BIOCONTROL OF

NEMATODES BY PASTEURIA spp.

^* Florida Agricultural experiment station Journal Series No. ......
^**Department of Entomology and Nematology, Institute of Food and Agricultural Sciences, Universtiy of Florida, Gainesville, FL 32611-0620, USA
^***Department of Plant Pathology, Universtiy of Hawaii at Manoa, 3190 Maile

Pasteuria spp., obligate parasites of plant-parasitic nematodes, have great potential as economically and environmentally friendly biological control agents. Field observations (Bird & Brisbane, 1988; Dickson et al., 1991; Dickson et al., 1994; Minton & Sayre, 1989), and glasshouse and microplot experiments (Brown & Smart, 1985; Channer & Gowen, 1988; Chen et al., 1996; Davies et al., 1988; Oostendorp et al., 1991) demonstrate that these organisms can provide effective control of plant-parasitic nematodes. Pasteuria spp. have been reported on a variety of nematode hosts and in many different climates and environments throughout the world (Hewlett et al., 1994; Sayre & Starr, 1988; Stirling, 1988).

Pasteuria spp. endospores are very small and difficult to see and they often go undetected during routine nematode observations. Several methods for detection, manipulation, propagation, and application of Pasteuria spp. endospores have been developed. This is a compilation of techniques used in working with Pasteuria spp. and they should aid researchers interested in studying these biological control organisms.

DETECTION OF PASTEURIA SPP.

Sites where Pasteuria spp. endospore soil densities are high can often be recognized by knowing the previous site history of nematode damage. Continuous or even sudden decline of Meloidogyne spp. and Heterodera spp. field populations, that were previously above economic thresholds, have been found to have high populations of Pasteuria spp. (Bird and Brisbane, 1988; Dickson et al., 1994; Dickson et al., 1994; Minton & Sayre, 1989; Nishizawa, 1987; Weibelzahl-Fulton et al., 1996). Pasteuria spp. infecting Belonolaimus longicaudatus and Meloidogyne spp. were located by sampling golf course putting greens and fairway turfgrass, and flower beds, that previously had been damaged by nematodes but had shown remarkable improvements in plant growth (unpublished). Pasteuria penetrans also was detected in fields that had ideal conditions for the development of severe disease from root-knot nematodes but continued to produce quality yields even though planted with root-knot nematode susceptible crops (Mankau, 1980)

Pasteuria spp. endospores are small, approximately 1.5 to 8.0 æm in diameter, thus they require high-power magnification to verify their presence on the cuticle of nematodes and (or) inside the nematode body. In most cases nematodes extracted from soil samples are identified using dissecting microscopes, which makes it difficult to see endospores attached to the nematode cuticle. Although Pasteuria spp. are visible under low power, they can be confused with fungal spores or soil debris. Often the first indication of the presence of Pasteuria spp. on nematodes extracted from soil in samples is the clumping of nematodes (Mankau & Prasad, 1977).

Inverted microscopes are a valuable tool in working with Pasteuria spp. because they enable the user to count nematodes extracted from soil samples at low power and also to detect or count endospores attached to the nematode cuticle at a magnification of 400 to 600. This eliminates the need to mount the specimens on slides for close examination to verify the presence of Pasteuria spp. with light microscopy under high power objectives or oil immersion.

Pasteuria penetrans endospores are round shaped with a dense central core. They have been described as having a flying saucer shape or hat shape. Endospores can be confused with lipid drops, debris, and nematode body parts. During work with Pasteuria spp., a permanent mounted slide of endospores is helpful in keeping the size and shape of the endospores in perspective.

Endospore filled nematode cadavers are often the sole source of Pasteuria spp. inoculum for experiments. Endospore filled females of Meloidogyne spp. are identified by their opaque dull creamy white to amber color when viewed by a dissecting microscope (light directed from above specimens), and absence of egg masses (Mankau & Prasad, 1977). This color often extends into the pharyngeal region of the females. Endospore filled ectoparasitic nematodes are generally dense, crystalline black (often including the pharyngeal region) when viewed under low power with a dissecting microscope (light directed from below specimens). Endospores can be detected inside the nematode body by viewing specimens under a high power microscope. Positive identification of Pasteuria spp. infected nematodes should be done by crushing the body under a coverslip in water and making observation of the nematode body contents at magnifications between 250 and 400.

STAINING TECHNIQUES

Vegetative stages of Pasteuria spp.: Pasteuria spp. vegetative stages developing inside Meloidogyne spp. J2 and females Fig. 1 can be stained dark blue by placing freshly dissected nematodes into a drop of lactophenol with 1% cotton blue, (v/w) (Serracin, et al., 1997) Fig. 2 . Nematode bodies can be crushed by applying pressure to the coverslip placed over the nematodes that have been placed in the staining solution. Pasteuria spp. are gram positive and vegetative structures and endospores can be stained dark blue using standard gram stain techniques, however, this technique is not good for quantitative studies because specimen parts are lost during slide washing.

Mature endospores: Brilliant Blue stain gives a satisfactory staining of endospores when applied as a 1 mg/ml solution in distilled water (Bird, 1988). Williams et al. (1989) found that P. penetrans endospores reacted to Schaffer-Fulton stain similarly to other endospore-forming organisms; the central core stained light green, whereas, the sporangial wall stained red. The sporangial wall of P. penetrans stained red, but the endospores do not stain using a modified Dorner stain. Using Mormak and Casida's staining procedure P. penetrans endospores stain purple (Williams et al., 1989). A fluorescent DNA stain (4'-6-diamidino-2-phenlindole) was used to stain P. penetrans endospores in M. javanica females (Bird, 1986). The fluorescent stain Acridine Orange (1% Acridine Orange in water) stained endospores of Pasteuria spp. attached to B. longicaudatus and Hoplolaimus galeatus, but did not stain endospores of P. penetrans (unpublished).

COLLECTION OF ENDOSPORES

Pasteuria penetrans endospores can be collected from infected females of Meloidogyne spp. by dissecting them from plant roots or by using a cytolase-sieving technique (Oostendorp et al., 1990). Plant roots are incubated in 12% cytolase PCL5 solution (Genencor, South San Francisco, CA) overnight. Roots are then washed vigorously over a 600-æm-pore sieve nested over a 150-æm-pore sieve. Females of M. arenaria are caught on the 150-æm-pore sieve, and the endospore-filled females, conspicuous by their opaque, white appearance, are collected (using forceps or a glass pipet) with a stereomicroscope. Specimens must be illuminated from above. A similar technique was used to collect the bacterium from Heterodera avenae endospore filled bodies (Davies et al., 1990).

Greater numbers of endospore filled vermiform endoparasitic and ectoparasitic nematodes, e.g., Pratylenchus scribneri, H. galeatus, and B. longicaudatus, can be collected from soil using centrifugal flotation with a sucrose solution with a specific gravity of 1.22 to 1.26 (Oostendorp et al., 1991b). The density of endospore filled nematode bodies of vermiform nematodes is increased because of the densely packed endospores within the diseased nematode body. By using a medium, such as sucrose, with a higher specific gravity than is normally used in the extraction of nematodes from soil, great improvements can be obtained in the identification of nematodes diseased by other microorganisms, e.g., fungal antagonists. The specific gravity of endospores of P. penetrans collected from Meloidogyne arenaria was shown to be 1.28 (Oostendorp et al., 1991b). Mesocriconema ornatum, Ditylenchus spp., and Aphelenchoides spp. endospore filled cadavers also have been extacted from soil with the use of centrifugal flotation with the higher density of sucrose solution (unpublished).

PREPARATION OF ENDOSPORE WATER SUSPENSIONS

Endospore water suspensions have been used as a method for continued propagation of Pasteuria spp. and to determine host preferences. Mankau and Prasad (1977) collected endospores from cadavers of endospore filled Meloidogyne spp. and used hand tools to crush the bodies in water. Oostendorp et al. (1990) macerated large numbers of Meloidogyne spp. spore-filled cadavers with the use of a glass tissue grinder to release endospores. Most of the nematode cuticle can be removed from water suspensions by hand picking with forceps or by passing endospore suspensions through a 24 æm-pore-sieve.

Endospore suspensions also may be attained from air-dried root systems containing Pasteuria spp. infected females of Meloidogyne spp.. Davies et al. (1988) air-dried root systems approximately 42 days before milling the roots. The root powder was mixed with water and the mixture was washed through a 30-æm-pore sieve to remove large root debris.

Collecting endospores from vermiform shaped ectoparasitic nematodes is much more difficult than collecting them from the larger pyriform-shaped nematodes, such as, Meloidogyne spp. Cadaver cuticles are not easily disrupted by tissue grinders because they float away from the grinding surface. Cutting cadavers with hand tools is time consuming and many endospores are not released. Also, the number of endospores released may be small, for example, B. longicaudatus averages approximately 6,000 endospores per spore-filled cadavar (unpublished). Endospores from cadavers of vermiform nematodes, such as, B. longicuadatus, M. ornatum, and H. galeatus can be collected by using glass beads, and mortar and pestle. Endospore-filled nematode cadavers are placed into 0.2 ml of distilled water in a 2.5 ml microcentrifuge tube and allowed to sink to the bottom of the tube. Glass beads (25 æm diam.) are added and a 14 cm long disposable plastic-tissue grinder (Fischer Scientific, Pittsburgh) is used to grind up the nematode bodies and release the endospores into the water. The tube is then shaken and the water-endospore suspension is pipetted from the centrifuge tube. The tube is filled with water two more times to collect all endospores (unpublished).

Endospore suspensions should be prepared and stored in containers that have water repellent properties. Endospores readily adhere to glass, plastic, and metal, which leads to the loss of large numbers of endospores and also poses as a threat for contamination of healthy nematode colonies. Glassware should be treated with water repellent chemicals, e.g., Repel Silane (LKB, Bromma, Sweden). Salinized pipettes and plastic containers are available, and soaking plastics with milk before use also works well.

The adhesive properties of mature Pasteuria spp. endospores can cause contamination problems in the laboratory. Pipette tips and plasticware used with Pasteuria spp. should not be reused when handling healthy nematode populations. Glass, hand tools, and sieves should be autoclaved to avoid contaminating clean cultures of nematodes.

ENDOSPORE ATTACHMENT

Attachment of Pasteuria spp. endospores to nematodes is necessary to acertain host range preferences, for efficacy tests, and for the propagation of Pasteuria spp. The most common method is to make a nematode-endospore water suspension that is left stationary or agitated at room temperature (ca. 25 C) for 1 day. Attachment rates of 1.6 endospores/nematode have been reached using 1.5 x 103 spores/ml in watch glasses with 20 Meloidogyne spp. second-stage juveniles (J2) (Channer & Gowen, 1988). Endospore attachment rates of 100% were obtained on Meloidogyne spp. J2 after 12 hours using a rotary shaker (90 revolutions/minute) with 200 J2 placed in staining blocks in 0.5 ml of a suspension of 106 endospores/ml (Davies et al., 1988). Meloidogyne spp. J2, bubbled in a water suspension with 3 x 103 endospores/ml for 18 hours, had a 78% attachment rate (Bird, 1986).

Meloidogyne spp. J2 were attached with endospores by placing them on a 1% water agar petri dish inoculated with 200 to 500 ml of a 5 x 105 endospores/ml suspension (Verdejo & Jaffee, 1988). The endospores attached to the J2 as they moved across the water agar. Endospore attachment to J2 also was accomplished by inoculating them into endospore infested soil and then extracting the nematodes via centrifugal flotation (Brown & Smart, 1985).

A rapid rate of endospore attachment to nematodes was achieved using a centrifuge technique (Hewlett & Dickson, 1993). Endospore suspensions were placed with 200 Meloidogyne J2 in 0.2 ml centrifuge tubes and centrifuged in a microfuge at 9,500g for 2 minutes. The rate of attachment of P. penetrans to M. javanica at 103, 104, and 105 endospore/0.1 ml/tube from two tests averaged 1.0, 5.76, and 28.3 endospores/nematod, respectively, with 100% of the J2 attached at the two higher rates. Attachment rates of 100% on 3 x 104 J2 of M. javanica were obtained with 5 x 105 endospores/ml, using a table top centrifuge.

Sonication causes the disruption of the sporangial coat of P. penetrans endospores and increases the rate of attachment to nematodes by several fold (Stirling et al., 1986). Endospores can be sonicated by placing endospore suspensions in tap water at pH 7, sonicated in an ice water bath for 10 minutes using a Kerry's ultrasound generator (Hawksley and Sons, LTD, London) fitted with a logarithmic probe (Davies et al., 1988) and using a IV ultrasound generator (Bronwill Scientific, Rochester, NY) supplying 20 kHz modulated electrical output with a power setting of 100 W (Stirling et al., 1986).

ENUMERATION OF ENDOSPORES

Endospore suspensions: The hemocytometer is the most common instrument used to estimate numbers of endospores per millimeter in water suspensions.

Endospore density in the soil: Presence of endospores in soil can be detected using a simple bioassay. Healthy, susceptible nematodes allowed to migrate through endospore-infested soil, and after a given time the nematodes are extracted from the soil sample and examined microscopically for the number of endospores attached to their cuticles (Sayre & Wergin, 1977). Estimations of endospore densities in soil have been made by creating a standard using three endospore-soil densities. Autoclaved field soil, 40 g placed in petri dishes, were inoculated with 10³ , 10 ⁴, and 10⁵ endospores. Healthy susceptible nematodes were inoculated into this soil and the nematodes extracted 3 days later using centrifugal flotation. Average number of endospores attached per J2 were then compared with the attachment of J2 placed into 40 g of air-dried field soil (Frietas, 1997).

Direct estimates of endospores in soil have been made using a centrifuge-sucrose gradient. An average of 59% of endospores present in soil were extracted with this technique, but it was concluded that the method was too laborious for routine use (Davies et al., 1988).

Estimations of endospore densities in roots used for increasing inoculum for field experiments has been made (Chen et al., 1996a). The highest estimates of endospore densities in root powder were obtained by grinding 0.5 g of root powder with a mortar and pestle. The slurry was diluted 200 times with water and the endospores concentration determined with a hemocytometer at a magnification of 450.

Endospores attached to nematode cuticles: Determining the number of endospore attached to nematodes is important for host attachment preference studies and for bioassay studies of endospore soil densities. Endospores can be difficult to detect because of their position of attachment on the nematode body. Endospores attached opposite the side observed with a microscope are difficult to see and intestine contents can hinder viewing. Also, in many cases large numbers of endospores are attached to nematodes, sometimes greater than 200 per nematode, which makes counting tedious. Chen (1996c) used tally thresholds T to determine the degree of endospore attachment of P. penetrans to Meloidogyne arenaria J2. This method decreases the need for precise counting of each endospore. The degree of endospore attachment is expressed by determing the proportion of J2 that has # T endospores attached.

CULTURING OF PASTEURIA PENETRANS

Currently the most efficient method of producing large quantities of P. penetrans endospores is to inoculate endospore infected Meloidogyne spp. J2 onto a suitable host plant. Plant roots were harvested and air dried for storage after 45 to 60 days (Stirling & Wachtel, 1980). Using this technique tomato and tobacco plants inoculated with 5,000 endospore encumbered J2 produced 1.86 x 109 and 54 x 106 endospores, respectively, per root system (Gowen & Ahmed, 1990).

Different methods of rearing Pasteuria spp. on nematodes have been studied (Sharma & Stirling, 1991). Results suggest that susceptible root systems have a maximum carrying capacity and that increasing the number of endospore attached per juvenile above a certain level does not increase the number of females in the roots and, therefore, the endospore concentration in them. In these experiments the main factors affecting the number of endospores produced were nematode inoculum density, plant species, and time of harvest.

Pasteuria penetrans has been grown on M. javanica placed in tissue culture with Agrobacterium rhizogenes transformed roots (Verdejo & Jaffee, 1988), but only one generation of endospores was produced. The cuticles of endospore filled females do not break down, thus the endospores are not released to initiate a new cycle.

Hydroponics was used to grow P. penetrans on Meloidogyne spp. with tomato (Serracin et al, 1997). Efficiency of this technique compared to growing Pasteuria spp. in potted plants, for large scale production, has not been tested. Hydroponics stimulates frequent root tip production, thus inoculations and infection of nematodes on roots can be optimized.

Population densities of Pasteuria spp. introduced or indigenous in fields can be increased by cultural practices. Significant control of root-knot nematodes has been achieved after four cropping cycles, by reincorporating the root systems of each crop containing Meloidogyne spp. infected with P. penetrans into the soil (Channer & Gowen, 1990). Drying the roots on the soil surface before reincorporation, should kill root-knot nematode J2 and eggs but not damage the endospores. This enabling the bacterium to accumulate and remain as infective inoculum in the field (Davies et al., 1991).

Pasteuria spp. cultures should be kept away from Pasteuria free nematode cultures by placing them in a separate room or greenhouse, if possible, to reduce the risk of contaminating Pasteuria free nematode cultures.

STORAGE OF PASTEURIA SPP.

Pasteuria spp. endospores can survive prolonged periods under dry conditions (Oostendorp et al., 1990; Stirling & Wachtel, 1980). A common practice for storage of P. penetrans is to dry root systems that have endospore filled root-knot nematode females. The roots may be stored intact or ground up into a powder. Endospore laden root powder that has been in dry storage must be rehydrated for 24 hours or more before endospores will begin to attach to root-knot nematode J2. P. penetrans endospores do not begin attaching to M. arenaria J2 (in 40 g of soil with 8 ml of water), until 72 hours after inoculation of 1 g of root powder containing an estimated 8 x 106 endospores/g. (unpublished).

Pasteuria spp. collected from ectoparasitic nematodes requires hand picking endospore filled bodies. The endospore filled cadavers can be stored for future use by placing them on glass slides and allowing them to dry. These endospore filled cadavers can be recovered at a later date by rehydrating their bodies with a drop of water after which they can be hand picked from the slide (unpublished).

Aqueous endospore suspensions in distilled or tap water can be refrigerated (4 C) for several months and remain infective. P. penetrans endospore suspensions also have been stored frozen, and upon thawing remained viable and able to attach to root-knot nematode J2 (unpublished). Endospore laden soil can be stored easily for years by air drying the soil and placing it in an air-tight container stored at room temperature.

INOCULATION TECHNIQUES

Acceptable control of root-knot nematodes, using applications of P. penetrans has been demonstrated in greenhouse and microplot experiments . Methods of delivery to soil included applications of endospore water suspensions (Chen et al., 1995), endospore-laden root powder (Stirling and Wachtel, 1980), endospore-laden soil (Mankau & Prasad, 1977) , and newly hatched root-knot J2 encumbered with P. penetrans endospores (Chen et al., 1996b).

The number of P. penetrans endospores needed to control root-knot nematodes depends on the susceptibility of the nematode to the Pasteuria spp., number of endospores that are viable, and the number of endospores attached to J2. Davies, et al. (1988) found that when Meloidogyne spp. J2 were attached with endospores before inoculation on to plants, attachment of at least five endospores per juvenile was found to be necessary to ensure infection and M. incognita J2 root penetration was reduced by 86% when J2 were attached with 15 or more endospores. Root-knot nematode J2 encumbered with 40 or more endospores were prevented from penetration into roots and nematode populations were controlled when at least 80% of the J2 were each encumbered with at least 10 endospores (Stirling, 1984).

The densities of endospores in soil needed to control plant-parasitic nematodes are dependent on several factors including soil texture and temperature, susceptibility of nematodes to Pasteuria spp. isolates, distance and time nematodes travel through endospore infested soil, and nematode population densities.

Several studies have demonstrated endospore densities of between 104 to 105 endospores/g of soil are needed to give significant reductions of damage caused by Meloidogyne spp. populations (Chen et al., 1995; Stirling, 1988; Stirling et al., 1990). M. arenaria J2 inoculated into soil previously inoculated with P. penetrans at 10³, 10, 10 ⁵ , and 10⁶ endospores/g of soil gave an average of 36, 58, 92, and 100% J2 attached with endospores with an average of 1.2, 3.4, 9.6, and 51 endospores/J2, respectively, after 3 days in the soil (unpublished).

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