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Tobacco parasitic nematode

Tobacco

1.0 Introduction

Tobacco (Nicotina tabacum L.) is one of the most important non- food crop and widely grown commercially (Akerhust, 1981). This plant had a high economic value and widely demanded throughout the world for the usage of the nicotine, cigarettes, cigars and other tobacco product (Akerhust, 1981).

Nowadays, in Malaysia, Tobacco industry is very crucial in uplifting the socio-economic status of farmer in Kelantan, Terengganu, Kedah and Perlis. There were 20,524 farm families, 355 tobacco curers, 1300 grower and 25384 station workers. This industry generates about 150 million in income per year. 38% of the income goes to the farmers and 18% to the curers (http://www.malaysiayellowpages.net/mpi/details/TOBACCO.htm). This plant can give a stable income and therefore increased the income of farmer (Wells, 1987). Tobacco was cultivated as a rotation with the paddy for a side income (Anon, 1981). Tobacco plant can be infected by bacteria, fungus, virus, and parasitic nematode. Disease infection lower the tobacco yield and also quality. This research focused on effect of plant parasitic nematode on tobacco.

Plant parasitic nematode can be found wherever tobacco is grown. The severity of the damage they caused depended on climate and soil type (Luc, Sikora et al. 2005). Nematode infection may lower the quality and yields. Annual report from North Carolina in 2008, showed Meloidogyne spp. it self cause loses around $2,505,126 in 2004, $1,596,452 in 2005, $ 1,772,819 in 2006, $1,542,864 in 2007, and $4,096,321 in 2008 while other nematodes cause $146,297 in 2004, $2281 in 2005, $529,188 in 2006 and $208,612 in 2008 (www.dowagro.com/soil/products/tobacco/economic.htm). In Malaysia, the effect of nematode on tobacco yield reduction has not been fully understand or revealed. Therefore, the objectives of this project were:

1. To observe the effect of nematode on tobacco.

2. To observe the relationship of soil physical properties on nematode population density and disease severity.

2.0 Literature review

2.1 Tobacco

Tobacco was one of the most important non- food crop and widely grown commercially (Akerhust, 1981). This plant originated came from South America (Tso, 1972). However according to Gerstel (1961), Nicotina tabacum not occurring in wild state it was amphidiploids which come from hybridization of Nicotina sylvestris and Nicotina tomentosiformis. This plant has a high economic value and has been widely demanded throughout the world for the usage of the nicotine such as cigarettes, cigars and other tobacco product (Akerhust, 1981). This plant also important for the research purposes (Tso, 1972). Many researches have been done by using this plant mostly in Plant physiology and Genetics (Bateman & Millar, 1966; Albersheim et. al., 1969; Kosuge, 1969).

In Malaysia, Tobacco was first introduced in year 1959 by Malayan Tobacco Company (now known as Malaysia Tobacco Company, MTC) in Kelantan cultivation area for 8 hectares (Anon., 1976). Nowadays, in Malaysia, Tobacco industry has been very crucial in uplifting the socio-economic status of farmer in Kelantan, Terengganu, Kedah and Perlis. There were 20,524 farm families, 355 tobacco curers, 1300 grower and 25384 station workers. This industry generated about 150 million in income per year. 38% of the income goes to the farmers and 18% to the curers (Ministry of primary industry, 2010). This plant can give a stable income and therefore can increase the income of farmer (Wells, 1987). Tobacco is cultivated as a rotation with the paddy for a side income (Anon, 1981). However, product and quality of tobacco leaf are quite low due to encountering many problems, including diseases. For Tobacco cultivation, a deep and well drained soil is needed. This is where nematodes problem develop rapidly (Luc, Sikora et al. 2005).

2.2 Nematodes related with the Tobacco

Plant parasitic nematode can be found wherever tobacco is grown. The severity of the damage they caused may depended on climate and soil type (Luc, Sikora et al. 2005). Many tobacco producing countries are near or within the inter-tropical zone. The dominant nematodes that parasitize tobacco plant were Meloidogyne spp. (a root-knot nematode). Most of important species from this genus were M.arenaria, M.incognita, M.javanica, and M.hapla. M.incognita and M.javanica were important species in Malaysia. Other Meloidogyne spp., were rarely reported. Similarly, Pratylenchus spp. were also dominant species that parasitize tobacco plant (Kimpinski and Thompson 1990). Apart from Meloidogyne spp. and Pratylenchus spp., Tylenchorhynchus spp., Globodera spp., Ditylenchus dipsaci and Aphelenchus ritzemabosi were reported to parasitized tobacco plant in certain restricted area. Other nematodes such as Helicotylenchus, Rotylenchus, Scutellonema, Rotylenchulus sp., Tetylenchus and Crinomella sp. have been found to infect tobacco plant but not normally associated with losses. Some nematode species such as Xiphinema, Longidorus, Trichodorus, and Paratrichodorus have been reported to transmit viruse to tobacco (Luc, Sikora et al. 2005). Nematodes also may cause disease complex. For example Meloidogyne spp. a root-knot nematodes has been proved to increase the incident of Fusarium wilt even when their population were incapable to cause direct damage to the tobacco plant (Webster, 1972). Another example was interaction between Pratylenchus brachyurus (lesion nematode) and Phyptopthora parasitica var. nicotianae (cause black shank disease). Inagaki and Powell (1969) found that P. brachyurus caused more severe and rapid diseased development of black shank symptom than when the fungus alone.

2.3 Root-knot nematodes, Meloidogyne spp.

2.3.1 Distribution

Meloidogyne spp. are always important parasites in tobacco cultivation, wherever the climate favours them (Nusbaum, 1960; Daulton, 1964; Barker et al., 1981; Rich et al., 1982). There were 61 species and two subspecies in this genus at the end of 1988 (Eisenback, 1985; Eisenback & Hirschmann, 1991). Nowadays until year 2000 there were 80 species have been describing (Carneiro et al., 2000). Parasitism of Meloidogyne spp. was first reported by Tisdale (1922) in Florida. This genus was also a serious pest in Southern Africa in the late 1920's (Jack, 1927; Naudѐ, 1929). Meloidogyne incognita and M. javanica were mostly found parasitize the tobacco plant. Their infection was very relying on the climate, since M.javanica had a higher tolerance towards drought and high temperature compared with M.incognita (Daulton & Nusbaum, 1969, 1962; Taylor et al., 1982). Meloidogyne arenaria and M.hapla were the next mostly found to cause infection on tobacco plant. Meloidogyne hapla was reported to be found in the cooler parts of the world. Report from fields' survey in Florida showed M.javanica was found in 65% of fields' survey area, M.incognita 33% and M.arenaria was rarely found (Rich &Garcia, 1985). Report from North Carolina showed M.arenaria population had increased gradually although M.incognita was the predominant species there. This observation also showed the same in South Carolina (Fortnum et al., 1984; Schmitt & Barker, 1988). Apart from that, M.javanica and M.hapla was reported to be found in North Carolina. Reported showed that there were 64% of M.incognita and 29% of M.javanica to be found in Philippines (Madamba, 1981). Meloidogyne incognitagraham, M.microcephala, M.mayaguensis, M.cruciani, M.enterolobii, M.ethiopica, M.platani, M. themesi were also reported to parasitize reproduce tobacco plant but their importance was very restricted (Cliff & hirschmann, 1984; Jepson, 1987; Rammah 1988; Rammah and Hirshmann, 1988).

2.3.2 General morphology

The morphology of this genus were almost all same the except for some characteristic which usually were very useful for species identification. They were usually sexually dimorphic. Adult female have swollen, saccate bodies (pear shape like body). The size of female ranged in median length 0.44-1.30 mm and width about 0.33-0.70 mm (Eisenback, 1985). They have protrudes neck anteriorly while vulva and anus were located terminally. The female of this genus have pearly white body with moderately thick cuticle. Stylet were short, moderately sclerotized and protrusibly hollow. The stylet size was 10-24µm in length which consists of cone, shaft and knobs. The morphology of the stylet was quite varying between species in this genus. The morphology of stylet should be one of the supplemental characteristic to be observed for species identification. The stylet functions like hypodermic needle which was moved by protractor muscles. The shaped of the cone, shaft, and knobs also differ among female species in this genus. At the posterior of stylet knobs, there was dorsal esophageal gland orifices (DEGO). DEGO was the two sub ventral gland orifices open into the esophagus lumen. DEGO had a varied distance among species which also can be supplemental character for species identification. The excretory pore of the Meloidogyne spp. female situated anterior to median bulb valve plat and usually near stylet base. They also have two convoluted genital tracts. The major part of the total body content consists of two gonads which were very long and greatly convoluted. There were ovary with germinal zone and growth zone, narrow oviduct, globular spermatotheca and long uterus in each gonad. Spermatotheca were differing among species. Therefore this character can also be use for species identification. Apart from that, the cuticle in the perineal region of female from this genus forming a finger print-like pattern (the perineal pattern) which also had been use for species identification. This is because, the perineal pattern hold most characteristic of female such as tail terminus, phasmids, lateral lines, anus, and vulva which surrounded by cuticular striae or folds. They also have six large unicellular rectal glands situated in the posterior body region. These rectal glands were connected to the rectum. This gland produce very large amount of gelatinous matrix material. This material was excreted through the rectum and act as protective egg sac (Nickle, 1991).

Different with the female, male of Meloidogyne sp. are vermiform. The size of the body vary between species which are about 700-2,000 µm (Eisenback, 1985). This is because the varying environmental condition existing during their development. Body of the male usually twisted through 180̊ upon heat relaxation. The male stylet vary in size which are about 13-30 µm. The stylet and head of male from this genus are robust. Apart from that, size and shape of the stylet cone, shaft, and knobs can be use for species identification (Eisenback and Hirschmann, 1981). The location of DEGO is 2-13 µm posterior to the stylet knob base. The isthmus is short and most of the species have ventrally two overlapping gland lobe instead of normally three esophageal nuclei. The hemizonid located at the front to excretory pore. However some species the hemizonid located at the posterior of excretory pore. In normal male there is only one gonad while in sex-reversed males have two gonads. There is long vas deferens packed with developing sperm in the gonad. Among the species, the size of the spicules range from 19 to 40 µm. The spicules usually robust and the bursa are absent. Tail is short (hemispherical shape). There is also variation of tail shape between species (Nickle, 1991).

A second stage juvenile was the infective stage of Meloidogyne sp. It has varied body length from 290 to 912µm (Eisenback, 1985). The head of second stage juvenile basically just same with the male. It has a delicate stylet with 8 to 18µm in length. The DEGO distance are varied among species with the distance mostly 2 to 8µm. The esophagus of the second stage juvenile is narrow with faintly outline procorpus. The median bulb is well defined. Median bulb has a large valve plate and three long ventrally overlapping glands that are use for molting and feeding. The second stage juvenile has a varied position of excretory pore. The hemizoid located posteriorly to the pore. The tail length of second stage juvenile varied among species. Usually the length is 15 to 100µm. At the end of the tail there is hyaline terminus. In this genus, second stage juveniles are group base on the tail length and tail shape (Whitehead, 1968; Jepson, 1984). Jepson (1987) showed that differences in either mean tail and or mean hyaline terminus are very large. These vast differences can be very useful to distinguish species within groups (Nickle, 1991).

2.3.3 Life cycle

Meloidogyne sp. shows sexually dimorphism, which is the female are pyriform or saccate, while the male's vermiform (Eisenback, 1987). The differences in body shaped between female and male occurred during the postembryonic development of Meloidogyne sp.. From the embryonic development, the egg hatched once to become first-stage juvenile and then molted as a second stage juvenile. The second-stage juvenile was infective stage. It moved into the soil and entered the root of suitable host plant. This second-stage juvenile then formed host-parasites relationship with the plant when it find preferred feeding site. The morphology of second-stage juvenile changed to flask-shape as it feeds on the special nurse cell. Then, without further feeding it molted three times into the third and fourth stage juvenile, and finally become an adult. The saccate adult female resumed feeding on the special nurse cell shortly after the last molt and continued to do so for the remainder of her life. The reproductive system of both female and male of this genus developed into functional gonads during the postembryonic development (Triantaphyllou and Hirschmann, 1960). From the number of the gonad, we can differentiate the sexes. Females always have two gonads while males usually have one. During fourth-stage juvenile, the shape of saccate male juvenile changed to the vermiform adult males. The metamorphosis occurred in which the body elongates from saccate to a vermiform shape. Fully developed male emerges after the final molt of enclosed fourth-stage male which enclosed within the cuticles of second-stage and third-stage. The adult male leaved the root and move freely through the soil and it does not feed. The mode of reproduction determined the function of the male for mating. Depending on particular species reproduction whether amphimixis or parthenogenesis, the male enters the root searching for the female to mate or just remain in the soil and die. Temperature plays a vital role for the length of the life cyle. For example, the first adult female of M.incognita on Tomato appear 13-15 days after root penetration at temperature approximately 29 ̊C, the female laid the first egg about 19-21 days after penetration (Triantaphyllou and Hirschmann,1960). The life span of female is much longer than the male from 2 to 3 month.

2.3.4 Effect of Meloidogyne spp. on Tobacco plant

Meloidogyne sp. caused formation of galls on Tobacco root. Usually, second stages juvenile entered via behind the root cap which involves mechanical penetration by using stylet (Linford, 1942). According to Bird et.al, (1975), the penetration also involve some enzymatic action (cellulolytic or pectolytic) which secreted by esophageal gland. Then, the second-stage juvenile moved through the cortex to the region of cell differentiation. This differentiation cell was the feeding site for them which later transformed into highly specialized feeding cells called "giant cells". This cell was the permanent feeding site for them (Hussey at al., 1994). According to Dropkin (1972) and Hussey (1987), the multinucleate "giant cell" was the result of the introduction of secretion produced by subventral esophageal gland cells of the feeding second stage juvenile. Giant cells serve as sourced of food. The nutrient from giant cells was transferred to the nematode (Jones and Northcote, 1972). According to McClure (1977) these cells act as metabolic sink. These giant cells affected the function of the root as it caused extensive distortion and blocked of the vascular tissue which slowed water and nutrient transport. Therefore, the absorption of nutrient and water greatly reduced. Plant growth and yield may be suppressed as photosynthates were mobilized to the giant cells. Above- ground symptoms showed chlorosis of foliage and temporary wilting (premature wilting) when water stress occurred usually during drought or sunny day. Plant was stunted and the leaves were yellow and thin. The formation of gall was due to the root tissues around nematode and giant cells undergo hyperplasia and hyperthrophy. The worse was when secondary larval invasion occurred which caused the gall to coalesce and finally the root begins to decay (Nickle, 1991). Nematode also had the ability to form disease complex with other plant pathogens. The giant cell produced by root-knot nematode was highly suitable for development of Fusarium wilt ( Porter and Powell, 1967).

2.4 Root lesion, Pratylenchus spp.

2.4.1 Distribution

Pratylenchus spp. is migratory endoparasites root-lesion nematodes. This genus was just slightly less economic important compare with Meloidogyne spp. in the tropical and subtropical regions. However, some species from this genus were responsible for significant yield loss in some tobacco cultivation area. Pratylenchus pratensis, P.negletus, P.brachyurus and P.zae have been reported to parasitized tobacco in North America while in South Africa P.hexincisus, P.thornei, P.vulnus, P.brachyurus, P.minyus, and P.zae have recorded on tobacco (Milne, 1961; Honey, 1967). In Hungary, P.pratensis had been reported to parasitize tobacco cultivation. Pratylenchus penetrans was responsible to cause yield loss in Iraq. In some region in Canada, P.penetrans, P.crenatus, and P.neglectus were mostly found in tobacco fields (Mountain, 1954; Kimpinski et. al., 1976). Canter-Vissher (1969) had found Pratylenchus penetrans in New Zealand while Singh (1974) has found Pratylenchus zae in Trinidad. In general Pratylenchus brachyurus and P.zae are mostly found in tropical areas while P.penetrans, P.thornei, and P.minyus are common species in temperate regions (Webster, 1972). In Malaysia, this Pratylenchus sp. was locally important. However their distribution were not clearly report (Luc, Sikora et al. 2005).

2.4.2 General morphology

In general the morphology of species in this genus was very similar. There was no marked sexually dimorphism in form of anterior region. Adults have body length range from 0.3 to 0.9 mm. Their body was rather stout. Because increasing of uterus volume and the presence of eggs, the gravid females were stouter than nongravid ones. The cuticle of this genus generally thin and shows fine transverse striation. There were four longitudal lines marking the lateral field. However, additional longitudal line may be present in the central zone. Because of cuticle of gravid female were quite stretch, the lateral field was indistinct.

The head of this genus was low and flattened with lip region divided into two,three, or four annules. This annules was continuous with the body countour. Cephalic framework of Pratylenchus sp. was heavily sclerotized. The apical anule among most species were round except for P.brachyurus which was angular. There were three types of head structure that can be found under SEM (Corbett and Clark, 1983). The stylet of Pratylenchus sp. were quite short around 11-25 µm. The stylet was stout with well-developed basal knobs. There was tapering procorpus in the pharynx which was usually roundish median bulb. The isthmus was short which overlapped with the anterior end of the mid-intestine on the ventral side. There were three unicellular glands in the lobe. The length of the ventrosublateral was unequal (Seinhorst, 1971). At 2-4 µm behind the stylet base, there was orifice of the dorsal pharyngeal gland duct. There was no deirids in this genus. The oesophagus of both male and female was equally developed. The tail of male was short and dorsally convex-conoid.

Female of Pratylenchus spp. are monoprodelph. The genital branch of most species in this genus occurred as a short sac which usually undifferentiated. The uterus of female often tricolumellar (Nickle, 1991). Different with male, female tail usually two to three anal body diameter long. The bisexual species in this genus, have oval or round spermatheca which was filled with sperm (Luc, Sikora et al. 2005).

2.4.3 Life cycle

Some species in this genus reproduced sexually while most of them parthenogenetic. This migratory endoparasitic root lesion nematode fed and laid eggs in the root cortex. Most of them can be found in roots, rhizomes, or tubers and somehow can also be found in stem or fruits. Usually after penetrate the root; this endoparasitic nematode will multiply to very large numbers (10,000-35,000 specimens per 10 g of root). All the stage starting from second stage juvenile entered the root. However with unknown reason, they moved in the soil for some time and goes for a new host root. The female laid the eggs in the root and starting from there their whole life cycle is in that root. Usually, the life cycle was completed in 50-60 days (Nickle, 1991).

2.4.4 Effect of Pratylenchus sp. on Tobacco plant

Pratylenchus sp. usually moved and fed on the root cortex. This activity caused disintegration of root cortex and leading to browning of the root tissue. This was known as "brown root rot" (Mountain, 1954). Symptoms of this disease were pruning-root, water soaked, and lesion on the root. If the infection occurred under aseptic conditions the symptoms showed less severe in the certain experimental condition (Mountain, 1954). The above ground symptoms showed that the stunted plant wilt prematurely and in worse condition died. Inagaki and Powell (1969) reported that this genus caused disease complex with the other plant pathogens. Pratylenchus.brachyurus showed to increase infection of Blackshank by wounding the root which served as entry site.

3.0 Material and method:

3.1 Soil sample:

24 soil samples were collected from Terengganu, Perlis and Kelantan state. Collected soil sample were naturally infested with nematodes and Fusarium spp. Soil samples were store in polyethylene bags. Soil sample were kept in moist condition and out of direct sunlight.

3.2 Tobacco seedling preparation:

Sterilized seeds were sown to sterile sandy soil. (River sand). After sown, seedlings were kept out of direct sunlight. Fertilizer applied for twice a week via foliar application. After 30 days of nursery tobacco seedlings were transferred to each soil.

3.3 Inoculation of tobacco seedlings:

6 kg of soil samples (naturally infested) were transferred into plastic container (33x22x10 cm) with drains. Then, 30 days of healthy Tobacco seedlings were transferred to each soil container. Each soil samples were planted with 10 Tobacco seedlings. Fertilizer was applied twice a week via foliar application. Ground symptoms were observed everyday. Tobacco plants were all harvested after 6 weeks.

3.4 Plant observation:

Harvested Tobacco plants were observed for the disease symptoms, size of the plant, number of leaf, leaf area, plant weight and disease severity index. Wet weight of Tobacco was measured by using a weigher. Plant size was determined by using ruler. Size of the plant was measured from crown up until shoots. Number of leaf was counted including the number of undeveloped leaf. Root gall disease severity index was determined by using following scale:

0= no root galls

1= 1-25% root galls

2= 26-50% root galls

3= 51-75% root galls

4= 75-100% root galls

Disease severity index for root lesion was determined by using following index:

0= no root lesion

1= 1-25% root lesions

2= 26-50% root lesions

3= 51-75% root lesions

4= 75-100% root lesions

Root then was stored in the FAA (Formaldehyde 100ml, Glacial acetic acid 50ml, Distilled water 850ml) suspension.

3.5 Isolation of nematode from soil samples:

Isolation of nematode and soil inhabiting forms were extracted from soil samples by using Modified Baerman Funnel Technique (Hooper,1968; Viglierchio and Schmitt,1983).This was the simplest technique to isolate nematode and soil inhabiting forms. By using this technique we can avoid lack of oxygen and possibility of nematode lodging on the sloping funnel sides due to instead of using funnel we used a shallow dish. For this experiment instead of funnel a round shallow plastic container was used. A supporting gauze was put onto the plastic container with 0.5cm space between them. A milk filter paper with 50cc soil was put on the supporting gauze. Distilled water was added until the material was almost awash. After 5 days, the content of the dish was transfer into test tube. FAA was added to prevent population changes during storage.

3.6 Nematode counting:

Nematode suspension collected via Modified Baerman Funnel was shaked. Then, 1ml was taken and transfer onto disposable plastic Petri dish. The number of all nematodes and parasitic nematodes were counted under a dissecting microscope by 5x to 10x magnification. Counting was repeated for three times. Percentage of parasitic nematodes was calculated.

3.7 Isolation of nematode from root:

Nematode from root part was isolated by direct isolation. For root-knot nematodes especially female, the root tissue was carefully tease away with forceps and a fine needle to release the head and neck. Infected plant part was put onto slide and squash to check for the existence of nematode. The nematode then was stain with Phyloxine 1%.

3.8 Isolation of Fusarium spp.

The root part was washed with running tap water to eliminate remaining soil particle. Then, the root was cut including healthy part (0.5cm). After that, the pieces of root were dipped in 70% ethanol for 1 minute. Then, the pieces were transferred into 5% sodium Hypochlorite solution to sterilize its surface for 3 to 5 minutes. The pieces then were transferred to sterilized distill water to rinse the pieces for 3 times each for 1 minute. After that, the plant pieces was put on sterile filter paper to eliminate excess water and then, were put on the acidified water agar medium. Finally, the dishes were sealed with parafilm and were incubated for a few days. Growing colonies were observed.

3.9 Soil pH:

The soil pH was determined using a soil suspension (Rowell, 1994). 10 ± 0.1 gram of air dry soil sample was used in this experiment. 25 ml of water was added to the soil sample. Then, soil suspension was shacked occasionally by hand over 15 minute's period. The pH meter was calibrated at pH 4 and then pH 7 consistent reading. The soil suspensions were stirred and insert the electrodes. The pH was recorded after 30 second.

3.10 Soil moisture percentage:

The water content of soils was determined by drying soil samples at 105 ̊c (Rowell, 1994). For this experiment, soil samples were air dry for two days. Then, weight air dry soil samples for 10±0.1 gram (W1). Instead of using a moisture can, aluminums foil was used. The aluminum foil was weighed (Wo). Then, weighed soil samples were put on the aluminum foil and placed them in an oven at 105 ̊C for 24 hour. The weigh of soil sample with aluminum foil was weighed (W2). To calculate the weight of soil samples after oven dry the following formulae was applied:

Weight oven dry soil (W3) = (W1+Wo)-W2

To calculate moisture percentage of soil samples, the following formulae was use:

Moisture percentage (%) = W3/ (W2-Wo) x 100

3.11 Soil particle density:

Determination of soil particle density involves the measurement of the volume of a known mass of particles. The soil is dispersed in water and all the air is expelled from the suspension. In a known volume of suspension the volume occupied by the particles is then found (Rowell, 1994). A clean and dry 50ml volumetric flask including stopper was weigh (Wo). Ten grams of oven dry soil samples were added into the volumetric flask. The volumetric flask was filled with distilled water until one-half full. The volumetric flask (without stopper) then was put in boiling water heating with a water bath for 30 minutes and gently agitated the content to prevent loss of soil by foaming. The volumetric flask and its content then cooled to room temperature. Distilled water was added up to the 50 ml mark. Water drop on the outer-side of the volumetric flask was wiped, insert the stopper and weighed (W2). The soil particle density was determined by using the following formulae:

Soil particle density (Dp) = Soil mass/Particle volume

Particle volume = Conical flask volume - volume of water in flask

Volume of water in flask = mass of suspension -mass of soil

Mass of suspension = W2-W0

3.12 Soil texture analysis:

Texture of soil samples were determined by using Hydrometer method (Bouyoucos, 1962; Page, 1982). Then, texture of soil samples determined by referring to USDA Textural triangle after calculation of the percentage of each particle (Brady, 1984). For this experiment, 50g of soil samples were placed into 600 ml beaker. Then, 100 ml of 6% hydrogen peroxide was added to decompose the organic matter. The mixture was kept remaining at room temperature overnight. After that, the beaker was placed on a hot plate at 90 ̊ C for 10 minutes. Then, 50ml of 1N Sodium hydroxide (NaOH) (dispersing agent) was added to the suspension and increase the volume to 400 ml with distilled water. The suspension was left for 20 minutes. Then, beaker was placed on a stirrer and stirred thoroughly for 10 minutes. The suspension was transferred to 1000 ml measuring cylinder. Then, distilled water was added to 1000 ml mark. Suspension was allowed to equilibrate thermally and the temperature was recorded. Mouth of the measuring cylinder was covered with a parafilm and inverted for several times until the contents are thoroughly mixed. Mixture was left in a cool, shaded place. Then, the hydrometer was immediately into the suspension and reading was taken after 40 seconds until consistent reading. Hydrometer was removed and cleaned. The temperature of the suspension was recorded with thermometer. The thermometer was removed and remixes the suspension. Then, let the cylinder sit for 2 hours. At exactly 2 hours later, the hydrometer was again placed into the suspension and data was read. The temperature of the suspension was also seconded with thermometer. The actual reading must be corrected in order to get revised value depending upon the actual temperature.

a. Add 0.36 g/L to hydrometer reading for each degree >20 ̊C

b. Subtract 0.36 g/L from hydrometer reading for each <20 ̊C

c. Density reading should also be corrected from the density of the dispensing solution (NaOH+ distilled water) without soil. These reading are must be subtract with the soil solution density reading.

Finally, after calculating the percentage of each particle, use the USDA Textural triangle to determine the textural class of soil samples.

Readings from specific gravity hydrometer was converted to soil g/l by using converting table (http://classic.globe.gov/fsl/html/templ.cgi?conversion&lang=ar).

Table 3.1 Hydrometer converting table

Specific

Gravity

Grams Soil/L

Specific

Gravity

Grams Soil/L

Specific

Gravity

Grams Soil/L

1.0024

0.0

1.0136

18.0

1.0247

36.0

1.0027

0.5

1.0139

18.5

1.0250

36.5

1.0030

1.0

1.0142

19.0

1.0253

37.0

1.0033

1.5

1.0145

19.5

1.0257

37.5

1.0036

2.0

1.0148

20.0

1.0260

38.0

1.0040

2.5

1.0151

20.5

1.0263

38.5

1.0043

3.0

1.0154

21.0

1.0266

39.0

1.0046

3.5

1.0157

21.5

1.0269

39.5

1.0049

4.0

1.0160

22.0

1.0272

40.0

1.0052

4.5

1.0164

22.5

1.0275

40.5

1.0055

5.0

1.0167

23.0

1.0278

41.0

1.0058

5.5

1.0170

23.5

1.0281

41.5

1.0061

6.0

1.0173

24.0

1.0284

42.0

1.0064

6.5

1.0176

24.5

1.0288

42.5

1.0067

7.0

1.0179

25.0

1.0291

43.0

1.0071

7.5

1.0182

25.5

1.0294

43.5

1.0074

8.0

1.0185

26.0

1.0297

44.0

1.0077

8.5

1.0188

26.5

1.0300

44.5

1.0080

9.0

1.0191

27.0

1.0303

45.0

1.0083

9.5

1.0195

27.5

1.0306

45.5

1.0086

10.0

1.0198

28.0

1.0309

46.0

1.0089

10.5

1.0201

28.5

1.0312

46.5

1.0092

11.0

1.0204

29.0

1.0315

47.0

1.0095

11.5

1.0207

29.5

1.0319

47.5

1.0098

12.0

1.0210

30.0

1.0322

48.0

1.0102

12.5

1.0213

30.5

1.0325

48.5

1.0105

13.0

1.0216

31.0

1.0328

49.0

1.0108

13.5

1.0219

31.5

1.0331

49.5

1.0111

14.0

1.0222

32.0

1.0334

50.0

1.0114

14.5

1.0226

32.5

1.0337

50.5

1.0117

15.0

1.0229

33.0

1.0340

51.0

1.0120

15.5

1.0232

33.5

1.0343

51.5

1.0123

16.0

1.0235

34.0

1.0346

52.0

1.0126

16.5

1.0238

34.5

1.0350

52.5

1.0129

17.0

1.0241

35.0

1.0353

53.0

1.0133

17.5

1.0244

35.5

1.0356

53.5

1.0359

54.0

1.0362

54.5

1.0365

55.0

4.0 Result:

4.1 Soil sampling

24 soil samples were collected from different place in three different regions.

Table 4.1: Table locations of collected soil samples and dates of collection

Code

Location

Region

Collection date

P1

Mata Ayer

Perlis

19 May 2009

P2

Simpang Empat (3)

Perlis

19 May 2009

P3

Simpang Empat (2)

Perlis

19 May 2009

P4

Simpang empat (1)

Perlis

19 May 2009

K5

Pantai Bisikan Bayu

Kelantan

24 May 2009

K6

Telong

Kelantan

6 July 2009

K7

Labok,Machang

Kelantan

5 July 2009

K8

Labok,Machang

Kelantan

5 July 2009

K9

Labok,Machang

Kelantan

5 July 2009

K10

Tek Desoh, Bachok (4)

Kelantan

6 July 2009

K11

Tek Desoh, Bachok (5)

Kelantan

6 July 2009

K12

Tek Desoh, Bachok (6)

Kelantan

6 July 2009

K13

Pantai Melawi (7)

Kelantan

6 July 2009

K14

Pantai Melawi (8)

Kelantan

6 July 2009

T15

Kg. Abu Ketitir, Lot 20 (1)

Terengganu

25 May 2009

T16

Kg. Abu Ketitir, Lot 20 (2)

Terengganu

25 May 2009

T17

Kg. Alor Ketitir, Marang

Terengganu

25 May 2009

T18

Kg. Alor Ketitir, Marang III (1)

Terengganu

25 May 2009

T19

Kg. Alor Ketitir, Marang III (2)

Terengganu

25 May 2009

T20

Kg. Alor Ketitir, Marang III (3)

Terengganu

25 May 2009

T21

Kg. Alor Ketitir, Marang IV (1)

Terengganu

25 May 2009

T22

Kg. Alor Ketitir, Marang IV (2)

Terengganu

25 May 2009

T23

Kg. Alor Ketitir, Marang V (1)

Terengganu

25 May 2009

T24

Kg. Alor Ketitir, Marang VI (6)

Terengganu

25 May 2009

Collected soil samples were naturally infested with nematodes and Fusarium spp. However from observation, there were no parasitic nematodes in soil K5, K9, K11, and K12. In soil K10, there was no Fusarium spp. isolated.

Table 4.2: Growth and yield characteristic (mean value) of Tobacco as affected by root gall and root lesion at various soil samples in

2009.

Soil

Plant size (cm)

Leaf area

(cm)

No.of leaf

Weight (gram)

Disease severity

Nematode density

Parasitic

(%)

Fusarium solani

Lesion

Gall

P1

10.72

25.44

3.56

2.99

1.6

1.0

639.9

17.22

+

P2

20.53

63.50

3.40

7.42

1.0

1.0

459.9

17.42

+

P3

17.60

44.88

3.80

5.56

1.0

1.0

489.9

18.37

+

P4

8.48

13.50

4.00

2.35

1.0

1.0

530.1

15.11

+

K5

K6

11.95

18.68

29.90

59.00

4.90

5.25

3.09

5.59

0.0

1.8

0.0

1.0

470.1

489.9

0.00

26.56

+

+

K7

25.93

95.60

4.00

15.02

1.5

1.0

710.1

32.44

+

K8

15.53

48.40

3.20

5.15

1.8

1.0

810.0

37.00

+

K9

8.97

19.30

4.00

2.80

0.0

0.0

819.9

0.00

+

K10

20.20

61.50

3.70

7.56

2.3

1.0

720.0

36.13

˗

K11

20.21

69.60

3.86

13.09

0.0

0.0

459.9

0.00

+

K12

15.12

46.70

3.83

5.62

0.0

0.0

560.1

0.00

+

K13

19.39

68.90

4.00

8.05

1.0

1.0

519.9

24.99

+

K14

17.54

48.40

4.70

7.54

1.5

1.0

609.9

34.43

+

T15

12.03

36.13

3.38

3.42

1.6

1.0

770.1

35.00

+

T16

11.82

35.88

3.13

3.91

2.1

1.0

669.9

40.00

+

T17

13.07

40.83

4.17

3.94

3.2

1.0

780.0

30.16

+

T18

16.11

54.60

4.20

4.02

2.2

1.0

740.0

33.82

+

T19

21.66

79.29

5.14

7.93

2.0

1.0

570.0

28.05

+

T20

21.18

72.22

4.78

5.56

1.3

1.0

530.1

30.16

+

T21

22.91

69.00

3.75

7.43

1.6

1.0

570.0

38.58

+

T22

20.47

68.71

4.14

5.44

1.4

1.0

530.1

43.00

+

T23

11.40

31.90

3.40

3.15

1.9

1.0

609.9

36.11

+

T24

16.06

63.00

4.25

4.84

1.4

1.0

510.0

29.41

+

+ : Existence of Fusarium solani - : No Fusarium solani isolated

Table 4.3 Growth and yield characteristic (mean value) of Tobacco as affected by root gall and root lesion at various soil samples in

2010.

Soil

Plant size (cm)

Leaf area

(cm)

No.of leaf

Weight (gram)

Disease severity

Nematode density

Parasitic

(%)

Fusarium solani

Lesion

Gall

P1

22.60

95.60

4.0

16.83

1.0

1.0

390.0

15.38

+

P2

23.90

88.00

4.1

16.33

1.0

1.0

380.1

21.05

+

P3

P4

25.20

24.45

81.14

75.10

3.9

4.2

18.11

16.16

1.0

1.0

1.0

1.0

470.1

489.9

21.27

18.37

+

+

K5

24.20

69.60

3.6

12.69

0.0

0.0

360.0

0.00

+

K6

20.97

81.90

4.0

12.50

1.4

1.0

249.9

24.01

+

K7

23.40

65.30

3.9

14.99

1.7

1.0

549.9

27.28

+

K8

24.30

81.00

3.7

13.15

1.8

1.0

600.0

21.67

+

K9

18.98

56.90

3.6

11.40

0.0

0.0

350.0

0.00

+

K10

30.81

85.70

4.3

21.06

1.5

1.0

459.9

21.74

˗

K11

22.48

85.90

3.7

15.13

0.0

0.0

380.1

0.00

+

K12

23.60

81.70

4.0

17.19

0.0

0.0

360.0

0.00

+

K13

22.20

70.90

3.1

12.63

1.0

1.0

270.0

25.93

+

K14

26.60

87.30

4.0

14.65

1.0

1.0

489.9

38.78

+

T15

25.16

86.67

4.0

14.41

1.4

1.0

350.1

27.71

+

T16

26.97

76.20

3.8

17.48

1.5

1.0

279.9

32.15

+

T17

21.39

75.90

4.2

14.36

2.1

1.0

459.9

34.29

+

T18

17.43

55.40

3.7

10.25

1.4

1.0

470.1

36.16

+

T19

16.97

52.00

4.0

12.2

1.7

1.0

420.0

28.57

+

T20

19.44

68.00

4.1

13.4

1.2

1.0

309.9

38.72

+

T21

19.42

75.30

3.6

10.32

1.6

1.0

630.0

43.59

+

T22

11.98

28.70

3.7

16.60

1.9

1.0

390.0

26.98

+

T23

21.23

89.30

3.5

15.13

1.5

1.0

300.0

40.00

+

T24

24.30

95.80

3.7

13.78

1.3

1.0

309.9

35.49

+

4.2 Effect of plant parasitic nematode on tobacco disease severity

In 2009 (Table 4.2, figure 4.1), the highest percentage of parasitic nematode was soil T22 with 43.00% while the lowest was soil P4 with 15.11%. Comparing average plant size from both of these soils, plant from the soil T22 (20.47 cm) was higher than those of from the soil P4 (8.48 cm). Plant size from P4 was the lowest compare to the others. The average weight of tobacco in for the soil P4 also lower comparing with the soil T22. The soil T17 showed highest root lesion disease severity comparing with the other soil. There was 30.16% of parasitic nematode population density in the soil T17 yet it showed higher disease severity comparing with the soil T22.

In 2010 (Table 4.3, figure 4.2), the soil T21 showed highest infestation of plant parasitic nematode which is 43.59% and the soil P1 was the lowest with 15.38%. The average plant size for the soil P1 (22.60cm) was higher comparing with the soil T21 (19.42 cm). The average weight of the soil P1 was also higher than the soil T21. The soil T21 with high parasitic nematode population showed higher disease severity for root lesion comparing with the soil P1. The result for root gall disease severity index did not showed any different between parasitic nematode population density and disease severity index.

4.3 Soil analysis:

Table 4.4: Soil physical analysis

Soil

pH

Moisture

(%)

Particle density (g/cm³)

Texture

Sand

(%)

Clay

(%)

Silt

(%)

Type

P1

5.97

25.36

2.2019

19.28

56.72

24.00

Clay

P2

6.28

19.26

2.1328

16.28

64.72

19.00

Clay

P3

6.21

28.87

2.0676

16.28

62.72

21.00

Clay

P4

6.27

28.34

2.2222

15.28

63.72

21.00

Clay

K5

5.68

11.22

2.0566

89.28

10.00

0.72

Sand

K6

5.71

11.13

2.0850

89.28

10.00

0.72

Sand

K7

6.12

19.24

2.0212

10.28

51.00

38.72

Silt clay loam

K8

5.87

18.29

2.1285

10.28

51.00

38.72

Silt clay loam

K9

5.84

19.53

2.0794

19.28

55.72

25.00

Clay

K10

5.84

17.97

2.1973

9.28

16.72

74.00

Loam

K11

6.23

18.19

2.2072

8.28

15.72

76.00

Loam

K12

5.98

20.27

2.1654

10.28

18.72

71.00

Loam

K13

6.92

22.17

2.2371

10.28

16.72

73.00

Loam

K14

5.93

20.68

2.1994

10.28

16.72

73.00

Loam

T15

6.19

7.73

2.1375

89.28

1.72

9.00

Sand

T16

5.77

11.29

2.0386

88.28

0.72

9.00

Sand

T17

5.67

8.76

2.0343

91.72

0.72

8.28

Sand

T18

5.75

11.34

2.1379

89.28

1.72

9.00

Sand

T19

5.76

9.55

2.0596

89.28

0.72

10.00

Sand

T20

5.31

7.65

1.7403

88.28

0.72

11.00

Sand

T21

5.96

10.07

2.1599

89.28

1.72

9.00

Sand

T22

5.57

8.81

2.2044

89.28

1.72

9.00

Sand

T23

5.73

8.70

2.1399

89.28

1.72

9.00

Sand

T24

5.83

11.46

2.18129

89.28

1.72

9.00

Sand

Table 4.5: Nematode population and disease severity as affected by different soil

properties in 2009

Soil

pH

Moisture (%)

Particle density (g/cm³)

Texture

Nematode

(%)

Disease severity

Lesion

Gall

P1

5.97

25.36

2.2019

Clay

17.22

1.6

1.0

P2

6.28

19.26

2.1328

Clay

17.42

1.0

1.0

P3

6.21

28.87

2.0676

Clay

18.37

1.0

1.0

P4

6.27

28.34

2.2222

Clay

15.11

1.0

1.0

K5

5.68

11.22

2.0566

Sand

0.00

0.0

0.0

K6

5.71

11.13

2.0850

Sand

26.56

1.8

1.0

K7

6.12

19.24

2.0212

Silt clay loam

32.44

1.5

1.0

K8

5.87

18.29

2.1285

Silt clay loam

37.00

1.8

1.0

K9

5.84

19.53

2.0794

Clay

0.00

0.0

0.0

K10

5.84

17.97

2.1973

Loam

36.13

2.3

1.0

K11

6.23

18.19

2.2072

Loam

0.00

0.0

0.0

K12

5.98

20.27

2.1654

Loam

0.00

0.0

0.0

K13

6.92

22.17

2.2371

Loam

24.99

1.0

1.0

K14

5.93

20.68

2.1994

Loam

34.43

1.5

1.0

T15

6.19

7.73

2.1375

Sand

35.00

1.6

1.0

T16

5.77

11.29

2.0386

Sand

40.00

2.1

1.0

T17

5.67

8.76

2.0343

Sand

30.16

3.2

1.0

T18

5.75

11.34

2.1379

Sand

33.82

2.2

1.0

T19

5.76

9.55

2.0596

Sand

28.05

2.0

1.0

T20

5.31

7.65

1.7403

Sand

30.16

1.3

1.0

T21

5.96

10.07

2.1599

Sand

38.58

1.6

1.0

T22

5.57

8.81

2.2044

Sand

43.00

1.4

1.0

T23

5.73

8.70

2.1399

Sand

36.11

1.9

1.0

T24

5.83

11.46

2.18129

Sand

29.41

1.4

1.0

4.4 Effect of soil physical properties on nematode population and disease severity

Result (Table 4.5) showed that the highest population of parasitic nematode was at pH 5.57 and the lowest at pH 6.27 (Table 4.5, figure 4.3). Root lesion disease severity was highest at pH 5.67 and lowest at pH within 6.21 to 6.92 (figure 4.4). Root gall disease severity index did not show any different with different soil pH.

Result (Table 4.5) showed the highest moisture percentage was 28.87% (Soil P3). However, the highest root lesion disease severity was at 8.76% moisture percentage (soil T17) (Figure 4.6).Root gall disease severity index did not show any different with different soil moisture percentage. Population density of parasitic nematodes was highest at 8.81% (soil T22) while in the lowest at 28.34% (soil P4) (Figure 4.5).

The highest disease severity of root lesion was occurring at soil with particle density 2.0343 g/cm³ (Table 4.5, figure 4.8). The highest population density of parasitic nematode was at 2.2044 g/cm³ and the lowest population density was at 2.2222 g/cm³ (Figure 4.7). Root gall disease severity index seem did not showed any different with different soil particle density.

Table 4.5 showed the highest root lesion disease severity was in sand soil (91.72% sand, 8.28%, silt, 0.72% clay) and the lowest disease severity was in clay soil (15.28% sand, 21% silt, 63.72% clay) (Figure 4.9). The highest parasitic nematode population percentage was in sandy soil (89.28% sand, 9% silt, 1.72% clay) and the lowest was in clay soil (15.28% sand, 21% silt, 63.72% clay) (Figure 4.10).

Table 4.6: Nematode population and disease severity as affected by different soil

properties in 2010

Soil

pH

Moisture (%)

Particle density (g/cm³)

Texture

Nematode

(%)

Disease severity

Lesion

Gall

P1

5.97

25.36

2.2019

Clay

15.38

1.0

1.0

P2

6.28

19.26

2.1328

Clay

21.05

1.0

1.0

P3

6.21

28.87

2.0676

Clay

21.27

1.0

1.0

P4

6.27

28.34

2.2222

Clay

18.37

1.0

1.0

K5

5.68

11.22

2.0566

Sand

0.00

0.0

0.0

K6

5.71

11.13

2.0850

Sand

24.01

1.4

1.0

K7

6.12

19.24

2.0212

Silt clay loam

27.28

1.7

1.0

K8

5.87

18.29

2.1285

Silt clay loam

21.67

1.8

1.0

K9

5.84

19.53

2.0794

Clay

0.00

0.0

0.0

K10

5.84

17.97

2.1973

Loam

21.74

1.5

1.0

K11

6.23

18.19

2.2072

Loam

0.00

0.0

0.0

K12

5.98

20.27

2.1654

Loam

0.00

0.0

0.0

K13

6.92

22.17

2.2371

Loam

25.93

1.0

1.0

K14

5.93

20.68

2.1994

Loam

38.78

1.0

1.0

T15

6.19

7.73

2.1375

Sand

27.71

1.4

1.0

T16

5.77

11.29

2.0386

Sand

32.15

1.5

1.0

T17

5.67

8.76

2.0343

Sand

34.29

2.1

1.0

T18

5.75

11.34

2.1379

Sand

36.16

1.4

1.0

T19

5.76

9.55

2.0596

Sand

28.57

1.7

1.0

T20

5.31

7.65

1.7403

Sand

38.72

1.2

1.0

T21

5.96

10.07

2.1599

Sand

43.59

1.6

1.0

T22

5.57

8.81

2.2044

Sand

26.98

1.9

1.0

T23

5.73

8.70

2.1399

Sand

40.00

1.5

1.0

T24

5.83

11.46

2.18129

Sand

35.49

1.3

1.0

Result (Table 4.6) showed the highest population density of parasitic nematodes was at pH 5.96. The lowest population was at pH 5.97 (Figure 4.3). Disease severity was the lowest at pH 5.93, 5.97, 6.21, 6.27, 6.28, and 6.92 (Figure 4.4). Root gall disease severity index seem did not showed any different with different soil pH.

Result showed (Table 4.6) showed the highest root lesion disease severity was occurring at the soil T17 with 8.76% moisture (Figure 4.6). Highest population density was from the soil T21 with 10.07% moisture (Figure 4.5).

In 2010 (table 4.6), the highest root lesion disease severity index was also occurred at soil with particle density 2.0343 g/cm³. Root gall disease severity index seem did not showed any different with different soil particle density. The highest population density of parasitic nematode was at 2.1599 g/cm³. The lowest population density was at 2.2019 g/cm³.

From the result (table 4.6), highest root lesion disease severity was in sand soil (91.72% sand, 8.28%, silt, 0.72% clay) and the lowest disease severity was in clay soil (15.28% sand, 21% silt, 63.72% clay). The highest population percentage was in sandy soil (89.28% sand, 9% silt, 1.72% clay) and the lowest in clay soil (19.28% sand, 24% silt, 56.72% clay).

4.5 Identification:

Meloidogyne spp.

Pratylenchus sp.

4.6 Disease ranking:

Root lesion:

Root gall:

5.0 Discussion

5.1 Effect of parasitic nematodes population density (%) on tobacco growth and disease severity

Plant parasitic nematodes caused direct and indirect damaged to tobacco. At higher level of population, this nematodes showed higher yield lost. However, not all plant parasitic nematodes that reproduce rapidly on tobacco cause high damage to the plant (Barker, Todd et al. 1981). Apart from that, disease complex between nematodes and other plant parasite such as fungus further affect the tobacco plant. From the result, we can see there were three soil samples shows no infestation of plant parasitic nematode which were soil K5, K9, K11, and K12 (table 4.1). However, although there was no infestation of parasitic nematode the growth of tobacco plant were stunted due to infestation of Fusarium solani which cause root rot (Table 4.2).

In 2009(Table 4.2, figure 4.1), the highest percentage of parasitic nematode was soil T22 with 43.00% while the lowest was soil P4 with 15.11%. Comparing average plant size from both of these soils, plant from the soil T22 (20.47 cm) was higher than those of from the soil P4 (8.48 cm). Plant size from P4 was the lowest compare to the others. The average weight of tobacco in for the soil P4 also lower comparing with the soil T22. This shows that, apart from nematodes, other factor such as soil properties affects growth of tobacco. The soil T17 showed highest root lesion disease severity comparing with the other soil. There was 30.16% of parasitic nematode population density in the soil T17 yet it showed higher disease severity comparing with the soil T22. These result showed that at high population density, their effects on tobacco have been suppressed due to competition. From Pearson correlation (r), there is positive relationship (r= 0.810253) between parasitic nematode percentage and root lesion disease severity (figure 4.1). However, different with root gall disease, there was low disease severity index for all soil and showed no different between the soils. The existence of Fusarium solani seems to cause further damage to tobacco plant.

In 2010 (Table 4.3, figure 4.2), the soil T21 showed highest infestation of plant parasitic nematode which is 43.59% and the soil P1 was the lowest with 15.38%. The average plant size for the soil P1 (22.60cm) was higher comparing with the soil T21 (19.42 cm). The average weight of the soil P1 was also higher than the soil T21. These showed that in general the higher the population density of plant parasitic nematode the lower growth and yield of tobacco would be (Olthof, Marks et al. 1973; Barker, Todd et al. 1981; Barker 1989). Disease severity index also showed same result. The soil T21 with high parasitic nematode population showed higher disease severity for root lesion comparing with the soil P1. There is positive correlation (figure 4.2) between disease severity and percentage of parasitic nematode (r= 0.794495). However, the result for root gall disease severity index did not showed any different between parasitic nematode population density and disease severity index. Different with report by Barker (1989), showed at high population density Meloidogyne sp. which cause root gall were higher especially aggressive species such as Meloidogyne javanica.

In conclusion, at high population density, plant parasitic nematodes suppress growth of tobacco and increase disease severity of root lesion and root gall. However if the population were to high, disease severity were lower due to competition (Barker and Weeks 1991). It was thought that in this experiment plant and nematode response to the environmental variable were responsible for the data variation. For example, early season soil temperatures were higher in 2010. Apart from that, other factors may seem to affect the result as the soil was naturally infested. Fusarium root rot seems to affect result for root lesion disease severity index and also lead to data variable. Moreover, different soil properties may affect to the nematode behavior.

5.2 Effect of soil physical properties on nematode population density (%) and disease severity.

5.2.1 Effect of soil pH

Gammmon & Kincaid (1957) state that number of parasitic nematodes in the root decreased with increasing soil pH. They also reported that disease incidence caused by Pratylenchus spp., increased with increasing soil alkalinity within range of pH 5.2 to 6.2. Egg hatching of Meloidogyne spp., was highest within pH 6.4 to 7.0 ( Loewenberg & Sullivan, 1960). It was also reported that optimum pH for Pratylenchus penetrants movement was 6.0 while P.crenatus within pH 5.0 to 7.0 (Kimpinski and Willis 1981).

In 2009 result shows that the highest population of parasitic nematode was at pH 5.57 and the lowest at pH 6.27 (table 4.5, figure 4.3). Root lesion disease severity was highest at pH 5.67 and lowest at pH within 6.21 to 6.92 (figure 4.4). Root gall disease severity index did not show any different with different soil pH.

In 2010, result showed the highest population density of parasitic nematodes was at pH 5.96. The lowest population was at pH 5.97 (table 4.6, figure 4.3). Same with 2009, disease was most severe at pH 5.67. However, different with results in 2009, disease severity was the lowest at pH 5.93, 5.97, 6.21, 6.27, 6.28, and 6.92. Same with 2009, root gall disease severity index seem did not showed any different with different soil pH.

In conclusion, increasing pH seems to affect the parasitic nematode population density and hence affect the disease severity index. There was weak negative correlation (r= -0.2419) between pH and nematode population density and between pH and disease severity (r= -0.30097). Optimum pH for nematode reproduction was at soil pH 5.2 and 6.4 (Gammon & Kincaid, 1957). Result for root gall was different from the previous result. Times of infestation of root gall nematode seem to affect the result.

5.2.2 Effect of soil moisture (%)

High soil moisture seems to increase the disease severity. Report from Wheeler and Barker (1991) showed that movement of infective juveniles stages (J2) of Meloidogyne incognita was high at high soil water content. From the result in 2009 (table 4.5), the highest moisture percentage was 28.87% (Soil P3). However, the highest root lesion disease severity was at 8.76% moisture percentage (soil T17).Root gall disease severity index did not showed any different with different soil moisture percentage. From table 4.5, population density of parasitic nematodes was highest at 8.81% (soil T22) while in the lowest at 28.34% (soil P4).

Result in 2010 (table 4.6) showed the highest root lesion disease severity was occur at the soil T17 with 8.76% moisture. Highest population density was from the soil T21 with 10.07% moisture.

As a conclusion, there was weak negative correlation (r= -0.38872) between soil moisture percentage and disease severity (figure 4.6) and weak positive correlation (r= 0.244654) between nematode population density and soil moisture percentage (figure 4.5). This result also affected by other factors such as soil texture. However there was no previous report showed effect of soil moisture percentage on nematode population density.

5.2.3 Effect of soil particle density (g/cm³)

Particle density is a measured of a soil mass in a given volume of particle. Soil particle density usually refers to soil particles itself not the volume they occupy in soil. Heavy texture soils have 2.60 g/cm³ particle density, grassland and woodland 2.4 g/cm³ and peats soil 1.4 g/cm³. From the result in 2009 (table 4.5), the highest disease severity of root lesion was occurring at soil with particle density 2.0343 g/cm³. In 2010 (table 4.6), the highest root lesion disease severity index was also occurred at soil with particle density 2.0343 g/cm³. Root gall disease severity index seem did not showed any different with different soil particle density.

Particle density can be used to indicate soil porosity. The movement of plant parasitic nematodes was high at high soil porosity which has low particle density. The highest population density of parasitic nematode was at 2.2044 g/cm³ in 2009 (table 4.5) and 2.1599 g/cm³ in 2010 (table 4.6). The lowest population density was at 2.2222 g/cm³ in 2009 (table 4.5) and 2.2019 g/cm³ in 2010 (table 4.6).

As a conclusion, there is weak positive correlation (r= 0.391908) between particle density and root lesion disease severity and almost no correlation (r= 0.016162) between particle density and nematode population density (Figure 4.6 & 4.7). However there was no previous report on the effect of soil particles density on nematode population density.

5.2.4 Effect of soil texture

Soil texture gave different effect on different nematode species activity (Barker and Weeks 1991). For examples Meloidogyne hapla activity was higher in sandy loam soil (Santo and Bolander, 1979) while ectoparasite nematodes on coarse-textured soils (Jones and Larbey, 1969).

From the result in 2009 and 2010, the highest root lesion disease severity was in sand soil (91.72% sand, 8.28%, silt, 0.72% clay) and the lowest disease severity was in clay soil (15.28% sand, 21% silt, 63.72% clay). Data for root gall disease severity index seems to be inaccurate (table 4.5 & 4.6). More time thought to be needed for the infection to be more viable. Apart from that, in 2009 (table 4.5), the highest parasitic nematode population percentage was in sandy soil (89.28% sand, 9% silt, 1.72% clay) and the lowest was in clay soil (15.28% sand, 21% silt, 63.72% clay). In 2010 (table 4.6), result also showed the highest population percentage was in sandy soil (89.28% sand, 9% silt, 1.72% clay) and the lowest in clay soil (19.28% sand, 24% silt, 56.72% clay). Barker and Weeks (1991) reported M.incognita caused greater yield losses in sandy soil while lower in clay loam soil.

As a conclusion many other factors affect the result due to soil samples were naturally infested. Existence of Fusarium root rot also affect the disease rank for root lesion cause by nematode. More time for cultivation should be done for the root gall to be more viable. However, further research should be done to see the effect of soil chemical properties on nematode and proper experimental design can be done to know effect of soil and nematode on tobacco plant

6.0 Conclusion:

From 24 soil samples, 20 soils were naturally infested with both Meloidogyne spp. and Pratylenchus spp. These soil samples also naturally infested with Fusarium root rot except for soil K10. Meloidogyne spp. led to formation of root gall and Pratylenchus spp. caused rot lesion on tobacco. From Pearson correlation (r) analysis, there was positive relationship (r= 0.810253) between parasitic nematode percentage and root lesion disease severity. Plant parasitic nematodes suppressed growth of tobacco and increased disease severity of root lesion and root gall. There was weak negative correlation (r= -0.2419) between pH and nematode population density and between pH and disease severity (r= -0.30097). Soil moisture percentage and disease severity showed weak negative correlation (r= -0.38872). However, there was weak positive correlation (r= 0.244654) between nematode population density and soil moisture percentage. There was weak positive correlation (r= 0.391908) between particle density and root lesion disease severity and almost no correlation (r= 0.016162) between particle density and nematode population density. Result showed the highest root lesion disease severity was in sand soil (91.72% sand, 8.28%, silt, 0.72% clay) and the lowest disease severity was in clay soil (15.28% sand, 21% silt, 63.72% clay). The highest population density was in sandy soil (89.28% sand, 9% silt, 1.72% clay) and the lowest was in clay soil (15.28% sand, 21% silt, 63.72% clay) in 2009 and clay soil (19.28% sand, 24% silt, 56.72% clay) in 2010. Other factors have affected the result such as environmental condition, and infestation of Fusarium root rot. Further research on effect of soil chemical properties on should be conducted.

7.0 References:

1. Akerhust, B.C. (1981) .Tobacco, 2nd Edition. London New York, Longman, 746 p.

2. Albersheim, P., Jones, T.M. & English, P.D. (1969). Biochemistry of the cell wall in relation to infective process. Annual review of Phytopathology. 7: 171-194.

3. Anon. (1976). A Report on Establishment of Tobacco Research Centre in Malaysia. MARDI. 25 ms.

4. Barker, K.R., Todd, F.A., Shane, W.W. & Nelson, L.A. (1981). Interelationships of Meloidogyne species with flue-cured tobacco. Journal of Nematology, 13:67-79.

5. Barker, K. (1989). "Yield Relationships and Population Dynamics of Meloidogyne spp, on Flue-cured Tobacco." Journal of Nematology.

6. Barker, K., F. Todd, et al. (1981). "Interrelationships of Meloidogyne species with flue-cured tobacco." Journal of Nematology

13

(1): 67.

7. Barker, K. and W. Weeks (1991). "Relationships between soil and levels of Meloidogyne incognita and tobacco yield and quality." Journal of Nematology

23

(1): 82.

8. Bateman, D.F. & Millar, R.L. (1966). Pectic enzyme in tissue degradation. Annual Review of Phtopathology 4: 119-146.

9. Bird, A.F., Downton, W.S.J., and Hawker, J.S. 1975. Cellulose secretion by second stage larvae of the root-knot nematode (Meloidogyne javanica). Marcellia 38: 165-169.

10. Bouyoucos, G.J. (1962). Hydrometer method improved for making particle size analyes of soil. Agronomy journal 54: 464-465.

11. Canter-Visscher, T.W. (1969). The association of root-lesion nematodes with reduced growth of tobacco in the Nelson district. New Zealand Journal of Agriculture Research, 12:423-426.

12. Carneiro, R.M.D.G. ,M.R.A. Almeida, Quénéhervé. Enzyme phenotypes of Meloidogyne spp. Populations. Nematology, 2(6), 2000, 645-654.

13. Cliff, G.M. & Hirshmann, H. (1984). Meloidogyne microcephala n.sp. (Meloidogynidae), a root-knot nematode from Thailand. Journal of Nematology, 16:183-193.

14. Corbett, D.C.M. and Clark, S.A. 1983. Surface features in the taxonomy of Pratylenchus species. Rev. Nematol. 6:85-98.

15. Daulton, R.A.C. & Nusbaum, C.J. (1961). The effect of soil temperature on the survival of the root-knot nematodes Meloidogyne javanica and M.hapla. Nematologica, 6:280-294.

16. Daulton, R.A.C. & Nusbaum, C.J. (1962). The effect of soil moisture and relative humidity on the root-knot nematode Meloidogyne javanica. Nematologica, 8:157-168.

17. Daulton, R.A.C. (1964). Effect of soil fumigant on tobacco in Southern Rhodesia. Biokemia, 5:10-15.

18. Dropkin, V.H. 1972. Pathology of Meloidogyne - galling, giant cell formation effects on host physiology. EPPO Bull. No. 6:23-32.

19. Eisenback, J. D & Hirschmann, H.. Root-knot nematodes: Meloidogyne species and races. In: Nickle, W. R. (Ed.). Manual of agricultural nematology, New York, NY, USA, Marcel Dekker, 1991, 191-274.

20. Eisenback, J.D. 1985. Detailed morphology and anatomy of second- stage juveniles, males, and females of the genus Meloidogyne (root-knot nematodes). In An advances treatise on Meloidogyne, Vol. 1, Biology and control, J.N. Sasser and C.C. Carter, eds., North Carolina State Univ. Graphics, Raleigh.

21. Eisenback, J.D., and Hirschmann, H. 1981. Identification of Meloidogyne species on the basis of head shape and stylet morphology of the male. J. Nematol. 13:413-521.

22. Fortnum, B.A., Krausz, J.P. & Conrad, N.G. (1984). Increasing incidence of Meloidogyne arenaria in flue-cured tobacco in South Carolina. Plant disease, 68:244-245.

23. Gerstel, D.u. 1961.Essay on the origin of tobacco. Tobacco science 5:15-17.

24. Honey, A. de.S. (1967). Studies on the genus Pratylenchus and a preliminary assessment of its importance to tobacco in South Africa. M.sc. thesis (unsubmitted)\

25. Hussey, R.S. 1987. Secretions of esophageal glands of Tylenchida nematodes. In Vistas on Nematology, J.A. Veech and D.W. Dickson, eds. Soc. Nematologists, Hyattsville, MD.

26. Hussey, R.S., E.L. Davis, C. Ray. Meloidogyne stilet secretions. In: Lamberti, F., C. De Giorgi, D. M. Bird (eds). Advances in Molecular Plant Nematology, Plenum Press, New York, 1994, 233-249.

27. Hooper, D.J. (1968). Extraction and handling of plant and soil nematodes. (In nematodes of tropical crops, edited by J.E. Pearchery , Tech. Bull. No.40, Cabb, pp.36.

28. Inagaki and H. and Powell, N.T. (1969). Influence of the root -lesion nematode on Blackshank symptom development in flue-cured tobacco. Phytopathology 59, 1350-1355.

29. Jepson, S.B. (1987). Identification of root-knot nematodes (Meloidogyne species). Wallingford, United Kingdom, C.A.B. international, 256 p.

30. Jepson, S.B. 1987. Identification of the root-knot nematodes (Meloidogyne species). Commonwealth Agr. Bereaux, Farnham Royal.

31. Jones, F.G.W., D.W. Larbey, and D.M. Parrot, 1969. The influence of soil structure and moisture on nematodes, especially Xiphinema, Longidorus, Trichodorus, and Heterodera spp. Soil Biology and Biology chemistry 1:153-165.

32. Jones, M.G.K., and Northcote, D.J. 1972. Multinucleate transfer cell induced in coleus roots by the root-knot nematode, Meloidogyne arenaria. Protoplasma 75: 381-395.

33. Kimpinski, J. and C. Willis (1981). "Influence of soil temperature and pH on Pratylenchus penetrans and P. crenatus in alfalfa and timothy." Journal of Nematology

13

(3): 333.

34. Kimpinski, J., Lelacheur, K.E., Marks, C.F., Thompson, L.S. & Willis, C.B. (1976). Nematodes in tobacco in the Maritime Provinces of Canada. Canadian journal of Plant Sciences, 56:357-364.

35. Kimpinski, J. and L. Thompson (1990). "Plant parasitic nematodes and their management in the Maritime provinces of Canada." Phytoprotection

71

: 45-54.

36. Kincaid, R.R., and N.Gammon, Jr.1966. Effect of soil pH on the incidence of three soil-borne disease of tobacco.Pl. Dis. Rep. 41:177-179.

37. Kosuge, T. (1969). The role of phenolics in host response to infection. Annual review of Phytopathology 7: 195-222

38. Lindford, M.B. 1942. The transient feeding of root-knot nematode larvae. Phytopathology 32:580-589.

39. Loewenberg, J.R., T. Sullivan, and M.L. Schuster, 1960. The effect of pH and minerals on the hatching and survival of Meloidogyne incognita larvae. Phytopathology D 50:215-217.

40. Luc, M., R. Sikora, et al. (2005). Plant parasitic nematodes in subtropical and tropical agriculture, CABI.

41. Madamba, C.P. (1981). Distribution and identification of Meloidogyne spp. in the Philippines and five other Asian countries. Philippines Agriculturist, 64:21-39.

42. McClure, M.A. 1977. Meloidogyne incognita: A metabolic sink. J.Nematol. 9:88-90.

43. Milne, D.L. (1961). A preliminary survey of nematodes present in South African tobacco soils. South Africa. Journal of agriculture Science,. 4: 217-223.

44. Mountain, W.B. (1954). Studies of nematode in relation to brown root rot of Tobacco in Ontario. Canadian. Journal of Botany, 32: 737-759.

45. Naudѐs, T.J. (1929). Insect pest of cotton and tobacco in South Africa. Pan-African Agricultural and Veterinary Conference. Agricultural Section. Paper No. 52:255-256.

46. Nickle, W. (1991). Manual of agricultural nematology, CRC.

47. Nusbaum, C.J. (1960). Soil fumigation for nematode control in flue-cured tobacco. Down to Earth, 16:15-17.

48. Olthof, T., C. Marks, et al. (1973). "Relationship between population densities of Pratylenchus penetrans and crop losses in flue-cured tobacco in Ontario." Journal of Nematology

5

(2): 158.

49. Page, A.L. (ed) (1982) Methods of Soil Analysis, Part I and II. Agronomy No. 9. American Society of Agronomy, Madison.

50. Porter, D.M. and Powell, N.T. (1967).Influence of certain Meloidogyne species on Fusarium wilt development in flue-cured tobacco. Phytopathology, 57: 282-285.

51. Rammah, A. & Hirschmann, H. (1988). Meloidogyne mayaguensis n. sp. (Meloidogynidae), a root-knot nematode from Puerto Rico. Journal of Nematology, 20:58-69.

52. Rammah, A. (1988). Morphological and taxonomic studies in certain populations of root-knot nematodes Meloidogyne arenaria and M.javanica. Ph.D. Thesis, North Carolina State University, Raleigh, U.S.A.

53. Rich, J.R. & Garcia, M.R. (1985). Nature of the root-knot disease in Florida tobacco. Plant disease, 69:972-974.

54. Rowell, D. L. (1994) Soil Science: Method and Application, Department of Soil Science, University of Reading, Longman.

55. Santo, G.S., and W.J. Bolander, 1979. Interacting effects of soil temperature and type on reproduction and pathogenicity of Heterodera schachtii and Meloidogyne hapla on sugarbeets. Journal of Nematology 11:289-291.

56. Schmitt, D.P. & Barker, K.R. (1988). Incidence of plant-parasitic nematodes in the coastal plains of North Carolina. Plant Disease, 72:107-110.

57. Seinhorst, J.W. 1971. The structure of the glandular part of the esophagus of Tylenchidae. Nematologica 17:431-443.

58. Singh, N.D. (1974). Preliminary investigation on the parasitic nematodes associate with tobacco in Trinidad. Nematropica, 4:11-16.

59. Tisdale, W.B. (1922). Tobacco disease in Gadsden County in 1922. University of Florida Agricultural Experimental Station, Bulletin 166.

60. Triataphyllou, A.C., and Hirshmann, H. 1960. Post-infection development of Meloidogyne incognita Chitwood 1949 (Nematoda: Heteroderidae). Ann. Inst. Phytopathol., Benaki 3: 3-11.

61. Tso, T.c. (1972). Physiology and biochemistry of Tobacco plants. Dowden, Hutchinson & Ross. Inc., Strousdbury, P. 3 ms.

62. Viglierchio, D.R. and R.V. Schmitt, 1983. On The methodology of nematode extraction from field samples: Comparison of methods for soil extraction. J. Nematol., 15: 438-444.

63. Webster, J. (1972). Economic nematology, Academic Press.

64. Wells, R. J. G. (1987). The Malaysian tobacco industry. Agricultural international 39: 46-48.

65. Whitehead, A.G. 1968. Taxonomy of Meloidogyne (Nematodea: Heteroderidae) with descriptions of four new species. Trans. Zoo. Soc. London 31:263-401.

66. Wheeler, T., K. Barker, et al. (1991). "Yield-loss models for tobacco infected with Meloidogyne incognita as affected by soil moisture." Journal of Nematology

23

(4): 365.

Web Page:

1. Dow agroScience: 2008 Tobacco Disease Annual Report (Online). (Accessed 3 April 2010) Available from World Wide Web: www.dowagro.com/soil/products/tobacco/economic.htm

2. Ministry Of Primary Industries Malaysia (Online). (Acessed 3 April 2010). Available from World Wide Web: http://www.malaysiayellowpages.net/mpi/details/TOBACCO.htm

3. The Globe Program (Online). (Acessed 25 March 2010). Available from World Wide Web: http://classic.globe.gov/fsl/html/templ.cgi?conversion&lang=ar

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