidogyne spp. ranged from 0.5 to 111 and PD from 10 to 1112 with RKN J2 occurring in 25% to 100% of the samples collected during 2010 (Table 2.9). Individuals of Pratylenchus spp. were present in soil from 22 of the 31 fields sampled with lower PV (0.5 to 34) and PD (10 to 340), and occurring in 25% to 100% of the samples collected. Individuals of the Heteroderidae family were present in only two of the 31 fields sampled with very low PV (1 to 3), PD (20 to 32) but FO values of 25% and 100%. During 2011 PV for Meloidogyne spp. ranged from 0.5 to 344, PD from 10 to 3442 with RKN J2 occurring in 25% to 100% of the samples collected (Table 2.9). Pratylenchus spp. was present in 21 of the 31 fields sampled with relatively low PV (0.5 to 14) and PD (10 to 137). Individuals of the latter genus occurred in 25% to 100% of the samples collected. Pratylenchus spp. identified during both seasons were P. zeae (fields 8, 9, 10, 18, 20, 23, 25, 26, 29 and 30) and P. neglectus (field 27). No mature male or female specimens were present in soil samples obtained from the other fields where Pratylenchus individuals occurred (Table 2.9) and could not be identified to species level using morphological characteristics. 82 Table 2.10. Data for semi- endo-/ecto- and ectoparasitic nematode genera/family in fields of five producers in the Christiana area sampled during the 2010 growing season as represented by prominence values (PV), frequency of occurrence (FO; %) and population density (PD) per 200g soil. Trichodoridae Hoplolaimidae Tylenchorhynchus spp. Criconematidae Xiphinema spp. Longidorus spp. Field no PV PD FO Field no PV PD FO Field no PV PD FO Field no PV PD FO Field no PV PD FO Field no PV PD FO 25 5 52 100 6 14 142 100 21 59 587 100 4 7 67 100 26 6 55 100 4 0.5 10 25 21 5 45 100 30 12 122 100 28 20 197 100 14 4 35 100 28 2 30 25 12 0.5 10 25 7 4 37 100 19 9 90 100 26 11 112 100 29 2 20 75 30 1 17 75 18 0.5 10 25 29 4 37 100 4 6 60 100 22 7 67 100 31 2 20 75 4 1 20 25 19 0.5 10 25 30 4 37 100 17 3 30 75 24 7 65 100 5 1 15 50 19 0.5 10 25 23 0.5 10 25 10 3 33 75 28 2 27 75 2 4 50 75 30 1 15 50 22 0.5 10 25 26 0.5 10 25 22 2 22 100 14 2 20 75 19 4 42 100 7 0.7 10 25 29 0.5 10 25 19 2 23 75 16 2 25 50 18 2 27 75 2 0.5 10 25 16 2 20 75 12 2 15 100 17 1 20 25 6 0.5 10 25 8 2 18 100 7 1 20 50 6 0.5 10 25 12 0.5 10 25 3 2 30 25 22 1 17 75 10 0.5 10 25 19 0.5 10 25 20 1 12 100 26 1 20 50 25 0.5 10 25 17 1 20 25 31 1 13 75 24 1 20 25 2 0.7 10 50 12 1 15 50 8 0.5 10 25 26 1 15 50 10 0.5 10 25 6 1 13 75 31 1 13 75 11 1 10 100 4 0.8 10 75 2 0.7 10 50 9 0.5 10 25 23 0.5 10 25 27 0.5 10 25 28 0.8 10 75 83 Nematode species identification could only be done for fields of which 5g root and their corresponding soil (200g) samples contained mature specimens of both ectoparasitic and semi-endo-/ectoparasitic genera and/or families listed in Table 2.10. For the other fields only juvenile specimens were present and in some cases very low numbers of these genera and/or families occurred, which complicated complete identification of individuals to species level. Semi-endo-/ectoparasitic genera present in 200g soil samples from the 31 fields surveyed during 2010 were individuals belonging to the Hoplolaimidae family (Table 2.10). Ectoparasites were represented by individuals from the Trichodoridae and Criconematidae families as well as individuals of the Longidorus, Tylenchorhynchus, and Xiphinema genera. Tylenchorhynchus spp. was the predominant genus and was present in 12 of the 31 fields sampled, followed by Hoplolaimidae, Criconematidae, Xiphinema, Trichodoridae and Longidorus spp. PV for Tylenchorhynchus spp. ranged from 0.5 to 59 and PD from 10 to 587. Individuals of this genus were present in 25% to 100% of the samples collected and represented T. brevilineatus and T. ventralis (field 28). Mature specimens of Tylenchorhynchus spp, occurring in the soil from the other fields could not be found for morphological identification. Individuals of the Hoplolaimidae family were present in 16 of the 31 fields sampled with PV ranging from 0.5 to 14, PD from 10 to 142, occurring in 25% to 100% of the samples collected. Individuals of this family were represented by Scutellonema brachyurus (field 8) and S. truncatum (field 30), Rotylenchus unisex (field 30) as well as R. caudaphasmidis (field 30). Mature specimens of Hoplolaimidae occurring in the soil from the other fields could not be found for morphological identification. Individuals belonging to the family Criconematidae occurred in 11 of the 31 fields sampled with PV ranging from 0.5 to 7, PD from 10 to 67 and FO from 25% to 100%. The Criconematidae family was represented by Criconema mutabile (field 7), Criconemoides sphaerocephalus (fields 4, 6, 7, 11, 14 and 15) and Hemicriconemoides brachyurus (fields 29 and 31). Individuals from the Trichodoridae family represented the genera Paratrichodorus and Nanidorus and occurred in 25 of the 31 fields sampled. For Trichodoridae (data pooled for the two genera) PV ranged from 0.8 to 5, PD from 10 to 52 and FO from 75% to 100%. The latter genera were represented by Nanidorus minor (fields 10, 19, 23 and 30) and Paratrichodorus lobatus (field 25). Xiphinema and Longidorus spp. occurred in seven and six of the 31 fields sampled, respectively. PV for Xiphinema spp. ranged from 0.5 to 6, PD from 10 to 55 and FO from 25% to 100%, while PV for Longidorus spp. was 0.5, PD 10 and the FO 25%. 84 Table 2.11. Data for semi- endo-/ecto- and ectoparasitic nematode genera/family in fields of five producers in the Christiana area sampled during the 2011 growing season as represented by prominence values (PV), frequency of occurrence (FO; %) and population density (PD) per 200g soil. Trichodoridae Criconematidae Tylenchorhynchus spp. Hoplolaimidae Xiphinema spp. Psilenchus spp. Field no PV PD FO Field no PV PD FO Field no PV PD FO Field no PV PD FO Field no PV PD FO Field no PV PD FO 14 105 104 100 6 2 20 100 21 6 62 100 26 2 30 50 27 1 13 75 21 6 66 75 16 8 75 100 3 1 16 75 3 6 55 100 22 2 30 25 6 0.5 10 25 13 4 50 50 23 5 57 75 7 1 20 50 29 3 27 100 2 1 15 50 12 0.5 10 25 10 5 50 100 14 1 20 50 5 3 30 75 5 0.7 10 50 16 0.5 10 25 5 5 45 100 15 1 15 50 14 2 20 100 3 0.5 10 25 17 0.5 10 25 1 2 20 100 19 1 20 25 27 2 23 75 6 0.5 10 25 19 0.5 10 25 3 2 25 50 5 0.7 10 50 31 1 20 50 15 0.5 10 25 21 0.5 10 25 29 2 25 50 2 0.5 10 25 9 1 20 25 16 0.5 10 25 18 2 15 100 29 0.5 10 25 23 0.5 10 25 20 0.5 10 25 2 1 20 50 31 0.5 10 25 26 0.5 10 25 7 1 20 50 25 1 20 50 8 1 15 50 19 1 13 75 27 1 20 25 6 0.8 10 75 9 0.8 10 75 20 0.8 10 75 31 0.8 10 75 21 0.7 10 50 15 0.5 10 25 85 Nematode species identification could only be done for fields of which 5g root and their corresponding soil (200g) samples contained mature specimens of both ectoparasitic and semi-endo-/ectoparasitic genera and/or families listed in Table 2.11. For the other fields, only juvenile specimens were present and in some cases very low numbers of these genera and/or families occurred, which complicated complete identification of individuals to species level. Individuals from the Hoplolaimidae family were the only semi-ectoparasitc group identified from 200g soil samples of the 31 fields surveyed during 2011 and were present in 9 of the 31 fields sampled (Table 2.11). Ectoparasitic nematodes present were individuals belonging to the Criconematidae and Trichodoridae families as well as those from the genera Psilenchus and Xiphinema. Trichodoridae spp. (numbers pooled) were predominant, followed by Psilenchus spp., Tylenchorhynchus spp., Hoplolaimidae, Criconematidae and Xiphinema spp. Trichodoridae individuals were present in 21 of the 31 fields sampled, with PV ranging from 0.5 to 105, PD from 10 to 104 and FO from 25% to 100% (Table 2.11). Psilenchus spp. individuals were present in only 2 of the 31 fields with PV ranging between 4 and 6, PD between 50 and 66 and occurred in 50% to 75% of the samples collected. Tylenchorhynchus spp. occurred in 10 of the 31 fields sampled with PV ranging from 0.5 to 6, PD from 10 to 62 and individuals being present in 25% to 100% of the samples collected. Individuals of the Criconematidae family also occurred in 10 of the 31 fields sampled, with PV ranging from 0.5 to 2, PD from 10 to 20 and FO from 25% to 100%. Individuals of the Hoplolaimidae family occurred in 9 of the 31 fields sampled, with PV ranging from 0.5 to 2, PD from 10 to 30 and FO from 25% to 50%. Xiphinema spp. occurred in seven of the 31 fields sampled, with PV ranging from 0.5 to 1, PD from 10 to 13 and FO from 25% to 75%. 86 Table 2.12. Data for bacterivorous nematode genera in fields of five producers in the Christiana area sampled during the 2011 growing season as represented by prominence values (PV), frequency of occurrence (FO; %) and population density (PD) per 200g soil. Rhabditis spp. Acrobeles spp. Cephalobus spp. Eucephalobus spp. Prismatolaimus spp. Acrobeloides spp. Field no PV PD FO Field no PV PD FO Field no PV PD FO Field no PV PD FO Field no PV PD FO Fiel d no PV PD FO 7 1104 11042 100 10 24 282 75 12 13 131 100 16 8 150 25 25 3 39 50 11 1 20 25 12 661 6610 100 12 23 225 100 7 13 129 100 10 6 70 75 28 180 2084 75 29 21 205 100 15 8 118 50 8 1 24 25 30 159 1594 100 7 19 193 100 5 6 75 75 13 1 24 25 3 80 798 100 3 14 135 100 10 4 38 100 17 68 681 100 9 13 133 100 3 4 41 75 2 61 611 100 20 13 150 75 9 3 50 25 15 54 541 100 2 13 150 75 4 1 24 25 4 49 488 100 23 12 175 50 1 1 22 25 22 41 408 100 8 9 100 75 22 0.9 18 25 1 38 382 100 21 7 83 75 6 0.5 11 25 13 38 534 50 31 6 126 25 10 28 277 100 18 4 63 50 11 24 236 100 22 4 61 50 27 22 223 100 16 4 44 75 6 20 200 100 25 4 76 25 21 18 208 75 11 3 56 25 16 17 165 100 15 3 32 75 26 12 150 75 17 3 37 50 23 10 100 100 6 3 50 25 8 10 138 50 30 3 50 25 18 7 83 75 14 2 45 25 14 7 79 75 5 2 24 75 19 7 66 100 4 1 24 25 5 6 66 75 13 1 24 25 31 6 56 100 24 1 22 25 24 3 36 75 1 1 20 25 9 2 26 50 19 1 20 25 20 1 24 25 2.4.1.2 Non-parasitic nematode (NPN) data • Soil (200g) 87 NPN assemblages were identified to genus and family level (Tables 2.12 and 2.13) only during the 2011 survey. Non-parasitic, bacterivorous nematodes in 200g soil samples from the 31 fields sampled during 2011 were represented by Acrobeles, Acrobeloides, Cephalobus, Eucephalobus, Prismatolaimus and Rhabditis spp. (Table 2.12). Rhabditis spp. was the predominant genus of which individuals were present in 29 of the 31 fields sampled, followed by Acrobeles, Cephalobus, Eucephalobus, Prismatolaimus and Acrobeloides spp. (Table 2.12). The PV for Rhabditis spp. was substantially higher than that for any of the other bacterivorous genera and ranged from 1 to 1104 and PD from 24 to 11042 (Table 2.12). Individuals of the latter genera were present in 25% to 100% of the samples collected. Acrobeles spp. individuals occurred in 28 of the 31 fields sampled with PV that ranged from 1 to 24, PD from 20 to 282 and individuals being present in 25% to 100% of the samples. Individuals of Cephalobus and Eucephalobus spp. occurred in 11 and 4 of the 31 fields sampled respectively. Cephalobus spp. had PV that ranged from 0.5 to 31, PD from 11 to 131 and FO from 25% to 100%, while PV for Eucephalobus spp. ranged from 1 to 8, PD from 24 to 150 and FO from 25% to 75%. The least dominant bacterivorous genera were Prismatolaimus and Acrobeloides spp. with low PV (3 and 1), PD (39 and 20) and FO (50% and 25%), respectively. 88 Table 2.13. Data for fungivorous and predator nematode genera/family in fields of five producers in the Christiana area sampled during the 2011 growing season as represented by prominence values (PV), frequency of occurrence (FO; %) and population density (PD) per 200g soil. Fungivores Predators Tylenchus spp. Aphelenchus spp. Aphelenchoides spp. Dorylaimidae Mononchidae Field no PV PD FO Field no PV PD FO Field no PV PD FO Field no PV PD FO Field no PV PD FO 18 37 369 100 10 12 175 50 29 13 127 100 10 21 300 50 2 13 251 25 16 14 157 75 3 5 73 50 18 16 163 100 9 5 52 100 27 13 130 100 30 3 50 25 11 15 145 100 15 9 91 100 6 2 19 75 2 14 167 75 11 8 167 25 28 1 24 25 29 12 122 100 3 7 143 25 23 11 113 100 23 7 100 50 15 9 103 75 20 6 67 75 16 7 105 50 10 5 100 25 30 7 100 50 13 5 100 25 3 6 85 50 7 4 53 50 22 6 57 100 14 3 49 50 5 6 75 50 12 3 50 25 20 5 58 75 8 3 37 50 21 4 43 100 25 3 37 50 13 4 76 25 1 1 21 50 28 4 76 25 9 1 27 25 31 4 76 25 5 1 24 25 4 3 50 25 6 1 24 25 8 3 50 25 17 1 24 25 9 3 50 25 22 1 24 25 17 3 50 25 19 3 33 50 1 2 45 25 14 2 24 50 25 2 24 50 24 1 22 25 89 Non-parasitic, fungivorous nematode genera that were present in 200g soil samples collected from 31 fields during 2011 were Aphelenchus, Tylenchus and Aphelenchoides species. Predator nematodes in the same samples were represented by the Dorylaimidae and Mononchidae families (Table 2.13). Tylenchus was the predominant fungivore genus of which individuals were present in 21 of the 31 fields sampled, followed by Aphelenchus and Aphelenchoides spp. (Table 2.12). Tylenchus spp. had PV that ranged from 1 to 37, PD from 24 to 369 and occurred in 25% to 100% of the samples collected. Individuals of Aphelenchoides occurred in only one of the 31 fields sampled and had a PV of 13, PD of 127 with individuals being present in all four samples (100%) from this field. Aphelenchus spp. was the least dominant and occurred in five of the 31 fields sampled with PV that ranged from 1 to 12, PD from 24 to 175 and FO from 25% to 50%. For the predator nematode families identified individuals of the Dorylaimidae family were predominant and occurred in 26 of the 31 fields sampled with PV ranging from 1 to 21, PD form 22 to 300 and FO form 25% to 100% (Table 2.13). Individuals of the Mononchidae family only occurred in 2 of the 31 fields sampled with PV of 5 and 13, PD of 52 and 251 and FO of 100% and 25%, respectively. 90 2.4.2 Parametric and non-parametric statistics 2.4.2.1 Plant-parasitic nematode (PPN) data • Roots (50g and 5g) Table 2.14. Population levels [log10(x+1) transformed] for Meloidogyne species per 50g roots for cultivation practices, nematicides and crop rotation sequences when data were pooled for 31 fields of five producers for the 2011 growing season (means in parenthesis represents the values that were back transformed and not the real means). a1=plough :2=rip & plough; b1=EDBฎ only :2=nematicides other than EDBฎ :3=EDBฎ & other nematicides (combined); c1=grass and potato :2=grass, maize & potato :3=maize, wheat & potato :4=maize and/or grass and potato Nematode genus Cultivation practicesa Nematicidesb Crop rotation sequencesc 1 2 1 2 3 1 2 3 4 Meloidogyne spp. 1.6 ฑ3 (4 ฑ19) 1 ฑ2 (2 ฑ6) 0 2.2 ฑ4.4 (8 ฑ80) 1.3 ฑ2.2 (3 ฑ8) 0.7 ฑ1.5 (1 ฑ3) 1.7 ฑ2.7 (5 ฑ14) 2.2 ฑ4.4 (8 ฑ80) 0 P values Mann-Whitney U: 0.7668 Kruskal-Wallis: 0.6653 Kruskal-Wallis: 0.5481 T-test: 0.531 Anova: 0.5925 Anova: 0.5155 91 Table 2.15. Population levels [log10(x+1) transformed] for endo- and semi-endoparasitic nematode genera/family per 5g roots as well as for cultivation practices, nematicides and crop rotation sequences when data were pooled for 31 fields of five producers for the 2010 growing season (means in parenthesis represents the values that were back transformed and not the real means). Nematode genus/family Cultivation practicesa Nematicidesb Crop rotation sequencesc 1 2 1 2 3 1 2 3 4 Meloidogyne spp. 1 ฑ1.8 (2 ฑ5) 1.4 ฑ2 (3 ฑ6) 2 ฑ2.9 (6 ฑ17) 2.1 ฑ2.5 (7 ฑ11) 1 ฑ1.8 (2 ฑ5) 0.4 ฑ1.3 (1 ฑ3) 1.9 ฑ2 (6 ฑ6) 1.1 ฑ2.3 (2 ฑ9) 1.4 ฑ2.4 (3 ฑ10) P values Mann-Whitney U: 0.678 Kruskal-Wallis: 0.4839 Kruskal-Wallis: 0.2498 T-test: 0.521 Anova: 0.4486 Anova: 0.3264 Hoplolaimidae 0.14 ฑ0.6 (0.1 ฑ0.8) 0 0 0 0.09 ฑ0.4 (0.09 ฑ0.5) 0 0.1 ฑ0.6 (0.1 ฑ0.8) 0 0 P values Mann-Whitney U: 0.7670 Kruskal-Wallis: 0.8869 Kruskal-Wallis: 0.7273 T-test: 0.3090 Anova: 0.8936 Anova: 0.7476 Total PPNd 1.1 ฑ1.8 (2 ฑ5) 1.4 ฑ2 (3 ฑ6) 2 ฑ2.9 (6 ฑ17) 2.1 ฑ2.5 (7 ฑ11) 1.1 ฑ1.8 (2 ฑ5) 0.4 ฑ1.3 (0.5 ฑ3) 2.1 ฑ1.9 (7 ฑ6) 1.1 ฑ2.3 (2 ฑ9) 1.4 ฑ2.4 (3 ฑ10) P values Mann-Whitney U: 0.8278 Kruskal-Wallis: 0.5570 Kruskal-Wallis: 0.1572 T-test: 0.6684 Anova: 0.5016 Anova: 0.2206 a1=plough :2=rip & plough; b1=EDBฎ only :2=nematicides other than EDBฎ :3=EDBฎ & other nematicides (combined); c1=grass and potato :2=grass, maize & potato :3=maize, wheat & potato :4=maize and/or grass and potato; dTotal PPN=Meloidogyne spp. + Hoplolaimidae 92 Table 2.16. Population levels [log10(x+1) transformed] for endoparasitic nematode genera/family per 5g roots as well as for cultivation practices, nematicides and crop rotation sequences when data were pooled for 31 fields of five producers for the 2011 growing season (means in parenthesis represents the values that were back transformed and not the real means). Nematode genera Cultivation practicesa Nematicidesb Crop rotation sequencesc 1 2 1 2 3 1 2 3 4 Meloidogyne spp. 0.7 ฑ1.6 (1 ฑ4) 0.3 ฑ1 (0.3 ฑ2) 0 1.4 ฑ3 (3 ฑ19) 0.4 ฑ1 (0.5 ฑ2) 0.2 ฑ0.6 (0.2 ฑ0.8) 0.6 ฑ1.2 (0.8 ฑ2) 1.4 ฑ3 (3 ฑ19) 0 P values Mann-Whitney U: 0.7072 Kruskal-Wallis: 0.7144 Kruskal-Wallis: 0.6955 T-test: 0.4667 Anova: 0.3176 Anova: 0.4255 Pratylenchus spp. 0.6 ฑ1.1 (0.8 ฑ2) 0.1 ฑ0.3 (0.1 ฑ0.3) 0 0.6 ฑ1.2 (0.8 ฑ2) 0.3 ฑ0.8 (0.3 ฑ1) 0.3 ฑ1.2 (0.3 ฑ2) 0.4 ฑ0.6 (0.5 ฑ0.8) 0.6 ฑ1.2 (0.8 ฑ2) 0 P values Mann-Whitney U: 0.4177 Kruskal-Wallis: 0.7343 Kruskal-Wallis: 0.6047 T-test: 0.0957 Anova: 0.7328 Anova: 0.8615 Ditylenchus spp. 0 0.3 ฑ1 (0.3 ฑ2) 0 0 0.2 ฑ0.7 (0.2 ฑ1) 0.5 ฑ1 (0.6 ฑ2) 0 0 0 P values Mann-Whitney U: 0.5665 Kruskal-Wallis: 0.7804 Kruskal-Wallis: 0.2470 T-test: 0.1710 Anova: 0.7946 Anova: 0.2602 Total PPNd 1.2 ฑ1.8 (2 ฑ5) 0.7 ฑ1.2 (1 ฑ2) 0 1.4 ฑ3 (3 ฑ19) 1 ฑ1.3 (2 ฑ3) 1.1 ฑ1.4 (2 ฑ3) 1 ฑ1.2 (2 ฑ2) 1.4 ฑ3 (3 ฑ19) 0 P values Mann-Whitney U: 0.7518 Kruskal-Wallis: 0.5199 Kruskal-Wallis: 0.5185 T-test: 0.4315 Anova: 0.5472 Anova: 0.6469 a1=plough: 2=rip & plough; b1=EDBฎ only :2=other nematicides than EDBฎ :3=EDBฎ & other nematicides; c1=grass and potato :2=grass, maize & potato :3=maize, wheat & potato :4=maize and/or grass and potato ; dTotal PPN=Meloidogyne spp. + Pratylenchus spp. + Ditylenchus spp. 93 The adapted NaOCl method was only used to extract eggs and J2 from 50g roots of crops and grasses sampled during 2011 (Table 2.14). When RKN numbers were pooled for the 31 fields, none of the respective cultivation practices, nematicides or crop rotation sequences applied had a significant (P?0.05) effect on RKN numbers/50g roots of crops and grasses sampled (Table 2.14). This trend was true when the log10(x+1) transformed data were subjected to both non-parametric (Kruskal-Wallis) and parametric (Mann-Whitney U, T-test and Anova) statistical analyses. Only individuals of the Meloidogyne genus and Hoplolaimidae family were present in 5g root samples collected from crops planted on 31 fields of five producers during 2010 (Table 2.15). The respective cultivation practices, nematicides and crop rotation sequences applied on the 31 fields did not have a significant (P?0.05) effect on population levels of either Meloidogyne or Hoplolaimidae or the total PPN, represented by pooled population levels [log10(x+1) transformed] of the latter genus and family when data were subjected to parametric and non-parametric statistics. During 2011 individuals of Ditylenchus, Meloidogyne and Pratylenchus spp. were present in 5g root samples from crops planted on 31 fields of five producers (Table 2.16). No significant (P?0.05) effect(s) were obtained for the cultivation practices, nematicides or crop rotation sequences applied in terms of nematode numbers of the three endoparasitic nematode genera as well as total PPN when data were subjected to both non-parametric (Kruskal-Wallis) and parametric (Mann- Whitney U, T-test and Anova) statistical analyses. 94 • Soil (200g) Table 2.17. Population levels [log10(x+1) transformed] for endoparasitic nematode genera/family per 200g soil as well as cultivation practices, nematicides and crop rotation sequences when data were pooled for 31 fields of five producers during the 2010 growing season (means in parenthesis represents the values that were back transformed and not the real means). Nematode genera/family Cultivation practicesa Nematicidesb Crop rotation sequencesc 1 2 1 2 3 1 2 3 4 Meloidogyne spp. 2 ฑ2.1 (6 ฑ7) 2.4 ฑ2.5 (10 ฑ11) 6 ฑ1.5a (402 ฑ3) 4 ฑ1.8ab (53 ฑ5) 1.6 ฑ2b (4 ฑ6) 2.1 ฑ2.2ab (7 ฑ8) 1 ฑ1.8a (2 ฑ5) 5 ฑ1.9b (147 ฑ6) 4 ฑ0.8ab (53 ฑ1) P values Mann-Whitney U: 0.9056 Kruskal-Wallis: 0.0128 Kruskal-Wallis: 0.0105 T-test: 0.6849 Anova: 0.0073 Anova: 0.0125 Pratylenchus spp. 2.7 ฑ1.6 (14 ฑ4) 1.7 ฑ2 (4 ฑ6) 5.3 ฑ0.7a (199 ฑ1) 3.2 ฑ1.8ab (23 ฑ5) 1.8 ฑ1.6b (5 ฑ4) 1.4 ฑ1.7 (3 ฑ4) 1.9 ฑ1.6 (6 ฑ4) 4.1 ฑ2 (59 ฑ6) 3.7 ฑ1.2 (39 ฑ2) P values Mann-Whitney U: 0.1138 Kruskal-Wallis: 0.0232 Kruskal-Wallis: 0.0580 T-test: 0.1368 Anova: 0.0154 Anova: 0.0625 Heteroderidae 0 0.3 ฑ1 (0.3 ฑ2) 0 0 0.2 ฑ0.8 (0.2 ฑ1) 0.5 ฑ1.2 (0.6 ฑ2) 0 0 0 P values Mann-Whitney U: 0.5665 Kruskal-Wallis: 0.7804 Kruskal-Wallis: 0.3619 T-test: 0.1913 Anova: 0.8106 Anova: 0.4391 a1=plough :2=rip & plough; b1=EDBฎ only :2=nematicides other than EDBฎ :3=EDBฎ & other nematicides (combined); c1=grass and potato :2=grass, maize & potato :3=maize, wheat & potato :4=maize and/or grass and potato 95 Table 2.18. Population levels [log10(x+1) transformed] for endoparasitic nematode genera/family per 200g soil as well as cultivation practices, nematicides and crop rotation sequences when data were pooled for 31 fields of five producers during the 2011 growing season (means in parenthesis represents the values that were back transformed and not the real means). 2011 Survey Nematode genera Cultivation practicesa Nematicidesb Crop rotation sequencesc 1 2 1 2 3 1 2 3 4 Meloidogyne spp. 1.9 ฑ2.5 (6 ฑ11) 1.6 ฑ1.8 (4 ฑ5) 2.5 ฑ0.5 (11 ฑ0.6) 3.8 ฑ3.4 (44 ฑ29) 1.3 ฑ1.8 (3 ฑ5) 1 ฑ1.1 (2 ฑ2) 1.5 ฑ2.3 (3 ฑ9) 4.4 ฑ2.7 (80 ฑ14) 1.5 ฑ1.4 (3 ฑ3) P values Mann-Whitney U: 0.8899 Kruskal-Wallis: 0.8650 Kruskal-Wallis: 0.9540 T-test: 0.7507 Anova: 0.0788 Anova: 0.0597 Pratylenchus spp. 1.6 ฑ1.8 (4 ฑ5) 1.9 ฑ1.3 (6 ฑ3) 3.8 ฑ0.9 (44 ฑ1) 1.9 ฑ2.3 (6 ฑ9) 1.6 ฑ1.4 (4 ฑ3) 1.5 ฑ1 (3 ฑ2) 1.7 ฑ1.6 (4 ฑ4) 2.3 ฑ2.7 (9 ฑ14) 1.6 ฑ1.6 (4 ฑ4) P values Mann-Whitney U: 0.5799 Kruskal-Wallis: 0.1929 Kruskal-Wallis: 0.9524 T-test: 0.4530 Anova: 0.1529 Anova: 0.8383 a1=plough :2=rip & plough; b1=EDBฎ only :2=nematicides other than EDBฎ :3=EDBฎ & other nematicides (combined); c1=grass and potato :2=grass, maize & potato :3=maize, wheat & potato :4=maize and/or grass and potato 96 Individuals of the Heteroderidae family as well as from the Meloidogyne and Pratylenchus genera were present in 200g soil samples obtained from five producers in the Christiana area during 2010 (Table 2.17). Significant differences (P?. were obtained for both Meloidogyne and Pratylenchus spp. in terms of nematicides applied and only for Meloidogyne spp. in terms of crop rotation sequences implemented (Table 2. 17). Nematicides application of EDBฎ combined with other nematicides (category 3) resulted in significantly lower (P?. 5K1 individuals/5g roots compared to EDBฎ application only (category 1), but it did not differ significantly (P?. from Fategor\ , whiFh inFlXded appliFation of nematicides other than EDBฎ. Categories 1 and 2 as well as 2 and 3 did, however, not differ significantly from each other with regard to nematicide application for both RKN and Pratylenchus spp. For cropping sequences, grass, maize and potato included together in a cycle (category 2) resulted in the lowest RKN population levels/200g soil and differed significantly (P?. from mai]e, wheat and potato (category 3) but not from cycles including grass and potato (category 1) and maize and/or grass and potato (category 4). No significant differences were obtained for RKN between categories 1 and 4 with regard to cropping sequences. No significant correlation (P?. , were obtained for the cultivation practices, nematicides or crop rotation sequences applied on the 31 fields using both non-parametric (Kruskal-Wallis) and parametric (Mann-Whitney U, T-test and Anova) statistical analyses for Heteroderidae (Table 2.17). During the 2011 survey Meloidogyne and Pratylenchus spp. were identified from 200g soil samples (Table 2.18). No significant effect (P?. was obtained for both Meloidogyne and Pratylenchus spp. numbers with regard to the different cultivation practices, nematicides or crop rotation sequences applied on the 31 fields. The latter trend was evident for data subjected to both non-parametric (Kruskal-Wallis) and parametric (Mann-Whitney U, T-test and Anova) statistical analyses. 97 Table 2.19. Population levels [log10(x+1) transformed] for semi- endo-/ecto- and ectoparasitic nematode genera/family per 200g soil for cultivation sequences, nematicides, crop rotation sequences when data were pooled for 31 fields of five producers for the 2010 growing season (means in parenthesis represents the values that were back transformed and not the real means). Nematode genera/families Cultivation practicesa Nematicidesb Crop rotation sequencesc 1 2 1 2 3 1 2 3 4 Hoplolaimidae 2.9 ฑ2 (17 ฑ6) 2.5 ฑ2.1 (11 ฑ7) 6 ฑ1.5a (402 ฑ3) 4 ฑ1.8ab (53 ฑ5) 2.2 ฑ1.8b (8 ฑ5) 1.8 ฑ1.9b (5 ฑ6) 2.2 ฑ1.8b (8 ฑ5) 5.1 ฑ1.9a (163 ฑ6) 4 ฑ0.8b (53 ฑ1) P values Mann-Whitney U: 0.4291 Kruskal-Wallis: 0.0318 Kruskal-Wallis: 0.0508 T-test: 0.5992 Anova: 0.0108 Anova: 0.0373 Trichodoridae 2 ฑ1.1 (6 ฑ2) 2 ฑ1.4 (6 ฑ3) 3.3 ฑ1 (26 ฑ2) 2.5 ฑ1.2 (11 ฑ2) 1.8 ฑ1.2 (5 ฑ2) 2 ฑ1.3 (6 ฑ2) 1.6 ฑ1.3 (4 ฑ3) 2.5 ฑ1.2 (11 ฑ2) 2.6 ฑ0.2 (12 ฑ0.2) P values Mann-Whitney U: 0.7220 Kruskal-Wallis: 0.2015 Kruskal-Wallis: 0.3843 T-test: 0.9121 Anova: 0.2304 Anova: 0.4204 Tylenchorhynchus spp. 1 ฑ1.6 (2 ฑ4) 1.6 ฑ2.2 (4 ฑ8) 0.6 ฑ0.9a (0.8 ฑ1) 3.7 ฑ2.6b (39 ฑ12) 1 ฑ1.7a (2 ฑ4) 1.1 ฑ2 (2 ฑ6) 1.5 ฑ2 (3 ฑ6) 2.4 ฑ2.1 (10 ฑ7) 0 P values Mann-Whitney U: 0.4890 Kruskal-Wallis: 0.0984 Kruskal-Wallis: 0.3428 T-test: 0.3544 Anova: 0.0285 Anova: 0.5426 Criconematidae 0.8 ฑ1.2 (1 ฑ2) 0.8 ฑ1.3 (1.2 ฑ2) 0 0 1 ฑ1.3 (1.7 ฑ2) 0.7 ฑ1.2 (1 ฑ2) 1.1 ฑ1.4 (2 ฑ3) 0 0 P values Mann-Whitney U: 0.8743 Kruskal-Wallis: 0.1547 Kruskal-Wallis: 0.1835 T-test: 0.9791 Anova: 0.2227 Anova: 0.3357 98 Nematode genera/families Cultivation practicesa Nematicidesb Crop rotation sequencesc 1 2 1 2 3 1 2 3 4 Xiphinema spp. 0.2 ฑ0.5 (0.2 ฑ0.6) 0.7 ฑ1.2 (1 ฑ2) 0 0.3 ฑ0.6 (0.3 ฑ0.8) 0.5 ฑ1 (0.6 ฑ2) 1 ฑ1.4 (1.7 ฑ3) 0.2 ฑ0.6 (0.2 ฑ0.8) 0.3 ฑ0.6 (0.3 ฑ0.8) 0 P values Mann-Whitney U: 0.3632 Kruskal-Wallis: 0.7387 Kruskal-Wallis: 0.4406 T-test: 0.1581 Anova: 0.7425 Anova: 0.3298 Longidorus spp. 0.2 ฑ0.5 (0.2 ฑ0.6) 0.2 ฑ0.5 (0.2 ฑ0.6) 0 0.3 ฑ0.6 (0.3 ฑ0.8) 0.2 ฑ0.5 (0.2 ฑ0.6) 0.2 ฑ0.4 (0.2 ฑ0.5) 0.4 ฑ0.6 (0.5 ฑ0.8) 0.3 ฑ0.6 (0.3 ฑ0.8) 0 P values Mann-Whitney U: 0.9684 Kruskal-Wallis: 0.7596 Kruskal-Wallis: 0.6267 T-test: 0.9327 Anova: 0.7717 Anova: 0.6543 Total PPNd 4.3 ฑ1.5 (72 ฑ3) 4.6 ฑ2 (98 ฑ6) 6.4 ฑ1.2ab (601 ฑ2) 6 ฑ0.7a (402 ฑ1) 4 ฑ1.7b (53 ฑ4) 4 ฑ2 (53 ฑ6.4) 4.1 ฑ1.8 (59 ฑ5) 6.1 ฑ1 (445 ฑ2) 4.7 ฑ0.8 (109 ฑ1) P values Mann-Whitney U: 0.3529 Kruskal-Wallis: 0.0083 Kruskal-Wallis: 0.1969 T-test: 0.6308 Anova: 0.0275 Anova: 0.3292 a1=plough :2=rip & plough; b1=EDBฎ only :2=nematicides other than EDBฎ :3=EDBฎ & other nematicides (combined); c1=grass and potato :2=grass, maize & potato :3=maize, wheat & potato :4=maize and/or grass and potato; dTotal PPN=Meloidogyne spp. + Pratylenchus spp.+ Heteroderidae + Hoplolaimidae + Trichodoridae + Tylenchorhynchus spp. + Criconematidae + Xiphinema spp. + Longodorus spp. 99 For the semi-endo-/ectoparasitic nematode genera/family per 200g soil, individuals of the Hoplolaimidae, Criconematidae and Trichodoridae families as well as those from the Tylenchorhynchus, Xiphinema and Longidorus genera were present (Table 2.19). There was a significant (P?. effect evident for Hoplolaimidae, Tylenchorhynchus spp. and total PPN with regard to the different nematicides applied. Hoplolaimidae as well as total PPN numbers were the lowest in soil of fields where EDBฎ and other nematicides were used in combination (category 3) and differed significantly (P?. from those where only EDBฎ (category 1) was applied, but not from those where nematicides other than EDBฎ (category 2) were applied. The latter scenario was true for both the non-parametric Kruskal-Wallis and parametric Anova tests. Tylenchorhynchus spp. numbers were significantly lower in fields where EDBฎ only (category 1) as well as where EDBฎ and other nematicides were applied in combination (category 3) and differed significantly from those for fields treated with nematicides other than EDBฎ (category 2). The latter trend was, however, evident for the parametric Anova and not for the non-parametric Kruskal-Wallis test. The latter implies that this significance should rather be ignored since the latter non-parametric test is more strict than the Anova and caters for uneven distributed data such as those generally obtained for nematodes. For crop rotation practices, significant (P?. differenFes were obtained among the varioXs sequences practiced and Hoplolaimidae numbers/200g soil, but this was only true for the Anova test and is thus ignored as explained above. In terms of cultivation practices, no significant correlation (P?. were obtained for the 31 fields either for Xiphinema or Longidorus spp. as well as for individuals from the Trichodoridae and Criconematidae families and total PPN using both non-parametric (Kruskal-Wallis) and parametric (Mann-Whitney U, T-test and Anova) statistical analyses. 100 Table 2.20. Population levels [log10(x+1) transformed] for semi-endo-/ecto- and ectoparasitic nematode genera/family per 200g soil for cultivation sequences, nematicides, crop rotation sequences when data were pooled for 31 fields of five producers for the 2011 growing seasons (means in parenthesis represents the values that were back transformed and not the real means). Nematode genera/families Cultivation practicesa Nematicidesb Crop rotation sequencesc 1 2 1 2 3 1 2 3 4 Trichodoridae 1.9 ฑ1.5 (6 ฑ3) 1.9 ฑ1.8 (6 ฑ5) 2.2 ฑ0.2 (8 ฑ0.2) 1.4 ฑ1.8 (3 ฑ5) 1.9 ฑ1.7 (6 ฑ4) 1.4 ฑ1.3 (3 ฑ3) 2.5 ฑ1.9 (11 ฑ6) 1.5 ฑ1.8 (3 ฑ5) 1.4 ฑ1.2 (3 ฑ2) P values Mann-Whitney U: 0.7072 Kruskal-Wallis: 0.6889 Kruskal-Wallis: 0.3657 T-test: 0.9329 Anova: 0.7919 Anova: 0.4324 Tylenchorhynchus spp. 0.7 ฑ1.3 (1 ฑ3) 1 ฑ1.5 (2 ฑ3) 0 1.3 ฑ2 (3 ฑ6) 0.8 ฑ1.4 (1 ฑ3) 1.1 ฑ1.3 (2 ฑ3) 1.1 ฑ1.7 (2 ฑ4) 0.3 ฑ0.6 (0.3 ฑ0.8) 0 P values Mann-Whitney U: 0.5799 Kruskal-Wallis: 0.4821 Kruskal-Wallis: 0.4857 T-test: 0.5274 Anova: 0.5619 Anova: 0.5001 Criconematidae 0.7 ฑ1.1 (1 ฑ2) 0.5 ฑ0.9 (0.6 ฑ1) 0 0 0.8 ฑ1 (1 ฑ2) 0.4 ฑ0.8 (0.5 ฑ1) 1 ฑ1.2 (2 ฑ2) 0 0 P values Mann-Whitney U: 0.7668 Kruskal-Wallis: 0.1951 Kruskal-Wallis: 0.1830 T-test: 0.8505 Anova: 0.2254 Anova: 0.1724 Hoplolaimidae 0.5 ฑ.0.8 (0.6 ฑ1) 0.4 ฑ0.8 (0.5 ฑ1) 0.6 ฑ0.9 (0.8 ฑ1) 0.5 ฑ1 (0.6 ฑ2) 0.4 ฑ0.8 (0.5 ฑ1) 0.4 ฑ0.9 (0.5 ฑ1) 0.6 ฑ0.8 (0.8 ฑ1) 0.5 ฑ1 (0.6 ฑ2) 0.4 ฑ0.7 (0.5 ฑ1) P values Mann-Whitney U: 0.6494 Kruskal-Wallis: 0.9016 Kruskal-Wallis: 0.8884 T-test: 0.5137 Anova: 0.9621 Anova: 0.9599 101 Nematode genera/families Cultivation practicesa Nematicidesb Crop rotation sequencesc 1 2 1 2 3 1 2 3 4 Xiphinema spp. 0.1 ฑ0.4 (0.1 ฑ0.5) 0.4 ฑ0.7 (0.5 ฑ1) 0 0.3 ฑ0.6 (0.3 ฑ0.8) 0.3 ฑ0.6 (0.3 ฑ0.8) 0.2 ฑ0.7 (0.2 ฑ1) 0.5 ฑ0.6 (0.6 ฑ0.8) 0 0 P values Mann-Whitney U: 0.3869 Kruskal-Wallis: 0.7413 Kruskal-Wallis: 0.1174 T-test: 0.2316 Anova: 0.7694 Anova: 0.2677 Psilenchus spp. 0 0.4 ฑ1.2 (0.5 ฑ2) 0 1 ฑ2 (2 ฑ6) 0.1 ฑ0.6 (0.1 ฑ0.8) 0.3 ฑ1 (0.3 ฑ2) 0.3 ฑ1.1 (0.3 ฑ2) 0 0 P values Mann-Whitney U: 0.5665 Kruskal-Wallis: 0.2466 Kruskal-Wallis: 0.8836 T-test: 0.1685 Anova: 0.2037 Anova: 0.9009 Total PPNd 6 ฑ0.7a (402 ฑ1) 6.4 ฑ0.6b (600 ฑ0.8) 6.2 ฑ0.2 (492 ฑ0.2) 6.6 ฑ0.4 (734 ฑ0.5) 6.2 ฑ0.7 (492 ฑ1) 6.3 ฑ0.5 (543 ฑ0.6) 6.3 ฑ0.8 (543 ฑ1.2) 6.6 ฑ0.5 (734 ฑ0.6) 5.4 ฑ0.7 (220 ฑ1) P values Mann-Whitney U: 0.0328 Kruskal-Wallis: 0.2537 Kruskal-Wallis: 0.1938 T-test: 0.0893 Anova: 0.5548 Anova: 0.2007 a1=plough :2=rip & plough; b1=EDBฎ only :2=nematicides other than EDBฎ :3=EDBฎ & other nematicides (combined); c1=grass and potato :2=grass, maize & potato :3=maize, wheat & potato :4=maize and/or grass and potato; dTotal PPN= Meloidogyne spp. + Pratylenchus spp.+ Trichodoride + Tylenchorhynchus spp. + Criconematidae + Hoplomaimidae + Xiphinema spp. + Psilenchus spp. 102 During the 2011 survey, for the semi-endo-/ectoparasitic nematodes Tylenchorhynchus, Xiphinema and Psilenchus spp. as well as Trichodoridae, Criconematidae and Hoplolaimidae were present in 200g soil samples (Table 2.20). A significant difference (P?. was evident only between the total PPN and cultivation practices, but only for the Mann-Whitney U test and is therefore ignored and not discussed further (Table 2.20). No significant (P?. effects were evident with regard to any of the cultivation practices, nematicides or cropping sequences applied for any of the PPN present in soil samples obtained during the 2011 season. 103 2.4.2.2 Non-parasitic nematode (NPN) data • Soil (200g) Table 2.21. Population levels [log10(x+1) transformed] for non-parasitic nematode genera/families per 200g soil for cultivation sequences, nematicides and crop rotation sequences applied, data were pooled for 31 fields of five producers for the 2010 growing season (means in parenthesis represents the values that were back transformed and not the real means). Nematode genera/families Cultivation practicesa Nematicidesb Crop rotation sequencesc 1 2 1 2 3 1 2 3 4 Total NPN 6.2 ฑ0.5 (492 ฑ0.6) 6.2 ฑ0.4 (492 ฑ0.5) 6 ฑ0.7 (402 ฑ1) 6.3 ฑ0.4 (543 ฑ0.5) 6.2 ฑ0.4 (491 ฑ0.5) 6 ฑ0.3a (402 ฑ0.3) 6.5 ฑ0.3b (664 ฑ0.3) 6.4 ฑ0.4ab (601 ฑ0.5) 5.7 ฑ0.2a (298 ฑ0.2) P values Mann-Whitney U: 0.6073 Kruskal-Wallis: 0.8981 Kruskal-Wallis: 0.0046 T-test: 0.7509 Anova: 0.7800 Anova: 0.0015 a1=plough :2=rip & plough; b1=EDBฎ only :2=nematicides other than EDBฎ :3=EDBฎ & other nematicides (combined); c1=grass and potato :2=grass, maize & potato :3=maize, wheat & potato :4=maize and/or grass and potato 104 Table 2.22. Population levels [log10(x+1) transformed] for non-parasitic nematode genera/families per 200g soil for cultivation sequences, nematicides and crop rotation sequences applied, data were pooled for 31 fields of five producers for the 2011 growing season (means in parenthesis represents the values that were back transformed and not the real means). Bacterivores Nematode genera/families Cultivation practicesa Nematicidesb Crop rotation sequencesc 1 2 1 2 3 1 2 3 4 Rhabditis spp. 5.6 ฑ1.8 (269 ฑ5) 1.5 ฑ2.2 (3 ฑ8) 1 ฑ1.3a (2 ฑ3) 1.7 ฑ1.1a (4 ฑ2) 5.4 ฑ1.9b (220 ฑ6) 4.9 ฑ2.2 (133 ฑ8) 5.5 ฑ1.4 (243 ฑ3) 3.5 ฑ2.5 (32 ฑ11) 3.9 ฑ1.8 (48 ฑ5) P values Mann-Whitney U: 0.2279 Kruskal-Wallis: 0.0591 Kruskal-Wallis: 0.3511 T-test: 0.1297 ANOVA: 0.0092 ANOVA: 0.2257 Acrobeles spp. 3.8 ฑ 1.3 (43 ฑ3) 2.8 ฑ1.4 (15 ฑ2) 3.8 ฑ1.2 (43 ฑ2) 3.5 ฑ1.1 (32 ฑ2) 3.2 ฑ1.5 (23 ฑ3) 3 ฑ1.9 (19 ฑ6) 3.3 ฑ1.2 (26 ฑ2) 3.2 ฑ1 (23 ฑ2) 3.9 ฑ1 (48 ฑ2) P values Mann-Whitney U: 0.1490 Kruskal-Wallis: 0.8429 Kruskal-Wallis: 0.8992 T-test: 0.0745 Anova: 0.8147 Anova: 0.8137 Cephalobus spp. 2 ฑ1.8a (6 ฑ5) 0.2 ฑ1b (0.2 ฑ2) 0 0.4 ฑ0.9 (0.5 ฑ1) 1.3 ฑ1.8 (2.7 ฑ5) 1 ฑ1.7 (2 ฑ4) 1.3 ฑ1.8 (3 ฑ5) 0.4 ฑ0.9 (0.5 ฑ1) 0 P values Mann-Whitney U: 0.0063 Kruskal-Wallis: 0.4144 Kruskal-Wallis: 0.4719 T-test: 0.0020 Anova: 0.4147 Anova: 0.5212 105 Bacterivores cont. Nematode genera/families Cultivation practices Nematicides Crop rotation sequences 1 2 1 2 3 1 2 3 4 Eucephalobus spp. 0.4 ฑ1.1 (0.5 ฑ2) 0.3 ฑ1 (0.3 ฑ2) 0 0 0.4 ฑ1.1 (0.5 ฑ2) 0.6 ฑ1.3 (0.8 ฑ3) 0.3 ฑ1 (0.3 ฑ2) 0 0.6 ฑ1.1 (0.8 ฑ2) P values Mann-Whitney U: 0.9684 Kruskal-Wallis: 0.5883 Kruskal-Wallis: 0.5517 T-test: 0.9066 Anova: 0.6375 Anova: 0.7659 Prismatolaimus spp. 0 0.4 ฑ1 (0.5 ฑ2) 1.5 ฑ2.1ab (3 ฑ7) 0a 0.1 ฑ0.6b (0.1 ฑ0.8) 0.3 ฑ1 (0.3 ฑ2) 0 0.7 ฑ1.5 (1 ฑ3) 0 P values Mann-Whitney U: 0.5665 Kruskal-Wallis: 0.0371 Kruskal-Wallis: 0.3386 T-test: 0.1674 Anova: 0.0310 Anova: 0.3521 Acrobeloides spp. 1.3 ฑ0.5 (3 ฑ0.6) 0 0 0 0.08 ฑ0.4 (0.08 ฑ0.5) 0a 0a 0a 0.7 ฑ1.2b (1 ฑ2) P values Mann-Whitney U: 0.7668 Kruskal-Wallis: 0.8869 Kruskal-Wallis: 0.0293 T-test: 0.3096 Anova: 0.8936 Anova: 0.0199 Fungivores Tylenchus spp. 2.2 ฑ1.3 (8 ฑ3) 2.4 ฑ2.1 (10 ฑ7) 3.4 ฑ0.6 (29 ฑ0.8) 1.4 ฑ1.9 (3 ฑ6) 2.4 ฑ1.8 (10 ฑ5) 2.1 ฑ2 (7 ฑ6) 2.2 ฑ1.9 (8 ฑ6) 2.2 ฑ1.7 (8 ฑ4) 3.6 ฑ.0.5 (35 ฑ0.7) P values Mann-Whitney U: 0.7518 Kruskal-Wallis: 0.3859 Kruskal-Wallis: 0.5114 T-test: 0.8245 Anova: 0.4182 Anova: 0.6606 106 Fungivores cont. Nematode genera/families Cultivation practices Nematicides Crop rotation sequences 1 2 1 2 3 1 2 3 4 Aphelenchus spp. 0.7 ฑ1.5 (1 ฑ3) 0.3 ฑ0.8 (0.3 ฑ1) 0 0 0.6 ฑ1.3 (0.8 ฑ3) 1 ฑ1.6 (2 ฑ4) 0.5 ฑ1.2 (0.6 ฑ2) 0 0 P values Mann-Whitney U: 0.6494 Kruskal-Wallis: 0.5038 Kruskal-Wallis: 0.4845 T-test: 0.3249 Anova: 0.5480 Anova: 0.5385 Aphelenchoides spp. 0 0.3 ฑ1.2 (0.3 ฑ2) 0 0 0.2 ฑ1 (0.2 ฑ2) 0.5 ฑ1.5 (0.6 ฑ3) 0 0 0 P values Mann-Whitney U: 0.7820 Kruskal-Wallis: 0.8869 Kruskal-Wallis: 0.5724 T-test: 0.3414 Anova: 0.8936 Anova: 0.5949 Predators Dorylaimidae 2.9 ฑ1.8 (17 ฑ5) 3.2 ฑ1.2 (23 ฑ2) 3.2 ฑ0.9 (23 ฑ1) 3.6 ฑ1.2 (35 ฑ2) 3 ฑ1.6 (19 ฑ4) 3.2 ฑ1.4 (23 ฑ3) 2.9 ฑ1.5 (17 ฑ3) 3.3 ฑ1.3 (26 ฑ3) 3.8 ฑ1.1 (43 ฑ2) P values Mann-Whitney U: 0.6212 Kruskal-Wallis: 0.7757 Kruskal-Wallis: 0.7913 T-test: 0.5238 Anova: 0.7286 Anova: 0.8194 Mononchidae 0.5 ฑ1.4 (0.6 ฑ3) 0 0 0 0.3 ฑ1.1 (0.3 ฑ2) 0.4 ฑ1.2 (0.5 ฑ2) 0.3 ฑ1.1 (0.3 ฑ2) 0 0 P values Mann-Whitney U: 0.5400 Kruskal-Wallis: 0.7804 Kruskal-Wallis: 0.8836 T-test: 0.1400 Anova: 0.7917 Anova: 0.8938 107 Predators cont. Nematode genera/families Cultivation practicesa Nematicidesb Crop rotation sequencesc 1 2 1 2 3 1 2 3 4 Total NPNd 6.2 ฑ1.4 (491 ฑ3) 5.8 ฑ1 (329 ฑ2) 4.8 ฑ0.8 (120 ฑ1) 5.3 ฑ1.1 (199 ฑ2) 6.2 ฑ1.2 (491 ฑ2) 6.1 ฑ1 (444 ฑ2) 6.1 ฑ1.1 (444 ฑ2) 5 ฑ1.1 (147 ฑ2) 5.5 ฑ0.4 (243 ฑ0.5) P values Mann-Whitney U: 0.3845 Kruskal-Wallis: 0.1117 Kruskal-Wallis: 0.2770 T-test: 0.3021 Anova: 0.1228 Anova: 0.1938 a1=plough :2=rip & plough; b1=EDBฎ only :2=nematicides other than EDBฎ :3=EDBฎ & other nematicides (combined); c1=grass and potato :2=grass, maize & potato :3=maize, wheat & potato :4=maize and/or grass and potato; dTotal NPN= Rhabditis spp. + Acrobeles spp. + Cephalobus spp. + Eucephalobus spp. + Prismatolaimus spp. + Acrobeloides spp. + Tylenchus spp. + Aphelenchus spp. + Aphelenchoides spp. + Dorylaimidae + Mononchidae 108 During the 2010 survey only the total number of NPN were obtained since no identification was made to genus and/or family level during counts (Table 2.21). A significant (P?. effect was evident for crop rotation sequences and total NPN numbers/200g soil (Table 2.21). Cropping sequences that included grasses, maize and potato (category 2) resulted in the highest total NPN/200g soil, followed by maize, wheat and potato sequences from which it did not differ significantly (P?. Fategor\  . ,t did, however, differ significantly (P?. from Fropping seTXenFes represented b\ Fategories  grass and potato and 4 (maize and/or grass and potato). For the 2011 season NPN were, however, counted to genera and/or family level with Rhabditis, Acrobeles, Cephalobus, Eucephalobus, Prismatolaimus and Acrobeloides spp. being identified from 200g soil samples (Table 2.22). A significant difference was evident (P?. between Rhabditis spp. and nematicides applied with significantly (P?. higher numbers of these genera being present in soil of fields where EDBฎ and other nematicides (category 3) were applied compared to those where only EDBฎ (category 1) and nematicides other than EDBฎ (category 2) were applied. This trend was, however, only evident for the parametric Anova test and not for the non-parametric Kruskal-Wallis test. Significant differences (P?. were also obtained between Prismatolaimus spp. and nematicides applied with significantly (P?. higher numbers of this genus for fields where only EDBฎ (category 1) were applied compared to those where nematicides other than EDBฎ (category 2) and EDBฎ in combination with other nematicides (category 3) were applied (Table 2.22). Although the latter was evident of data subjected to both parametric and non-parametric statistical tests, population levels of this genera were very low. For Acrobeloides spp., significant (P?. differenFes were obtained for crop rotation sequences implemented but since population levels for this genus were very low and ranged between zero and 0.7 this data are not discussed further. For Cephalobus spp. a significant difference (P?. was obtained for cultivation practices with numbers of this genus being higher for fields that were only ploughed compared to those that were ripped and ploughed (Table 2.22). The latter was evident for both the parametric and non- parametric tests. Fungivore nematodes were represented by Tylenchus, Aphelenchus and 109 Aphelenchoides spp. No significant differences (P?. were, however, obtained between either of the latter genera and cultivation practices, nematicides or crop rotation sequences applied. Predator nematodes were represented by individuals of the Dorylaimidae and Mononchidae families (Table 2.22). No significant correlation (P?. was obtained for the respective cultivation practices, nematicides or crop rotation sequences applied on the 31 fields using both non-parametric (Kruskal-Wallis) and parametric (Mann-Whitney U, T-test and Anova) statistical analyses either for Dorylaimidae, Mononchidae or total NPN. In terms of associations between soil type characteristics such as percentage soil, clay and organic material and PPN as well as NPN no significant (P?. effects were evident and data are thus not shown. 2.4.3 Faunal analyses for non-parasitic nematodes (NPN) Table 2.23. Functional guilds, to which non-parasitic nematode genera/families (identified in soil samples from the 31 fields of five producers in Christiana area), were assigned according to their feeding habits and life stage characteristics (Bongers, 1990). Genus Guild Rhabditis Ba1 Acrobeles Ba2 Acrobeloides Ba2 Cephalobus Ba2 Chiloplacus Ba2 Eucephalobes Ba2 Prismatolaimus Ba3 Aphelenchus Fu2 Tylenchus* Fu2 Aphelenchoides Fu2 Doryliamidae Om4 Mononchidae Ca4 *Personal communication, Dr. A. Swart, Nematode Taxonomist, ARC-PPRI, Pretoria, 15 May 2010 ba: bacterivores, ca: carnivore, fu: fungivores, om: predator (omnivore). Suffix numbers represent cp values for the various taxa. 110 The proposed faunal profile into which soil can be categorised according to the presence, abundance and diversity of NPN, soil nematodes (Ferris et al., 2001)(Figure 2.4) was also consulted continuously throughout the results section of this study. Figure 2.4. Functional guilds of non-parasitic, soil nematodes characterized by their respective feeding habit (trophic group) and by life history characteristics as expressed along a colonizer– persister (cp) scale (after Bongers & Bongers, 1998) (Graphical illustration from Ferris et al., 2001). Quadrant A Stress Enriched Quadrant B Stable Enriched Quadrant C Stable Depleted Quadrant D Stress Depleted 111 Figure 2.5. (A & B). Enrichment (EI) and structural indices (SI) for non-parasitic nematodes identified from producer 1 & 2’s fields in the Christiana area during the 2011 growing season categorised in functional guilds according to their feeding habits and life history characteristics. A B Producer 1 Producer 2 112 Figure 2.6. (A & B). Enrichment (EI) and structural indices (SI) for non-parasitic nematodes identified from producer 3 & 4’s fields in the Christiana area during the 2011 growing season categorised in functional guilds according to their feeding habits and life history characteristics. Producer 3 Producer 4 A B 113 Figure 2.7. Enrichment (EI) and structural indices (SI) for non-parasitic nematodes identified from producer 5’s fields in the Christiana area during the 2011 growing season categorised in functional guilds according to their feeding habits and life history characteristics. Four (1, 2, 4 and 5) of the six fields sampled for producer 1 were plotted in the “Stable Enriched” and two (6 and 3) in the “Stress Enriched” quadrants of the simplified food web representation for NPN (Figure 2.5A). These results reflect that for the “Stable Enriched” quadrant both high EI and SI for fields 1, 2 and 4, but lower SI and EI for field 5. On the Enrichment Trajectory this indicates that soil of both fields 1, 2 and 4 were dominated by enrichment opportunistic bacterivores (cp1), while abundant nematode taxa such as bacterivores and fungivores (cp2) were also present but in lower proportions. For the Structural Trajectory, results showed that bacterivores (cp4-5), carnivores (cp4-5), fungivores (cp4-5) and omnivores (cp 4&5) dominated. However, individuals from Ba3, Ca3 and Fu3 were also present in lower proportions, while none/very low numbers of the Om5 group were present. In contrast, for the “Stress Enriched” quadrant field 6 had a high EI but a zero SI, indicating the presence of abundant bacterivorous and fungivorous nematode taxa (cp2) in combination with enrichment opportunistic bacterivores (cp1) that dominates. However, no bacterivorous (Ba), carnivorous (Ca) or fungivorous (Fu) nematodes in cp scales 3-5 (reflected on the Structural Trajectory) as well as omnivorous/predator (Om) for cp scales 4 and 5 were present in this field. Producer 5 114 For field 3, NPN assemblages present had medium EI and SI levels. For the Enrichment Trajectory the latter indicates that abundant nematode taxa such as bacterivores and fungivores (cp2) dominated, while enrichment opportunistic bacterivores (cp1) also occurred but in low proportions. The SI Trajectory indicated that bacterivores (cp 3&4), carnivores (cp2) and fungivores (cp 3&4) as well as omnivores (cp4) dominated in this field, with very low proportions/if any of the latter guilds for cp 5 were present in this field. The same general scenario was true for categorisation of fields of producer 2 compared to those for producer 1. Only subtle differences existed for NPN assemblages categorised along EI and SI trajectories of a faunal profile. Four (8, 9, 10 and 11) of the fields were plotted in the “Stable Enriched” quadrant and two (7 and 12) in the “Stress Enriched” quadrant (Figure 2.5B). Refer to explanation of the particular NPN groups grouped according to their cp values as have been done for producer 1 with regard to these two specific faunal quadrants. All six fields sampled for producer 3 were plotted in the “Stable Enriched” quadrant (Figure 2.6A). Refer to explanations of the status of NPN present in these fields to those supplied for producer 1 above. For producer 4, four (21, 22, 23 and 24) of the six fields were plotted in the “Stable Enriched” quadrant (see explanation for producer 1) with two (20 and 25) in the “Stable Depleted” quadrant (Figure 2.6B). Fields plotted in the latter quadrant had low (field 25) to medium (field 20) EI and SI. EI for these fields indicated that NPN assemblages represented by Ba2 and Fu2 dominated, while Ba1 also occurred. In terms of the SI, Ba4, Ca4, Fu4 and Om4 dominated. NPN from the latter guilds with cp values of 5 were, generally not present in soil from these fields or in very low numbers. For producer 5 three (28, 30 and 31) of the six fields were plotted in the “Stable Enriched” (see explanation for producer 1) two (26 and 27) in the “Stress Enriched” (see explanation for producer 1) and one in the “Stable depleted” (see explanation for producer 4) quadrants (Figure 2.7). None of the fields sampled from the five producers, were plotted in the “Stressed Depleted” quadrant, which indicates the presence of and dominance by bacterivores, carnivores, fungivores and omnivores of cp values 2-4. 115 2.5 Discussion Thirteen PPN genera and 16 nematode species have been identified from both root and soil samples from the 31 fields in which potato is included as a rotation crop every eight years in the Christiana area. In addition, although not identified to species level, individuals of the Aphelenchoides genus of which only A. ritzemabozi is an obligate PPN (Hunt, 1993), have been found and preliminary been categorised as fungus feeders until species identification can be done. Of the genera found, only one genus (Psilenchus) and six species (Criconemoides sphaerocephalus, Hemicriconemoides brachyurus, Paratrichodorus lobatus, Rotylenchus caudaphasmidis and Tylenchorhynchus ventralis) have never been associated with maize (Keetch & Buckley, 1984; De Waele & Jordaan, 1988; Kleynhans et al., 1996; Riekert, 1996; Riekert & Henshaw, 1998), potato (Kleynhans, 1978; Keetch & Buckley, 1984; Kleynhans et al., 1996), sunflower (Keetch & Buckley, 1984; Kleynhans et al., 1996; Bolton et al., 1989), onion (Coetzee, 1968; Keetch & Milne, 1982; Meyer, 1984) or grasses (Keetch & Buckley, 1984; Kleynhans et al., 1996; Ntidi et al., 2012) in cropping systems in SA. In terms of NPN assemblages nine genera (six bacterivorous and three fungivorous) as well as two families (omnivores/predators) have been identified from soil samples from these fields. Also, assessment of NPN populations and classification of fields accordingly in terms of their soil health is reported for the first time in annual crops such as potato-based cropping systems in SA as a result of this study. Endoparasitic Meloidogyne spp. were the predominant PPN group in both soil and root samples collected during this survey from the 31 fields that are part of potato-based rotation systems in the Christiana area. These results support data that RKN are widely distributed in the Christiana potato production areas of SA (Kleynhans et al., 1996; Jones et al., 2011), where it causes severe damage to the crop (G. Posthumus, pers comm., Maart 2010). Results from this study also coincides with reports from other authors that Meloidogyne spp. are considered economically one of the most important PPN genera that are locally associated with crops that are planted in rotation with potato (Kleynhans, 1978; Kleynhans et al., 1996; Jones et al., 2011) such as maize (Keetch & Buckley, 1984; De Waele & Jordaan, 1988; Kleynhans et al., 1996; Riekert, 1996; Riekert & Henshaw, 1998), sunflower (Keetch & Buckley, 1984; Bolton et al., 1989; Kleynhans et al., 1996), onion (Coetzee, 1968; Keetch & Milne, 1982; Meyer, 1984) as well as grasses. The latter is also true for weeds (Keetch & Buckley, 1984; Kleynhans et al., 1996; Ntidi et al., 2012) that occur in such crop fields. Also, on a global scale Meloidogyne spp. are considered as one of the economically most important and predominant PPN that attack a 116 wide range of crops (Moens et al., 2009) such as potato (Scurrah et al., 2005; Castagnone- Sereno, 2006), maize (Mc Donald & Nicol, 2005; Bridge & Starr, 2007), sunflower (Korayem et al., 2011), onion (Bridge et al., 2005; Mc Donald & Nicol, 2005; Sikora et al., 2005) where it causes severe quality and yield losses (Moens et al., 2009). Both M. incognita and M. javanica were identified during this study but not M. acronea, M. arenaria, M. chitwoodi and M. hapla that are also reported as pests of potato in SA (Keetch & Buckley, 1984; Kleynhans et al., 1996; Jones et al., 2011) and in other countries (Scurrah et al., 2005; Bridge & Starr, 2007). Meloidogyne incognita and M. javanica have also been identified as the predominant RKN in the maize- (Riekert, 1996; Riekert & Henshaw, 1998) and sunflower- producing (Bekker et al., 2007) areas of SA. The same scenario applies for the occurrence of these two RKN species in weeds and grasses (Keetch & Buckley, 1984) sampled in commercial as well as small-scale fields of farmers in SA (Ntidi et al., 2012). Inclusion of crops such as maize, sunflower, wheat and onion as well as grass cover crops in this potato-producing area on which Meloidogyne species survive and reproduce implies that their population levels can build up to reach damaging levels when potato and/or other susceptible crops are planted on such fields. Following Meloidogyne spp., other predominant endoparasitic nematode genera and species recorded during this study from both soil and root samples were individuals from the endoparasitic Pratylenchus (P. zeae, P. neglectus and P. scribneri) and Ditylenchus spp. as well as Trichodoridae (P. lobatus and N. minor) and Hoplolaimidae (Rotylenchus unisex, R. caudaphasmidis, Scutellonema brachyurus and S. truncatum). Pratylenchus brachyurus, which is regarded as an economically important parasite of potato in SA (Keetch & Buckley, 1984; Kleynhans et al., 1996; Jones et al., 2011) and other countries (Scurrah et al., 2005) in particular, was not found during this study. However, all three Pratylenchus spp. that were identified during this study have been associated with potato in SA earlier (Keetch & Buckley, 1984; Kleynhans et al., 1996). Pratylenchus neglectus and P. scribneri are also known to damage potato in temperate, tropical and subtropical regions of the world (Scurrah et al., 2005). Important is that P. neglectus and P. zeae are also associated with maize, grasses, wheat and onions (Kleynhans et al., 1996) that are all used as rotation crops with potato in the Christiana area. Not only does Pratylenchus spp. cause damage to the potato crop, they are also associated with the fungus (Verticillium) that can cause Verticillium-wilt in potato (Jones et al., 2011). 117 Although Ditylenchus destructor and D. dipsaci are regarded as major parasites of potato in countries other than SA (Scurrah et al., 2005; Bridge & Starr, 2007), the Ditylenchus sp. found in this study could not be identified since only juvenile specimens were encountered in root samples and their population levels were very low. However, in SA only D. africanus has been found to be of economical importance only infecting and reducing quality of groundnut kernel consignments (De Waele et al., 1989) and therefore, poses no risk to potato (Basson et al., 1990). However, high yield losses have been reported from other parts of the world where climatological conditions favour the establishment of D. destructor and D. dipsaci as a pest of potato (Scurrah et al., 2005). Also members of the Hoplolaimidae family are not regarded as economically important parasites of potato (Scurrah et al., 2005; Jones et al., 2011) with various species such as Helicotylenchus cavenessi, H. dihystera, H. pseudorobustus, Rotylenchus brevicaudatus, Scutellonema brachyurum and S. cavenessi being associated with potato in SA (Keetch & Buckley, 1984; Kleynhans, 1991; Kleynhans et al., 1996) in particular. Although J2 of the family Heteroderidae (other than Meloidogyne spp.) were also present in samples from two fields during the 2010 survey, no cysts could be isolated. In SA G. rostochiensis causes damage to potato in the Bon Accord, Ceres and Sandveld areas, but have not been reported in the Christiana area where this study was conducted (Jones et al., 2011). It is, however, important to obtain more soil samples from these specific fields to enable extraction and identification of cysts to make sure of the status of members of the cyst families in this important potato-production area. Other PPN genera identified during this study such as Longidorus, Nanidorus, Paratrichodorus, Tylenchorhynchus and Xiphinema have also been associated with potato in SA (Keetch & Buckley, 1984; Kleynhans et al., 1996; Jones et al., 2011) and in other parts of the world (Scurrah et al., 2005) but are generally not regarded as important pests of the crop. The presence of N. minor and P. lobatus in particular in the Christiana potato production area is, however, important since these ectoparasites are vectors of the tobacco rattle virus that causes tuber spraing in potato (Manzanilla-L๓pez et al., 2004). Also these seemingly lesser important PPN identified in fields from this area have been associated with maize (Keetch & Buckley, 1984; De Waele et al., 1988), sunflower (Keetch & Buckley, 1984; Bolton et al., 1989), onion (Coetzee, 1968; Keetch & Milne, 1982; Keetch & Buckley, 1984; Kleynhans et al., 1996) and grasses (Keetch & Buckley, 1984; Van Biljon, 2004; Ntidi et al., 2012) that are planted in rotation with 118 potato in this area and should be monitored to prevent high build-up of their population levels which may lead to quality and quantity losses. In terms of NPN assemblages present in the 31 fields sampled in the Christiana area, their total population levels were generally higher during both growing seasons compared to that of the PPN. Bacterivore nematode genera were represented by a wide spectrum with, individuals of the Rhabditis (Ba1) genus dominating, followed by Cephalobus spp. (Ba2). The latter genera as well as the other bacterivorous genera found during this study are commonly found in soils in SA (Heyns, 1971). Bacterivores are regard as the beneficial nematodes of which the population levels increase substantially and relatively quick after the addition of organic material (Heyns, 1971) or any kind of disturbance (Bongers, 1990; Ferris et al., 2001). The fungivore nematode community were represented by a smaller variety of genera, with Tylenchus (Fu2) being predominant. In terms of the predator nematodes, only individuals of the Dorylaimidae (Om4) and Mononchidae (Ca4) families were present in relatively low numbers. Of particular interest is that the majority of the 31 fields were plotted in the ‘Stable Enriched” quadrant that indicates that the soils of such fields were relatively healthy, maturing and stable in terms of the beneficial nematodes as indicated by the simplified food web representation (Bongers, 1990; Bongers & Ferris, 1999; Ferris et al., 2001). This scenario is surprising and could probably be explained by the inclusion of grass cover crops on these fields for five to six years of the eight-year cropping cycles practiced by these producers. The dominant contribution of particularly bacterivores from the cp 1, 4 and 5 scales, carnivores (Ca 4&5), fungivores (cp 4&5) and omnivores (Om4) contributed substantially to classification of such fields in the “Stable Enriched” quadrant. Generally, the only common factors shared for all fields categorised in the “Stable Enriched” quadrants are the inclusion of grasses (Eragrostis, Digitaria and/or Panicum spp.) and potato. This is, however, also true for the other fields that were classified in either the “Stress Enriched” and/or “Stable Depleted” quadrants. Planting of rotation crops, particularly grass cover crops and potato as well as ploughing generally were the three common factors generally applied to all 31 fields during both years. Also the organic content of these fields were generally low and ranged from 0.15% to 0.48% when determined during 2010 when this study commenced. Therefore, reasons why the majority of these fields were classified as stable and enriched are currently unknown. Although more research is needed to conclude on this scenario, results obtained during this study already represents baseline information for fields in this specific potato-producing area. 119 Investigation of relationships between both PPN and NPN rendered interesting results but it must be remembered that data for these groups were pooled for all 31 fields thus resulting in low mean population levels for the various parameters, i.e. cultivation practices, nematicides used and crop rotation sequences. Although a few significant relationships between the various PPN as well as NPN nematode genera and cultivation practices, nematicide(s) used as well as cropping sequences were revealed it has to be verified with additional results. Collection of data of this kind for more than one growing season needs to be done and will add substantial value to the science of Nematology as well as to producers and the related crop industries. Of particular interest are findings that application of EDBฎ in combination with other nematicides rendered significantly lower population levels of some of the endo- and ectoparasitic nematode genera identified during the study. This trend indicates that application of a variety of nematicides rather than one specific product seems to be the best way to reduce population levels of PPN in this area. Concurrently, indications that some NPN, i.e. Rhabditis and Prismatolaimus spp. had higher population levels in fields where EDBฎ combined with other nematicides and EDBฎ alone were applied, respectively, also indicate that interesting and important trends could be revealed should this type of research be extended over more growing seasons. The same applies for crop rotation sequences where RKN population levels were significantly lower in fields where grass, maize and potato were planted compared to those that excluded grass. This may be explained by the poor host status of grasses for RKN and may add substantial value in such cropping systems to manage these parasites. Validation of such information is, however, crucial and can have pertinent implications for designing and planning of management strategies for PPN to fit the needs of not only the Christiana, but also other local producers. No explanation could at this stage be given for no significant effects being evident found for either PPN or NPN with regard to soil characteristics such as percentages of sand, silt, clay and organic material during this study. It is, however, important to bear in mind that soil from all these fields were all sandy with percentages ranging from 91.6%to 96%. The latter substantiates the predominant occurrence of RKN in sandy soils, such as those occurring in the Christiana area, as been noted by other authors (Greco & Di Vito, 2009). 120 2.6 Conclusions Although not present in soil and or roots from crops grown in all 31 fields, RKN were identified as the predominant PPN group in the Christiana potato-producing area where it poses a real problem to potato producers. These parasites may also impact adversely on other crops and grass cover crops planted by these producers since they have a wide host range. In terms of NPN assemblages, characterisation of the majority of potato fields in the “Stable Enriched” quadrant was unexpected but is most probably due to the inclusion of grasses during six to seven years of the eight-year cropping cycle. Results obtained during this study are valuable to serve as a baseline study with regard to prevailing community structures, species composition and population levels of both PPN and NPN present in the 31 fields sampled. More such studies should be done in important potato-production areas such as Christiana where life-sustaining grain crops including maize and wheat, oilseed crops such as sunflower and high-cash vegetable crops are produced. This way important correlations between these parasites and factors such as cultivation practices, nematicides used as well as crop rotation sequences can be obtained to assist producers and the industry to minimise the adverse effect of PPN on their crops. 121 2.7 References Basson, S., De Waele, D. & Meyer, A.J. 1990. An evaluation of crop plants as hosts for Ditylenchus destructor isolated from peanut. Nematropica 20: 23-29. Bekker, S., Venter, G.A. & Fourie, H. 2007. Nematode diagnostic service at the ARC-Grain Crops Institute. 18th Symposium of the Nematological Society of Southern Africa. Port Elizabeth, South Africa, 6-10 May 2007. (Poster Presentation). Black, V. 2008. Potatoes in South Africa. In: GM potatoes in South Africa – a critical analysis. The African Centre for Biosafety, Johannesburg, South Africa. pp. 12-24. Bolton, C., De Waele, D. & Loots, G.C. 1989. Plant-parasitic nematodes on field crops in South Africa. 3. Sunflower. Revue de N้matology 12: 69-76. Bongers, T. 1990. The maturity index: an ecological measure of environmental disturbance based on nematode species composition. Oecologia 83: 14-19. Bongers, T. & Ferris, H. 1999. Nematode community structure as a bioindicator in environmental monitoring. Trends in Evolution and Ecology 14: 224-228. Bridge, J., Plowright, R.A. & Peng, D. 2005. Nematode parasites of rice. In: Luc, M., Sikora, R.A. & Bridge, J. (eds) Plant parasitic nematodes in subtropical and tropical agriculture. CAB International, London, UK. pp. 87-130. Bridge, J. & Starr, J.L. 2007. Plant nematodes of agricultural importance A color handbook. Academic Press, Boston, USA. 152 pp. Castagnone-Sereno, P. 2006. Genetic variability and adaptive evolution in pathogenetic root- knot nematodes. Heredity 96: 282-289. Coetzee, V. 1968. The distribution of the family Heteroderidae (Filipjev, 1934) in South Africa and some host records of Meloidogyne species. South African Journal of Agricultural Sciences 11: 775-788. 122 Coolen, W.A. & D’Herde, C.J. 1972. A method for the quantitative extraction of nematodes from plant tissue. Ministry of Agriculture, Belgium. p. 77. De Grisse, A. 1965. A labour-saving method for fixing and transferring eelworms to anhydrous glycerine. Offset, University of Gent, p. 3. De Waele, D. & Jordaan, E.M. 1988. Plant-parasitic nematodes on field crops in South Africa. 1. Maize. Revue de N้matology 11: 65-74. De Waele, D., Jones, B.L., Bolton, C. & Van den Berg, E. 1989. Ditylenchus destructor in hulls and seeds of peanut. Journal of Nematology 21: 10-15. Ferris, H., Bongers, T. & De Goede, R.G.M. 2001. A framework for soil food web diagnostics: extension of nematode faunal analysis concept. Applied Soil Ecology 18: 13-29. Fourie, H., Leswifi, C., Mc Donald, A.H. & De Waele, D. 2007. Host suitability of vetiver grass to Meloidogyne incognita and M. javanica. Nematology 9: 49-52. Greco, N. & Di Vito, M. 2009. Population dynamics and damage levels. In: Perry, R.N., Moens, M. & Starr, J.L. (eds) Root-knot nematodes. CABI Publishing, Wallingford, UK. pp. 246-274. Heyns, J. 1971. A guide to the plant and soil nematodes of South Africa. Content Solutions, Pretoria, South Africa. Hooper, D.J., Hallman, J. & Subbotin, S.A. 2005. Methods for extraction, processing and detection of plant and soil nematodes. In: Luc, M., Sikora, R.A. & Bridge, J. (eds) Plant parasitic nematodes in subtropical and tropical agriculture. CAB International, Wallingford, UK. pp. 53-86. Jones, R.K., Marias, M. & Swart, A. 2011. Aalwurms (Nematodes). In: Denner, F. & Venter, S. (eds) Handleiding vir aartappelproduksie in Suid-Afrika. Landbounavorsingsraad-Roodeplaat Instituut vir Groente en Sierplante. Pretoria, Suid-Afrika. pp. 195-201. Keetch, D.P. & Buckley, 1984. A check-list of the plant parasitic nematodes of Southern Africa. Technical Communication No. 795. Department of Agricultural Development, South African Government, Republic of South Africa. 123 Keetch, D.P. & Milne, D.L. 1982. The control of plant-parasitic nematodes. In: Keetch, D.P. & Heyns, J. (eds) Nematology in southern Africa. Science Bulletin Department of Agriculture and Fisheries Republic of South Africa. Science Bulletin No. 400. pp. 113-129. Kleynhans, K.P.N. 1978. Nematode pests in potatoes. Leaflet of the Department of Agriculture and Technical Services of the Republic of South Africa Potato Series No. G. 13. Kleynhans, K.P.N., Van den Berg, E., Swart, A., Marais, M. & Buckley, N.H. 1996. Plant nematodes in South Africa. Plant Protection Research Institute Handbook No 8, Pretoria, Business Print. p. 165. Korayem, A.M., Youssef, M.M.A., Ahmed, M.M. & Mohamed, M.M.M. 2011. Distribution and association of plant parasitic nematodes with some oil crops in Egypt. Pakistan Journal of Nematology 29: 79-91. Manzanilla-L๓pez, R.H., Evans, K. & Bridge, J. 2004. Plant diseases caused by nematodes. In: Chen, Z.X., Chen, S.Y. & Dickson, D.W. (eds) Nematology advances and perspectives. Volume 2. Nematode management and utilization. CABI Publishing, Wallingford, UK. pp. 637-716. Mc Donald, A. & Nicol, J. 2005. Nematode parasites of cereals. In: Luc, M., Sikora, R.A. & Bridge, J. (eds) Plant parasitic nematodes in subtropical and tropical agriculture. CAB International, London, UK. pp. 131-192. Meyer, A.J. 1984. Damage to onion by Nanidorus minor in the Koue Bokke veld. Phytophylactica 16:331. Moens, M., Perry, N. & Starr, J.L. 2009. Meloidogyne species – a diverse group of novel and important plant parasites. In: Perry, R.N., Moens, M. & Starr, J.L. (eds) Root-knot nematodes. CAB International, Wallingford, UK. pp. 1-17. Mucina, L. & Rutherford, M.C. 2006. The vegetation of South Africa, Lesotho and Swaziland. Strelitzia 19. South African National Biodiversity Institute, Pretoria, South Africa. p. 516. 124 Ntidi, K.N., Fourie, H., Mc Donald, A.H., De Waele, D. & Mienie, M.S. 2012. Plant-parasitic nematodes associated with weeds in developing agriculture in South Africa. Nematology (in press). Riekert, H.F. 1995. An adapted method for extraction of root-knot nematode eggs from maize root samples. African Plant Protection 1: 41-43. Riekert, H.F. 1996. Economic feasibility of nematode control in dryland maize in South Africa. African Crop Science Journal 4: 477-481. Riekert, H.F. & Henshaw, G.E. 1998. Effect of soybean, cowpea and groundnut rotations on RKN build-up and infestation of dryland maize. African Crop Science Journal 6: 377-383. Scurrah, M.I., Niere, B. & Bridge, J. 2005. Nematode parasites of solanum and sweet potatoes. In: Luc, M., Sikora, R.A. & Bridge, J. (eds) Plant parasitic nematodes in subtropical and tropical agriculture. CAB International, London, UK. pp. 193-219. Seinhorst, J.W. 1962. Modifications of the elutriation method for extracting nematodes from soil. Nematologica 8:117-128. Sellschop, J.P.F. & Du Toit, J.J. 1948. The production of potatoes. Farming in South Africa 23: 645-654. Sikora, R.A., Bridge, J. & Starr, J.L. 2005. Management practices: an overview of integrated nematode management technologies. In: Luc, M., Sikora, R.A. & Bridge, J. (eds) Plant parasitic nematodes in subtropical and tropical agriculture. CAB International, London, UK. pp. 793-824. Taylor, C.E. & Martin, G.C. 1960. Potato production. 8. Pests. Rhodesia Agricultural Journal 57: 60-63. Van Biljon, E.R. 2004. Crop rotation as part of an integrated pest management programme for the control of plant parasitic nematodes. Proceedings of the Soilborne Plant Diseases Interest Group Symposium, Stellenbosch, 14-16 September 2004. 125 Yeates, G.W., Ferris, H., Moens, T. & Van der Putten, W.H. 2009. The role of nematodes in ecosystems. In: Wilson, M.J. & Kakouli-Duarte, T. Nematodes as environmental indicators. pp. 1- 44. Zijlstra, C. 2000. Identification of Meloidogyne chitwoodi, M. fallax and M. hapla based on SCAR-PCR: a powerful way of enabling reliable identification of populations or individuals that share common traits. European Journal of Plant Pathology 106: 283-290. Zijlstra, C., Donkers-Venne, D. H. & Fargette, M. 2000. Identification of Meloidogyne incognita, M. javanica, and M. arenaria using sequence characterized amplified region (SCAR) based PCR assays. Nematology 2: 847-853. 126 CHAPTER 3 MOLECULAR IDENTIFICATION OF ROOT-KNOT NEMATODE SPECIES USING SEQUENCE CHARACTERIZED AMPLIFIED REGION BASED POLYMERASE CHAIN REACTION ASSAYS 3.1 Introduction Accurate identification of organisms is fundamental to our understanding and communication about their role in soil ecology (Abebe et al., 2011). Knowledge of a pest, in this case, soilborne root-knot nematodes (RKN; Meloidogyne spp.), necessitates identification of the species present in a specific field (Moens et al., 2009). Because RKN is one of the most economically important pests of a wide variety of agricultural and horticultural crops (Luc et al., 2005) correct identification of their juvenile and adult stages is crucial (Powers, 2004) for quarantine and plant protection purposes (Sikora et al., 2005). Species identification of these organisms is also essential where more than one crop is included in cropping cycles, such as the Christiana area where this study was conducted. This is also applicable where control of RKN is planned using rotation of susceptible and resistant crops (Manzanilla-L๓pez et al., 2004). Several new molecular technologies such as sequence characterized amplified region based polymerase chain reaction (SCAR-PCR) assays have been developed and provide tools to assist in accurate and rapid identification of a wide range of plant-parasitic nematodes (PPN), particularly RKN (Moens et al., 2009). An increasing number of species have been described during the last decades within the genus Meloidogyne. It is, therefore, important that nematode species descriptions conform to a general standard in order to facilitate accurate comparisons and differential diagnoses of such individuals (Eisenback & Hunt, 2009). Due to the relatively conserved morphology within the RKN group there is an increasing emphasis on using techniques such as molecular sequences to accurately identify these species (Eisenback & Hunt, 2009). On the other hand, due to the nature and morphology of Meloidogyne species the differences that exist between them are very small, with certain features being influenced by the host plant as well as a range of environmental factors (Jepson, 1987). Thus, correct identification of RKN is difficult and no single morphological approach or single developmental-stage feature can be used to distinguish among all species belonging to this genus. Therefore, different approaches have been investigated to resolve problems experienced with the identification of RKN species (Hartman & Sasser, 1985; Jepson, 1987), including molecular techniques (Manzanilla-L๓pez et al., 2004), to supplement morphological identification. 127 Modern identification methods used to distinguish between RKN species must provide accuracy, speed, reliability, affordability and enable characterisation of specimens new to science (Powers, 2004). A major benefit of using molecular techniques to identify RKN species is that individual J2, which is the most common and infective stage of the RKN found in the soil, can be used. This way, the need to use mature females from roots and/or other parts of host plants, which are not always present during sampling, or establishment of cultures in growth-regulated glasshouses to obtain females, are eliminated. Molecular techniques based on characterisation of the deoxy-ribonucleic acid (DNA) can be used for every stage of the nematode’s life cycle (Devran & S|๗?t, 2009). This approach, however, requires some expensive equipment such as a PCR thermocycler and a furnished molecular laboratory. None the less, the use of molecular techniques to identify RKN are likely to play an ever increasing role in species identification, especially by those not well trained or experienced in classical morphometrics (Moens et al., 2009). Moreover, problems are experienced with morphological identification of RKN such as their sheer abundance and small size (Floyd et al., 2002), conservative morphology, the presence of considerable variation within populations of these parasites due to indistinct species boundaries or species complexes and the lack of experts trained as nematode taxonomists (Blok et al., 1997; Zijlstra et al., 2000; Blok & Powers, 2009). Therefore, it is preferable (although not always possible) to use molecular techniques to complement morphological identification to ensure accurate identification of the RKN species involved. The main objective for this part of the study was to accurately identify RKN species from fields where various crops and cover crops (i.e. grasses) were included in potato-based cropping systems in the Christiana area of South Africa (SA) both during the 2010 and 2011 growing seasons. 128 3.2 Material and methods 3.2.1 Deoxy-ribonucleic acid (DNA) extraction Soil from each of the 31 fields sampled during 2010 and 2011 was (see Chapter 2, paragraph 2.2.6) transferred to individual 4L pots. One tomato seedling (cv. Rodade) was planted in each pot to enable in vivo rearing of the respective RKN species present in soil sampled from each of these fields. Plants were watered three times a week using equal amounts of tap water. Fifty-six days after seedling transplanting the root system of each tomato plant was removed, washed under a gentle stream of tap water to remove excess debris and mature RKN females removed for identification purposes. Ten RKN females, if available, from each of the 31 fields were randomly removed with a scalpel from galls present on each of the respective individual tomato root systems. This procedure was conducted using a stereo microscope at 100x magnification. Single, mature RKN females from each infected tomato root system were placed in an individual microcentrifuge tube (1ml capacity) and crushed with a disposable pipette tip. DNA was isolated following a method by Zijlstra et al. (1995) that was adapted for the purpose of this study since mature RKN females were used instead of J2. This was done by extracting DNA from each RKN female by adding 50ตl cell lysis buffer (50mM TrisHCl pH 8.0; 100mM NaCl; 10mM EDTA; 1% SDS (v/v)) to each tube as well as 2ตl of Proteinase K (10mg/ml). The tube containing the crushed female suspended within the latter solution was incubated at 65บC for 1 hour. The DNA from each RKN female was extracted using a GeneClean Kit (Bio101) according to the manufacturer’s instructions. Three volumes of sodium iodide (NaI) solution (150ตl) and 2ตl glass milk were added to each tube containing the RKN DNA, mixed and incubated at room temperature (25บC- 27ฐC) for five minutes. The solution was then centrifuged at 10 000 rpm for five seconds and the supernatant decanted. The pellet, consisting of glass milk with bound RKN DNA remaining at the bottom of the tube was resuspended in 200ตl NEW Wash and centrifuged at 10 000rpm for another five seconds. This process was repeated two more times. The liquid supernatant was removed and the pellet containing the RKN DNA dried at room temperature for 12 hours, resuspended in 20ตl TE buffer and centrifuged for 30 seconds at 10 000rpm. Two microlitres of RKN DNA was subsequently used for SCAR-amplification reactions. 129 3.2.2 Sequence characterized amplified region (SCAR) amplification PCR was conducted in a total volume of 20ตl, containing 2ตl DNA, 1x GoTaq Green Master Mix (Promega) containing GoTaq DNA polymerase, dNTPs, MgCl2 and reaction buffer at optimal concentrations and 5pmol each of M. incognita, M. javanica, M. arenaria, M. chitwoodi, M. fallax and M. hapla forward and reverse primers respectively. DNA amplification for M. incognita was done in a ThermoHybaid Thermal Cycler programmed as follows: denaturation at 94บC for two minutes (one cycle), followed by 40 cycles of denaturation at 94บC for 30 seconds, annealing at 54บC for 30 seconds and extension at 72บC for one minute, followed by a final extension step at 72บC for five minutes. The same programme was used for M. javanica with the exception of the annealing temperature at 64บC (Zijlstra, 2000; Zijlstra et al., 2000). Female RKN DNA products from the field sites of which mature RKN were reared successfully, resulting from the amplification processes, were subsequently analysed by electrophoresis in a 3% (m/v) agarose gel with 1x TBE running buffer (89mM Tris-borate, 2.5mM EDTA, pH 8.3). Ethidium bromide (1ตg/ml) was added to the gel and samples were visualized under UV-light. One blank reaction (no DNA), one reference sample of M. incognita and M. javanica monoculture populations that have been reared in vivo in separate greenhouses, respectively, and identified by means of morphological characteristics earlier served as the positive control. However, no mature females and subsequently no DNA could be obtained for other RKN species such as M. arenaria, M. hapla, M. chitwoodi or M. fallax, which are also economically important species that infects potato in SA (Jones et al., 2011). Ultimately, Lambda DNA restricted with HindIII and EcoRI as molecular weight markers were also loaded on both sides of the gel comb to establish the size of the DNA bands for each of the RKN species that were present in the fields sampled. Gels containing the DNA products of the respective RKN females were electrophoresed for 2 hours at 80V and banding patterns were visualised with ultraviolet (UV) illumination. A photograph was taken of each gel containing the RKN female DNA-banding patterns. 130 3.3 Results Table 3.1. Meloidogyne species identified from the 31 fields of the five producers in the Christiana area, indicating their respective deoxy-ribonucleic acid (DNA) fragment sizes (bp). Source/Field no. DNA fragment size(s) (bp) Meloidogyne spp. identified 29 (UK1) 1 200 and 670 M. incognita and M. javanica 3 (CG3) 1 200 M. incognita 9 (LH13) 1 200 M. incognita 11 (LH20) 1 200 M. incognita 14 (A6) 1 200 M. incognita 18 (SP11) 1 200 M. incognita 19 (S5) 1 200 M. incognita 20 (P1) 1 200 M. incognita 22 (VH1) 1 200 M. incognita 23 (VH2) 1 200 M. incognita 25 (VH5) 1 200 M. incognita 5 (T6T5) 670 M. javanica 16 (B2) 670 M. javanica 26 (D4) 670 M. javanica 28 (GH17) 670 M. javanica Standard RKN populations used Greenhouse – in vivo reared 1 200 M. incognita Greenhouse – in vivo reared 670 M. javanica PCR with the M. incognita specific SCAR-primers resulted in amplification of the M. incognita 1 200bp SCAR-fragment for RKN females reared from 11 of the fields sampled during this study (Table 3.1). DNA from the latter females were similar to those of the standard M. incognita race 2 population used, indicating that M. incognita were present on 11 (29, 3, 9, 11, 14, 18, 19, 20, 22, 23 and 25) of the 31 fields sampled. 131 On the other hand, M. javanica was identified from RKN females reared in vivo from soil of five of the 31 fields sampled (Table 3.1). The 670bp M. javanica SCAR fragment was amplified during PCR- reactions with the M. javanica-specific SCAR marker with RKN DNA from the five fields (29, 5, 16, 26 and 28) corresponding with that of the standard M. javanica population used. The combined occurrence of M. incognita and M. javanica in one (29) of the 15 fields were further revealed when both the M. incognita (1 200bp) and M. javanica (670bp) SCAR fragments were amplified as a result of PCR reactions with both these primers (Table 3.1). No 420bp, 525bp, 515bp and 610bp SCAR fragments were amplified during any of the PCR reactions when the respective primers for M. arenaria, M. chitwoodi, M. fallax and M. hapla-specific were added to the DNA products of the mature females from the fields sampled during this study (Table 3.1). As indicated earlier, no mature females of the latter species could be found to enable DNA extraction and use thereof as respective standards during this study. 3.4 Discussion DNA-based identification was successfully used as a tool during this study for verifying and confirming the identity of economically important RKN species that were present in 15 of the 31 fields sampled in the Christiana area (North-West Province of SA). Only M. incognita and M. javanica were identified as a result of this study. Meloidogyne incognita was identified as the predominant RKN species present on 11 of the fields sampled, while M. javanica was identified from five of the fields sampled. In addition results from SCAR-PCR reactions indicated that monospecific populations of M. incognita race 2 and M. javanica, respectively, were mass-reared in vivo and subsequently used as standards to complement the use of primers of these two RKN species during molecular identification done during this study. Identification and verification of M. incognita and M. javanica species in fields of producers in the Christiana area will contribute substantially in terms of crop selection based on resistance of commercially available maize and grass genotypes that could be included in their current cropping systems. Information on the latter crops exists in terms of their host suitability (Van Biljon, 2004; Ngobeni et al., 2010) for these RKN respective species. In this way producers can optimise crop production strategies for sustainable production of high-cash crops such as potato. Value adding in 132 terms of crop production in this particular agricultural area thus emanated as a result of this part of this study. Although only M. incognita and M. javanica were identified from fields using the SCAR-PCR technique, it cannot be concluded beyond debate that only these species are present in these fields and in this area. A shortcoming of this particular part of the study is that mature females could not be obtained after in vivo rearing of such populations due to the presence of low J2 population levels for some of the fields sampled, i.e. fields 1, 2, 4, 6, 7, 8, 10, 12, 13, 15, 17, 21, 24, 27, 30 and 31. Endeavours will, however, be made to continue with in vivo rearing of RKN populations from the soil obtained during the surveys in order to obtain a more complete picture on species identification that represents the majority of the fields sampled in this particular area. No specimens of other economically important RKN species such as M. arenaria, M. chitwoodi, M. fallax and M, hapla could be obtained to ensure that their presence could be detected in DNA samples from the fields included in this study. However, use of primers of the latter species was included as the second best option to try and detect their presence should they be prevalent. It should be noted that endeavours will be made to obtain DNA samples of all RKN species that are regarded as economically important parasites of potato and other rotation crops included in such rotation systems. This way exclusion of RKN species that occurs in fields in potato-producing areas will be prevented and interesting information with regard to the occurrence of these parasites in such areas could be obtained. Such knowledge will benefit producers, the industry and the scientific arena. 3.5 Conclusions Molecular identification of mature RKN females enabled accurate identification of two of the economically most important species in 15 of the 31 fields sampled in the study area. Individual J2 from each field was not used during this study since problems were experienced to isolate such small amounts of DNA. These results will be to the advantage of producers since they can now select poor host cultivars of these respective RKN species identified of crops such as maize as well as grasses to be planted in such fields. This way RKN populations can be reduced to minimise damage to high-value crops such as potato. 133 The SCAR-PCR method used in this study has proved to be a reliable tool for routine diagnostic identification of RKN and was also used with success by various other researchers (Blok et al., 1997; Zijlstra, 2000; Zijlstra et al., 2000; Fourie et al., 2001; Berry et al., 2008). 134 3.6 References Abebe, E., Mekete, T. & Thomas, W.K. 2011. A critique of current methods in nematode taxonomy. African Journal of Biotechnology 10: 312-323. Berry, S.D., Fargette, M., Spaull, V.M., Morand, S. & Cadet, P. 2008. Detection and quantification of root-knot nematode (Meloidogyne javanica), lesion nematode (Pratylenchus zeae) and dagger nematode (Xiphinema elongatum) parasites of sugarcane using real-time PCR. Molecular and Cellular Probes 22: 168-176. Blok, V.C., Phillips, M.S. & Fargette, M. 1997. Comparison of sequences from the ribosomal DNA intergenic region of Meloidogyne mayaguensis and other major tropical root-knot nematodes. Journal of Nematology 29: 16-22. Blok, V.C. & Powers, T.O. 2009. Biochemical and molecular identification. In: Perry, R.N., Moens, M. & Starr, J.L. (eds) Root-knot nematodes. CAB International, Wallingford, UK. pp. 98-118. Devran, Z. & S|๗?t, M.A. 2009. Distribution and identification of root-knot nematodes from Turkey. Journal of Nematology 41: 128-133. Eisenback, J.D. & Hunt, D.J. 2009. General morphology. In: Perry, R.N., Moens, M. & Starr, J.L. (eds) Root-knot nematodes. CAB International, Wallingford, UK. pp. 18-54. Floyd, R., Abebe, E., Papert, A. & Blaxter, M. 2002. Molecular barcodes for soil nematode identification. Molecular Ecology 11: 839-850. Fourie, H., Zijlstra, C. & Mc Donald, A.H. 2001. Identification of Root-knot nematode species occurring in South Africa using the SCAR-PCR technique. Nematology 3: 675-680. Hartman, K.M. & Sasser, J.N. 1985. Identification of Meloidogyne species on the basis of differential host treatise and perineal-pattern morphology. In: Barker, K.R., Carter, C.C. & Sasser, J.N. (eds) Advanced treatise on Meloidogyne. Volume II. Methodology. North Carolina University Graphics, Raleigh, North Carolina, USA. pp. 69-78. 135 Jepson, S.B. 1987. Identification of root-knot nematodes (Meloidogyne species). CAB International, Wallingford, UK. 265 p. Jones, R.K., Marias, M. & Swart, A. 2011. Aalwurms (Nematodes). In: Denner, F. & Venter, S. (eds) Handleiding vir aartappelproduksie in Suid-Afrika. Landbounavorsingsraad-Roodeplaat Instituut vir Groente en Sierplante. Pretoria, Suid-Afrika. pp. 195-201. Luc, M., Sikora, R.A. & Bridge, J. 2005. (eds) Plant parasitic nematodes in subtropical and tropical agriculture. CAB International, London, UK. 871 p. Manzanilla-L๓pez, R.H., Evans, K. & Bridge, J. 2004. Plant diseases caused by nematodes. In: Chen, Z.X., Chen, S.Y. & Dickson, D.W. (eds) Nematology advances and perspectives. Volume 2. Nematode management and utilization. CABI Publishing, Wallingford, UK. pp. 637-716. Moens, M., Perry, N. & Starr, J.L. 2009. Meloidogyne species – a diverse group of novel and important plant parasites. In: Perry, R.N., Moens, M. & Starr, J.L. (eds) Root-knot nematodes. CAB International, Wallingford, UK. pp. 1-17. Ngobeni, L., Fourie, H. & Mc Donald, A.H. 2010. Host suitability of selected South African maize genotypes to the root-knot nematode species Meloidogyne incognita race 2 and Meloidogyne javanica: A preliminary study. South African Journal of Plant and Soil 28: 49-54. Powers, T. 2004. Nematode molecular diagnostics: from bands to barcodes. Annual Review of Phytopathology 42: 367-383. Sikora, R.A., Bridge, J. & Starr, J.L. 2005. Management practices: an overview of integrated nematode management technologies. In: Luc, M., Sikora, R.A. & Bridge, J. (eds) Plant parasitic nematodes in subtropical and tropical agriculture. CAB International, London, UK. pp. 793-824. Van Biljon, E.R. 2004. Crop rotation as part of an integrated pest management programme for the control of plant parasitic nematodes. Proceedings of the Soilborne Plant Diseases Interest Group Symposium, Stellenbosch, 14-16 September 2004. 136 Zijlstra, C. 2000. Identification of Meloidogyne chitwoodi, M. fallax and M. hapla based on SCAR-PCR: a powerful way of enabling reliable identification of populations or individuals that share common traits. European Journal of Plant Pathology 106: 283–290. Zijlstra, C., Donkers-Venne, D.T.H.M. & Fargette, M. 2000. Identification of Meloidogyne incognita, M. javanica and M. arenaria using sequence characterised amplified region (SCAR) based PCR assays. Nematology 2: 847-853. Zijlstra, C., Lever, A.E.M., Uenk, B.J. & Van Silfhout, C.H. 1995. Differences between ITS regions of solates of the root-knot nematodes Meloidogyne hapla and M. chitwoodi. Phytopathology 85: 1231- 1237. 137 CHAPTER 4 THE EFFECT OF COVER/GREEN MANURE CROPS ON PLANT-PARASITIC AND NON-PARASITIC NEMATODE ASSEMBLAGES IN A POTATO FIELD 4.1 Introduction Plant-parasitic nematodes (PPN), particularly root-knot nematodes (RKN) are recognized as a serious constraint to intensive crop production systems worldwide (Nyczepir & Thomas, 2009). After decades of high input farming, more sustainable practices based on ecologically sound management techniques are being developed for the agricultural sector. These changes involve reduced pesticide use, integrated pest management (IPM), increased use of biological control products and maintenance and improvement of the quality of ground water and soil (Halbrendt, 1996). However, increased levels of control are likely to be needed in nematode-susceptible, high-value crops such as potato (Nyczepir & Thomas, 2009). For the last 50 years, potato producers have used methyl bromide, a broad-spectrum fumigant, highly efficient in killing fungi, nematodes, insects and weeds. This fumigant has effectively controlled PPN, particularly in potato-based production systems (Monfort et al., 2007). However, methyl bromide was identified as a contributor to the depletion of the stratosphere ozone layer in 1992 and was scheduled for worldwide phase out by 2005 (UNEP, 1992; Noling, 2002; Schneider et al., 2003). The same scenario exist for other highly efficient synthetic nematicides, including fumigants (e.g. ethylenedibromide – EDBฎ) other than methyl bromide, contact as well as systemic products that have been/are registered for use on potato, particularly in South Africa (SA) (Nel et al., 2007; CropLife, 2011). Although aldicarb (Temikฎ) has, for example been phased out for use on potato in SA during 2010 already, it will only be announced in the Government Gazette during June/July 2012 (D. Uys, pers comm., March 2012). Threats about the phasing out of fenamiphos also exist (D. Uys, pers comm., March 2012). With the decreasing number of synthetic nematicides available for use on potato and increasing pressure towards environmental, human and animal safety alternatives for managing these parasites on potato are becoming quintessential. One potential cultural alternative to synthetic nematicides for management of RKN pests in potato fields is the use of cover/green manure crops with biofumigation characteristics (Monfort et al., 2007). Such crops, particularly species from the Brassicaceae family are used as green manure amendments prior to planting (Monfort et al., 2007) of crops such as potato. The presence of glucosinolates (GL) in 138 Brassicaceae plant organs and sufficient activity of the enzyme myrosinase (MYR) to catalyse their hydrolysis have indicated the practical possibility of using such crops for biofumigation of various soilborne organisms (Rosa et al., 1997). Incorporating Brassicaceae residues into soil is known to suppress a range of pest and disease-causing organisms, including fungi (Chan & Close, 1987), nematodes (Mojtahedi et al., 1991), insects (Brown & Morra, 1997, Mathiessen & Kirkegaard, 1998) bacteria (Akiew et al., 1996; Gouws & Mienie, 2000; Gouws & Wehner, 2004) and weeds (Brown & Morra, 1995). When used as a cover/green manure crop, research has demonstrated that many Brassicaceae species have a nematicidal effect against multiple PPN species especially the RKN species M. incognita and M. javanica (Mojtahedi et al., 1991; Lazzeri et al., 2004) and the lesion nematode Pratylenchus neglectus (Potter et al., 1998; Monfort et al., 2007). Glucosinolates, contained by these crops, themselves are not biologically active (McLeod & Steel, 1999). When Brassicaceae tissues are disrupted, however, the GL react with MYR, thioglucosidase and generates biologically active compounds including nitriles, thiocyanates and, most notably isothiocyanates (McLeod & Steel, 1999). To achieve the maximum benefit from utilizing Brassicaceae species as cover and/or green manure crops for the control of soilborne PPN pathogens, a species and cultivar (cv.) with a high potential of GL production and also one that produces large quantities of biomass in the selected geographic location or environment should be selected (Kirkegaard & Sarwar, 1998; Morra & Kirkegaard, 2002; Zasada et al., 2003). The objective of this study was to evaluate the effect of selected Brassicaceae species that are commonly available in SA as cover/green manure crops, on a M. incognita population as well as soilborne, non-parasitic nematode (NPN) assemblages that are present in a potato field in the Christiana area. 4.2 Material and methods 4.2.1 Field trial The field trial was conducted during the 2010/2011 cropping season on a quarter (8ha) of a potato field (included in the survey as field 14; see Chapter 2) on the farm of Mr. Johan Greyling (Figure 4.1), which is situated in the Christiana area (See Figure 2; Chapter 2; North-West Province of SA). This field has a crop rotation history of Digitaria grass and maize for the past twenty years. High M. incognita population levels (ranging between 1 000-10 000 eggs and J2/20g tubers) prevailed at this particular field during the 2009/2010 growing season when potato was planted there. Although it is not normal practice to plant potato during two consecutive seasons, it has been done for the purpose of this experiment since 139 it was known that potato planted during the 2009/2010 season suffered quality losses due to high population levels of RKN. Before planting of the cover/green manure crops, soil samples from all plots were obtained (Sampling interval 1; Table 4.1) for baseline purposes. The cover/green manure crops, all belonging to the family Brassicaceae, planted before the potato follow-up crop in 2011 represented two mustard cultivars viz. Nemat (Brassica juncea) at a planting density of 6kg seeds/ha and Calient้ (Eruca sativa) at 11kg seeds/ha. In addition two cultivars of oilseed radishes (Raphanus sativus) Terranova and Doublet were planted at a rate of 20kg seeds/ha. These planting densities are recommended for commercial purposes for these particular cultivars (J. Fourie & B. Schoeman pers comm., September 2010). An untreated control as well as a synthetic nematicide ethylenedibromide (EDBฎ) at 40?/ha (a.s. 1 800g/?) treatment were also included in the trial, but were left fallow during planting of the Brassicaceae crops. Figure 4.1. The trial site where four Brassicaceae cultivars and a follow-up potato crop were planted at Mr. Johan Greyling’s farm in the Christiana area during the 2010/2011 growing season. The concentration of GL in the leaves and shoots of Brassicaceae crops decreases as the plant matures (Uppstrom, 1983). Since it is suggested that the breakdown products of GL are responsible for nematicidal activities, the age of incorporation of its aerial parts may significantly influence the suppression of nematode populations (Mojtahedi et al., 1991). Therefore, at the 50% flowering stage, which was eight weeks after planting of the Brassicaceae crops, soil and root samples were obtained from all Brassicaceae crops while only soil samples were obtained from plots that were designated for the untreated control and EDBฎ treatment. This represented sampling interval 2. Subsequently, the 140 aerial parts of the four Brassicaceae cultivars were slashed, chopped and ploughed into the soil using commercial implements. After being left in the soil for seven weeks to enable biofumigation and decomposition of organic material, potato tubers of cv. Mondial were planted on all plots including the untreated control and EDBฎ-fumigated ones. Before planting of the potato crop, twenty nematode samples were taken randomly per replicate for each treatment and combined. Subsequently one representative sub-sample for each treatment was taken for nematode extraction purposes. Nematode sampling dates and types of samples taken are indicated in Table 4.1. Two rows of potato (cv. Mondial) were planted in each plot (including those designated for the untreated control and EDBฎ treatments) after the cover/green manure crops were incorporated and left for seven weeks to enable biofumigation. Before planting of the potato crop, soil samples were obtained (Sampling Interval 3) from all plots for nematode extractions. The trial site was then ploughed and 800kg/ha fertiliser [N:P:K (3:1:5)] was incorporated across the site. Seven weeks after planting, at potato tuber initiation (Sampling Interval 4), root and tuber as well as soil samples were obtained for nematode extractions. The prevailing air temperatures at the trial site ranged between 15บC–30บC for the duration of the trial, while rainfall figures between 100mm–300mm were measured for the same period. The trial layout was a randomised complete block design (RCBD; Figure 4.2) with five replicates and six treatments (see paragraph above). Each treatment represented a 5m wide plot with a buffer zone of 200mm on each side. Furthermore, each plot differed in length ranging from 300m-150m. Therefore, samples were only taken inside a 100m long sampling site that stretched across the trial site (Figure 4.3) for the purpose of this study. Soil at this trial site consisted of 91.8% sand, 1.2% silt, 0.31% organic C and had a pH(H2O) of 6.55. 141 Figure 4.2. The trial layout where the Brassicaceae and potato crops were planted in the Christiana area during the 2010/2011 growing season represented a randomised complete block design (RCBD). 142 Figure 4.3. An aerial illustration (Google Earth, 2012) of the field where the Brassicaceae and potato crop trial was planted during the 2010/2011 growing season in the Christiana area. 143 Table 4.1. Nematode sampling dates for four sampling intervals, dates of planting of Brassicaceae and potato crops as well as type of samples collected during the 2010/2011 growing season for the trial planted in the Christiana area. Date when general production practices were conducted General practices conducted for trial purposes Date of nematode sampling Type of nematode sample(s) collected 15 October 2010 Planting of cover/green manure crops 15 October 2010 (Sampling interval 1) Soil samples only 15 November 2010 Application of fertiliser N:P:K 50% flowering of cover/green manure crops (8 weeks after planting) 14 December 2010 (Sampling interval 2) Soil and root/tuber samples 15 December 2010 Incorporation of aerial parts of cover/green manure crops Six weeks after incorporation of cover/green manure crops 24 January 2011 (Sampling interval 3) Soil samples only 29 January 2011 Application of EDBฎ on plots reserved for fumigation treatment 2 February 2011 Potato crop planted on all plots of trial site (7 weeks after incorporation of aerial parts of Brassicaceae crops) Tuber initiation of potato crop (seven weeks after planting) 14 March 2011 (Sampling interval 4) Soil, roots and tuber samples 4.2.2 Nematode extractions 4.2.2.1 Root (50g and 5g) and tuber (20g) samples The adapted NaOCl method was used for the extraction of RKN eggs and females from the 50g root sub-samples (see Chapter 2; Paragraph 2.2.3.2). Nematodes were extracted from 5g root as well as 20g potato tuber sub-samples of each treatment and replicate of both the Brassicaceae and potato crops using the adapted sugar-flotation method (Hooper et al., 2005), which has originally been described by (Coolen & D’Herde, 1972) (see Chapter 2; Paragraph 2.2.3.2). 144 4.2.2.2 Soil samples (200g) Extraction of nematodes, both PPN and NPN, from 200g of each soil sub-sample collected for each treatment and replicate was done using the adapted decanting and sieving method (Hooper et al., 2005), which has originally been described by Cobb, 1918. The latter was followed by the sugar centrifugal-flotation method (Caveness & Jensen, 1955). • Decanting and sieving method Principle This method is based upon the density and size of nematodes. Soil samples were mixed with water in a 5L bucket, stirred manually using a kitchen spoon and allowed to settle for 30 seconds. During this time about 90% of the nematodes stay in suspension. About 20% of the nematodes originally present in the soil sample will be lost using this method (Cobb, 1918). Procedure A 200g soil sample was soaked in tap water where soil particles and debris with a diameter of more than mm were removed b\ passing the sample throXgh a ?m-mesh sieve, which was nested on a 5L plastic bucket. The soil residue on the sieve was washed for about 2 minutes, discarded and the bucket filled up to 5 litres with tap water. The soil within the bucket was then thoroughly mixed with the water and the mixture allowed to settle down for about 30 seconds. The sediment that had settled to the bottom of the bucket was discarded when the dissolved soil-water mixture was decanted through a ?m-mesh sieve. The nematodes, however, that were contained within the soil-water mixture were left behind on the latter sieve. The entire procedure was repeated again by adding tap water to the sediment present at the bottom of the 5L bucket. The process was repeated as described above until the nematodes and fine soil partiFles retained on the ?m-mesh sieve were washed into 50ml centrifuge tubes. • Water centrifugation The soil within the centrifuge tubes for each sample was centrifuged for 5 minutes with a Relative Centrifugal Force (RCF) of 3000rpm using a Hettich Zentrifugen Roto fix 32 A centrifuge. After centrifugation, the supernatant that contained debris was carefully decanted and discarded. Nematodes were at this stage present at the bottom of the centrifuge tubes due to the centrifugation process. From 145 this step onwards, the nematodes were subjected to the sugar centrifugal-flotation method for final extraction from each of the soil samples. • Sugar centrifugal-flotation method Principle This method is based on the specific gravity of nematodes. Terrestrial nematodes have a specific gravity of about 1.08, while that of marine nematodes are approximately 1.13. After the centrifugation process in water that was done during the decanting and sieving method, only organic material with a specific gravity lower than 1 remains in suspension and is discarded from each of the centrifuge tubes. When centrifuged in a sugar solution with a higher specific gravity (1.15) than that of the nematodes, these parasites will remain in suspension and this way being separated from soil particles with a specific gravity larger than 1.15. Procedure a) A sucrose solution (specific gravity of 1.15) was added to each of the centrifuge tubes containing the nematodes. The sucrose solution was prepared by adding 624g sugar to 1L of tap water. b) The sugar solution and the sediment containing the nematodes in each of the centrifuge tubes were thoroughly stirred with a spatula and centrifuged for 1 minute at a RCF of 4000rpm. The spatula was rinsed in clean tap water every time after it was used to stir the solution in the separate centrifuge tubes to prevent transfer of nematodes from one sample to the next. c) After the sugar centrifugation process was completed, the supernatant that contained the nematodes was deFanted on a ?m-mesh sieve and immediately rinsed with tap water. This was done to limit the time that the nematodes were exposed to the sugar solution to less than four minutes. This way plasmolysis of the nematodes was prevented. d) The nematodes were finally washed into a 100ml plastic sample bottle for counting of RKN J2. For NPN counting and identification to family/genus level were done. 146 4.2.3 Identification of Meloidogyne species from the Christiana trial site 4.2.3.1 Morphological identification • Identification of Meloidogyne females from infected tomato roots The Meloidogyne sp. present at the trial site was reared in vivo to obtain mature females both for morphological and molecular identification (see Chapter 2, paragraph 2.2.6). For morphological identification, galls containing mature females were selected on root fragments of tomato (cv. Rodade) plants. Root tissue was gently forced open with a forceps and scalpel to remove adult females. Each individual female was transversely dissected behind the region where the body was swollen and the body tissues pushed out. The head of each female was transferred to a drop of glycerin/lactophenol on a glass microscope slide. The area on the cuticle containing the perineal pattern was removed and placed on a perspex cutting block in a drop of glycerin/lactophenol with the dome-like posterior end facing upwards. The perineal pattern was trimmed to a flat square and transferred next to the female’s head on the microscope slide, exterior side facing upwards (Hartman & Sasser, 1985). Twenty-one heads and perineal patterns of mature females (Kleynhans, 1991) were cut and transferred, next to each other, on a single microscope slide that was sealed with a cover slip and colourless Cutex. This enabled an accurate indication of the sp. or spp. present for this trial site. The slide was properly labelled and sp. identification was done under a light microscope 4.2.3.2 Molecular identification The same procedure was followed for identification of RKN using molecular techniques as described in Chapter 3, paragraph 3.2. 4.2.4 Data analysis Data for both PPN and NPN were subjected to an analysis of variance (ANOVA) using Statistica Version 10 for Windows. Means was separated using the Tukey Test where P?.. Nematode data was transformed to log10(x+1) to remove the inherent variability before analyses. In addition, nematode faunal analyses were performed for NPN as indicators of the food web structure, status and functionality, as well as resource availability (Ferris et al., 2001; Ferris & Matute, 2003) at this particular trial site as described in Chapter 2, paragraph 2.4.3. 147 4.3 Results 4.3.1 Identification of root-knot nematode (RKN) species The RKN species present at the Christiana trial site was identified as M. incognita using molecular techniques (Zijlstra et al., 2000) as well as by using morphological characteristics (Kleynhans, 1991). Morphological identification of this species was also confirmed by Dr. Mariette Marais (Nematode Taxonomist: ARC-PPRI, Pretoria). Figure 4.4. A perennial pattern of a mature M. incognita female reared in vivo from J2 present in soil samples obtained from the site at Christiana where the Brassicaceae and potato trial was planted during the 2010/2011 growing season. Trial site 148 Figure 4.5 (A & B). Amplification products (1 200bp) of mature root-knot nematode females (lanes 2-7), present in fields in the Christiana area where the Brassicaceae trial was conducted during the 2010/2011 growing season, identified by means of the sequence characterised amplified region- polymerase chain reactions (SCAR-PCR) method. [lane 1 (A) = positive standard from in vivo reared Meloidogyne incognita populations earlier positively identified; lane 1 (B) = positive standard from in vivo reared Meloidogyne javanica populations earlier positively identified; lane 2 = field 13 (A1), lane 3 = field 16 (B2); lane 4 = field 28 (GH17); lane 5 = field 19 (S5); lane 6 = field 26 (D4); lane 7 = field 14 (A6 – trial site where Brassicaceae trial was conducted); lane 8 = field 5 (T6T5); lane 9 = blank tap water control and lane 10 = 1kb DNA marker]. PCR with the M. incognita specific SCAR-primer resulted in amplification of the M. incognita 1 200bp SCAR-fragment for the ten mature RKN females that were pooled after they have been obtained from in vivo-reared tomato roots (cv. Rodade) that were planted in soil containing RKN eggs and J2 from the site (represented by lane 7; Figure 4.5A) where the trial was conducted. These results agreed with and were substantiated by those for the positive standard used during this part of the study which represented M. incognita from a monoculture, in vivo reared population. 1 200bp 670bp 1 2 3 4 5 6 7 8 9 10 Meloidogyne incognita Meloidogyne javanica 1 2 3 4 5 6 7 8 9 10 Trial site Trial site A B 149 4.3.2 Root and tuber data for Brassicaceae and potato crops Root (Calient้ and Nemat) and tuber (Doublet and Terranova) samples were obtained during 50% flowering of these cover/green manure crops (Sampling interval 2), while root and tuber samples of potato were obtained during tuber initiation of the potato crop (Sampling interval 4). During the other two sampling intervals (1 and 3) no crops were present and only soil samples were collected. 4.3.2.1 Roots and tubers (50g and 5g): Brassicaceae crops Table 4.2. Meloidogyne incognita egg and J2 numbers [log10(x+1)] in root/tuber samples of Brassicaceae crops planted before potato at a trial site in the Christiana area during the 2010/2011 growing season (means in parenthesis represents the values that were back transformed and not the real means). Sampling interval Untreated control (T 1) Terranova (T 2) Doublet (T 3) Nemat (T 4) Calient้ (T 5) EDBฎ (T 6) P value F ratio 50g roots/tubers (Brassicaceae crops) 2 Left fallow 3.9 (48) 4.7 (109) 3.4 (29) 5.2 (180) Left fallow 0.8184 0.31 5g roots/tubers (Brassicaceae crops) 2 Left fallow 1.8 (5) 1.1 (2) 1.5 (3) 2.7 (14) Left fallow 0.3114 1.34 Only M. incognita J2, were present in root (Calient้ and Nemat) and tuber (Doublet and Terranova) samples collected from the various cover/green manure cultivars (Sampling interval 2) planted before a potato crop at this trial site. For M. incognita eggs and J2 numbers/50g roots/tubers collected during sampling interval 2 (50% flowering stage), no significant differences (P?. e[isted between the foXr Fovergreen manXre crops (Table 4.2). Meloidogyne incognita population levels in 50g roots/tubers ranged from 29 (Nemat) to 180 (Calient้). Both the untreated control and EDBฎ plots were left fallow during this stage. No significant difference (P?. e[isted among the different %rassiFaFeae treatments with regard to M. incognita J2 numbers/5g roots/tubers for sampling interval 2 (Table 4.2). J2 population levels were, 150 however, very low and ranged from 2 (Doublet) to 14 (Calient้). Both the control and EDBฎ plots were left fallow and, therefore, no root/tuber data were obtained during sampling interval 2. 4.3.2.2 Roots (50g and 5g) and tubers (20g): Potato crop Table 4.3. Meloidogyne incognita egg and J2 numbers [log10(x+1)] in root and tuber samples of potato crop planted at a trial site in the Christiana area during the 2010/2011 growing season (means in parenthesis represents the values that were back transformed and not the real means). Sampling interval Untreated control (T 1) Terranova (T 2) Doublet (T 3) Nemat (T 4) Calient้ (T 5) EDBฎ (T 6) P value F ratio 50g roots (Potato crop) 4 10.2a (26902) 9.8ab (18032) 10a (22025) 10.2a (26902) 11a (59873) 8.2b (3639) 0.0140 6.27 5g roots (Potato crop) 4 7a (1095) 6.2ab (492) 5.5ab (244) 5.6ab (269) 7.7a (2207) 4.2b (66) 0.0025 5.58 20g tubers (Potato crop) 4 1.7 (4) 1.5 (3) 0.9 (1) 1 (2) 1 (2) 2.4 (10) 0.0600 2.59 Meloidogyne incognita was the only PPN present in root and potato tuber samples obtained during sampling interval 4. During tuber initiation (Sampling interval 4) of the potato crop M. incognita population levels were substantially higher than during the 50% flowering stage (Sampling interval 2) of the Brassicaceae crops for both the 50g and 5g roots (Tables 4.2 & 4.3). No significant difference (P?. in egg and - numbers/50g roots was, however, evident among the plots where the four cover/green manure crop treatments were planted before the potato crop and the untreated control contained the lowest egg and J2 population levels. EDBฎ treated plots, however, maintained significantly (P?. higher M. incognita egg and J2 numbers/50g roots compared to the three Brassicaceae treatments Doublet, Nemat and Calient้ as well as the untreated control plots, but not compared to Terranova. Population levels of this RKN species were, however, still high (3 639 eggs and J2/50g roots) for the EDBฎ treatment. Plots 151 planted with Calient้ had the highest number of eggs and J2 numbers/50g roots (59 873 eggs and J2/50g roots), followed by Nemat (26 902), Doublet (22 025) and Terranova (18 032). Untreated control plots had similar egg and J2 population levels than plots planted with Nemat. For the 5g root samples, EDBฎ treated plots maintained significantly (P?. less J2/5g roots than the untreated control and Calient้ treated plots (Table 4.3). Population levels of M. incognita J2 ranged from 66/5g roots (EDBฎ) to 2 207/5g roots (Calient้). No significant (P?. difference was obtained among the six treatments in terms of M. incognita eggs and J2 numbers/20g tubers for sampling interval 4 (Table 4.3). Plots that were treated with EDBฎ had the highest number of J2/20g tubers (10), Nemat and Calient้ (2) and plots that were planted with Doublet (1) had the lowest number of J2/20g tubers. 4.4 Soil data 4.4.1 Plant-parasitic nematodes (PPN) PPN that were present in 200g soil samples at this trial site included the endoparasitic M. incognita as well as Pratylenchus spp., semi-endo/ectoparasitic Helicotylenchus spp., Rotylenchus spp. and Scutellonema spp. and ectoparasitic Criconema and Tylenchorhynchus spp. Since population levels of all these PPN, except those for M. incognita, were very low throughout the duration of the trial (Table 1; Appendix 1), only those for M. incognita will be discussed further. 152 Table 4.4. Meloidogyne incognita J2 numbers/200g soil samples [log10(x+1)] for four sampling intervals from a trial site in the Christiana area where Brassicaceae crops were planted before potato during the 2010/2011 growing season (means in parenthesis represents the values that were back transformed and not the real means). Sampling interval Untreated control (T 1) Terranova (T 2) Doublet (T 3) Nemat (T 4) Calient้ (T 5) EDBฎ (T 6) P value F ratio 1 0.7 (2) 0.3 (1) 0.3 (1) 0.3 (1) 2 (9) 1.5 (11) 0.1437 1.88 2 1.2a (3) 0.3ab (0.6) 0.3ab (0.4) 0.2b (0.4) 0.3ab (0.4) 0.3ab (0.4) 0.0324 3.07 3 1.5a (4) 0.2b (0.4) 0.3b (0.6) 0b 0.1b (0.2) 0.4b (0.6) 0.0001 9.01 4 3 (55) 3 (92) 1.7 (22) 1 (5)a 5 (300)b 2.4 (20) 0.0521 2.68 Although M. incognita J2 numbers were generally very low in soil samples at this trial site throughout the duration of the trial, it increased substantially in plots of all treatments included from sampling intervals 1 to 4 (Table 4.4). Significant (P?. differenFes were onl\ evident among the various treatments during sampling intervals 2 and 3, with the untreated control containing significantly (P?. lower - nXmbersg soil than those for Nemat and the other treatments included (the three Brassicaceae cultivars as well as EDBฎ). 153 4.4.2 Non-parasitic nematodes (NPN) Table 4.5. Functional guilds, to which non-parasitic nematode genera/families (identified in soil samples from the Christiana trial site where Brassicaceae and potato crops were planted), were assigned according to their feeding habits and life stage characteristics (Bongers, 1990). Genus Guild Cephalobus Ba1 Cruznema Ba1 Diploscapter Ba1 Mesorhabditis Ba1 Panagrolaimus Ba1 Rhabditis Ba1 Acrobeles Ba2 Acrobeloides Ba2 Chiloplacus Ba2 Eucephalobes Ba2 Aphelenchus Fu2 Aphelenchoides Fu2 Tylenchus Fu2 Doryliamidae Om4 ba: bacterivores, fu: fungivores, om: predator (omnivore/carnivore). Suffix numbers represent cp values for the various taxa. NPN present at this trial site were categorised into different non-parasitic nematode guilds, representing bacterivores (cp 1&2), fungivores (cp2) and omnivores [cp4 (Table 4.5)]. 154 Figure 4.6. Enrichment (EI) and structural indices (SI) of non-parasitic nematode (NPN) assemblages for four sampling intervals for the various treatments applied at a trial where Brassicaceae and potato crops were planted in the Christiana area during the 2010/2011 growing season. Categorisation of both the different treatments and sampling intervals included in the Brassicaceae and potato trial in the Christiana area differed substantially with regard to its placement in the simplified food web (Figure 4.6). These results substantiate the typical spot occurrence of nematodes in soils (Been & Schomaker, 2006) and represent the inherent variation in nematode communities in terms of their distribution before the trial commenced. 155 • Sampling interval 1 (before commencement of the trial) NPN assemblages in soil samples obtained from plots designated for Doublet, Nemat and, untreated control treatments obtained during the first sampling interval 1 (indicated by green dots) were generally categorised within the “Stress Enriched” quadrant (Figure 4.6), which represents a disturbed food web condition (Ferris et al., 2001). Those obtained for plots designated for Terranova were plotted in the “Stable Enriched” quadrant, indicating a maturing food web while those designated for Calient้ were plotted in the “Stress Depleted” quadrant that represents a degraded food web. Plots designated for EDBฎ applications were shared between the “Stress Enriched” and “Stress Depleted” quadrants and are thus classified as disturbed and/or degraded. Soil, NPN communities for plots designated for planting of Doublet, untreated control and Nemat were dominated by Ba1 on the Enrichment Trajectory (medium-high EI), while those for Nemat plots were dominated by Fu2 (medium EI) (Figure 4.6). On the Structure Trajectory plots designated for Doublet treatments were dominated by Ca2, Ca3, Fu3 and Ba3 (low SI). Plots designated for Nemat, untreated control and EDBฎ plots were, however, dominated by Om4, Ca4, Fu4 and Ba4 (medium SI) in terms of the Structure Trajectory. Soil, NPN communities for plots designated for planting Terranova were dominated by Ba1 (medium- high EI) and Om4, Ca4, Fu4 and Ba4 (medium-high SI) in terms of the Structure Trajectory (Figure 4.6). Plots designated for planting Calient้ were, however, dominated by Fu2 (low-medium EI) on the Enrichment Trajectory and Om4, Ca4, Fu4 and Ba4 (low-medium SI) on the Structure Trajectory. • Sampling Interval 2 (50% flowering of the Brassicaceae crops) During the second sampling interval (indicated by brown dots) changes were evident in terms of categorisation of the various treatments in terms of their NPN assemblages regarding their food web status (Figure 4.6). Food web conditions for EDBฎ (medium EI and low-medium SI) and untreated control (medium-high EI and medium SI) plots generally remained disturbed (“Stress Enriched”; quadrant A), while those for Doublet improved from disturbed (“Stress Depleted”; quadrant A) with high EI but low SI to structured (“Stable Depleted”; quadrant C) with low-medium EI but medium-high SI. Food web conditions for plots planted with Calient้ improved from degraded (“Stress Depleted”; quadrant D) with low-medium EI and low SI to maturing (‘Stable Enriched”; quadrant B) with medium- high EI and SI. However, plots planted with Terranova changed from maturing food web conditions 156 (“Stable Enriched”; quadrant B) with high EI and medium-high SI to degraded (“Stress Depleted; quadrant D) with low-medium EI and SI. Sampling interval 3 (eight weeks after incorporation of aerial parts of Brassicaceae cultivars – indicated by orange dots) • Sampling interval 3 (incorporation of cover/green manure crops) During the third sampling interval (indicated by orange dots) changes in terms of food web conditions were again observed for the six treatments (Figure 4.6). Plots designated for EDBฎ treatments changed from disturbed (medium EI and low-medium SI)] to maturing (medium-high EI and SI) food web conditions, while those for the untreated control changed from disturbed (medium to high EI and medium SI) to degraded (medium EI and low SI). Food web conditions for Doublet remained structured but decreased in terms of its EI and increased in terms of its SI. Those for Calient้ changed from matured (medium-high EI and SI) to degraded (medium EI and low-medium SI). For Nemat, soil food web conditions changed from structured (low-medium EI and medium SI) to mature (medium-high EI and SI) and for Terranova from degraded (low-medium EI and SI) to matured (medium-high EI and SI). • Sampling interval 4 (tuber initiation of potato crop) For the fourth sampling interval (indicated by red dots), soil food web conditions for all four Brassicaceae treatments were indicated as maturing (high EI and medium-high SI), while those for EDBฎ (high EI and low-medium SI) and untreated control (high EI and medium SI) plots were indicated as disturbed (Figure 4.6). In general a substantial improvement in terms of soil food web conditions for three of the four Brassicaceae cultivars from disturbed (for Doublet and Nemat) and degraded (for Calient้) were evident from Sampling Intervals 1-4. In contrast, soil food web conditions for the EDBฎ and untreated control plots, except that the EI of both the latter treatments increased during the duration of the experiment, generally remained in disturbed quadrant. While an increase in Ba1 was the main trend visible for both the latter treatments during the duration of this trial, the same were evident for the four Brassicaceae treatments as well as in increase in Om4, Ca5, Fu5 and Ba5. 157 4.5 Discussion Second-stage juveniles (J2) of the predominant endoparasitic species Meloidogyne incognita present in soil at the Christiana trial site were able to infect and develop to mature, egg-producing females in roots/tubers of all four of the Brassicaceae cultivars evaluated during this study. The build-up of relatively high population levels of this local M. incognita population in tubers of the two oilseed radish (Doublet and Terranova) and roots of the two mustard (Calient้ and Nemat) cultivars at this trial site was not expected. Various reports (Mojtahedi et al., 1993; Kirkegaard & Sarwar, 1998; Morra & Kirkegaard, 2002; Zasada et al., 2003; Lazzeri et al., 2004; Curto et al., 2005) suggested that Brassicaceae crops generally are poor hosts to PPN, particularly Meloidogyne spp., which was not the case in this study. Furthermore, the four Brassicaceae cultivars used in this study were unable to reduce M. incognita population densities at the Christiana trial site by means of their biofumigation effects. The latter trend was reflected in the high population levels of this RKN sp. in roots of a follow-up potato crop sampled during tuber initiation. At this stage the approach towards the objective of the Brassicaceae trial had to be changed and a registered synthetic nematicide was applied to limit potential RKN damage to potato tubers and this way minimise quality losses. Although it is not normal practice for producers to plant potato during successive growing seasons on the same field, it was only done for the purpose of this study, as mentioned earlier. In addition to data obtained on the plant-parasitic species M. incognita during this study, novel information about the NPN assemblages and their substantial changes in terms of community structures as a result of Brassicaceae amendments were gathered for annual crops, in this case potato. Previous studies in SA in this regard for PPN and NPN assemblages have only been done on perennial crops such as sugar cane and vineyard (Berry & Wiseman, 2003; Berry & Rhodes, 2006; Kruger et al., 2011) as well as for common scab on seed potatoes (Gouws & Wehner, 2004). Valuable information, indicating pronounced shifts of NPN assemblages in plots where the four Brassicaceae cultivars were planted from the “Stress Depleted” (degraded) and/or “Stressed Enriched” (disturbed) to the “Stable Enrich” (maturing) quadrants obtained during this study indicate the positive effect of these crops in terms of soil health. In terms of the maintenance of M. incognita in roots/tubers of Brassicaceae spp. used all four cultivars generally maintained relatively high RKN eggs and J2. Although cv. Nemat maintained the lowest egg and J2 numbers, it did not differ significantly from the other three cultivars. In addition, crop roots of the follow-up potato crop planted in plots seven weeks after aerial plant parts of cv. Nemat were 158 incorporated maintained similar, very high M. incognita population levels compared to those in plots where the other three Brassicaceae cultivars were planted and their aerial parts incorporated. Lazerri et al. (2004) and Curto et al. (2005), however, reported that cv. Nemat was a poor host to the M. incognita populations used in their studies. Results obtained in terms of the host suitability and poor biofumigation effects of Brassicaceae species evaluated during this study with regard to the local M. incognita population is in contradiction with those by Kirkegaard and Sarwar (1998), Morra and Kirkegaard (2002), Zasada et al. (2003), Lazzeri et al. (2004) and Curto et al. (2005). The latter authors evaluated different Brassicaceae species, namely E. sativa, B. juncea and R. sativus for their high active GL content and biocidal activity against M. incognita J2. All varieties screened during the latter studies were recommended for use in pest management programmes based on their ability to reduce M. incognita infestation levels in the soil due to their high GL content and subsequent good nematicidal activity. Mojtahedi et al. (1993) also reported that rapeseed of cultivar Jupiter (Brassica napus) was effective in reducing M. chitwoodi population densities on field-grown potato. However, other studies in which the effects of Brassicaceae species on M. incognita were evaluated were similar to results obtained during this study. For example, Monfort et al. (2007) found that M. incognita was capable of reproducing on a majority of the Brassicaceae species they screened. Li้banas and Castillo (2004) further reported that all the Brassicaceae varieties they tested were hosts to M. arenaria, M. incognita and M. javanica. Also, Johnson et al. (1992) found no significant reduction of nematode numbers after incorporation of rapeseed green manures to control M. incognita and M. javanica on squash (Cucurbita pepo) in Georgia, North America. Furthermore, studies with M. javanica with regard to various Brassicaceae species suggested that although Brassicaceae crops are hosts to these RKN, mechanism(s) responsible for their poor host status become effective only after invasion by J2 (McLeod et al., 2001). Although Brassicaceae crops are hosts to RKN, they are poorly invaded by J2 and suppress their concurrent development, reducing their risk to increase in numbers in roots/tubers of Brassicaceae crops before their incorporation into soil as green manures (McLeod et al., 2001). The latter scenario did not seems to be true for the four Brassicaceae cultivars used during this study since relative high populations levels, except for cv. Nemat, were recorded for the M. incognita population in roots/tubers at 50% flowering of these crops. In addition, the high M. incognita population pressure that existed at the Christiana trial site is expected to have contributed to the poor performance of these cultivars in terms of their inability to reduce 159 population levels of these parasites during this study in such a short period of time. Another possible hypothesis is that the persistence and the activity of some soil micro-organisms, of which no information was obtained during this study, could have played an additional role in isothiocynate loss or degradation of the Brassicaceae cultivars (Rumberger & Marschner, 2003). Also, green manure cultivars that have high leaf concentrations of GL are needed before the potential offered by Brassicaceae crops in terms of reducing economically important PPN are fully realized (McLeod & Steel, 1999). GL concentrations for the four cultivars used in this study as well as that of other Brassicaceae cultivars that have potential as cover/green manure crops to be used under local conditions should be determined to indentify superior ones in this regard. It should also be noted that for future research, more than one incorporation of aerial amendments of Brassicaceae crops should be considered and that trials should be conducted over longer periods of time and over more than one growing season using different planting densities of different Brassicaceae crops, in order to better conclude about their efficacy in terms of biofumigation. The latter reasons may be responsible for the lack/reduced nematicidal activity of the four Brassicaceae cultivars used against the local M. incognita during this study. These recommendations are supported by Zasada and Ferris (2004) who reported that suppression of different PPN species is dependent on the rate of green manure application in soil as well as Brassicaceae GL concentrations within aerial plant parts. In terms of the NPN assemblages, pronounced changes in terms of their categorisation within the simplified soil food web (Bongers & Ferris, 1999; Ferris et al., 2001) were evident for plots where the Brassicaceae cultivars were planted compared to those treated with EDBฎ as well as the untreated control ones. However, although the diversity of NPN increased at this trial site as a result of the amendment of Brassicaceae crops, population levels of M. incognita in roots of potato at tuber initiation were still high in all plots including Brassicaceae-amended ones. In general results for soil, NPN assemblages gained during this study showed that substantial increases in the diversity (genera and/or families) and population levels of particularly bacterial- and fungus-feeding NPN were evident for all six treatments during the duration of the trial. The presence of enrichment and general opportunistic NPN guilds, representing bacterivores, carnivores and fungivores with cp values of 1 & 2 thus resulted in high EI for all treatments at the end of the Brassicaceae trial (Sampling Interval 4). However, higher NPN diversity represented by guilds with cp values (3-5) higher than 2 was also present in Brassicaceae-amended plots. The predominant bacterivores (Ba1) that were present in EDBฎ as well as the untreated control plots are regarded as entry-level indicators of enrichment, multiplying rapidly by feeding on other soilborne 160 micro-organisms that decompose organic matter (Ferris et al., 2001). Predominance by Ba1 communities in particular resulted in the increased and high EI for the latter treatments during fourth sampling interval. The presence of enrichment and general opportunistic NPN guilds, representing bacterivores, carnivores and fungivores with cp values of 1 & 2 thus resulted in high EI for all treatments at the end of the Brassicaceae trial (Sampling Interval 4). However, higher NPN diversity represented by guilds with cp values (3-5) higher than 2 was also present in Brassicaceae-amended plots, indicating the extent to which decomposition of organic material present in soil has occurred (Ferris & Bongers, 2006; Ferris & Bongers, 2009). Disruption of the soil as a result of cultivation practices could have contributed to the latter scenario for these treatments. The absence of NPN guilds with cp values ranging between 3 and 5 in EDBฎ and untreated control plots, however, indicated that soil food webs in these plots were disturbed and unstable (Ferris et al., 2001; Ferris & Bongers, 2009). The concurrent occurrence of bacterivores, fungivores, carnivores as well as omnivores with higher cp values (4-5) in Brassicaceae-amended plots indicates that the NPN communities in the latter plots benefited from and responded positively to such amendments. Although enrichment opportunists (Ba1) built up and had high EI values for all Brassicaceae-amended plots during sampling interval 4, general opportunists (Fu2) that are not necessarily dependent on the same resources were replenished by persisters during further succession. The latter represented bacterivores, fungivores, carnivores and omnivores with cp values ranging between 3-5. Amendment using the four Brassicaceae crops thus resulted in an increase in the diversity of the latter NPN guilds, resulting in soil food webs being stable, complex, structured and mature (Ferris et al., 2001; Ferris & Bongers, 2009). The pronounced shifts in NPN community structures in Brassicaceae-amended plots obtained during this study coincide with those from other studies where the effects of Brassicaceae amendments were evaluated on NPN assemblages in soil environments (Osler et al., 2000; Stirling & Stirling, 2003). NPN nematodes are a food source for natural enemies, representing other soilborne micro-organisms such as bacteria, fungi, mites, protozoa and others, of nematodes (Stirling, 1991). These NPN assemblages play an important role in nutrient cycling and plant nutrition (Ingham et al., 1985; Griffiths, 1994). Therefore, information gained on improvement of soil health using Brassicaceae amendments during this study is important, particularly in SA where no official reports on this topic has been published, from a soil health perspective. It also serve as a reminder that as we strive to increase the efficacy of biofumigation, we need to conserve beneficial organisms, which is an achievable goal (Stirling & Stirling, 2003). Since NPN nematodes are less sensitive than PPN to the toxins produced by Brassicaceae crops (Halbrendt & Jing, 1996) and because their generation times are relatively short, 161 populations tend to recover quickly following treatment with crop residues of Brassicaceae species that have nematicidal characteristics (Sturz & Kimpinski, 1999). In addition, proof that organisms such as PPN differ markedly in their sensitivity to ITC’s (Sarwar et al., 1998) and that GL profiles vary between Brassicaceae species and are influenced by the environment (Kirkegaard & Sarwar, 1998; Sarwar & Kirkegaard, 1998), suggests that it may eventually be possible to maximise the effect of Brassicaceae amendments on key pathogens such as RKN, while minimising their impact on non-target, beneficial species (Stirling & Stirling, 2003). 4.6 Conclusions According to the authors’ knowledge, this is the first study on the effects of Brassicaceae crops both for PPN (M. incognita in this case) and NPN on potato in SA. Although none of the Brassicaceae crops used in this study resulted in a significant reduction of population levels of M. incognita (predominant PPN) in roots and/or tubers of Brassicaceae and potato crops, pronounced improvement in terms of soil health has been demonstrated using categorisation of NPN assemblages according to a simplified food web. To optimise the effect of Brassicaceae crops in potato-based cropping systems in SA, the following recommendations are proposed: • Prevent the build-up of RKN to very high population levels in potato fields before planting of a green manure crops such as Brassicaceae. • Screening of all Brassicaceae cultivars available in SA for their host status to various economically important RKN that occur in local potato-producing areas. • Experimenting with higher cover crop planting densities. • Slashing and incorporation of aerial parts should be done more than once during the growth period. • Similar trials should be conducted over a longer period during which cover crops are planted on a more frequent basis. • Glucosinolate content should be determined for local Brassicaceae crops. • Combination of biofumigation with other environmentally-friendly nematode control strategies, such as soil solarisation, steam sterilisation, natural nematicidal products. 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Identification of Meloidogyne incognita, M. javanica and M. arenaria using sequence characterised amplified region (SCAR) based PCR assays. Nematology 2: 847-853. 168 CHAPTER 5 GENERAL CONCLUSIONS AND FUTURE PROSPECTS 5.1 Conclusions The two nematode surveys conducted during the 2010 and 2011 growing seasons resulted in one plant-parasitic nematode (PPN) family, 13 genera and 16 nematode species being identified from both root and soil samples from the 31 fields in which potato is included as a rotation crop every eight years in the Christiana area of South Africa (SA). Other rotation crops included in these fields are maize, sunflower, onion, as well as grass cover crops. Root-knot nematode (RKN) species were the predominant genera with two species being identified using the molecular SCAR-PCR method, namely M. incognita and M. javanica. These two species generally also are the predominant ones, which cause problems to other rotation crops included in potato fields in the Christiana area. Results obtained during this study substantiated data that RKN are widely distributed in the Christiana potato-production area of SA where it causes severe damage to the crop. Furthermore, it is also in agreement with reports that RKN have a wide host range and are considered economically one of the most important PPN genera that are locally associated with crops planted in rotation with potato such as maize, sunflower, onion, as well as grasses and weeds. In terms of the other economically important endo- and semi endo-/ecto- as well as ectoparasitic nematodes present in the 31 fields sampled, Pratylenchus spp. generally ranked second with individuals of the Hoplolaimidae and Trichodoridae families following. Ditylenchus spp. and J2 individuals of the Heteroderidae family (other than Meloidogyne spp.) were also present, but in low numbers. Also of significance is that one genus (Psilenchus) and six species (Criconemoides sphaerocephalus, Hemicriconemoides brachyurus, Paratrichodorus lobatus, Rotylenchus caudaphasmidis and Tylenchorhynchus ventralis) have never before been associated with maize, potato, sunflower, onion and grasses in cropping systems in SA and are thus new records. Concerning non-parasitic nematode (NPN) assemblages, this is the first official report for SA on categorisation of fields planted to annual crops in terms of their soil health status and thus represents baseline information in this regard. Nine NPN genera of which six were bacterivores and three fungivores as well as two predator families have been identified from soil samples obtained from the 31 fields. Bacterivore nematode genera were represented by a wide spectrum with individuals of the Rhabditis genus belonging to the Ba1 guild that dominated and resulted in high EI values, followed by 169 Cephalobus spp. (Ba2). The latter genera as well as the other bacterivorous genera identified from these fields commonly occur in soils in SA. These NPN are regarded as the beneficial nematodes of which population levels increase substantially and relatively quickly after the addition of organic material or any kind of disturbance such as cultivation practices. The fungivore nematode communities of the 31 fields sampled were represented by a smaller variety of genera, with Tylenchus spp. (Fu2) being predominant. In terms of the omnivore/predator and carnivore nematodes, individuals of the Dorylaimidae (Om2-5) and Mononchidae (Ca2-4) families were respectively present and are responsible for the high SI values obtained for these fields. In terms of soil food web conditions, the majority of the 31 fields were plotted in the “Stable Enriched” quadrant that indicates that the soils of such fields were maturing and thus relatively healthy in terms of the beneficial nematodes as indicated by the simplified food web. This scenario is surprising and could probably be explained by the inclusion of grass cover crops on these fields for five to six years of the eight-year cropping cycles practised by these producers. The dominant contribution of bacterivore nematodes (Ba1) contributed substantially to high EI values, while the presence of bacterivore, fungivore, omnivore/predator as well as carnivore guilds with higher cp values ranging, between 3-5, contributed to the medium to higher SI values for such fields. The presence of a few significant relationships between the various PPN as well as NPN nematode genera with regard to cultivation practices, nematicide use as well as cropping sequences revealed interesting trends, but have to be verified by additional results before any conclusions can be made in this regard. Only then will it add substantial value to producers, the related crop industries and the science of Nematology. However, of particular interest are findings that application of EDBฎ in combination with other nematicides rendered significantly lower population levels of some of the predominant PPN genera such as Meloidogyne and Pratylenchus spp. identified. Concurrently, indications that some NPN, i.e. Rhabditis and Prismatolaimus spp., had higher population levels where EDBฎ combined with other nematicides than EDBฎ alone, respectively, were applied also indicate that interesting and important trends could be revealed should this type of research be extended over more growing seasons. The same applies for some of the PPN and NPN that reacted significantly with regard to specific crop rotation sequences as well as cultivation practices. Validation of such information can have important implications for designing and planning of nematode management strategies. Unfortunately no significant effects were, however, found for either PPN as well as NPN with regard to soil characteristics such as percentages of sand, silt, clay and organic material. 170 Concerning the use of Brassicaceae amendments as an environmentally-safe alternative to synthetic nematicides, M. incognita were identified as the predominant PPN present at the field site where the trial was conducted during the 2010/2011 growing seasons. J2 of the latter RKN species were able to infect and develop to mature, egg-producing females in roots/tubers of all four the Brassicaceae cultivars evaluated. The presence of relatively high population levels of this local M. incognita population in tubers of the two oilseed radish (Doublet and Terranova) and roots of the two mustard (Calient้ and Nemat) cultivars at this trial site were, however, not expected and is not in agreement with the majority of reports related to this topic. The subsequent build-up of high population levels of this RKN species in roots of a follow-up potato crop sampled during tuber initiation, further substantiated results that the Brassicaceae cultivars used was not successful in reducing numbers of this parasite effectively at this specific trial site. As a result, the approach had to be reconsidered during tuber initiation and a registered synthetic nematicide was applied to limit RKN damage to potato tubers and this way minimise quality losses. Although results in terms of the host suitability showed that Nemat was the poorest host of the four Brassicaceae cultivars used, it still supported development of individuals of this RKN population. Also, no significant differences in terms of RKN numbers in roots of the follow-up potato crop in plots amended with Nemat compared to those for the other three Brassicaceae cultivars were recorded. Although it is not normal practice for producers to plant potato during successive growing seasons, it was only done for the purpose of this study. In addition to data obtained on the plant-parasitic species M. incognita during this study, novel information about the NPN assemblages and their substantial changes in terms of community structures as a result of Brassicaceae amendments were gathered for annual crops, in this case potato. Previous studies in SA in this regard have only been done for perennial crops such as sugar cane and vineyard as well as for common scab on seed potatoes. Valuable information, indicating pronounced shifts of NPN assemblages in plots where the four Brassicaceae cultivars were planted from mainly the “Stress Depleted” (degraded) and/or “Stressed Enriched” (disturbed) to the “Stable Enriched” (maturing) quadrants indicate the positive effect of these crops in terms of soil health. In general substantial increases in the diversity (genera and/or families) and population levels of particularly bacterial- and fungus-feeding NPN were evident for all six treatments during the duration of the trial. The presence of enrichment and general opportunistic NPN guilds, representing bacterivores and fungivores with cp values of 1 and 2 thus resulted in high EI values for all treatments at the end of the Brassicaceae trial (Sampling interval 4). However, higher NPN diversity represented by guilds with cp values (3-5) higher than 2 that was also present in Brassicaceae-amended plots and resulted in the higher SI values. The latter indicate that such fields were maturing since it represents stability, food 171 web complexity and connectance in terms of the prevailing nematode assemblages. High EI values were most probably mainly due to disruption of the soil as a result of cultivation practices before and during planting of the crops, while the presence of Brassicaceae amendments most probably contributed predominantly to the presence of nematode diversity in terms of the higher guilds present in soil of such plots. The outcome of this study, which included the two nematode surveys as well as the evaluation of the effect of Brassicaceae cultivars in terms of their host suitability to and biofumigation effects on a prevailing M. incognita population, provided valuable information to producers in the Christiana area. Such information can add value since it forms baseline studies that can already be build upon in efforts to manage RKN in particular but also NPN assemblages in their potato-based cropping systems. 5.2 Future prospects The current study can be improved and value added to the results obtained by addressing the following issues: ? Obtaining of more soil and crop root/tuber samples from the 31 fields in the Christiana area to i) identify PPN present in each of the fields to species level and ii) increase the probability of discovering significant effects of various cultivation practices, nematicides as well as crop rotation sequences both for PPN and NPN. ? Screening of all Brassicaceae cultivars available in SA to determine i) glucosinolate content and ii) their host status to economically important RKN species that occur in potato-producing areas. ? Experimenting with Brassicaceae cultivars by planting higher densities to optimise their use in terms of amendment and biofumigation. ? Slashing and incorporation of aerial parts of Brassicaceae cultivars more than once during the growth period to determine the minimum application(s) in this regard. ? Conducting similar Brassicaceae trials over a longer period during which such crops are planted on a more frequent basis in potato-based cropping systems. ? Combination of biofumigation using Brassicaceae cultivars with other environmentally-friendly nematode control strategies, such as soil solarisation, steam sterilisation, natural products with nematicidal activity. ? Recording of pH and rainfall data to determine whether there could be correlations between the pH or amount of rainfall during the trial and degrading of parts of Brassicaceae crops in soil. 172 Appendix A PPN (excluding M. incognita) and NPN J2 numbers/200g soil Legend Treatments: T1 = Treatment 1 (Untreated control) T2 = Treatment 2 (Terranova) T3 = Treatment 3 (Doublet) T4 = Treatment 4 (Nemat) T5 = Treatment 5 (Calient้) T6 = Treatment 6 (EDBฎ treatment) 173 Table 1. PPN (excluding M. incognita) J2 numbers/200g soil samples [log10(x+1)] for four sampling intervals from a trial site in the Christiana area where Brassicaceae crops were planted before potato during the 2010/2011 growing season (means in parenthesis represents the values that were back transformed and not the real means). Sampling interval 1 Nematode genera/family/spp. Untreated control (T 1) Terranova (T 2) Doublet (T 3) Nemat (T 4) Calient้ (T 5) EDBฎa (T6) P value F ratio Criconema spp. 1.1 (2) 0.8 (1) 0.8 (1) 0.3 (0.3) 0.7 (1) 1.3 (2) 0.8518 0.39 Hoplolaimidae spp. 0.3 (0.3) 0 0 0.3 (0.3) 0 0 0.5878 0.76 Trichodoridae 0.3 (0.3) 0 0 0 0 0 0.4430 1 Pratylenchus spp. 0.3 (0.3) 0.3 (0.3) 0 0 0.5 (0.6) 0 0.6543 0.66 Tylenchorhynchus spp. 0 0 0 0 0 0.3 (1) 0.4430 1 Total PPN 1.7 (1) 1 (2) 1.2 (2) 0.8 (1) 2.2 (8) 2.1 (7) 0.5585 0.81 Sampling interval 2 Criconema spp. 0 0.2 (0.2) 0 0 0 0.1 (0.1) 0.4430 1 Hoplolaimidae spp. 0 0.1 (0.1) 0 0.1 (0.1) 0.3 (0.3) 0 0.2057 1.6 Trichodoridae 0 0 0 0.1 (0.1) 0.3 (0.3) 0.1 (0.1) 0.6111 0.73 Tylenchorhynchus spp. 0 0 0 0 0 (0) 0.3 (0.3) 0.0528 2.67 Total PPN 1.2 (2) 0.6 (1) 0.3 (0.3) 0.5 (1) 0.8 (1) 0.6 (1) 0.2249 1.53 174 Sampling interval 3 Nematode genera/family/spp. Untreated control (T 1) Terranova (T 2) Doublet (T 3) Nemat (T 4) Calient้ (T 5) EDBฎa (T6) P value F ratio Criconema spp. 0 0.3 (0.3) 0 0.1 (0.1) 0.1 (0.1) 0 0.2057 1.6 Hoplolaimidae spp. 0.1 (0.1) 0.3 (0.3) 0.1 (0.1) 0.1 (0.1) 0 0.1 (0.1) 0.9154 0.29 Trichodoridae 0 0 0.2 (0.2) 0 0 0.1 (0.1) 0.4430 1 Tylenchorhynchus spp. 0.3 (0.3) 0 0 0.1 (0.1) 0 0.3 (0.3) 0.4633 0.96 Total PPN 1.6b (2) 0.7ab (1) 0.5ab (1) 0.3a (0.3) 0.3a (0.3) 0.8ab (1) 0.0334 3.04 Sampling interval 4 Criconema sp. 0.4 (0.5) 0.5 (0.6) 0 0 0 0 0.5856 0.77 Hoplolaimidae sp. 0 0.7 (1) 0.4 (0.5) 0 0 0 0.5697 0.79 Total PPN 3ab (19) 3.6ab (35) 1.8ab (5) 1a (2) 5b (147) 2.4ab (10) 0.0364 2.97 aEDBฎ= ethylenedibromide 175 Table 2. Numbers [log10(x+1)] of non-parasitic nematodes J2 numbers/200g soil samples for four sampling intervals from a trial site in the Christiana area where Brassicaceae crops were planted before potato during the 2010/2011 growing season (means in parenthesis represents the values that were back transformed and not the real means). Sampling interval 1 Nematode genera/family Untreated control (T 1) Terranova (T 2) Doublet (T 3) Nemat (T 4) Calient้ (T 5) EDBฎa (T 6) P value F ratio Acrobeles spp. 2.4 (10) 2.3 (9) 2.7 (14) 1.5 (3) 2.5 (11) a 2 (6) 0.5390 0.84 Acrobeloides spp. 0 0.3 (0.3) 0 0 0 0.3 (0.3) 0.5878 0.76 Aphelenchus spp. 0.7 (1) 0.5 (0.6) 0.8 (1) 0.3 (0.3) 0 1.2 (2) 0.6559 0.66 Aphelenchoides spp. 0.3 (0.3) 0.3 (0.3) 0 0.5 (0.6) 0.3 (0.3) 0 0.8351 0.41 Cephalobus spp. 1.5 (3) 1.4 (3) 1.1 (2) 1.5 (3) 2.1 (7) 2.2 (8) 0.8737 0.35 Chiloplacus spp. 0 0 0.7 (1) 0.3 (0.3) 1.3 (2) 0.5 (0.6) 0.2964 1.32 Cruznema spp. 0. 5 (0.6) 0 0 0 0.7 (1) 0 0.2933 1.33 Diploscapter spp. 0 0.3 (0.3) 0 0 0 0 0.4430 1 Doryliamidae 1.3 (2) 2.3 (9) 0.3 (0.3) 1.2 (2) 1.6 (4) 1.4 (3) 0.2012 1.62 Eucephalobes spp. 1 (2) 0 0.3 (0.3) 0.8 (1) 0.3 (0.3) 0.8 (1) 0.5482 0.82 Mesorhabditis spp. 0.3 (0.3) 0 0.5 (0.6) 0 0.3 (0.3) 0.8 (1) 0.5680 0.79 Panagrolaimus spp. 0.3 (0.3) 0.7 (1) 0.8 (1) 0.8 (1) 0.5 (0.6) 0.8 (1) 0.9599 0.2 Rhabditis spp. 2.1 (7) 1.4 (3) 1.3 (2) 2.1 (7) 1.5 (3) 0.5 (0.6) 0.2321 1.51 Tylenchus spp. 0 0.3 (0.3) 0 0 0 0.3 (0.3) 0.5878 0.76 Total NPN 3.8 (43) 3.8 (43) 3.6 (35) 3.1 (21) 4 (53) 3.9 (48) 0.7502 0.53 176 Sampling interval 2 Nematode genera/family Untreated control (T 1) Terranova (T 2) Doublet (T 3) Nemat (T 4) Calient้ (T 5) EDBฎ (T 6) P value F ratio Acrobeles spp. 0.4 (0.5) 0.1 (0.1) 0 0.3 (0.3) 0.3 (0.3) 0.4 (0.5) 0.5919 0.76 Acrobeloides spp. 0 0.3 (0.6) 0 0.1 (0.2) 0.1 (0.2) 0.2 (0.2) 0.4430 1 Cephalobus spp. 1 (2) 1 (2) 0.8 (1) 0.6 (1) 0.8 (1) 0.6 (1) 0.8566 0.38 Dorylaimidae spp. 0.4 (0.5) 0.4 (0.5) 0.6 (1) 0.2 (0.2) 0.5 (1.6) 0.2 (0.2) 0.8648 0.37 Eucephalobus spp. 0.4 (0.5) 0.4 (0.5) 0.5 (0.6) 0.1 (0.1) 0.2 (0.4) 0.3 (0.3) 0.8652 0.37 Mesorhabditis spp. 0.3 (0.3) 0.1 (0.1) 0.1 (0.1) 0.1 (0.1) 0.1 (0.1) 0.1 (0.1) 0.8990 0.31 Rhabditis spp. 0.4 (0.5) 0.1 (0.1) 0.1 (0.1) 0.1 (0.1) 0.5 (1) 0.4 (0.6) 0.5225 0.86 Total NPN 1.8 (5) 1.6 (4) 1.4 (3) 1.2 (2) 1.6 (4) 1.4 (3) 0.6724 0.64 Sampling interval 3 Acrobeles spp. 1.1 (2) 1 (2) 1.4 (3) 1.1 (2) 1 (2) 1.2 (2) 0.5246 0.86 Cephalobus spp. 1.1 (2) 1 (2) 1 (1.4) 0.3 (0.6) 1 (2) 0.5 (2) 0.5906 0.76 Dorylaimidae 0.1 (0.1) 0.5 (0.6) 1 (2) 1 (2) 0.5 (0.6) 1 (2) 0.6199 0.71 Eucephalobus spp. 0.2 (0.2) 0.4 (0.5) 0.1 (0.1) 0 1 (2) 0.3 (0.3) 0.1219 2 Mesorhabditis spp. 0.1 (0.1) 0.2 (0.2) 0 0 0 0.2 (0.2) 0.6278 0.7 Rhabditis spp. 0.8 (1) 0.7 (1) 0.3 (0.3) 0.7 (1) 0.8 (1) 1 (2) 0.7238 0.57 Total NPN 2.2 (8) 2 (6) 2 (6) 2 (6) 2 (6) 2.2 (8) 0.8624 0.37 177 Sampling interval 4 Nematode genera/family Untreated control (T 1) Terranova (T 2) Doublet (T 3) Nemat (T 4) Calient้ (T 5) EDBฎa (T 6) P value F ratio Acrobeles spp. 1.7 (4) 0 1.2 (2) 0 0.5 (0.6) 0.8 (1) 0.1393 1.9 Aphelenchoides spp. 1.8 (5) 1.5 (3) 0 0 0 0.4 (0.5) 0.2784 1.37 Cephalobus spp. 2.4 (10) 3.7 (39) 3 (19) 2.5 (11) 3 (19) 3.7 (39) 0.5381 0.84 Diploscapter spp. 0 1.2 (2) 0 0 0 0 0.4430 1 Dorylaimidae 3.4 (29) 2.6 (12) 3.4 (29) 1.5 (3) 3.4 (29) 1.4 (3) 0.1085 2.09 Rhabditis spp. 7.3 (1479) 5 (147) 6.8 (897) 5.3 (199) 7 (1095) 7.5 (1807) 0.2998 1.31 aEDBฎ= ethylenedibromide