DIY Organic Potting Mix's for Grass - Ace Spicoli

A Farmer’s Guide to Rearing Predatory Mites La Trinidad Benguet 2007​

Beverly S. Gerdeman1 and Rufino C. Garcia2
Lynell K. Tanigoshi1
1 Washington State University, USA
2 UPLB Laguna, Philippines
Basically the methods described follow two simple principles to rear predatory mites:

1. leaves are stacked.
2. the device is inverted.

With this in mind, you may be able to design a method that will better suit your own area or needs. You are encouraged to try!

I. Tile Method (developed by Glen Scriven and James McMurtry, UC Riverside)​

Shallow pan set up for the Tile Method.

Materials​

• Shallow pan filled with ½ inch of water.
• 1-inch mattress foam cut to fit pan. Squeeze in water until soaked.
• Black plastic or vinyl tile cut to fit foam. Leave a narrow margin of foam exposed.
• Tissue cut to fit around the edge of the black plastic. Drape it over the edge of the foam and touch the water.

Preparation​

• Place a handful of spider mite infested leaves on the black plastic.
• Place a leaf with predatory mites on the spider mite infested leaves.

Where to find a start of predatory mites​

(We are using Neoseiulus longispinosus.)

1. Search abandoned strawberry fields.
2. Look for poinsettia leaves infested with spider mites.
3. Look for spider mite infested cassava leaves.

Maintenance​

There is no exact schedule for maintenance. It depends on the rearing method you choose, where you place your colony and the weather. How often you add food depends on the number of predatory mites you begin with and how many spider mites you feed them.

• Maintain correct water level.
• If only a few live spider mites remain, add more infested leaves.
• Keep leaves in the center of the tile or plastic, to prevent moldy leaves.
• Remove moldy leaves.

Release​

(Refers to all the rearing methods presented.)

• When you can see predators on every leaf
• When you see only a few live spider mites on each leaf.

II. Clay Pot Method (developed by Carl Shanks, Washington State University)​

Materials​

The Clay Pot Method.

• Two shallow clay pots (you may use a single pot if you prefer).
• One shallow basin large enough to hold the clay pots

Preparation​

1. Wet but don’t soak the pots in water.
2. Invert one pot in the basin.
3. Fill the basin so that the water comes up to the middle of the pot.
4. Place a handful of spider mite infested on the clay pot.
5. Place a start of predators on the leaves.

Maintenance​

• Maintain proper water level.
• Remove moldy/black leaves.

III. Can Method (developed by Lynell K. Tanigoshi, Washington State University)​

Materials​

The Can Method

• Two cans with plastic snap-top lids (such as cans containing powdered milk for infants)
• small amount of chickenwire
• elastomeric sealant

Construction​

1. Cut out the bottom of the cans leaving a narrow ledge.
2. Cut a circle of chicken wire to fit between the two cans.
3. Run a bead of sealant along the ledge of one can and press the chickenwire in place.
4. Run a bead of sealant along the ledge of the other can and press the two cans together.
5. Run a bead of sealant outside where the cans join, to seal any gaps between the cans.
6. Allow to air dry before using.
7. Cut a hole in each snap-top lid and glue in a 120 mesh silkscreen window for ventilation.

Preparation​

• Fill one side of the can with dry, spider mite infested strawberry leaves.
• Set the cans upright with the leaves in the upper can.
• Place a start of predators inside the upper can with the spider mite infested leaves.
• Snap both lids on the cans.

Maintenance and Use​

• Check for moisture inside the cans. Dry them out.
• If fungus develops in the can or on the leaves, clean it out and start all over.
• When only a few live spider mites remain, fill the lower can with spider mite infested leaves and invert so the new leaves are now in the upper can (do not remove the old leaves yet).
• When there are only a few live spider mites in the upper can, invert and remove the oldest leaves (now in the upper can) and replace with fresh spider mite infested leaves.
• Repeat until you can see at least one predator on each leaf and there are few spider mites.

Remember the freshest leaves should be in the upper can because most predatory mites like to crawl up!

IV. Bag Method (developed by Beverly S. Gerdeman, Washington State University and Rufino C. Garcia, UPLB)​

Materials​

The Bag Method
You can sew any size but remember—the larger the bag, the farther the mites have to crawl to reach the fresh leaves.

• Silkscreen (at least 120 mesh)
• Netting
• Two bamboo sticks or flat Plexiglas strips. The length should match the width of your tube.
• Two hangers

Construction​

1. Sew the silkscreen into a tube. (Be careful and do not make any unnecessary holes with the needle or the predatory mites will escape.)
2. Cut a circle of netting and sew inside the tube, dividing it into two equal halves.

How to Use​

• Place spider mite infested leaves in one side of the bag.
• To close, roll the silkscreen ends of the bag on a bamboo stick and hang the bag in a shady, warm spot protected from rain.
• Check leaves and when only a few live spider mites remain, place new spider mite infested leaves in the empty side of the bag and invert but do not discard the old leaves yet because they contain eggs which haven’t hatched yet. (Remember, new leaves are always in the top portion of the can).
• When few live spider mites remain on the newest set of leaves, invert the bag, now remove the old leaves and fill the top portion with fresh spider mite infested leaves.
• Repeat the process, until you can see at least one predator on each leaf and only a few live spider mites remain.

---

Abstract: Coccinella novemnotata L., the ninespotted lady beetle, and Coccinella transversoguttata richardsoni Brown,
the transverse lady beetle, are predatory species whose abundance has declined significantly over the last few decades in
North America. An ex situ system for continuously rearing these two beetles is described here to aid conservation efforts
and facilitate studies aimed at determining factors in their decline and possible recovery. All rearing of lady beetles was
conducted in the laboratory at or near room temperatures and 16:8 L photoperiod. The two coccinellid species were
each reared separately, and different life stages were handled independently. Eggs were collected every 1 to 2 d and
placed in holding containers, and individual clutches were transferred to cages with prey when their eggs began to hatch.
Neonate larvae were fed live bird cherry-oat aphids [Rhopalosiphum padi (L.)] for 3 to 4 d, and second instars were trans-
ferred to different cages and fed live pea aphids [Acyrthosiphon pisum (Harris)]. Third and fourth instars were also fed pea
aphids, but reared individually in small cups to preclude cannibalism. Upon pupation, individuals were collectively trans-
ferred to fresh cups and placed in a different container for the duration of pupation. Newly emerged adults were collected
within containers about 2 d after eclosion. Adults were housed in cages stocked with live pea aphids, supplemental food,
and rumpled paper towels as oviposition substrate. Over 80% of egg clutches were deposited by beetles on rumpled paper
towels versus other surfaces within cages, and incidence of cannibalism of egg clutches was greatly reduced on rumpled
paper towels. Techniques for successful rearing of these two coccinellids and future research regarding adaptations to fur-
ther optimize their rearing methods are discussed.
Keywords: Coccinella novemnotata, Coccinella transversoguttata, Coccinella septempunctata, bird cherry-oat aphid, pea
aphid, insect conservation.

11$ lifetime warranty, machine washable

For lack of a better place to leave this... feel free to drop your organic diy soil equipment setup etc

Anyone else have pot size recommendations for outdoor and indoor ?

I go for the bigger the better with my water only indoor plants. They get 90l for up to 3 plants under a scrog screen. With more than 3 plants i got one big dominante plant(thickest stem) most of the time and the others stay a little behind. Outdoor i would say it depends on how big the plants can get. If size is not a problem put them in the ground and mulch.

Soil

Besides a good seed, any primo plant needs a fertile, living, organic soil to be really healthy. For a cubic yard of soil (enough to fill approximately 10-20 3ft deep by 3ft wide holes for plants) use 40% compost (or worm castings or horse shit), 30% aged or composted redwood sawdust and 30% sandy loam. Add 5 lb. greensand, 5 lb. rock phosphate, 5 lb. dolomite, 3 lb. hoof and horn meal (or 3 lb. fish meal), and 2 lb. kelp with no salt. Also; enough trace elements, with iron sulfur, for a yard of soil. Agricultural frit is a good source but don’t overdo it!

Mix well, and allow to “set” at least a few weeks. Then test for PH. With Dolomite, adjust the pH to between 7 and 8 – slightly alkaline). You might want to use sterilized potting soil for starting your seeds, if you’re short of good seeds and fear damping off. And a cat or mouse trap, if you have mice.

Sprouts and young plants, kept off the ground, say on a table with a 1/4″ wire mesh screen cage on top and around them will also keep out mice, yet admit light and air. crickets, slugs and other insects will eat young sprouts. It’s pretty discouraging to wake up one morning to find out that the one ten year old hash seed you’ve saved and carefully hand germinated was some bug’s dinner! Destroy any questionable bugs or use fine window screening, instead of 1/4″ mesh, to protect your ladies.

With enough room (5 feet between plants if possible) and a loose, aerated, organic soil, thousands of tiny hairlike roots will make available the nutrients that can help grow giant, potent, gooey ladies. A good sized healthy plant can easily have a 50 gallon volume of roots, mostly very fine and spread out, not so much deep down. Anyone who thinks Cannabis roots are all small just hasn’t seen a really healthy plant. If seeds are started in little 4″ peat pots, transplant before one month to avoid stunting. Larger container plants should be planted in the ground before flowering.



From, sun-soil-seeds-soul-1979

Thanks for the advice acespicoli! Although I recommend not digging as deep as 3 ft if thats what you mean. Plant roots are generally in the first foot of soil, and if you go too deep you bring up the subsoil. Its good to maybe loosen it up and dig deep the first time, but now, I just loosen the soil at the top about a shovel deep.

Potato dextrose agar - Wikipedia

en.wikipedia.org

Leaf Spot – Curvularia pseudobrachyspora – 26 June 2020​

This pathogen was first observed during a field trial in 2019. Leaf spot symptoms on some varieties were seen as high as 70%. The symptoms were typical of leaf spot with small yellow spots which became tan to brown with an accompanying yellow halo. The pathogen was observed on young and old leaves.


https://apsjournals.apsnet.org/doi/10.1094/PDIS-03-20-0546-PDN





Photo Credit - Marin, Coburn, Desaeger and Peres. UF-GCREC, Wimauma, 2020. Infection of Curvularia pseudobrachyspora on hemp leaf and pathogen morphology.

Cercospora Leaf Spot – Cercospora cf. flagellaris – 17 March 2020


This pathogen was first observed during a greenhouse and field trial in 2019. Leaf spot symptoms were seen on as high as 60% of the plant’s leaves. The symptoms were typical of leaf spot, beginning as small yellow flecks, developing into lesions which turned to light tan or white, with yellow halos, and in severe cases developing chlorosis, leading to defoliation. The infections began on older leaves and moved through the canopy.


https://apsjournals.apsnet.org/doi/10.1094/PDIS-11-19-2287-PDN





Photo Credit – Marin, Coburn, Desaeger and Peres. UF-GCREC, Wimauma, 2020. Infection of Cercospora cf. flagellaris on hemp leaf and pathogen morphology. https://doi.org/10.1094/PDIS-11-19-2287-PDN

Septoria Leaf Spot – Septoria sp.​

Septoria leaf spot is one of the most common leaf spot pathogens among the hemp leaf spot diseases. Like other leaf spot pathogens, the symptoms of this disease begin in the lower canopy as small brown spots with yellow margins. The leaf spots may continue to grow, turn brown and coalesce. More advanced symptoms may be observed, such as leaf yellowing, stunting, and plant death. This disease can progress quickly during the summer months when there is excess moisture, humidity, and the canopies are more dense.

Management Considerations for Leaf Spot Diseases​

At this time the best recommendation is to avoid susceptible cultivars. Research is ongoing to determine how to best manage these pathogens.

---

Powdery Mildew – Golovinomyces sp.​

This disease is characterized by its white powdery appearance on the upper surface of the leaf. The powdery growth is made up of masses of fungi that give it its distinct appearance. Additional symptoms can include leaf necrosis and distortion, as well as early defoliation. The disease is common in both outdoor and greenhouse growing environments. High humidity and free water increase susceptibility to this disease. The pathogen is spread by spores that originate from infected tissue. The most common means of spore dispersal are through air currents.

Management Considerations for Powdery Mildew​

At this time, cultural controls are the most effective way to manage Powdery Mildew. Specifically through the manipulation of the growing environment. The most effective cultural control strategy is to reduce humidity and moisture around and on the plant surface. This may be accomplished by increasing airflow around the plants to avoid areas of high humidity, and if possible, avoiding overhead irrigation.

---

Stem Canker – Diaporthe phaseolorum​

This pathogen was first observed during a field trial in 2019. The symptoms observed with this disease were stem cankers. The symptoms began on the main stem as light-to-dark brown lesions of varying formations, eventually coalescing into larger necrotic areas. In this trial, at least 60% of three-month-old plants showed symptoms of this disease.





Photo Credit – Marin, Wang, Coburn, Desaeger, and Peres. UF-GCREC, Wimauma, 2019. Symptoms in different stages of disease development of hemp stem canker, caused by Diaporthe phaseolorum.

Management Considerations for Stem Canker​

At this time, management of this disease may only be accomplished through resistant cultivars. Research is ongoing to identify more about management of this disease.

---

Gray Mold – Botrytis cinerea​

The gray mold pathogen is one of the most important pathogens, as it is typically considered to be always present. Since this pathogen is rarely ever absent, it relies on wounds or natural openings to infect its host. The symptoms of this pathogen are often observed on the flower buds that are generally observed as having a gray moldy appearance. This fungus is very opportunistic, so any openings caused by pruning or insect damage are sites for possible infection. Other symptoms associated with this pathogen depend upon the site of infection, i.e. girdling and breaking of the stem. This pathogen may also cause damping off of seedlings in greenhouses, and is easily spread through the air.

Management Considerations for Gray Mold​

While this pathogen can occur in field and greenhouse settings, it is favored by the high relative humidity inside greenhouses, which makes it a particularly important pathogen for indoor growers. This pathogen is very difficult to manage once it is found, but reducing air current after infection, sanitation afterwards are important to limiting the spread of the gray mold pathogen.

Municipal compost can be a source of soil borne disease, bacterial fungal and viral
Its a very good practice to dispose of infected material appropriately

The pathogens that infect cannabis plants cause different symptoms. (A)-(C) Fusarium causes yellowing of foliage and internal stem necrosis. (D)-(F) Pythium causes wilt and crown necrosis. (G)-(I) Golovinomyces causes powdery mildew on leaves and inflorescences. (J)-(L) Botrytis causes bud rot. Respective pathogen cultures are shown in (C), (F) and (L).​

Contexts in source publication​

Context 1
... Using symptomology, morphological criteria and molecular approaches for pathogen identification, these descriptions of cannabis and hemp diseases are the most comprehensive currently available. In the present review, further characterization of newly emerging pathogens of cannabis and hemp reported over the period 2017-2020 is summarized (see Table S1) and approaches to disease management are discussed (Table 1). ...
View in full-text
Context 2
... Using symptomology, morphological criteria and molecular approaches for pathogen identification, these descriptions of cannabis and hemp diseases are the most comprehensive currently available. In the present review, further characterization of newly emerging pathogens of cannabis and hemp reported over the period 2017-2020 is summarized (see Table S1) and approaches to disease management are discussed (Table 1). ...
View in full-text
Context 3
... the type of plant disease symptoms can provide a preliminary assessment of the causal agent involved. 26 In cannabis and hemp, descriptions of recently emerging diseases have begun to appear in the published literature (Table S1). The pathogens can be grouped by the tissues they infect: root and crowninfecting, foliar and stem-infecting, inflorescence-infecting, and postharvest pathogens (Fig. 2). ...
View in full-text
Context 4
... of the pathogens are fungi and oomycetes, followed by viruses or viroids. Bacterial pathogens are less commonly reported ( Table S1). The most destructive root pathogens are Fusarium and Pythium species (Fig. 3(A)-(F)), particularly when infections occur during the rooting phase or vegetative growth. ...
View in full-text
Context 5
... not yet been diagnosed on indoor cultivated cannabis plants (Table S1). The stem-infecting pathogens are varied and diverse in the symptoms they produce and occur both indoors and outdoors (Table S1), although outdoor grown plants have a higher prevalence of these diseases as they are difficult to manage. ...
View in full-text
Context 6
... not yet been diagnosed on indoor cultivated cannabis plants (Table S1). The stem-infecting pathogens are varied and diverse in the symptoms they produce and occur both indoors and outdoors (Table S1), although outdoor grown plants have a higher prevalence of these diseases as they are difficult to manage. Disease control measures for these pathogens rely on prevention of initial infection (exclusion) and reducing subsequent spread of secondary inoculum (spores). ...
View in full-text
Context 7
... infections can also lead to significant additional postharvest losses. The most damaging fungi are Botryis and Fusarium species (Table S1), as well as a number of other fungi that colonize foliar and flower tissues, including Penicillium and Golovinomyces species. These fungi produce large numbers of spores to ensure spread (Fig. S1). ...
View in full-text
Context 8
... fungi produce large numbers of spores to ensure spread (Fig. S1). Recently observed fungi that can cause bud rot include species of Diaporthe and Sclerotinia (Table S1). Hop latent viroid can also cause 'dudding' of the buds, which are essentially destroyed as a result of infection. ...
View in full-text
Context 9
... cannabis and hemp pathogen identification, the most widely used method is the polymerase chain reaction (PCR) of the ribosomal DNA region that includes the internal transcribed spacer (ITS) and intergeneric spacer regions (IGS) [34][35][36][37] (Fig. S2). PCR was used to identify most of the species listed in Table S1. In addition, the elongation factor 1 (EF-1) region was used to discriminate among Fusarium species. ...
View in full-text
Context 10
... For the powdery mildew pathogen Golovinomyces, however, the ITS region was insufficient to distinguish among species. 41 Additional molecular markers are needed to conclusively confirm or amalgamate the three species currently reported to cause powdery mildew on cannabis and hemp (Table S1). Following identification, demonstration of pathogenicity on cannabis and hemp plants is an essential requirement -reports of presence or recovery of fungi from these tissues is insufficient, since many saprophytes are found on cannabis and hemp plants. ...
View in full-text
Context 11
... or cannabina for pathogens recovered from cannabis or hemp plants, is discouraged until host range studies and molecular confirmations support these designations. The host of origin is insufficient to imply specific host preference since most of the pathogens affecting cannabis and hemp are known to have wide host ranges (Table S1). ...
View in full-text
Context 12
... To date, recent reports of wileyonlinelibrary.com/journal/ps bacterial pathogens affecting cannabis or hemp plants and the symptoms they cause are surprisingly few (Table S1); most appear to be saprophytes and incidental/secondary contaminants. 7,9,35,44,45 Whether or not this is due to innate resistance in the plant or because environmental conditions have not supported bacterial infection or spread remains to be determined. ...
View in full-text
Context 13
... observations indicate that cannabis inflorescences are highly susceptible to infection by fungi that may originate from neighboring fields with diseased plants. Similarly, hemp leaves and stems are susceptible to many leaf-spotting fungi as well as a range of canker and die-back pathogens (Table S1), some of which can devastate the crop. The spread of pathogens from hop plants to cannabis plants has also been reported for powdery mildew and Hop latent viroid. ...
View in full-text
Context 14
... planting hemp or cannabis in fields previously cropped to other hosts with residual inoculum can result in disease development. For example, fields under pasture followed by planting to hemp resulted in crown rot due to Fusarium avenaceum, F. graminearum and F. tricinctum (Table S1). Lettuce fields with Sclerotinia infection resulted in white mold development in a subsequent crop of hemp. ...
View in full-text
Context 15
... fields with Sclerotinia infection resulted in white mold development in a subsequent crop of hemp. 68 Lastly, inoculum of Sclerotium rolfsii in fields planted to peanuts or vegetable crops can cause southern blight on hemp the following season in many US southern states, especially under excessively hot conditions (Table S1). Knowledge of previous cropping history prior to selection of a site for cannabis or hemp cultivation should indicate the potential for disease occurrence. ...
View in full-text
Context 16
... of disease development requires disruption of the disease cycle, beginning with prevention of inoculum introduction, preventing infection or symptom development, reducing secondary inoculum production and spread, and reducing survival of the pathogen. These approaches are summarized in Table 1 and will be discussed with reference to specific pathogens that can be mitigated to prevent infection of cannabis or hemp plants. ...
View in full-text
Context 17
... greatest challenge remains in dealing with pathogens that infect cannabis inflorescences (Table S1) since economic losses from tissue destruction and a build-up of colony-forming units pre and post harvest can be as high as 20%. Research on this topic is limited by government regulations in Canada which restrict the cultivation of flowering cannabis plants to producers with approved licenses; researchers have limited capacity to grow such plants. ...
View in full-text

In Vitro Study of Biocontrol Potential of Rhizospheric Pseudomonas aeruginosa against Pathogenic Fungi of Saffron (Crocus sativus L.)​

Institute of Chinese Materia Medica, Shanghai University of Traditional Chinese Medicine, Shanghai 201203, China
*
Author to whom correspondence should be addressed.
Pathogens 2021, 10(11), 1423; https://doi.org/10.3390/pathogens10111423
Received: 10 August 2021 / Revised: 30 October 2021 / Accepted: 31 October 2021 / Published: 2 November 2021
(This article belongs to the Section Fungal Pathogens)
Download
Browse Figures

Versions Notes

Abstract​

Plant rhizosphere soil contains a large number of plant-growth promoting rhizobacteria, which can not only resist the invasion of pathogenic microorganisms and protect plants from damage, but also promote the growth and development of plants. In this study, Pseudomonas aeruginosa strain YY322, isolated and screened from the rhizosphere soil of saffron (Crocus sativus L.), was found through a plate confrontation experiment to show highly effectual and obvious antagonistic activity against the pathogens of saffron, including Fusarium oxysporum, Fusarium solani, Penicillium citreosulfuratum, Penicillium citrinum and Stromatinia gladioli. In addition, the volatile organic compounds of strain YY322 had great antagonistic activity against these pathogens. Observation under a scanning electron microscope and transmission electron microscope reflected that strain YY322 had a significant effect on the hyphae and conidia of F. oxysporum and F. solani. Through the detection of degrading enzymes, it was found that P. aeruginosa can secrete protease and glucanase. The plant growth promoting performance was evaluated, finding that strain YY322 had the functions of dissolving phosphorus, fixing nitrogen, producing siderophore and producing NH3. In addition, whole genome sequencing analysis indicated that the YY322 genome is comprised of a 6,382,345-bp circular chromosome, containing 5809 protein-coding genes and 151 RNA genes. The P. aeruginosa YY322 genome encodes genes related to phenazine (phzABDEFGIMRS), hydrogen cyanide(HCN) (hcnABC), surfactin (srfAA), salicylate (pchA), biofilm formation (flgBCDEFGHIJKL, motAB, efp, hfq), and colonization (minCDE, yjbB, lysC). These results collectively indicated the role of P. aeruginosa YY322 in plant growth enhancement and biocontrol mechanisms. All in all, this study provides a theoretical basis for P. aeruginosa as the PGPR of saffron, paving the way for the subsequent development and utilization of microbial fertilizer.
Keywords:
Pseudomonas aeruginosa; saffron; rhizosphere; antagonistic activity; biological control; PGPR

1. Introduction​

Saffron (Crocus sativus L.) is a well-known dye and spice, widely cultivated in India, Iran, China, Egypt, France, Greece, Israel, Italy, Spain, Turkey and other countries [1]. Saffron is an essential medicinal plant in China, purported to have the ability of promoting blood circulation, removing blood stasis, cooling blood, resolving toxins, relieving depression, and calming nerves. However, because of its low output and high values, saffron is costly and is referred to as “red gold” [2,3]. At present, saffron is widely cultivated on Chongming Island in Shanghai. However, many corms have rotted due to long-term continuous cropping and other factors, resulting in a decline in saffron yield and quality. Although using chemical fertilizers and pesticides reduces the occurrence and development of diseases, they inevitably contribute to environmental pollution and safety issues with medicinal materials.
The term rhizosphere, coined by German scientist Lorenz Hiltner in 1904, refers to a micro-domain environment influenced by plant root activities and differs from soil in physical, chemical, and biological properties. It is considered as the second genome of plants and is a hotspot for plant–soil–microbe interactions [4,5,6]. Due to unique environment of rhizosphere soil, numerous microorganisms gather in this area, where they play a vital role in the life history of plants [7,8]. Numerous plant growth-promoting rhizobacteria (PGPR) capable of strengthening plant nutrient absorption and immunological function have been isolated from rhizosphere soil, and a number of these have been converted into microbial fertilizers with favorable results [9,10,11].
Currently, the most extensively investigated PGPR includes Pseudomonas, Bacillus, Agrobacterium, Eriwinia, Flavobacterium, Pasteuria, Serratia, Enterobacter, etc. Pseudomonas is a type of obligate aerobic gram-negative rod-shaped bacteria widely distributed in nature and has garnered widespread interest due to its intimate association with human activities. Pseudomonas grows rapidly and can adapt to the natural environment. Aside from this, it can effectively antagonize the occurrence and development of plant pathogens by secreting a series of metabolites and can help plants to absorb and utilize nutrients [12]. In addition, it is easily selected and bred through genetic mutation or modification, then inoculated into plants’ rhizospheres using bacterialization of seeds to perform its function [13]. Currently, the main types of Pseudomonas classified as PGPR are P. putida [14], P. aeruginosa [15], P. fluorescens [16,17], P. chlororaphis [18], and P. Parafulva [19].
Herein, 85 strains of bacteria were isolated from the rhizosphere soil of saffron. Among them, strain YY322 exhibited a good antagonistic effect against the five pathogens of saffron. Combining physiological and biochemical tests and molecular biology identification, it was identified as P. aeruginosa. The strain YY322 could destroy the hyphae and conidia of pathogens, as observed under a scanning electron microscope (SEM) and transmission electron microscope (TEM). Moreover, hydrolytic enzymatic (chitinase, protease, cellulase, and glucanase) activity and plant growth-promoting (PGP) potential (phosphate solubilization, potassium dissolution, atmospheric nitrogen fixation, siderophore production, indole-3-acetic acid (IAA) and ammonia (NH3) production, 1-aminocyclopropane-1-carboxylate (ACC) deaminase activity) of the strain were tested, discovering that the strain possessed good performance. Finally, we performed whole-genome sequencing of the bacterium to further explore its antagonistic and growth-promoting mechanisms. This article provides evidence to prove that P. aeruginosa can be used as PGPR in the production of saffron, laying a theoretical foundation for the development and utilization of microbial fertilizer.

2. Results​

2.1. Isolation and Screening of Antagonistic Bacteria​

A total of 85 strains of bacteria were isolated from saffron rhizosphere soil. After several screenings, it was found that the antagonistic effect of strain YY322 was the best, with inhibitory rates for Fusarium oxysporum, Fusarium solani, Penicillium citreosulfuratum, Penicillium citrinum, and Stromatinia gladioli of 73.17%, 62.20%, 23.16%, 49.17%, and 79.76%, respectively (Figure 1). During the process of plate confrontation, not only was the growth of pathogenic fungi significantly inhibited, but the morphological characteristics of strain YY322 were also changed compared with the normal state, indicating that it may have produced some unknown antifungal substances. The strain is currently stored in the China Center for Type Culture Collection under the number CCTCC M 2021208 YY322.

Figure 1. Antagonistic effect of strain YY322 on five pathogens cultivated on PDA medium for one to two weeks. (A) F. oxysporum, (B) F. solani, (C) P. citreosulfuratum, (D) P. citrinum, (E) S. gladioli (1) control, (2) treatment.

2.2. Strain YY322: Fusarium spp. Interaction Study by SEM and TEM​

SEM indicated that the hyphae and conidia of Fusarium spp. under the action of strain YY322 were changed significantly compared with the control group. Hyphae were round and full in the control group, with complete morphology and structure, smooth surface, and uniform thickness, whereas conidia were normal spindle-shaped, moderate in size, and demonstrating a good growth state. F. oxysporum affected by strain YY322 had numerous broken hyphae, with an uneven surface, severe segmentation, abnormal apex development, and malformed conidia. F. solani affected by strain YY322 caused seriously damaged hyphae, with surface shrinkage and leakage of contents. In addition, it possessed only a few large conidia and no small conidia (Figure 2). Under TEM, it could be observed that compared with the control group, the internal structure of Fusarium spp. under the action of strain YY322 was severely damaged. In the control group, the evenly distributed hyphae were regularly round and exhibited clear boundaries. The cell wall and membrane remained intact and compact, and the cytoplasm was well-distributed, maintaining a normal growth state. The cell walls of the hyphae of F. oxysporum affected by strain YY322 ruptured, resulting in leakage of the contents. The number of vacuoles decreased, but their volume increased. In addition, the cytoplasm was turbid and chaotic. F. solani, affected by strain YY322, exhibited numerous immature conidia gathered together, and the shape of the hyphae section was abnormal. The cell wall was thickened, the cytoplasm was mixed and black, and the vacuole almost disappeared (Figure 3).

Figure 2. Observation of the morphological changes of hyphae and conidia under SEM. (A) F. oxysporum; (B) F. solani (1) control: the hyphae and conidia showed normal growth, (2) treatment: the surface of the hyphae is shrunk and broken, and its contents leak out; the conidia develop abnormally and decrease in number (arrowheads).

Figure 3. Observation of the changes in the internal structure of hyphae under TEM. (A) F. oxysporum; (B) F. solani (1,2) control: the evenly distributed hyphae have clear boundaries, the cell wall and cell membrane structure are complete, and the cytoplasm is in a normal state, (3,4) treatment: the section of hyphae is irregular, the cell wall is thickened, the cytoplasm is turbid and chaotic, and the vacuole develops abnormally (arrowheads).

2.3. Inhibitory Effect of VOCs of Strain YY322 on Mycelial Growth​

Since we discovered that the strain YY322 could produce a special smell when cultured, we tested the antifungal effect of its VOCs, and the results showed it demonstrated a pronounced inhibitory effect on F. oxysporum and F. solani, with inhibition rates of 90.24% and 89.02%, respectively. Intriguingly, a few hyphae of both pathogens could still be observed spreading to the entire plate (Figure 4).

Figure 4. Antifungal activity of VOCs: (A) F. oxysporum; (B) F. solani (1) control, (2) treatment.

H. bacteriaphora, S. carpocapsae, and S. feltiae​

Production and Storage Technology​

Entomopathogenic nematodes are mass-produced for use as biocontrols using in vivo or in vitro methods (Shapiro-Ilan and Gaugler 2002). In vivo production (culture in live insect hosts) requires a low level of technology, has low startup costs, and resulting nematode quality is generally high, yet cost efficiency is low. The approach can be considered ideal for small markets. In vivo production may be improved through innovations in mechanization and streamlining. A novel alternative approach to in vivo methodology is production and application of nematodes in infected host cadavers; the cadavers (with nematodes developing inside) are distributed directly to the target site and pest suppression is subsequently achieved by the infective juveniles that emerge. In vitro solid culture, i.e., growing the nematodes on crumbled polyurethane foam, offers an intermediate level of technology and costs. In vitro liquid culture is the most cost- efficient production method but requires the largest startup capital. Liquid culture may be improved through progress in media development, nematode recovery, and bioreactor design. A variety of formulations have been developed to facilitate nematode storage and application including activated charcoal, alginate and polyacrylamide gels, baits, clay, paste, peat, polyurethane sponge, vermiculite, and water-dispersible granules. Depending on the formulation and nematode species, successful storage under refrigeration ranges from one to seven months. Optimum storage temperature for formulated nematodes varies according to species; generally, steinernematids tend to store best at 4-8 °C whereas heterorhabditids persist better at 10-15 °C.\

Entomopathogenic Nematode Production and Application Technology​

David I. Shapiro-Ilan,1 Richou Han,2 and Claudia Dolinksi3
Author information Article notes Copyright and License information Disclaimer

Go to:

Abstract​

Production and application technology is critical for the success of entomopathogenic nematodes (EPNs) in biological control. Production approaches include in vivo, and in vitro methods (solid or liquid fermentation). For laboratory use and small scale field experiments, in vivo production of EPNs appears to be the appropriate method. In vivo production is also appropriate for niche markets and small growers where a lack of capital, scientific expertise or infrastructure cannot justify large investments into in vitro culture technology. In vitro technology is used when large scale production is needed at reasonable quality and cost. Infective juveniles of entomopathogenic nematodes are usually applied using various spray equipment and standard irrigation systems. Enhanced efficacy in EPN applications can be facilitated through improved delivery mechanisms (e.g., cadaver application) or optimization of spray equipment. Substantial progress has been made in recent years in developing EPN formulations, particularly for above ground applications, e.g., mixing EPNs with surfactants or polymers or with sprayable gels. Bait formulations and insect host cadavers can enhance EPN persistence and reduce the quantity of nematodes required per unit area. This review provides a summary and analysis of factors that affect production and application of EPNs and offers insights for their future in biological insect suppression.
Keywords: application technology, entomopathogenic nematode, Heterorhabditis, production, Steinernema

​

Entomopathogenic nematodes (EPNs) of the families Heterorhabditidae and Steinernematidae are obligate parasites of insects and are used as biological control agents of economically important insect pests. The two major genera are Heterorhabditis Poinar, 1976, and Steinernema Travassos, 1927, with 85 species described to date. These nematodes possess a symbiotic association with pathogenic bacteria from the Xenorhabdus and Photorhabdus genera, associated with Steinernema and Heterorhabditis respectively (Poinar, 1990). Infective juveniles (IJs), considered the only free-living stage of EPNs, enter the host insect through its natural apertures (oral cavity, anus and spiracles) or in some cases through the cuticle (Dowds and Peters, 2002). After penetrating the insect’s hemocoel, IJs release their symbiotic bacteria, which are the primary agents responsible for host death and also provide the nematodes with nutrition and defense against secondary invaders (Poinar, 1990). The nematodes complete their development and live for two or three generations inside their host. When food is depleted, IJs exit from host cadaver searching for new hosts (Grewal and Georgis, 1999).
Entomopathogenic nematodes are currently produced by different methods either in vivo or in vitro (solid and liquid culture) (Friedman, 1990). Each approach has its advantages and disadvantages relative to production cost, technical know-how required, economy of scale, and product quality, and each approach has the potential to be improved. Following production, a variety of formulation and application alternatives are also available (Grewal, 2002). The objectives of this review are to provide a summary and analysis of factors that affect production and application of EPNs, and provide insight toward enhancing approaches to achieve greater success in biological control.
Go to:

Production Methods​

In vivo culture method The general approach to in vivo culture is a two dimensional system that relies on production in trays and shelves (Friedman, 1990; Shapiro and Gaugler, 2002; Ehlers and Shapiro-Ilan, 2005). In vivo production methods for culturing EPNs have been reported by various authors (Dutky et al., 1964; Poinar, 1979; Woodring and Kaya, 1988; Lindegren et al., 1993; Flanders et al., 1996; Kaya and Stock, 1997; Shapiro-Ilan et al., 2002a). The systems described in all of these references (with some variation) entail a system that is based on the White trap (White, 1927), which takes advantage of the progeny IJ’s natural migration away from the host-cadaver upon emergence. The approach consists of inoculation, harvest, concentration, and (if needed) decontamination. Insect hosts are inoculated on a dish or tray lined with absorbent paper or another substrate conducive to nematode infection such as soil or plaster of Paris. After approximately 2-5 days, infected insects are transferred to the White traps; if infections are allowed to progress too long before transfer the chance of the cadaver rupturing and harm to reproductive nematode stages is increased (Shapiro-Ilan et al., 2001). White traps consist of a dish or tray on which the cadavers rest surrounded by water, which is contained by a large arena. As IJs emerge they migrate to the surrounding water trap where they are harvested. The scale of the White trap in size and number can be expanded to commercial levels.
Factors affecting yield for in vivo culture In vivo production yields depend on nematode dosage and host density (Zervos et al., 1991; Boff et al., 2000; Shapiro-Ilan et al., 2002a). A dosage that is too low results in low host mortality and a dosage that is too high may result in failed infections due to competition with secondary invaders (Woodring and Kaya, 1988). Thus, intermediate dosages can be used to maximize yield (Boff et al., 2000). For example, for infecting the greater wax moth Galleria mellonella, rates of approximately 25 to 200 IJs per insect are sufficient (depending on nematode species and method of inoculation) whereas higher rates are generally needed for the yellow mealworm, Tenebrio molitor (e.g., 100 to 600 IJs per insect). Crowding of hosts can lead to oxygen deprivation or buildup of ammonia (Shapiro-Ilan et al., 2000a; 2002a). In general, the number of hosts exhibiting patent signs of nematode infection increases with nematode concentration and decreases with host density per unit area (Shapiro-Ilan et al., 2002a). Thus, optimization of host density and inoculation rate for maximum yield is recommended (Shapiro-Ilan et al., 2002a).
In vivo production yields vary greatly among insect hosts and nematode species. Due to high susceptibility to most nematodes, wide availability, ease in rearing, and the ability to produce high yields, the most common insect host used for laboratory and commercial EPN culture is G. mellonella (Woodring and Kaya, 1988). Significant research (and commercial application) has also been achieved for production of EPNs in T. molitor (Blinova and Ivanova, 1987; Shapiro-Ilan et al., 2002a). Other hosts in which in vivo production has been studied include the navel orangeworm (Ameylois transitella), tobacco budworm (Heliothis virescens), cabbage looper (Trichoplusia ni), pink bollworm (Pectinophora gossypiella), beet armyworm (Spodoptera exigua), corn earworm (Helicoverpa zea), gypsy moth (Lymantria dispar), house cricket (Acheta domesticus) and various beetles (Coleoptera) (Lindegren et al., 1979; Blinova and Ivanova, 1987; Cabanillas and Raulston, 1994; Grewal et al., 1999; Elawad et al., 2001).
Generally nematode yield is proportional to host size (Blinova and Ivanova, 1987; Flanders et al.1996). Yet yield per mg insect (within host species) and susceptibility to infection is often inversely proportional to host size or age (Dutky et al. 1964; Blinova and Ivanova 1987; Shapiro et al. 1999a; Dolinski et al., 2007; Dias et al., 2008). Yield is also generally inversely proportional to nematode size (Grewal et al., 1994; Hominick et al., 1997; Shapiro and Gaugler, 2002). The choice of host species and nematode for in vivo production should ultimately rest on nematode yield per cost of insect and the suitability of the nematode for the pest target (Blinova and Ivanova, 1987; Shapiro-Ilan et al., 2002a).
Environmental factors including optimum temperature, and maintaining adequate aeration, and moisture can affect yield (Burman and Pye, 1980; Woodring and Kaya, 1988; Friedman, 1990; Grewal et al., 1994; Shapiro-Ilan et al., 2002a; Dolinski et al., 2007). Infection efficiency and yield can also be affected by inoculation method. In vivo can be accomplished by pipetting or spraying nematodes onto a substrate, immersion of insects in a nematode suspension, or (in some cases) applying the nematodes to the insect’s food. Immersion of hosts is generally more time efficient but requires more nematodes than other procedures (Shapiro-Ilan et al., 2002a). Blinova and Ivanova (1987) reported that infection of T. molitor by S. carpocapsae was increased using the feeding method relative to other methods. Feeding, however, requires an additional step of removing infected cadavers from food remnants (which may cause contamination); thus, before a method is decided upon, inoculation procedures must be included in a cost efficiency analysis.
In vitro solid culture method: In vitro culturing of EPNs is based on introducing nematodes to a pure culture of their symbiont in a nutritive medium. In earlier work, to create monoxenic cultures surface sterilized nematodes were added to a lawn of the bacterial symbionts (Akhurst, 1980; Wouts, 1981). However, Lunau et al. (1993) suggested that surface sterilization of IJs is insufficient to establish monxenicity because contaminating bacteria survive beneath the nematode’s cuticle. Thus, an improved method has been developed where axenic nematode eggs are placed on a pure culture of the symbiont (Lunau et al. 1993).
Solid culture was first accomplished in two dimensional arenas e.g, petri dishes, using various media (Hara et al., 1981, Wouts, 1981). Subsequently, in vitro solid culture advanced considerably with the invention of a three-dimensional rearing system involving nematode culture on crumbled polyether polyurethane foam (Bedding, 1981). A liquid medium is mixed with foam, autoclaved, and then inoculated with bacteria followed by the nematodes. Nematodes are then harvested within 2-5 weeks (Bedding, 1981; Bedding, 1984) by placing the foam onto sieves immersed in water. Media for this approach was initially animal product based (e.g., pork kidney or chicken offal) but was later improved and may include various ingredients including peptone, yeast extract, eggs, soy flour, and lard (Han et al., 1992; 1993). The approach developed by Bedding (1981) was expanded to autoclavable bags with filtered air being pumped in (Bedding, 1984). Large scale production was further advanced through several measures including using bags with gas permeable Tyvac ® strips for ventilation, automated mixing and autoclaving, simultaneous inoculation of nematodes and bacteria, sterile room technology, and automated harvest through centrifugal sifters (Gaugler and Han, 2002).
Factors affecting yield for in vitro solid culture: Nematode inoculum rate (IJs per unit of media) can affect yield in some nematode strains but not others (Han et al. 1992, 1993; Wang and Bedding, 1998). Culture time is inversely related to temperature and thus should be optimized for maximum yield on a species or strain basis (Dunphy and Webster, 1989; Han et al. 1992, 1993). Increasing inoculum size can increase nematode growth and decrease culture time (Han et al., 1992). Longer culture times can provide higher yields yet nematode mortality may also increase with time (Han et al., 1992, 1993) and culture time must be weighed against the cost of space and diminishing returns.
Media composition can have considerable effects on yield in solid culture. Increasing the lipid quantity and quality leads to increases in nematode yield (Dunphy and Webster, 1989; Han et al., 1992). Lipid components reflecting the nematode’s natural host composition are most suitable (Abu Hatab and Gaugler, 2001; Abu Hatab et al., 1998). Other media ingredients that may have an effect on nematode yield include proteins and salts (Dunphy and Webster, 1989).
In vitro liquid culture method: The development of monoxenic liquid culture of EPNs faces the opposing challenges of supplying enough oxygen while preventing excessive shearing of nematodes (Pace et al., 1986; Buecher and Popiel, 1989; Friedman et al., 1989, 1990). The issue was initially addressed in various ways such as relying on bubbling, e.g., with a downward sparger, coupled with limited agitation (Pace et al., 1986), or using an airlift fermenter coupled with a variable agitation regime (Friedman et al., 1989). Various innovations in mixing and aeration have been subsequently introduced including internal (Strauch and Ehlers, 2000) and external (Neves et al., 2001) bioreactors; internal loop vessels have baffles placed inside the single vessel, which creates the channels required for the circulation, whereas in external loop vessels circulation takes place through separate conduits.
Generally, in liquid culture, symbiotic bacteria are first introduced followed by the nematodes (Buecher and Popiel, 1989; Surrey and Davies, 1996; Strauch and Ehlers, 2000). Various ingredients for liquid culture media have been reported including soy flour, yeast extract, canola oil, corn oil, thistle oil, egg yolk, casein peptone, milk powder, liver extract and cholesterol (Surrey and Davies, 1996; Ehlers et al., 2000; Yoo et al., 2000). Culture times vary depending on media and species, and may be as long as three weeks (Surrey and Davies, 1996; Charvarria-Hernandez and de la Torre, 2001) though many species can reach maximum IJ production in two weeks or less (Friedman 1990; Ehlers et al., 2000; Neves et al., 2001; Strauch and Ehlers, 2000; Yoo et al., 2000). Once the culture is completed, nematodes can be harvested from media via centrifugation (Surrey and Davies, 1996).
Factors affecting yield for in vitro liquid culture: Both steinernematids and heterorhabditids share the requirements of adequate aeration (without shearing). Otherwise, the strategies for maximizing yield of the two genera in liquid culture differ due to their life cycles and reproductive biology. Steinernematids (except one species) occur only as males and females and are capable of mating in liquid culture (Strauch et al., 1994); thus, maximization of mating is paramount and can be achieved through bioreactor design and regulation of aeration (Neves et al., 2001). Maximization of mating, however, is not applicable for heterorhabditid production in liquid culture because the first generation is exclusively hermaphrodites and, although subsequent generations contain amphimictic forms, they cannot mate in liquid culture (Strauch et al., 1994). Thus, maximizing heterorhabditid yields in liquid culture depends on the degree of recovery (the developmental step when IJs molt to initiate completion of their life cycle). While levels of heterorhabditids recovery in vivo tend to be 100% (Strauch and Ehlers, 1998), recovery in liquid culture may range from 0-85% (Ehlers et al., 2000; Jessen et al., 2000; Strauch and Ehlers, 1998, 2000; Yoo et al. 2000). Recovery can be affected by nutritional factors, aeration, CO2, lipid content, and temperature (Ehlers et al., 2000; Jessen et al., 2000; Strauch and Ehlers, 1998, 2000; Yoo et al., 2000).
Yield from liquid culture may also be affected by other factors including media, nematode inoculum, and nematode species (Han, 1996; Ehlers et al., 2000). The central component of the liquid culture media is lipid source and quantity (Abu Hatab et al., 1998; Yoo et al., 2000). Other nutrients that have been reported to affect yield positively include the content of glucose (Jeffke et al., 2000) and yeast extract content (Chavarria-Hernandez and de la Torre, 2001). Generally nematode yield is inversely proportional to the size of the species (Ehlers et al., 2000). Maximum average yields reported include 300,000 and 320,000 IJs per ml for H. bacteriophora and S. carpocapsae, respectively (Han, 1996), 138,000 per ml for H. megidis (Strauch and Ehlers, 2000), 71,470 IJs per ml for S. feltiae (Chavarria-Hernandez and de la Torre, 2001), and 450,000 IJs per ml for H. indica (Ehlers et al., 2000).
Analysis of production methods and potential for improvement (the road to the future): A comparison of production methods is summarized in Table 1. In vivo EPN production offers several advantages relative to in vitro culture including requiring the least capital outlay and the least amount of technical expertise for start-up (Shapiro-Ilan and Gaugler, 2002); additionally, the quality of in vivo produced nematodes tends to be equal to or greater than EPNs produced with other approaches (Gaugler and Georgis, 1991; Yang et al., 1997). Furthermore, adapting production techniques to new or additional nematode species or strains is straightforward with in vivo production, whereas in vitro methods can require substantial adjustments in media or processing parameters. The primary challenge for in vivo EPN culture relative to in vitro methods is the costs of insects, which tends to make in vivo culture the least cost efficient approach.

Table 1​

A comparison of entomopathogenic nematode production approaches.
[IMG alt="An external file that holds a picture, illustration, etc.
Object name is 206tbl1.jpg"]https://www.ncbi.nlm.nih.gov/pmc/articles/PMC3578468/bin/206tbl1.jpg[/IMG]
The economics of in vivo production can be improved substantially by producing the insect hosts “in-house” and mechanizing the process thereby reducing labor. Several approaches to mechanization of nematode inoculation (Shapiro-Ilan et al., 2009) and harvest (Gaugler et al., 2002; Shapiro-Ilan et al., 2011a) have been introduced. Additionally, optimization of insect diets can lead to improved efficiency in insect host production and quality of nematodes (via tri-trophic interactions) (Morales et al., 2011a; Shapiro-Ilan et al., 2008, 2011c). Insect host production can also be mechanized, e.g., through automated sifting of insects for separation (Morales et al., 2011b). Despite shortcomings in economy of scale, in vivo production has managed to sustain itself as a cottage industry throughout the commercial development of in vitro enterprises (Shapiro-Ilan and Gaugler, 2002), and it is likely to continue and perhaps expand based upon advancements in mechanization described above.
In terms of capital outlay and economy of scale, the merits of in vitro solid culture are generally considered to be intermediate between in vivo and liquid culture. Similar to in vivo production, efficiency can be increased through mechanization (labor reduction) and media enhancement (Gaugler and Han, 2002; Shapiro and Gaugler, 2002). Several studies indicate quality to be similar to nematodes produced by in vivo methods (Abu Hatab et al., 1998; Abu Hatab and Gaugler, 1999; Gaugler and Georgis, 1991). Contrarily, Yang et al. (1997) reported reduced fitness in S. carpocapsae produced in solid culture compared with in vivo culture. Recently, advances have been made in expanding in vitro solid production of EPNs. For example, in China, a pilot factory was established for solid production of several EPN species based on the lower labor cost, improved media and mechanization process. The factors influencing the production efficiency were explored, including medium development, optimization of the culture parameters, recovery of the IJ inocula, formation of the IJs, extraction and harvest of IJs (Han et al., 1995, 1997). A company called Century Horse Development Ltd, under the guidance of Guangdong Entomological Institute, is currently in commercial production; products from the solid culture system are provided for field trials in China and for internal and international markets.
In vitro liquid culture is considered to be the most cost efficient process for producing entomopathogenic nematodes and thus accounts for the bulk of the world market in EPNs. Liquid culture of entomopathogenic nematodes has been accomplished in bioreactors of up to 80,000 liters (Georgis et al., 1995). Although liquid culture offers increased cost efficiency relative to other production methods, it also demands greater capital investment and a higher level of technical expertise.
Several reports indicated reduced quality or efficacy in in vitro liquid produced EPNs relative to those produced in solid culture or in vivo (Gaugler and Georgis, 1991; Cottrell et al., 2011) whereas other comparisons did not detect any differences (Georgis and Gaugler, 1991; Shapiro and McCoy, 2000a). Thus, it is clear that high quality EPNs can be produced using in vitro liquid culture, but factors such as media composition and environmental conditions in the bioreactor or downstream processing can reduce quality. Some recent advancements in liquid culture technology serve to increase quality and efficiency of production e.g., optimizing media and bioprocess kinetics through modeling (Chavarria-Hernandez et al., 2006, 2010), as well as improvements in inoculum and bacterial cell density (Hirao and Ehlers, 2010), timing of inoculation (Johnigk et al., 2004) and downstream processing (Young et al., 2002). Future research and development in liquid culture (focusing on media optimization and bioreactor design) are expected to lead to additional benefits such as higher yields and reduced costs.
A concern for both in vivo and in vitro production is strain deterioration. Repeated culturing of nematodes can result in reduction of beneficial traits such as virulence, environmental tolerance or reproductive capacity (Shapiro et al., 1996a; Wang and Grewal, 2002; Bai et al., 2005; Bilgrami et al., 2006); therefore, precautions against strain deterioration should be employed, e.g., minimization of serial passages, introduction of fresh genetic material, improved cryopreservation methods of stock cultures (Bai et al., 2004), or creation of homozygous inbred lines which are resistant to trait deterioration (Bai et al., 2005; Chaston et al., 2011).
Go to:

Application Technology And Factors That Affect Efficacy​

Standard application approaches: Entomopathogenic nematodes can be applied with nearly all agronomic or horticultural ground equipment including pressurized sprayers, mist blowers, and electrostatic sprayers or as aerial sprays (Georgis, 1990; Wright et al., 2005; Shapiro-Ilan et al., 2006a). The application equipment used depends on the cropping system, and in each case there are a variety of handling considerations including volume, agitation, nozzle type, pressure and recycling time, system environmental conditions, and spray distribution pattern (Grewal, 2002; Fife et al., 2003, 2005; Wright et al., 2005; Shapiro-Ilan et al., 2006a; Lara et al., 2008). It is important to ensure adequate agitation during application. For small plot applications, hand-held equipment (e.g., water cans) or back-pack sprayers may be appropriate. When nematodes are applied to larger plots, a suitable spraying apparatus such as a boom sprayer should be considered. Conceivably, applicators could also be using other methods such as through microjet irrigation systems, subsurface injection or baits (Wright et al., 2005; Lara et al., 2008). Various formulations for entomopathogenic nematodes may be used for applying EPNs in aqueous suspension including activated charcoal, alginate and polyacrylamide gels, clay, peat, polyurethane sponge, vermiculite, and water dispersible granules (WDG) (Georgis, 1990; Georgis et al., 1995).
Biotic factors affecting application success: A number of factors related to the nematode are critical for application success. Most importantly, the appropriate nematode must be matched with the particular target pest. Factors that must be considered in choosing the appropriate nematode include virulence, host finding, and environmental tolerance and in some cases persistence (Shapiro-Ilan et al., 2002b; Shapiro-Ilan et al., 2006b). Also of paramount importance, to be effective, EPNs usually must be applied to soil at minimum rates of 2.5 x 109 IJs/ha (=25/cm2) or higher (Georgis and Hague, 1991; Georgis et al., 1995; Shapiro-Ilan et al., 2002b). Depending on the target pests, a higher rate of application may be required (or in some rare cases lower rates may suffice) (Shapiro-Ilan et al., 2006a). Recycling potential should also be considered. Generally, as long as environmental conditions are conducive, nematode populations will remain high enough to provide effective pest control for 2 to 8 weeks after application (Kaya, 1990; Duncan and McCoy, 1996; Shapiro-Ilan et al., 2002b). Thus, seasonal re-application is often necessary. However, in some cases effective control has been reported over multiple seasons or years (Klein and Georgis, 1992; Parkman et al., 1994; Shields et al., 1999).
Biotic agents can have positive, negative, or neutral effects on EPN applications. Antagonists include nematode pathogens or predators such as phages, bacteria, protozoans, nematophagous fungi, predacious mites and nematodes, etc. (Kaya, 2002). Phoretic relationships have been indicated with other soil organisms such as mites, earthworms, and isopods (Epsky et al., 1988; Shapiro et al., 1995; Eng et al., 2005). Entomopathogenic nematodes have been reported to act synergistically with other entomopathogens such as Paenibacillus popilliae (Thurston et al., 1994), Bacillus thuringiensis (Koppenhöfer and Kaya, 1997), and Metarhizium anisopliae Sorokin (Ansari et al., 2004, 2006; Anbesse et al., 2008), yet other studies indicate antagonism, e.g., with Beauveria bassiana (Balsamo) Vuillemin (Brinkman and Gardner, 2000) or Isaria fumosorosea (Shapiro-Ilan et al., 2004). The relationship between nematodes and other entomopathogens (antagonism, additivity, synergism) can vary depending on the nematode species and relative timing or rate of application (Barbercheck and Kaya, 1990; Koppenhöfer and Kaya, 1997; Shapiro-Ilan et al., 2004).
Abiotic factors affecting application success: Successful application of EPNs depends on several critical factors including protection from ultraviolet radiation, adequate soil moisture/relative humidity, and temperature (Kaya, 1990; Shapiro-Ilan et al., 2006a). Indeed, EPN applications for aboveground pests have been severely limited due to environmental hindrances (e.g., UV radiation or desiccation) that reduce survival and efficacy (Begley, 1990; Grewal and Georgis, 1999; Arthurs et al., 2004; Shapiro-Ilan et al., 2006a), and thus, biocontrol success is most likely achieved when EPNs are applied to soil or cryptic habitats. Furthermore, because ultraviolet radiation is detrimental to nematodes (Gaugler and Boush, 1978), applications are best applied in the evening or early morning hours, or exposure to ultraviolet radiation avoided, through subsurface application (Cabanillas and Raulston, 1995). For soil applications, moisture for EPN survival and movement is required, but too much moisture may cause oxygen deprivation and restrict movement (Wallace, 1958; Kaya, 1990; Womersley, 1993; Koppenhöfer et al., 1995). Thus, irrigation is recommended for maintaining adequate moisture (Shetlar et al., 1988; Zimmerman and Cranshaw, 1991; Downing, 1994). Optimum moisture levels will vary by nematode species and soil type (Koppenhöfer et al., 1995). Optimum temperatures for infection and reproduction will also vary among nematode species and strains (Grewal et al., 1994). Some nematodes such as H. indica, S. glaseri, and S. riobrave are relatively heat tolerant whereas others, such as H. megidis, S. feltiae, and Heterorhabditis are more tolerant to cooler temperatures (Kung et al., 1991; Grewal et al., 1994; Berry et al., 1997; Shapiro and McCoy, 2000b).
Soil parameters can also be important for surface or below-ground applications. Soil texture affects nematode movement and survival (Kaya, 1990; Barbercheck, 1992). Generally, compared with lighter soils, soils with higher clay content restrict nematode movement and have potential for reduced aeration, which, in combination, can result in reduced nematode survival and efficacy (Georgis and Poinar, 1983; Molyneux and Bedding, 1984; Kung et al., 1990a). However, exceptions to this trend have been reported (Georgis and Gaugler, 1991; Shapiro et al., 2000b). Soil pH can affect natural EPN distributions (Kanga et al., 2012). A soil pH of 10 or higher is likely to be detrimental to EPN applications, whereas a range of 4-8, is not likely to have any significant effect on EPNs (Kung et al., 1990b).
Biocontrol success can also be impacted by fertilizers and chemical pesticides, which can have positive, neutral, or negative effects on entomopathogenic nematodes. In general, fertilizers that are applied at recommended rates have little impact on EPN efficacy (Shapiro et al., 1996b; Bednarek and Gaugler, 1997). However, fresh manure or high rates of chemical fertilizers (e.g., urea at 560 kg N per ha) can be detrimental to EPN persistence and efficacy (Shapiro et al., 1996b; Bednarek and Gaugler, 1997; Shapiro et al., 1999b). Some chemical pesticides are toxic to EPNs (e.g., abamectin, acephate, aldicarb, dodine, fenamiphos, methomyl, parathion, and Teflubenuron), whereas others tend to be compatible and in some cases may be synergistic when applied with EPNs (e.g., carbaryl, chlorpyrifos, dimethoate, endosulfan, fonofos, tefluthrin, imidicloprid) (Koppenhöfer and Kaya, 1998; Nishimatsu and Jackson, 1998; Alumai and Grewal, 2004; Koppenhöfer and Grewal, 2005; Koppenhöfer and Fuzy, 2008; Shapiro-Ilan et al., 2011b). Similar to interactions with other microbial agents, the relationship between chemical pesticides and EPNs varies based on the specific chemical and nematode species or strain, dosages, and timing of application (Benz, 1971; Koppenhöfer and Grewal, 2005); thus, combinations should be tested on a case by case basis.
Improved technology for EPN application (the road to the future): Enhanced efficacy in EPN applications can be facilitated through improved formulation. Substantial progress has been made in recent years in developing EPN formulations, particularly for aboveground applications, e.g., mixing EPNs with a surfactant and polymer (Schroer and Ehlers, 2005). Improved efficacy may also be achieved by relying on leaf flooding with the addition of surfactants to increase leaf coverage (Williams and Walters, 2000; Head et al., 2004). Additionally, S. carpocapsae applications for control of the lesser peachtree borer, Synanthedon pictipes, were greatly improved by a follow-up application of a sprayable gel (the gel is commonly used for protecting structures from fire) (Shapiro-Ilan et al., 2010a), and S. carpocapsae caused high levels of suppression (98% efficacy in a preventative treatment) in the red palm weevil, Rhynchophorus ferrugineus, when applied in a chitosan formulation (Llàcer et al., 2009). Furthermore, EPN applications to apple tree trunks for control of codling moth, Cydia pomonella (L.), were enhanced when the treatments included the sprayable fire-gel or wood flour foam as a protecting agent (Lacey et al., 2010).
Efficacy of EPN applications can also be enhanced through improved application equipment or approaches. Despite well-established procedures, equipment used for entomopathogen application can be improved further, e.g., optimizing spray systems (e.g., nozzles, pumps, spray distribution) for enhanced pathogen survival and dispersion (Shapiro-Ilan et al., 2006a; Brusselman et al., 2010). Bait formulations can enhance EPN persistence and reduce the quantity of microbial agents required per unit area (Grewal, 2002); though limited thus far, conceivably, baits can be developed further for wide applications. Another novel application approach that has gained attention is delivery of EPNs in their infected host cadavers (Jansson et al., 1993; Shapiro and Glazer, 1996; Del Valle et al., 2008). Advantages to the cadaver application approach relative to standard application in aqueous suspension have been reported such as increased nematode dispersal (Shapiro and Glazer, 1996), infectivity (Shapiro and Lewis, 1999), survival (Perez et al., 2003), and efficacy (Shapiro-Ilan et al., 2003), whereas other studies did not detect a benefit in the cadaver approach (Bruck et al., 2005). Application of cadavers may be facilitated through formulations that have been developed to protect cadavers from rupture and improve ease of handling (Shapiro-Ilan et al., 2001, 2010b; Del Valle et al., 2009), and development of mechanized equipment for field distribution (Zhu et al., 2011). The time period of six to ten days between infection and application on soil of Galleria mellonella cadavers resulted in higher emergence of IJs and was thus recommended when using the cadaver application approach (Del Valle et al., 2011). Recently, nematodes applied in host cadavers were effective and persistent when added to bags of potting media for subsequent distribution to target pest sites (Deol et al., 2011).
Finally, superior biocontrol applications with EPNs can also be achieved through strain improvement. Improved strains may include EPNs that possess enhanced levels of various beneficial traits such as environmental tolerance, virulence, reproductive capacity, etc. Methods to improve EPNs include strain or species discovery or genetic enhancement via selection, hybridization or molecular manipulation (Gaugler, 1987; Burnell, 2002; Grewal et al., 2005). Discovery of new strains and species that are superior to currently commercialized nematodes is a straightforward approach that can rapidly result in enhanced efficacy. The rate of EPN species discovery has been increasing considerably (Poinar, 1990; Adams and Nguyen, 2002; Lewis and Clarke, 2012). Since the time that the first EPN species was reported in 1923 (Steiner, 1923; Poinar, 1990) more than 85 EPN species have been described (Nguyen and Buss, 2011; Lewis and Clarke, 2012) and more than half the described species have been reported in the past 10 years. Additionally, the numerous new strains of existing species being discovered may offer enhanced virulence or other properties (e.g., Grewal et al., 2004; Stuart et al., 2004). The number of new strains and species discovered will likely continue to rise adding more potential for biocontrol applications. However, in order to leverage the advantages that strain/species offer, characterization of biocontrol potential in these new organisms must keep pace with the survey/discovery research. Currently, less than 20% of the species discovered since 2001 have been evaluated for biocontrol efficacy indicating there is substantial untapped potential.
If existing or newly discovered entomopathogen strains or species cannot achieve desired levels of biocontrol efficacy, strain enhancement might be achieved through genetic improvement approaches, which can include molecular or non-molecular methods. One non-molecular approach is selection for desired traits; directed selection has been demonstrated to be effective for improving various traits in EPNs such as host-finding (Gaugler et al., 1989, 1990) and nematicide resistance (Glazer et al., 1997). Hybridization (the transfer of beneficial traits from one strain to another) is another option for strain improvement that has shown promise (Shapiro et al., 1997; Shapiro-Ilan et al., 2005). Combination of the two non-molecular approaches (selection and hybridization) can also be effective for development of superior EPN strains (Mukaka et al., 2010). In addition to strain improvement approaches described above, molecular methods (e.g., transgenics) may offer potential for enhanced biocontrol (Gaugler et al., 1997), and we can expect that both molecular and non-molecular genetic approaches to strain improvement will be enhanced through the forthcoming advances in genomics (Bai and Grewal, 2007; Ciche, 2007; Bai et al., 2009).
Go to:

Conclusion​

Progress in developing large-scale production and application technology has led to the expanded use of EPNs. For laboratory use, small-scale field-testing, and niche markets, in vivo EPN production is the appropriate method requiring the least capital outlay and the least amount of technical expertise for start-up, but is hindered by the costs of labor and insect media. When it comes to commercial use for international markets, in vitro liquid culture is considered to be the most cost efficient process while in vitro solid culture is generally considered to be intermediate between in vivo and liquid culture. Although liquid culture offers increased cost efficiency relative to other production methods, it also demands greater capital investment and a higher level of technical expertise. Improvements in efficiency and scalability by producing the insect hosts “in-house” and mechanizing the process reduce labor, enabling in vivo production to play an expanded role in pest management programs. Similar to in vivo production, technical improvements will expand efficiency of in vitro solid production, but even so, neither approach may reach the scale-up potential of liquid culture technology.
EPNs are excellent biocontrol agents for insect pests. When an EPN is used against a pest insect, it is critical to match the right nematode species against the target pest. Biotic agents including nematode pathogens, predators and other soil organisms, as well as abiotic factors such as ultraviolet radiation, soil moisture/relative humidity, temperature, etc. can affect EPN application efficacy. Recently, improvement of nematode formulation, application equipment or approaches, and strain improvement have been made to enhance EPN application efficacy. Additional research toward lowering product costs, increasing product availability, enhancing ease-of-use, and improving efficacy and carryover effect will stimulate the extensive use of EPNs in biocontrol. With these advances EPNs will serve to reduce chemical insecticide inputs and contribute to the stabilization of crop yields and the environment.
Go to:

Literature Cited​

Abu Hatab M, Gaugler R. Lipids of in vivo and in vitro cultured Heterorhabditis bacteriophora. Biological Control. 1999;15:113–118. [Google Scholar]
Abu Hatab M, Gaugler R. Diet composition and lipids of in vitro-produced Heterorhabditis bacteriophora. Biological Control. 2001;20:1–7. [Google Scholar]
Abu Hatab M, Gaugler R, Ehlers R. Influence of culture method on Steinernema glaseri lipids. Journal of Parasitology. 1998;84:215–221. [PubMed] [Google Scholar]
Adams BJ, Nguyen KB. 2002. Taxonomy and systematics. Pp 1–34 in R. Gaugler, ed. Entomopathogenic Nematology. New York, NY: CABI. [Google Scholar]
Akhurst RJ. Morphological and functional dimorphism in Xenorhabdus spp., bacteria symbiotically associated with insect pathogenic nematodes Neoaplectana and Heterorhabditis. Journal of General Microbiology. 1980;121:303–309. [Google Scholar]
Alumai A, Grewal PS. Tank-mix compatibility of the entomopathogenic nematodes, Heterorhabditis bacteriophora and Steinernema carpocapsae, with selected chemical pesticides used in turfgrass. Biocontrol Science and Technology. 2004;14:725–730. [Google Scholar]
Anbesse SA, Adge BJ, Gebru WM. Laboratory screening for virulent entomopathogenic nematodes (Heterorhabditidis bacteriophora and Steinernema yirgalemense) and fungi (Metarhizium anisopliae and Beauveria bassiana) and assessment of possible synergistic effects of combined use against grubs of the barley chafer Coptognathus curtipennis. Nematology. 2008;10:701–709. [Google Scholar]
Ansari MA, Tirry L, Moens M. Interaction between Metarhizium anisopliae CLO 53 and entomopathogenic nematodes for the control of Hoplia philanthus. Biological Control. 2004;31:172–180. [Google Scholar]
Ansari MA, Shah FA, Moens TM. Field trials against Hoplia philanthus (Coleoptera: Scarabaeidae) with a combination of an entomopathogenic nematode and the fungus Metarhizium anisopliae CLO 53. Biological Control. 2006;39:453–459. [Google Scholar]
Arthurs S, Heinz KM, Prasifka JR. An analysis of using entomopathogenic nematodes against above-ground pests. Bulletin of Entomological Research. 2004;94:297–306. [PubMed] [Google Scholar]
Bai C, Shapiro-Ilan DI, Gaugler R, Yi S. Effect of entomopathogenic nematode concentration on survival during cryopreservation in liquid nitrogen. Journal of Nematology. 2004;36:281–284. [PMC free article] [PubMed] [Google Scholar]
Bai C, Shapiro-Ilan DI, Gaugler R, Hopper KR. Stabilization of beneficial traits in Heterorhabditis bacteriophora through creation of inbred lines. Biological Control. 2005;32:220–227. [Google Scholar]
Bai X, Grewal PS. Identification of two down-regulated genes in entomopathogenic nematode Heterorhabditis bacteriophora infective juveniles upon contact with insect hemolymph. Molecular and Biochemical Parasitology. 2007;156:162–166. [PubMed] [Google Scholar]
Bai X, Adams BJ, Ciche TA, Clifton S, Gaugler R, Hogenhout SA, Spieth J, Sternberg PW, Wilson RK, Grewal PS. Transcriptomic analysis of the entomopathogenic nematode Heterorhabditis bacteriophora TTO1. BMC Genomics 10, art. No. 205. 2009 [PMC free article] [PubMed] [Google Scholar]
Barbercheck ME. Effect of soil physical factors on biological control agents of soil insect pests. Florida Entomologist. 1992;75:539–548. [Google Scholar]
Barbercheck ME, Kaya HK. Interactions between Beauveria bassiana and the entomogenous nematodes Steinernema feltiae and Heterorhabditis heliothidis. Journal of Invertebrate Pathology. 1990;55:225–234. [Google Scholar]
Bedding RA. Low cost in vitro mass production of Neoaplectana and Heterorhabditis species (Nematoda) for field control of insect pests. Nematologica. 1981;27:109–114. [Google Scholar]
Bedding RA. Large scale production, storage, and transport of the insect-parasitic nematode Neoaplectana spp. and Heterorhabditis spp. Annals of Applied Biology. 1984;104:117–120. [Google Scholar]
Bednarek A, Gaugler R. Compatibility of soil amendments with entomopathogenic nematodes. Journal of Nematology. 1997;29:220–227. [PMC free article] [PubMed] [Google Scholar]
Begley JW. 1990. Efficacy against insects in habitats other than soil. Pp. 215–232 in R. Gaugler, and H. K. Kaya, eds. Entomopathogenic Nematodes in Biological Control. Boca Raton, FL: CRC Press. [Google Scholar]
Benz G. 1971. Synergism of micro-organisms and chemical insecticides. Pp. 327–355 in H. D. Burgess and N. W. Hussey eds. Microbial Control of Insects and Mites. London: Academic Press. [Google Scholar]
Berry RE, Liu J, Groth E. Efficacy and persistence of Heterorhabditis marelatus (Rhabditida: Heterorhabditidae) against root weevils (Coleoptera: Curculionidae) in strawberry. Environmental Entomology. 1997;26:465–470. [Google Scholar]
Bilgrami AL, Gaugler R, Shapiro-Ilan DI, Adams BJ. Source of trait deterioration in entomopathogenic nematodes Heterorhabditis bacteriophora and Steinernema carpocapsae during in vivo culture. Nematology. 2006;8:397–409. [Google Scholar]
Blinova SL, Ivanova ES. 1987. Culturing the nematode-bacterial complex of Neoaplectana carpocapsae in insects. Pp. 22–26 in M. D. Sonin, ed. Helminths of Insects. New Delhi: American Publishing Co. [Google Scholar]
Boff M, Wiegers GL, Gerritsen LJM, Smits PH. Development of the entomopathogenic nematode Heterorhabditis megidis strain NLH-E 87.3 in Galleria mellonella. Nematology. 2000;2:303–308. [Google Scholar]
Brinkman MA, Gardner WA. Possible antagonistic activity of two entomopathogens infecting workers of the red imported fire ant (Hymenoptera: Formicidae) Journal of Entomological Science. 2000;35:205–207. [Google Scholar]
Bruck DJ, Shapiro-Ilan DI, Lewis EE. Evaluation of application technologies of entomopathogenic nematodes for control of the black vine weevil, Otiorhynchus sulcatus. Journal of Economic Entomology. 2005;98:1884–1889. [PubMed] [Google Scholar]
Brusselman E, Beck B, Temmerman F, Pollet S, Steurbaut W, Moens M, Nuyttens D. The spray pattern of entomopathogenic nematodes. American Society of Agricultural and Biological Engineers Annual International Meeting 2010. 2010;6:5054–5067. [Google Scholar]
Buecher EJ, Popiel I. Liquid culture of the entomogenous nematode Steinernema feltiae with its bacterial symbiont. Journal of Nematology. 1989;21:500–504. [PMC free article] [PubMed] [Google Scholar]
Burman M, Pye AE. Neoaplectana carpocapsae: respiration of infective juveniles. Nematologica. 1980;26:214–219. [Google Scholar]
Burnell A. 2002. Genetics and genetic improvement. Pp 333–356 in Gaugler, R. (ed.), Entomopathogenic nematology. New York, NY: CABI. [Google Scholar]
Cabanillas HE, Raulston JR. Pathogenicity of Steinernema riobravis against corn earworm, Helicoverpa zea (Boddie) Fundamentals and Applied Nematology. 1994;17:219–223. [Google Scholar]
Cabanillas HE, Raulston JR. Impact of Steinernema riobravis (Rhabditida: Steinernematidae) on the control of Helicoverpa zea (Lepidoptera: Noctuidae) in corn. Journal of Economic Entomology. 1995;88:58–64. [Google Scholar]
Chaston JM, Dillman AR, Shapiro-Ilan DI, Bilgrami AL, Gaugler R, Hopper KR, Adams BJ. Outcrossing and crossbreeding recovers deteriorated traits in laboratory cultured Steinernema carpocapsae nematodes. International Journal of Parasitology. 2011;41:801–809. [PMC free article] [PubMed] [Google Scholar]
Chavarria-Hernandez N, de la Torre M. Population growth kinetics of the nematode, Steinernema feltiae, in submerged monoxenic culture. Biotechnology Letters. 2001;23:311–315. [Google Scholar]
Chavarria-Hernandez N, Espino-Garcia J-J, Sanjuan-Galindo R, Rodriguez-Hernandez A-I. Monoxenic liquid culture of the entomopathogenic nematode Steinernema carpocapsae using a culture medium containing whey kinetics and modeling. Journal of Biotechnology. 2006;125:75–84. [PubMed] [Google Scholar]
Chavarría-Hernández N, Ortega-Morales E, Vargas-Torres A, Chavarría-Hernández J, Rodríguez-Hernández A. Submerged monoxenic culture of the entomopathogenic nematode, Steinernema carpocapsae CABA01, in a mechanically agitated bioreactor: Evolution of the hydrodynamic and mass transfer conditions. Biotechnology and Bioprocess Engineering. 2010;15:580–589. [Google Scholar]
Ciche TA. 2007. The biology and genome of Heterorhabditis bacteriophora. (February 20, 2007), WormBook, ed. The C. elegans Research Community, WormBook, doi/10.1895/wormbook.1.135.1, http://www.wormbook.org. [Google Scholar]
Cottrell TE, Shapiro-Ilan DI, Horton DL, Mizell RF., III Laboratory virulence and orchard efficacy of entomopathogenic nematodes toward the lesser peachtree borer (Lepidoptera: Sesiidae) Environmental Entomology. 2011;104:47–53. [PubMed] [Google Scholar]
Del Valle EE, Dolinski C, Barreto ELS, Souza RM, Samuels RI. Efficacy of Heterorhabditis baujardi LPP7 (Nematoda: Rhabditida) applied in Galleria mellonella (Lepidoptera: Pyralidae) insect cadavers to Conotrachelus psidii, (Coleoptera: Curculionidae) larvae. Biocontrol Science and Technology. 2008;18:33–41. [Google Scholar]
Del Valle EE, Dolinksi C, Barreto ELS, Souza RM. Effect of cadaver coatings on emergence and infectivity of the entomopathogenic nematode Heterorhabditis baujardi LPP7 (Rhabditida: Heterorhabditidae) and the removal of cadavers by ants. Biological Control. 2009;50:21–24. [Google Scholar]
Del Valle EE, Dolinski C, Barreto ELS, Souza RM. Influence of time period between infection by Heterorhabditis baujardi LPP7 and soil application of Galleria mellonella over emergence of infective juvenile nematodes. Nematologia Brasileira 34: in press. 2011 [Google Scholar]
Deol YS, Jagdale GB, Cañas L, Grewal PS. Delivery of entomopathogenic nematodes directly through commercial growing media via the inclusion of infected host cadavers: A novel approach. Biological Control. 2011;58:60–67. [Google Scholar]
Dias PVC, Dolinski C, Molina JPA. Influence of infective juvenile doses and Galleria mellonella (Lepidoptera: Pyralidae) larvae weight in the in vivo production of Heterorhabditis baujardi LPP7 (Rhabditida: Heterorhabditidae) Nematologia Brasileira. 2008;32:317–321. [Google Scholar]
Dolinski C, Del Valle EE, Burla RS, Machado IR. Biological traits of two native brazilian entomopathogenic nematodes (Heterorhabditidae: Rhabditida) Nematologia Brasileira. 2007;31:180–185. [Google Scholar]
Dowds BCA, Peters A. Virulence mechanisms 2002. Pp. 79–98 in R. Gaugler, ed. Entomopathogenic nematology. New York, NY: CABI. [Google Scholar]
Downing AS. Effect of irrigation and spray volume on efficacy of entomopathogenic nematodes (Rhabditida: Heterorhabditidae) against white grubs (Coleoptera: Scarabaeidae) Journal of Economic Entomology. 1994;87:643–646. [Google Scholar]
Duncan LW, McCoy CW. Vertical distribution in soil, persistence, and efficacy against citrus root weevil (Coleoptera: Curculionidae) of two species of entomogenous nematodes (Rhabditida: Steinernematidae: Heterorhabditidae) Environmental Entomology. 1996;25:174–178. [Google Scholar]
Dunphy GB, Webster JM. The monoxenic culture of Neoaplectana carpocapsae DD136 and Heterorhabditis heliothidis. Revue de Nématologie. 1989;12:113–123. [Google Scholar]
Dutky SR, Thompson JV, Cantwell GE. A technique for the mass propagation of the DD-136 nematode. Journal of Insect Pathology. 1964;6:417–422. [Google Scholar]
Ehlers R-U, Shapiro-Ilan DI. Mass production. Pp. 65–79 2005 in P. Grewal, R-U Ehlers, and D. Shapiro-Ilan, eds. Nematodes as biological control agents. Wallingford, UK: CABI Publishing. [Google Scholar]
Ehlers RU, Niemann I, Hollmer S, Strauch O, Jende D, Shanmugasundaram M, Mehta UK, Easwaramoorthy SK, Burnell A. Mass production potential of the bacto-helminthic biocontrol complex Heterorhabditis indica-Photorhabdus luminescens. Biocontrol Science and Technology. 2000;10:607–616. [Google Scholar]
Elawad SA, Gowen SR, Hague NGM. Progeny production of Steinernema abbasi in lepidopterous larvae. International Journal of Pest Management. 2001;47:17–21. [Google Scholar]
Eng MS, Preisser EL, Strong DR. Phoresy of the entomopathogenic nematode Heterorhabditis marelatus by a non-host organism, the isopod Porcellio scaber. Journal of Invertebrate Pathology. 2005;88:173–176. [PubMed] [Google Scholar]
Epsky ND, Walter DE, Capinera JL. Potential role of nematophagous microarthropods as biotic mortality factors of entomogenous nematodes (Rhabditida: Steinernematidae, Heterorhabditidae) Journal of Economic Entomology. 1988;81:821–825. [Google Scholar]
Fife JP, Derksen RC, Ozkan HE, Grewal PS. Effects of pressure differentials on the viability and infectivity of entomopathogenic nematodes. Biological Control. 2003;27:65–72. [Google Scholar]
Fife JP, Ozkan HE, Derksen RC, Grewal PS, Krause CR. Viability of a biological pest control agent through hydraulic nozzles. Transactions of the ASAE. 2005;48:45–54. [Google Scholar]
Flanders KL, Miller JM, Shields EJ. In vivo production of Heterorhabditis bacteriophora ‘Oswego’ (Rhabditida: Heterorhabditidae), a potential biological control agent for soil inhabiting insects in temperate regions. Journal of Economic Entomology. 1996;89:373–380. [Google Scholar]
Friedman MJ. 1990. Commercial production and development. Pp 153–172 in R. Gaugler, and H. K. Kaya, eds. Entomopathogenic Nematodes in Biological Control. Boca Raton, FL: CRC Press. [Google Scholar]
Friedman MJ, Langston SL, Pollit S. Mass production in liquid culture of insect-killing nematodes. International Patent WO89/04602. 1989 [Google Scholar]
Gaugler R. 1987. Entomogenous nematodes and their prospects for genetic improvement. Pp. 457–484 in K. Maramorosch, ed. Biotechnology in invertebrate pathology and cell culture. San Diego, CA: Academic Press. [Google Scholar]
Gaugler R, Boush GM. Effects of ultraviolet radiation and sunlight on the entomogenous nematode Neoaplectana carpocapsae. Journal of Invertebrate Pathology. 1978;32:291–296. [Google Scholar]
Gaugler R, Georgis R. Culture method and efficacy of entomopathogenic nematodes (Rhabditida: Steinernematidae and Heterorhabditidae) Biological Control. 1991;1:269–274. [Google Scholar]
Gaugler R, Han R. 2002. Production technology. Pp. 289–310 in R. Gaugler, ed. Entomopathogenic Nematology. Wallingford, UK: CABI. [Google Scholar]
Gaugler R, Campbell JF, McGuire TR. Fitness of a genetically improved entomopathogenic nematode. Journal of Invertebrate Pathology. 1990;56:106–116. [Google Scholar]
Gaugler R, Brown I, Shapiro-Ilan DI, Atwa A. Automated technology for in vivo mass production of entomopathogenic nematodes. Biological Control. 2002;24:199–206. [Google Scholar]
Georgis R. 1990. Formulation and application technology. Pp. 173–194 in R. Gaugler, and H. K. Kaya, eds. Entomopathogenic Nematodes in Biological Control. Boca Raton, FL: CRC Press. [Google Scholar]
Georgis R, Gaugler R. Predictability in biological control using entomopathogenic nematodes. Journal of Economic Entomology. 1991;84:713–720. [Google Scholar]
Georgis R, Hague NGM. Nematodes as biological insecticides. Pesticide Outlook. 1991;3:29–32. [Google Scholar]
Georgis R, Poinar GO., Jr Effect of soil texture on the distribution and infectivity of Neoaplectana carpocapsae (Nematoda: Steinernematidae) Journal of Nematology. 1983;15:308–311. [PMC free article] [PubMed] [Google Scholar]
Georgis R, Dunlop DB, Grewal PS. 1995. Formulation of entomopathogenic nematodes. Pp. 197–205 in F. R. Hall, and J. W. Barry, eds. Biorational Pest Control Agents: Formulation and Delivery. Washington, DC: American Chemical Society. [Google Scholar]
Glazer I, Salame L, Segal D. Genetic enhancement of nematicide resistance in entomopathogenic nematodes. Biocontrol Science and Technology. 1997;7:499–512. [Google Scholar]
Grewal PS. 2002. Formulation and application technology. Pp. 265–288 in R. Gaugler, ed. Entomopathogenic Nematology. Wallingford, UK: CABI Publishing. [Google Scholar]
Grewal PS, Georgis R. 1999. Entomopathogenic nematodes. Pp. 271–299 in F. R. Hall, and J. J. Menn, eds. Methods in Biotechnology, vol. 5: Biopesticides: Use and Delivery. Totowa, NJ: Humana Press. [Google Scholar]
Grewal PS, Selvan S, Gaugler R. Thermal adaptation of entomopathogenic nematodes – niche breadth for infection, establishment and reproduction. Journal of Thermal Biology. 1994;19:245–253. [Google Scholar]
Grewal PS, Converse V, Georgis R. Influence of production and bioassay methods on infectivity of two ambush foragers (Nematoda: Steinernematidae) Journal of Invertebrate Pathology. 1999;73:40–44. [PubMed] [Google Scholar]
Grewal PS, Power KT, Grewal SK, Suggars A, Haupricht S. Enhanced consistency of white grubs (Coleoptera: Scarabaeidae) with new strains of entomopathogenic nematodes. Biological Control. 2004;30:73–82. [Google Scholar]
Grewal PS, Ehlers R-U, Shapiro-Ilan DI. 2005. Nematodes as Biological Control Agents. Wallingford, UK: CABI Publishing. [Google Scholar]
Han RC. The effects of inoculum size on yield of Steinernema carpocapsae and Heterorhabditis bacteriophora in liquid culture. Nematologica. 1996;42:546–553. [Google Scholar]
Han R, Cao L, Liu X. Relationship between medium composition, inoculum size, temperature and culture time in the yields of Steinernema and Heterorhabditis nematodes. Fundamental and Applied Nematology. 1992;15:223–229. [Google Scholar]
Han R, Cao L, Liu X. Effects of inoculum size, temperature, and time on in vitro production of Steinernema carpocapsae Agriotos. Nematologica. 1993;39:366–375. [Google Scholar]
Han R, Pang X, Li L. Optimization of the medium components for the solid culture of entomopathogenic Steinernema and Heterorhabditis nematodes. Natural Enemies of Insects. 1995;17:153–164. [Google Scholar]
Han R, Li L, Pang X. Modeling of the culture parameters for production of Steinernema carpocapsae and Heterorhabditis bacteriophora in solid culture. Natural Enemies of Insects. 1997;19:75–83. [Google Scholar]
Hara AH, Lindegren JE, Kaya HK. Monoxenic mass production of the entomogenous nematode Neoplectana carpocapsae Weiser on dog food/agar medium. USDA Advances in Agriculture W-16, 8 pp. 1981 [Google Scholar]
Head J, Lawrence AJ, Walters KFA. Efficacy of the entomopathogenic nematode, Steinernema feltiae, against Bemisia tabaci in relation to plant species. Journal of Applied Entomology. 2004;128:543–547. [Google Scholar]
Hirao A, Ehlers R-U. Influence of inoculum density on population dynamics and dauer juvenile yields in liquid culture of biocontrol nematodes Steinernema carpocapsae and S. feltiae (Nematoda: Rhabditida) Applied Microbiology and Biotechnology. 2010;85:507–515. [PubMed] [Google Scholar]
Hominick WM, Briscoe BR, del Pino FG, Heng J, Hunt DJ, Kozodoy E, Mracek Z, Nguyen KB, Reid AP, Spiridonov S, Stock P, Sturhan D, Waturu C, Yoshida M. Biosystematics of entomopathogenic nematodes: Current status, protocols, and definitions. Journal of Helminthology. 1997;71:271–298. [PubMed] [Google Scholar]
Jansson RK, Lecrone SH, Gaugler R. Field efficacy and persistence of entomopathogenic nematodes (Rhabditida: Steinernematidae, Heterorhabditidae) for control of sweetpotato weevil (Coleoptera: Apionidae) in southern Florida. Journal of Economic Entomology. 1993;86:1055–1063. [Google Scholar]
Jeffke T, Jende D, Matje C, Ehlers R-U, Berthe-Corti L. Growth of Photorhabdus luminescens in batch and glucose fed-batch culture. Journal of Applied Microbiology and Biotechnology. 2000;54:326–330. [PubMed] [Google Scholar]
Jessen P, Strauch O, Wyss U, Luttmann R, Ehlers R-U. Carbon dioxide triggers dauer juvenile recovery of entomopathogenic nematodes (Heterorhabditis spp. Nematology. 2000;2:310–324. [Google Scholar]
Johnigk S-A, Ecke F, Poehling M, Ehlers R-U. Liquid culture mass production of biocontrol nematodes, Heterorhabditis bacteriophora (Nematoda: Rhabditida): Improved timing of dauer juvenile inoculation. Applied Microbiology and Biotechnology. 2004;64:651–658. [PubMed] [Google Scholar]
Kanga FN, Waeyenberge L, Hauser S, Moens M. Distribution of entomopathogenic nematodes in Southern Cameroon. Journal of Invertebrate Pathology: In Press. 2012 [PubMed] [Google Scholar]
Kaya HK. 1990. Soil ecology. Pp. 93–116 in R. Gaugler and H. K. Kaya, eds. Entomopathogenic nematodes in biological control. Boca Raton, FL: CRC Press. [Google Scholar]
Kaya HK. 2002. Natural enemies and other antagonists. Pp. 189–204 in R. Gaugler, ed. Entomopathogenic Nematology. Wallingford, UK: CABI Publishing. [Google Scholar]
Kaya HK, Stock SP. 1997. Techniques in insect nematology. Pp. 281–324 in L. A. Lacey, ed. Manual of Techniques in Insect Pathology. San Diego, CA: Academic Press. [Google Scholar]
Klein MG, Georgis R. Persistence of control of Japanese beetle (Coleoptera: Scarabaeidae) larvae with steinernematid and heterorhabditid nematodes. Journal of Economic Entomology. 1992;85:727–730. [Google Scholar]
Koppenhöfer AM, Fuzy EM. Early timing and new combinations to increase the efficacy of neonicotinoid–entomopathogenic nematode (Rhabditida: Heterorhabditidae) combinations against white grubs (Coleoptera: Scarabaeidae) Pest Management Science. 2008;64:725–735. [PubMed] [Google Scholar]
Koppenhöfer AM, Grewal PS. 2005. Compatibility and interactions with agrochemicals and other biocontrol agents. Pp. 363–381in P. S. Grewal, R.-U. Ehlers, and D. I. Shapiro-Ilan, eds. Nematodes as Biological Control Agents. Wallingford, UK: CABI Publishing. [Google Scholar]
Koppenhöfer AM, Kaya HK. Additive and synergistic interactions between entomopathogenic nematodes and Bacillus thuringiensis for scarab grub control. Biological Control. 1997;8:131–137. [Google Scholar]
Koppenhöfer AM, Kaya HK. Synergism of imidacloprid and an entomopathogenic nematode: a novel approach to white grub (Coleoptera: Scarabaeidae) control in turfgrass. Journal of Economic Entomology. 1998;91:618–623. [Google Scholar]
Koppenhöfer AM, Kaya HK, Taormino SP. Infectivity of entomopathogenic nematodes (Rhabditida: Steinernematidae) at different soil depths and moistures. Journal of Invertebrate Pathology. 1995;65:193–199. [Google Scholar]
Kung S, Gaugler R, Kaya HK. Influence of soil pH and oxygen on persistence of Steinernema spp. Journal of Nematology. 1990a;22:440–445. [PMC free article] [PubMed] [Google Scholar]
Kung S, Gaugler R, Kaya HK. Soil type and entomopathogenic nematode persistence. Journal of Invertebrate Pathology. 1990b;55:401–406. [Google Scholar]
Kung S, Gaugler R, Kaya HK. Effects of soil temperature, moisture, and relative humidity on entomopathogenic nematode persistence. Journal of Invertebrate Pathology. 1991;57:242–249. [Google Scholar]
Lacey LA, Shapiro-Ilan DI, Glenn GM. Post-application of anti-desiccant agents improves efficacy of entomopathogenic nematodes in formulated host cadavers or aqueous suspension against diapausing codling moth larvae (Lepidoptera: Tortricidae) Biocontrol Science and Technology. 2010;20:909–921. [Google Scholar]
Lara JC, Dolinski C, Fernandes de Sousa E, Figueiredo Daher E. Effect of mini-sprinkler irrigation system on Heterorhabditis baujardi LPP7 (Nematoda: Heterorhabditidae) infective juvenile. Scientia Agricola. 2008;65:433–437. [Google Scholar]
Lewis EE, Clarke DJ. 2012. Nematode parasites and entomopathogens. Pp. 395–424 in F. Vega and H. K. Kaya, eds. Insect Pathology, 2nd ed. Amsterdam, The Netherlands: Elsevier. [Google Scholar]
Lindegren JE, Hoffman DF, Collier SS, Fries RD. Propagation and storage of Neoaplectana carpocapsae Weiser using Amyelois transitella (Walker) adults. USDA Advances in Agriculture. 1979;3:1–5. [Google Scholar]
Lindegren JE, Valero KA, Mackey BE. Simple in vivo production and storage methods for Steinernema carpocapsae infective juveniles. Journal of Nematology. 1993;25:193–197. [PMC free article] [PubMed] [Google Scholar]
Llàcer E, Martinez de Altube MM, Jacas JA. Evaluation of the efficacy of Steinernema carpocapsae in a chitosan formulation against the red palm weevil, Rhynchophorus ferrugineus, in Phoenix canariensis. BioControl. 2009;54:559–565. [PubMed] [Google Scholar]
Lunau S, Stoessel S, Schmidt-Peisker AJ, Ehlers RU. Establishment of monoxenic inocula for scaling up in vitro cultures of the entomopathogenic nematodes Steinernema spp. and Heterorhabditis spp. Nematologica. 1993;39:385–399. [Google Scholar]
Molyneux AS, Bedding RA. Influence of soil texture and moisture on the infectivity of Heterorhabditis sp. D1 and Steinernema glaseri for larvae of the sheep blowfly Lucilia cuprina. Nematologica. 1984;30:358–365. [Google Scholar]
Morales-Ramos JA, Rojas MG, Shapiro-Ilan DI, Tedders WL. Nutrient regulation in Tenebrio molitor (Coleoptera: Tenebrionidae): SELF-selection of two diet components by larvae and impact on fitness. Environmental Entomology. 2011a;40:1285–1294. [PubMed] [Google Scholar]
Morales-Ramos JA, Rojas MG, Shapiro-Ilan DI, Tedders WL. 2011b. Automated. Insect Separation System. Application No. 12/536,221. Patent Number: 8025027. [Google Scholar]
Mukaka J, Strauch O, Hoppe C, Ehlers R-U. Improvement of heat and desiccation tolerance in Heterorhabditis bacteriophora through cross-breeding of tolerant strains and successive genetic selection. BioControl. 2010;55:511–521. [Google Scholar]
Neves JM, Teixeira JA, Simoes N, Mota M. Effect of airflow rate on yield of Steinernema carpocapsae Az 20 in liquid culture in an external-loop airlift bioreactor. Biotechnology and Bioengineering. 2001;72:369–373. [PubMed] [Google Scholar]
Nguyen KB, Buss EA. Steinernema phyllophagae n. sp. (Rhabditida: Steinernematidae), a new entomopathogenic nematode from Florida, USA. Nematology. 2011;13:425–442. [Google Scholar]
Nishimatsu T, Jackson JJ. Interaction of insecticides, entomopathogenic nematodes, and larvae of the western corn rootworm (Coleoptera: Chrysomelidae) Journal of Economic Entomology. 1998;91:410–418. [PubMed] [Google Scholar]
Pace WG, Grote W, Pitt DE, Pitt JM. Liquid culture of nematodes. International Patent WO 86/01074. 1986 [Google Scholar]
Parkman JP, Frank JH, Nguyen KB, Smart GC., Jr Inoculative release of Steinernema scapterisci (Rhabditida: Steinernematidae) to suppress pest mole crickets (Orthoptera: Gryllotapidae) on golf courses. Environmental Entomology. 1994;23:1331–1337. [Google Scholar]
Perez EE, Lewis EE, Shapiro-Ilan DI. Impact of host cadaver on survival and infectivity of entomopathogenic nematodes (Rhabditida: Steinernematidae and Heterorhabditidae) under desiccating conditions. Journal of Invertebrate Pathology. 2003;82:111–118. [PubMed] [Google Scholar]
Poinar GO. 1979. Nematodes for biological control of insects. Boca Raton, FL: CRC Press. [Google Scholar]
Poinar GO. 1990. Biology and taxonomy of Steinernematidae and Heterorhabditidae. Pp. 23–62 in R. Gaugler and H. K. Kaya eds. Entomopathogenic nematodes in biological control. Boca Raton, FL: CRC Press. [Google Scholar]
Schroer S, Ehlers R-U. Foliar application of the entomopathogenic nematode Steinernema carpocapsae for biological control of diamondback moth larvae (Plutella xylostella) Biological Control. 2005;33:81–86. [Google Scholar]
Shapiro DI, Glazer I. Comparison of entomopathogenic nematode dispersal from infected hosts versus aqueous suspension. Environmental Entomology. 1996;25:1455–1461. [Google Scholar]
Shapiro DI, Lewis EE. Comparison of entomopathogenic nematode infectivity from infected hosts versus aqueous suspension. Environmental Entomology. 1999;28:907–911. [Google Scholar]
Shapiro DI, McCoy CW. Effect of culture method and formulation on the virulence of Steinernema riobrave (Rhabditida: Steinernematidae) to Diaprepes abbreviatus (Curculionidae) Journal of Nematology. 2000a;32:281–288. [PMC free article] [PubMed] [Google Scholar]
Shapiro DI, McCoy CW. Virulence of entomopathogenic nematodes to Diaprepes abbreviatus (Coleoptera: Curculionidae) in the laboratory. Journal of Economic Entomology. 2000b;93:1090–1095. [PubMed] [Google Scholar]
Shapiro-Ilan DI, Gaugler R. Production technology for entomopathogenic nematodes and their bacterial symbionts. Journal of Industrial Microbiology and Biotechnology. 2002;28:137–146. [PubMed] [Google Scholar]
Shapiro DI, Tylka GL, Berry EC, Lewis LC. Effects of earthworms on the dispersal of Steinernema spp. Journal of Nematology. 1995;27:21–28. [PMC free article] [PubMed] [Google Scholar]
Shapiro DI, Glazer I, Segal D. Trait stability and fitness of the heat tolerant entomopathogenic nematode Heterorhabditis bacteriophora IS5 strain. Biological Control. 1996a;6:238–244. [Google Scholar]
Shapiro DI, Tylka GL, Lewis LC. Effects of fertilizers on virulence of Steinernema carpocapsae. Applied Soil Ecology. 1996b;3:27–34. [Google Scholar]
Shapiro DI, Glazer I, Segal D. Genetic improvement of heat tolerance in Heterorhabditis bacteriophora through hybridization. Biological Control. 1997;8:153–159. [Google Scholar]
Shapiro DI, Obrycki JJ, Lewis LC, Abbas M. The effects of fertilizers on black cutworm, Agrotis ipsilon, (Lepidoptera: Noctuidae) suppression by Steinernema carpocapsae. Journal of Nematology. 1999b;31:690–693. [PMC free article] [PubMed] [Google Scholar]
Shapiro DI, Cate JR, Pena J, Hunsberger A, McCoy CW. Effects of temperature and host age on suppression of Diaprepes abbreviatus (Coleoptera: Curculionidae) by entomopathogenic nematodes. Journal of Economic Entomology. 1999a;92:1086–1092. [Google Scholar]
Shapiro DI, Lewis EE, Paramasivam S, McCoy CW. Nitrogen partitioning in Heterorhabditis bacteriophora-infected hosts and the effects of nitrogen on attraction/repulsion. Journal of Invertebrate Pathology. 2000a;76:43–48. [PubMed] [Google Scholar]
Shapiro DI, McCoy CW, Fares A, Obreza T, Dou H. Effects of soil type on virulence and persistence of entomopathogenic nematodes in relation to control of Diaprepes abbreviatus. Environmental Entomology. 2000b;29:1083–1087. [Google Scholar]
Shapiro-Ilan DI, Lewis EE, Behle RW, McGuire MR. Formulation of entomopathogenic nematode-infected-cadavers. Journal of Invertebrate Pathology. 2001;78:17–23. [PubMed] [Google Scholar]
Shapiro-Ilan DI, Gaugler R, Tedders WL, Brown I, Lewis EE. Optimization of inoculation for in vivo production of entomopathogenic nematodes. Journal of Nematology. 2002a;34:343–350. [PMC free article] [PubMed] [Google Scholar]
Shapiro-Ilan DI, Gouge DH, Koppenhöfer AM. 2002b. Factors affecting commercial success: Case studies in cotton, turf and citrus. Pp. 333–356 in R. Gaugler, ed. Entomopathogenic Nematology. Wallingford, UK: CABI Publishing. [Google Scholar]
Shapiro-Ilan DI, Lewis EE, Tedders WL, Son Y. Superior efficacy observed in entomopathogenic nematodes applied in infected-host cadavers compared with application in aqueous suspension. Journal of Invertebrate Pathology. 2003;83:270–272. [PubMed] [Google Scholar]
Shapiro-Ilan DI, Jackson M, Reilly CC, Hotchkiss MW. Effects of combining an entomopathogenic fungi or bacterium with entomopathogenic nematodes on mortality of Curculio caryae (Coleoptera: Curculionidae) Biological Control. 2004;30:119–126. [Google Scholar]
Shapiro-Ilan DI, Stuart RJ, McCoy CW. Targeted improvement of Steinernema carpocapsae for control of the pecan weevil, Curculio caryae (Horn) (Coleoptera: Curculionidae) through hybridization and bacterial transfer. Biological Control. 2005;34:215–221. [Google Scholar]
Shapiro-Ilan DI, Gouge DH, Piggott SJ, Patterson Fife J. Application technology and environmental considerations for use of entomopathogenic nematodes in biological control. Biological Control. 2006a;38:124–133. [Google Scholar]
Shapiro-Ilan DI, Stuart RJ, McCoy CW. A comparison of entomopathogenic nematode longevity in soil under laboratory conditions. Journal of Nematology. 2006b;38:119–129. [PMC free article] [PubMed] [Google Scholar]
Shapiro-Ilan DI, Guadalupe Rojas M, Morales-Ramos JA, Lewis EE, Tedders WL. Effects of host nutrition on virulence and fitness of entomopathogenic nematodes: Lipid and protein based supplements in Tenebrio molitor diets. Journal of Nematology. 2008;40:13–19. [PMC free article] [PubMed] [Google Scholar]
Shapiro-Ilan DI, Tedders WL, Morales-Ramos JA, Rojas MG. 2009. Insect Inoculation System and Method. Patent disclosure, Docket No. 0107.04. Submitted to the US patent and trademark office on 12/11/2009. Application No. 12636245. [Google Scholar]
Shapiro-Ilan DI, Cottrell TE, Mizell RF, Horton DL, Behle B, Dunlap C. Efficacy of Steinernema carpocapsae for control of the lesser peachtree borer, Synanthedon pictipes: Improved aboveground suppression with a novel gel application. Biological Control. 2010a;54:23–28. [Google Scholar]
Shapiro-Ilan DI, Morales-Ramos JA, Rojas MG, Tedders WL. Effects of a novel entomopathogenic nematode–infected host formulation on cadaver integrity, nematode yield, and suppression of Diaprepes abbreviatus and Aethina tumida under controlled conditions. Journal of Invertebrate Pathology. 2010b;103:103–108. [PubMed] [Google Scholar]
Shapiro-Ilan DI, Tedders WL, Morales-Ramos JA, Rojas MG. System and Method for Producing Beneficial Parasites. US Patent Application No. 1 13/217,956. DN 172.07. 2011a [Google Scholar]
Shapiro-Ilan DI, Cottrell TE, Wood BW. Effects of combining microbial and chemical insecticides on mortality of the pecan weevil (Coleoptera: Curculionidae) Journal of Economic Entomology. 2011b;104:14–20. [PubMed] [Google Scholar]
Shapiro-Ilan DI, Rojas G, Morales-Ramos JA, Tedders WL. Optimization of a host diet for in vivo production of entomopathogenic nematodes. Journal of Nematology: In Press. 2011c [PMC free article] [PubMed] [Google Scholar]
Shetlar DJ, Suleman PE, Georgis R. Irrigation and use of entomogenous nematodes, Neoaplectana spp. and Heterorhabditis heliothidis (Rhabditida: Steinernematidae and Heterorhabditidae), for control of Japanese beetle (Coleoptera: Scarabaeidae) grubs in turfgrass. Journal of Economic Entomology. 1988;81:1318–1322. [Google Scholar]
Shields EJ, Testa A, Miller JM, Flanders KL. Field efficacy and persistence of the entomopathogenic nematodes Heterorhabditis bacteriophora ‘Oswego’ and H. bacteriophora ‘NC’ on Alfalfa snout beetle larvae (Coleoptera: Curculionidae) Environmental Entomology. 1999;28:128–136. [Google Scholar]
Steiner G. Aplectana kraussei n. sp. einer in der Blattwespe Lyda sp. parasitierende Nematodenform, nebst Bemerkungen uber das Seitenorgan der para sitischen Nematoden. Zentralbl. Bakteriol. Parasitenkd. Infektionskr. Hyg. Abt. 1923;59:14–18. [Google Scholar]
Strauch O, Ehlers RU. Food signal production of Photorhabdus luminescens inducing the recovery of entomopathogenic nematodes Heterorhabditis spp. in liquid culture. Applied Microbiology and Biotechnology. 1998;50:369–374. [Google Scholar]
Strauch O, Ehlers RU. Influence of the aeration rate on the yields of the biocontrol nematode Heterorhabditis megidis in monoxenic liquid culture. Applied Microbiology and Biotechnology. 2000;54:9–13. [PubMed] [Google Scholar]
Strauch O, Stoessel S, Ehlers R-U. Culture conditions define automictic or amphimictic reproduction in entomopathogenic rhabditid nematodes of the genus Heterorhabditis. Fundamentals and Applied Nematology. 1994;17:575–582. [Google Scholar]
Strauch O, Niemann I, Niemann A, Schmidt AJ, Peters A, Ehlers R-U. Storage and formulation of the entomopathogenic nematodes Heterorhabditis indica and H. bacteriophora. Biocontrol. 2000;45:483–500. [Google Scholar]
Stuart RJ, Shapiro-Ilan DI, James RR, Nguyen KB, McCoy CW. Virulence of new and mixed strains of the entomopathogenic nematode Steinernema riobrave to larvae of the citrus root weevil Diaprepes abbreviatus. Biological Control. 2004;30:439–445. [Google Scholar]
Surrey MR, Davies RJ. Pilot scale liquid culture and harvesting of an entomopathogenic nematode, Heterorhabditis. Fundamentals and Applied Nematology. 1996;17:575–582. [Google Scholar]
Thurston GS, Kaya HK, Gaugler R. Characterizing the enhanced susceptibility of milky disease-infected scarabaeid grubs to entomopathogenic nematodes. Biological Control. 1994;4:67–73. [Google Scholar]
Wallace HR. Movement of eelworms. Annals of Applied Biology. 1958;46:74–85. [Google Scholar]
Wang JX, Bedding RA. Population dynamics of Heterorhabditis bacteriophora and Steinernema carpocapsae in in vitro solid culture. Fundamentals and Applied Nematology. 1998;21:165–171. [Google Scholar]
Wang X, Grewal PS. Rapid genetic deterioration of environmental tolerance and reproductive potential of an entomopathogenic nematode during laboratory maintenance. Biological Control. 2002;23:71–78. [Google Scholar]
White GF. A method for obtaining infective nematode larvae from cultures. Science. 1927;66:302–303. [PubMed] [Google Scholar]
Williams EC, Walters KFA. Foliar application of the entomopathogenic nematode Steinernema feltiae against leafminers on vegetables. Biocontrol Science and Technology. 2000;10:61–70. [Google Scholar]
Womersley CZ. 1993. Factors affecting physiological fitness and modes of survival employed by dauer juveniles and their relationship to pathogenicity. Pp. 79–88 in R. Bedding, R. Akhurst, and H. Kaya, eds. Nematodes for the biological control of insects. East Melbourne, Australia: CSIRO Press. [Google Scholar]
Woodring JL, Kaya HK. 1988. Steinernematid and heterorhabditid nematodes: A handbook of biology and techniques. Southern Cooperative Series Bulletin 331, 30 pp, Arkansas Agricultural Experiment Station, Fayetteville, AR. [Google Scholar]
Wouts WM. Mass production of the entomogenous nematode Heterorhabditis heliothidis (Nematode: Heterorhabditidae) on artificial media. Journal of Nematology. 1981;13:467–469. [PMC free article] [PubMed] [Google Scholar]
Wright DJ, Peters A, Schroer S, Fife JP. 2005. Application technology. Pp. 91–106 in P. S. Grewal, R.-U. Ehlers, and D. I. Shapiro-Ilan, eds. Nematodes as biocontrol agents. New York, NY: CABI. [Google Scholar]
Yang H, Jian H, Zhang S, Zhang G. Quality of the entomopathogenic nematode Steinernema carpocapsae produced on different media. Biological Control. 1997;10:193–198. [Google Scholar]
Yoo SK, Brown I, Gaugler R. Liquid media development for Heterorhabditis bacteriophora: Lipid source and concentration. Applied Microbiology and Biotechnology. 2000;54:759–763. [PubMed] [Google Scholar]
Young JM, Dunnill P, Pearce JD. Separation characteristics of liquid nematode cultures and the design of recovery operations. Biotechnology Progress. 2002;18:29–35. [PubMed] [Google Scholar]
Zervos S, Johnson SC, Webster JM. Effect of temperature and inoculum size on reproduction and development of Heterorhabditis heliothidis and Steinernema glaseri (Nematoda: Rhabditoidea) in Galleria mellonella. Canadian Journal of Zoology. 1991;69:1261–1264. [Google Scholar]
Zhu H, Grewal PS, Reding ME. Development of a desiccated cadaver delivery system to apply entomopathogenic nematodes for control of soil pests. Applied Engineering in Agriculture. 2011;27:317–324. [Google Scholar]
Zimmerman RJ, Cranshaw WS. Short-term movement of Neoaplectana spp. (Rhabditida: Steinernematidae) and Heterorhabditis “HP-88” strain (Rhabditida: Heterorhabditidae) through turfgrass thatch. Journal of Economic Entomology. 1991;84:875–878. [Google Scholar]

Vinegar and baking soda are not only useful cooking and cleaning agents but are also helpful in the garden. Vinegar can be used as a spray to kill weeds; a baking soda solution can be used to treat black spot fungus on roses, and both can be used to change the pH of soil.

Step 1​

Test the pH of soil by using a pH meter or a pH test kit. Both come with instructions, but generally, add 1 tbsp. of soil to a cup, add distilled water and stir. Insert the meter's prong or the test's litmus paper strip into the soil. The meter or litmus strip will provide a pH reading.

Step 2​

Conduct a homemade version of a pH test if you do not have access to a pH meter or kit. Collect two soil samples from the same area. Add distilled water to each and stir. Add 1 tbsp. of vinegar to the first and 1 tbsp. of baking soda to the other. If the cup with vinegar added starts to fizz and bubble, the soil is alkaline, since vinegar is acidic with a pH of 3.3. If the soil with the baking soda bubbles, the soil is acidic, since baking soda is alkaline with a pH of 8.2.

Step 3​

Add vinegar to your soil if you need to lower the pH or make the soil more acidic. Mix 1 gallon of water with 1 cup of vinegar. Pour the solution around the base of plants in the soil you are adjusting.

Step 4​

Add baking soda to the soil if you need to raise the pH or make the soil more alkaline. Mix 1 tbsp. of baking soda with 1 gallon of water and stir. Apply the solution to your soil.

Step 5​

Test your soil the next day and regularly to monitor pH levels.

Plant pathology is fun! Remember there are multiple stages to the life cycle of most pathogens, so when doing tissue examinations with a microscope, be aware a particular disease may appear differently depending on life stage of the pathogen.

Take tissue samples to examine the center of large leaf spots to find spores/conidia for identification.

pipeline said:

Plant pathology is fun! Remember there are multiple stages to the life cycle of most pathogens, so when doing tissue examinations with a microscope, be aware a particular disease may appear differently depending on life stage of the pathogen.

Take tissue samples to examine the center of large leaf spots to find spores/conidia for identification.

Leaf spot diseases of trees and shrubs

Leaf spots, cankers and blights caused by multiple pathogens have very similar management options.

extension.umn.edu

Click to expand...

Great point, appreciate you pointing that out. Thank you

Bio-tone Starter Plus is the ultimate starter plant food! It’s a rich blend of the finest natural & organic ingredients and enhanced with beneficial microbes, humates and mycorrhizae. No sludges or fillers are ever used. And because it’s made in our state-of-the-art solar powered manufacturing facility, consistency and quality are guaranteed.

• Enhanced with Beneficial Microbes & Mycorrhizae
• Helps Plants Establish Fast
• Grow Deeper Roots & Bigger Blooms
• 100% Natural & Organic Approved
• Safe for People, Pets & Planet

Looks like a great mix, I have seen that for sale before. I think its a little higher priced obviously. I have used Espoma products for years, but about 5 years ago it I seemed to have more burn issues when top dressing. Not sure if they changed their formulation and it was a little hot. Looks like this blend would be less prone to burn, just be careful with it and stick to the recommended rate.

Interesting study there! I haven't used innoculants at this plot, I should do that. Soil naturally has some of these microorganisms in certain amounts, but its good to innoculate to optimize colonization.