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Grower Notes and Pest News

Entomophthora planchoniana infecting the strawberry aphid, Chaetosiphon fragaefolii in California

Species of the genus Entomophthora cause epizootics in various insects around the world, but such infections are less common in California.  Burger and Swain (1918) reported Entomophthora chromaphidis infections in the walnut aphid, Chromaphis juglandicola, in Southern California.  With infections as high as 95%, the fungus was a significant mortality factor in aphid populations.  In 2011, nearly a century after the epizootics in the walnut aphid, a single strawberry aphid, Chaetosiphon fragaefolii, was found infected with a species of Entomophthora in an organic strawberry field (Eraud Farms) in Santa Maria.  The strawberry aphid is an occasional and minor pest in strawberry in California.

Entomophthora planchoniana infection in strawberry aphid

Attempts to in vitro culture the fungus were unsuccessful, but microscopic measurements of conidial size and shape indicate that the causal agent could be Entomophthora planchoniana.  Bell-shaped conidia measured 17.3 μm or micrometers (14.8-20.1) long and 14.6 μm (12.7-17.7) wide (based on the measurements of 100 conidia) and had a broad base (papilla) and a pointed apex.  Conidia also appeared to have 4-6 nuclei.

E. planchoniana and E. chromaphidis are closely related and were previously considered as synonymous species (MacLeod et al., 1976; Waterhouse and Brady, 1982).  Humber and Feng (1991) later described these two as separate species due to the variation in conidial size, geographic distribution, host range, in vitro culturing techniques, and other characteristics.  While E. planchoniana is commonly found in Europe, E. chromaphidis is reported elsewhere.

The following are the characters of E. chormaphidis and E. planchoniana from Keller's (2002) description of 22 species of Entomophthora.

Entomophthora chromaphidis

First found in walnut aphids in California.  Primary conidia are 11-14 μm long and 10-11 μm wide with 4-6 nuclei and contain a single large oil globule.  Resting spores are 30 μm.  Insect host is attached to the plant surface with fungal rhizoids (bundles of modified hyphal bodies).

Entomophthora planchoniana

First found on unidentified aphids on elder in western Europe.  Primary conidia are 15-20 μm long and 12-16 μm wide with 4-11 nuclei [species description has 4-11 and key has 5-9 nuclei in Keller (2002)].  Resting spores are 31-38 μm.  The fungus produces rhizoids to attach the host insect to the plant surface.

Keller (2002) noted an overlap in the number of nuclei between these two species and suggested the size of primary conidia as the distinguishing character.  Hence the fungus found in the strawberry study is considered E. planchoniana, which was first reported in 1948 in the strawberry aphid, then known as Pentatrichopus fragariae (Petch, 1948; Leatherdale, 1970).  Cédola and Greco (2010) reported E. planchoniana as a major mortality factor of the strawberry aphid in Argentina.

Life cycle of E. planchoniana

The infection process starts when a conidium (single spore) comes in contact with the insect cuticle, produces a germ tube and gains entry into the insect body with the help of a penetration peg and cuticle degrading enzymes.  The fungus multiplies inside the insect body as protoplasts (cells without a cell wall) and invade the tissues.  Vegetative growth stops when nutrients are depleted and the insect is dead.  The fungus then produces conidiophores that emerge from the cuticle, produce bell-shaped primary conidia that are forcibly discharged.  A halo of protoplasm (cellular contents) is often seen around the primary conidium, which may either produce the germ tube to cause infection or a secondary conidium.  Secondary conidia are smaller than primary conidia, have rounded or less pointed apices, and more rounded basal papillae.

This is the first report of the occurrence of E. planchoniana in the strawberry aphid in California.  Although strawberry aphid is not an important pest in California and only one infected aphid was found, this finding is important to record the distribution of E. planchoniana.


Burger, O. F. and A. F. Swain.  1918.  Observations on a fungus enemy of the walnut aphis in Southern California.  J. Econ. Entomol. 11: 278-289.

Cédola C. and N. Greco.  2010.  Presence of the aphid, Chaetosiphon fragefolii, on strawberry in Argentina.  J. Ins. Sci. 10: 1-9.

Humber, R. A. and M.-G. Feng.  1991.  Entomophthora chromaphidis (Entomophthorales): the correct identification of an aphid pathogen in the pacific northwest and elsewhere.  Mycotaxon 41: 497-504.

Leatherdale, D.  1970.  The arthropod hosts of entomogenous fungi in Britain.  Entomophaga 15: 419-435.

MacLeod, D. M., E. Müller-Kögler, and N. Wilding.  1976.  Entomophthora species with E. muscae-like conidia.  Mycologia 68: 1-29.

Petch, T.  1948.  A revised list of British entomogenous fungi.  Trans. Brit. Mycol. Soc. 31: 286-304.

Waterhouse, G. M. and B. L. Brady.  1982.  Key to the species of Entomophthora sensu lato.  Bull. Brit. Mycol. Soc. 16: 113-143.


Posted on Monday, September 25, 2017 at 9:18 AM

Evaluating beneficial microbe-based products for their impact on strawberry plant growth, health, and fruit yield

Various soilborne, fruit and foliar diseases can affect strawberry crop and fruit yields.  Chemical fumigants and a variety of fungicides are typically used for managing the disease issues.  In addition to the environmental and human health concerns with chemical control options there is a need to improve current disease management with alternatives that include beneficial microbes.  Previous studies showed some promise with some of the treatments, but additional studies are required to evaluate the efficacy, which is more evident especially when there is disease incidence. 

A study was conducted in summer-planted conventional strawberries in 2016 at Manzanita Berry Farms to evaluate the impact of various beneficial microbial treatments on plant growth, health, and fruit yield.  Untreated control and the grower standard practice (Healthy Soil treatment) were compared with MycoApply EndoMaxx (Glomus intraradices, G. aggregatum, G. mosseae, and G. etunicatum), Actinovate AG (Streptomyces lydicus WYEC 108), and Inocucor Garden Solution (Saccharomyces cerevisiae and Bacillus subtilis) applied in the following treatments:

1. Untreated control

2. Grower Standard-Healthy Soil; transplant dip in Switch 63 WG 5 oz in 100 gal

3. MycoApply EndoMaxx 2 gpa transplant dip (TD)

4. MycoApply EndoMaxx 2 gpa drip at planting (DrP)

5. MycoApply EndoMaxx 2 gpa transplant dip + 2 gpa drip at planting

6. MycoApply EndoMaxx 4 gpa transplant dip

7. MycoApply EndoMaxx 4 gpa drip at planting

8. MycoApply EndoMaxx 4 gpa transplant dip + 4 gpa drip at planting

9. Actinovate AG 6 oz/ac transplant dip + 6 oz drip at planting + 6 oz drip monthly (DrM)

10. Inocucor Garden Solution 1 gpa drip at planting + 1 gpa drip monthly

Transplanting was done on 21 May, 2016 with appropriate treatments administered at the time of planting and thereafter.  Study had two blocks of 10 strawberry beds (300' long) and treatments were randomly applied to a bed within each block.  Two 15' long plots were marked within each bed for sampling.  Canopy growth was measured on June 21, July 5 and 20; powdery mildew severity on August 3, September 1, October 10 and November 16; botrytis severity 3 and 5 days after harvest (DAH) for berries harvested on September 13 and 27, and October 11 and 18; and dead and dying plants were counted on September 16 and October 23.  Yield data were collected from August 20 to November 18.  Powdery mildew and botrytis fruit rot severity was measured on a scale of 0 to 4 where 0=No disease, 1=1-25%, 2=26-50%, 3=51-75%, and 4=76-100% severity.  Data were analyzed and means were separated using LSD test.

Strawberry field and plots on June 9 (above) and August 31 (below).

Two sampling plots were set up within each bed to collect plant growth, health, and yield data.

Canopy growth: MycoApply EndoMaxxat 2 gpa either as a transplant dip with or without drip application at planting appeared to promote significantly higher growth (P <0.0001) than MycoApply EndoMaxx at 2 and 4 gpa as drip at planting, untreated control, and grower standard.  Inoculating the entire transplant with Glomus spp. through a dip appears to be better than application through drip irrigation system.

Powdery mildew: Disease incidence and severity was low during the observation period.  When the average of four observations period was compared, the grower standard, MycoApply Endomaxx at 2 and 4 gpa as drip at planting, and the Actinovate treatments had the lowest incidence (P = 0.0271).

Botrytis fruit rot: There was no difference (P >0.05) among the treatments on botrytis when the mold growth on fruit was compared 3 and 5 days after harvest.

Unknown issue: Some wilting and dead plants were found throughout the field during the study.  Although symptoms suggested some kind of wilt, laboratory testing did not identify any pathogens.  The total number of dead and dying plants was the lowest in Actinovate treatment, but it was significantly different (P = 0.0429) only from the grower standard Healthy Soil treatment.

*Means followed by the same or no letter are not significantly different at the P value indicated in the table.

Fruit yield: There were no statistically significant difference among the treatments and the seasonal total of marketable yield varied between 66 lb/plot in the grower standard and about 76 lb/plot in MycoApply EndoMaxx applied as a transplant dip at 4 gpa.

Total and marketable berry yields and their proportion among different treatments.

We need to continue to evaluate beneficial microbial products and their potential benefit in improve crop health and yields.

Acknowledgements: Thanks to Chris Martinez and Tamas Zold for technical assistance, and Valent USA and Inocucor Technologies for the financial support of the study.

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Posted on Thursday, September 7, 2017 at 1:16 PM

Epizootics of an entomophthoralean fungus in spotted-wing drosophila populations on fig

Life stages of spotted-wing drosophila (Photos by Elizabeth Beers, Washington State University)

Spotted-wing drosophila (SWD), Drosophila suzukii is an invasive pest that attacks many cultivated and wild fruits.  With the help of a strong, saw-like ovipositor or egg laying appendage, SWD is able to deposit eggs in ripe and occasionally in unripe or developing fruit unlike other Drosophila spp., commonly known as vinegar flies or fruit flies, that attack ripe or fallen fruit.  Larvae develop in the fruit and pupation occurs either in or outside the fruit.  Blueberry, caneberries, cherry, peach, and strawberry are some of the commercially important fruit crops that are at a risk of SWD damage.

Monitoring with lures, application of pesticides, use of exclusion netting, and sanitation are some of the control practices currently adopted in organic and conventional crops.  Among the microbial control options, entomopathogenic fungi such as Beauveria bassiana, Isaria fumosorosea, and Metarhizium brunneum (=M. anisopliae) against adults and entomopathogenic nematodes such as Heterorhabditis spp. and Steinernema spp. against pupae in the soil can be potential choices.  A few lab studies that evaluated these options showed limited efficacy of the most except for B. bassiana treatments in Italy and M. brunneum in Oregon that appeared promising (Gargani et al., 2013; Cuthbertson et al., 2014; Woltz et al., 2015).  Biocontrol potential with predators and parasitoids is also limited based on current research data (Haye et al., 2015; Renkema et al, 2015; Woltz et al., 2015).

In light of limited microbial and biocontrol control agents, a recent outbreak of fungal epizootics in SWD on fig offers a potential natural control option.  SWD populations in a small fig orchard in Clinton, Mississippi were infected by a fungus in June, 2017.  Unusually cool and wet conditions caused epizootics of a fungus, which was later identified by Connecticut Agricultural Experiment Station scientists as Entomophthora muscae or a closely related species.  SWD first appeared in blueberry, blackberry, and mulberry plots of this orchard in 2012 and infestations on figs were noticed only in 2017.  Other SWD hosts that are grown at this orchard include grapes, pears, and strawberries.  Having a variety of hosts with extended availability of fruits could have supported SWD populations at this location.

Dead or immobilized SWD (above) and sporulating cadavers (below) from Entomophthora muscae infections.  Photos by Tom Mann, Mississippi Museum of Natural Science, Jackson, MS

Conidia at different stages of development on conidiphores (left) and discharged conidia (right).  Photos by DeWei Li, Connecticut Agricultural Experiment Station, Windsor, CT

Unlike B. bassiana, I. fumosorosea, and M. brunneum (Phylum Ascomycota: Class Sordariomycetes: Order Hypocreales), Entomophthora spp. belong to a different fungal group (Phylum Zygomycota: Class Entomophthoromycetes: Order Entomophthorales).  Entomophthora spp. cause disease outbreaks in their host populations when environmental conditions are favorable with high humidity and low temperature aided by high host densities.  Entomophthora muscae is considered to be a species complex infecting a variety of dipteran families including Drosophilidae (Goldstein, 1927; MacLeod et al., 1976; Gryganskyi et al., 2013).  However, it appeared to be less pathogenic to the common fruit fly, Drosophila melanogaster compared to other dipteran species (Steinkraus and Kramer, 1987).

Entomopathogenic fungi typically take 3-5 days to kill their hosts.  Infection process typically starts when host insect comes in contact with the conidia (asexual spores) of the fungus (Brobyn and Wilding, 1983).  Primary conidia either produce a germ-tube that penetrates through the host cuticle or produce secondary conidia (which later produce germ-tubes) or hyphae.  Both enzymatic degradation of cuticle and mechanical pressure by the penetration peg of the germ-tube aid in fungus gaining entry into the host body.  Hyphal bodies are formed inside the host, invade the fat bodies and other tissues, and eventually cause death of the host insect.  The fungus later emerges from the intersegmental membranes and conidiophores or spore bearing structures produce conidia that are dispersed to continue the infection cycle.  Infected flies become sluggish and typically fly to the higher parts of the plant canopy where they become attached to plant surfaces with rhizoids (peg like structures that emerge from the ventral or lower side of the insect body) and sticky secretions (Steinkraus and Kramer, 1987).  This process increases the chances of disease spread as insect cadavers are securely attached to plant surfaces and infective conidia are dispersed from a higher elevation in the canopy.  When host populations diminish and during the winter months, entomophthoralean fungi may produce environmentally resilient resting spores to survive cold winters (Eilenberg and Michelsen, 1999) or survive as hyphal bodies in the dead (Keller, 1987) or winter hosts (Klingen et al., 2008).  Other overwintering options for these fungi include infections in their host insects on winter crops (Dara and Semtner, 2001) or infections in alternative host insects (Eilenberg et al., 2013).

Entomophthoralean fungi are difficult to culture in vitro and do not have the biopesticide potential as the hypocrealean fungi.  However, they can be significant mortality factors in some areas and bring down high host populations.  Neozygites fresenii epizootics in cotton aphid, Aphis gossypii (Steinkraus et al., 1995), Entomophaga maimaiga in gypsy moth, Lymantria dispar (Hajek and Elkinton, 1991), and Pandora neoaphidis in green peach aphid, Myzus persicae populations (Dara and Semtner, 2007) are some of the examples for the natural control of insects by entomophthoralean fungi.

Anecdotal reports indicated outbreaks of a possible entomophthoralean fungus in aphids on some vegetables in California, but there are no published reports of fungal outbreaks except for a study in the 1980s.  Mullens et al. (1987) reported E. muscae epizootics in house fly (Musca domestica), little house fly (Fannia canicularis), and predatory fly (Ophyra aenescens) populations in Southern California poultry facilities.  Similarly, E. muscae infections in adult onion fly (Delia antiqua) and seed corn maggot (D. platura) caused significant population reductions in Michigan (Carruthers et al., 1985).  In a recent study in North Carolina, E. muscae infected both cabbage maggot (D. radicum) and a predatory fly (Coenosia tigrina). 

The extent of E. muscae epizootics in Mississippi populations of SWD show promise for the natural control of this pest.  While large scale in vitro production of the fungus may not be practical at this moment, small scale production in vivo or a specialized culture medium is possible for laboratory and greenhouse studies.  In vivo culturing of entomophthoralean fungi and releasing infected live arthropods was successful for a large scale release of Neozygites tanajoa for controlling the cassava green mite, Mononychellus tanajoae in West Africa (Hountondji et al., 2002) and a small scale release of P. neoaphidis for controlling M. persicae in Virginia (Dara and Semtner, 2006).  Future studies will shed light on the potential of E. muscae in SWD integrated pest management. 


Brobyn, P. J. and N. Wilding.  1983.  Invasive and developmental processes of Entomophthora muscae infecting houseflies (Musca domestica).  Trans. Br. Mycol. Soc. 80: 1-8.

Carruthers, R., D. L. Haynes, and D. M. MacLeod.  1985.  Entomophthora muscae (Entomophthorales: Entomophthoraceae) mycosis in the onion fly, Delia antiqua (Diptera: Anthomyiidae).  J. Invertebr. Pathol. 45: 81-93.

Cuthbertson, A.G.S., D. A. Collins, L. F. Blackburn, N. Audsley, and H. A. Bell.  2014.  Preliminary screening of potential control products against Drosophila suzukii.  Insects 5: 488-498.

Dara, S. K. and P. J. Semtner.  2001.  Incidence of Pandora neoaphidis (Zygomycetes: Entomophthorales) in the Myzus persicae (Sulzer) complex (Homoptera: Aphididae) on three species of Brassica in the fall and winter.  J. Entomol. Sci. 36: 152-161.

Dara, S. K. and P. J. Semtner.  2006.  Introducing Pandora neoaphidis (Zygomycetes: Entomophthorales) into populations of Myzus persicae ss. nicotianae (Homoptera: Aphididae) on flue-cured tobacco.  J. Agric. Urban Entomol. 22: 173-180.

Dara, S. K. and P. J. Semtner.  2007.  Within-plant distribution of Pandora neoaphidis (Zygomycetes: Entomophthorales) in populations of the tobacco-feeding form of Myzus persicae (Homoptera: Aphididae) on flue-cured tobacco.  J. Agric. Urban Entomol. 23: 65-76.

Eilenberg, J. and V. Michelsen.  1999.  Natural host range and prevalence of the genus Strongwellsea (Zygomycota: Entomophthorales) in Denmark.  J. Invertebr. Pathol. 73: 189-198.

Eilenberg, J., L. Thomsen, and A. B. Jensen.  2013.  A third way for entomophthoralean fungi to survive the winter: slow disease transmission between individuals of the hibernating host.  Insects 4: 392-403.

Gargani, E., F. Tarchi, R. Frosinini, G. Mazza, and S. Simoni.  2013.  Notes on Drosophila suzukii Matsumura (Diptera Drosophiliae): field survey in Tuscany and laboratory evaluation of organic products.  Redia 96: 85-90.

Goldstein, B.  1927.  An Empusa disease of Drosophila.  Mycologia 19: 97-109.

Gryganskyi, A. P., R. A. Humber, J. E. Stajich, B. Mullens, I. M. Anishchenko, and R. Vilgalys.  2013.  Sequential utilization of hosts from different fly families by genetically distinct, sympatric populations within the Entomophthora muscae species complex.  PLoS ONE 8(8): e71168.  Doi:10:1371/journal.pone.0071168.

Hajek, A. E. and J. S. Elkinton.  1991.  Entomophaga maimaiga panzootic in northeastern gypsy moth populations.  In: Gottschalk, Kurt W.; Twery, Mark J.; Smith, Shirley I., eds. Proceedings, U.S. Department of Agriculture interagency gypsy moth research review 1990; East Windsor, CT. Gen. Tech. Rep. NE-146. Radnor, PA: U.S. Department of Agriculture, Forest Service, Northeastern Forest Experiment Station: 45.

Haye, T., P. Girod, A.G.S. Cuthbertson, X. G. Wang, K. M. Daane, K. A. Hoelmer, C. Baroffio, J. P. Zhang, and N. Desneux.  2016.  Current SWD IPM tactics and their practitcal implementation in fruit crops across different regions around the world.  J. Pest Sci. 89: 643-651.

Hountondji, F.C.C., C. J. Lomer, R. Hanna, A. J. Cherry, and S. K. Dara.  2002.  Field evaluation of Brazilian isolates of Neozygites floridana (Entomophthorales: Neozygitaceae) for the microbial control of cassava green mite in Benin, West Africa. Biocon. Sci. Tech. 12: 361-370. 

Keller, S.  1987.  Observations on the overwintering of Entomophthora planchoniana.  J. Invertebr. Pathol. 50: 333-335.

Klingen, I., G. Waersted, and K. Westrum.  2008.  Overwintering and prevalence of Neozygites floridana (Zygomycetes: Neozygitaceae) in hibernating females of Tetranychus urticae (Acari: Tetranychidae) under cold climatic conditions in strawberries.  Exp. Appl. Acarol. 46: 231-245.

MacLeod, D. M., E. Müller Kögler, and N. Wilding.  1976.  Entomophthora species with E. muscae-like conidia. Mycologia 68: 1–29.

Mullens, B. A., J. L. Rodriguez, and J. A. Meyer.  1987.  An epizootiological study of Entomophthora muscae in muscoid fly populations on Southern California poultry facilities.  Hilgardia 55: 1-41.   

Renkema, J. M., Z. Tefer, T. Gariepy, and R. H. Hallett.  2015.  Dalotia coriaria as a predator of Drosophila suzukii: functional responses, reduced fruit infestation and molecular diagnostics.  Biol. Control 89: 1-10.

Steinkraus, D. C., R. Hollingsworth, and P. H. Slaymakeh.  1995.  Prevalence off Neozygites fresenii (Entomophthorales: Neozygitaceae) on cotton aphids (Homoptera: Aphididae) in Arkansas cotton.  Environ. Entomol. 24: 465-474.

Steinkraus, D. C. and J. P. Kramer.  1987.  Susceptibility of sixteen species of Diptera to the fungal pathogen Entomophthora muscae (Zygomycetes: Entomophthoraceae).  Mycopathologia 100: 55-63.

Woltz, J. M., K. M. Donahue, D. J. Bruck, and J. C. Lee.  2015.  Efficacy of commercially available predators, nematodes, and fungal entomopathogens for augmentative control of Drosophila suzukii.  J. Appl. Entomol. 139: 759-770.


Posted on Wednesday, September 6, 2017 at 3:16 PM
  • Author: Surendra K. Dara, UC Cooperative Extension
  • Author: Tom Mann, Mississippi Museum of Natural Science, Jackson, MS
  • Author: De-Wei Li, Connecticut Agricultural Experiment Station, Windsor, CT
  • Author: Blake Layton, Mississippi State University, Mississippi State, MS
  • Author: Richard Cowles, Connecticut Agricultural Experiment Station, Windsor, CT
  • Author: Blair Sampson, USDA-ARS, Poplarville, MS

IPM-based production for food safety, sustainability, and security

Santa Maria strawberry grower, Dave Peck

Different people have defined sustainable agriculture or food production in different ways.  In general, sustainable food production refers to the farming systems that maintain productivity indefinitely through ecologically balanced, environmentally safe, socially acceptable, and economically viable practices.  It is a system that ensures food security for the growing population of the world by taking science, economics, human and environmental health, and social aspects into consideration.

Agriculture has evolved over thousands of years from subsistence farming meeting the needs of individual families to agribusiness catering to the needs of consumers around the world.  Arthropod pests, diseases, and weeds (hereafter referred to as pests) have been an issue all along, but their management went through cyclical changes.  Pest management initially started by using naturally available materials such as sulfur or plant-based pyrethrums that gradually evolved into using toxic natural or synthetic compounds.  While pesticide use improved farm productivity and food affordability, indiscriminate use of synthetic broad-spectrum pesticides in the mid-1900s led to serious environmental and human health issues.  Pesticide use regulations, the discovery of safer pesticides, and new non-chemical alternatives, in the past few decades, have improved pest management practices to some extent.  Newer pesticides are also relatively less toxic to the environment. However, large quantities of synthetic chemical pesticides are still used in conventional farms along with other control options for managing a variety of pests to prevent yield losses and optimize returns.  Lack of good agricultural practices or IPM awareness has also contributed to the excessive use of chemicals and the associated risk of resistance in pests and environmental contamination in some areas.  For example, in some developing countries, or countries where pesticide use is not strictly regulated, highly toxic pesticides are used very close to the harvest date, causing serious health risks for consumers. 

Under these circumstances, in recent years, consumer preference for chemical-free food gave impetus for organic production; thus, the acreage of organically produced fruits, vegetables, and nuts has been gradually increasing.  Many stores now promote and sell fresh or processed organic foods, at premium prices, to those who can afford them.  While organic farming is generally considered more challenging and less productive, growers are willing to take the risk as they try to meet the market demand and produce organically.  However, managing weeds in organic farms continues to be a labor-intensive and expensive part of production.  The labor shortage in many areas exacerbates manual weed control.  In some crop and pest situations, control of pests with organically acceptable tools is not sufficient.  Unmanaged pest populations can spread to other areas and/or crops, cause higher yield losses, and indirectly contribute to higher pesticide use on neighboring conventional farms.

Jimmy Klick (Driscoll's) and Sanjay Kumar Rajpoot (Rajpoot Industries and FarmX) with Todd Fitchette (Western Farm Press) in the background at the Santa Maria Strawberry Field Day in 2016

On the other hand, IPM offers an effective, practical, and sustainable solution where excessive use of chemical pesticides is limited, pest populations are effectively managed, and returns are optimized without having a negative impact on the environment.  IPM is an approach where host plant resistance (selection of resistant cultivars), modification of planting dates, crop density, irrigation and nutrient management or use of trap crops (cultural control), conservation or augmentation of natural enemies (biological control), pheromones for mating disruption or to attract and kill (behavioral control), traps, netting, and vacuums (mechanical control), chemicals from various mode of action groups (chemical control), plant extracts (botanical control), and entomopathogens or their derivatives (microbial control) are used in a balanced manner.  It is a comprehensive approach where all available strategies are considered to achieve pest control with minimal impact on the ecosystem.  However, many consumers are not aware of the difference between organic and conventional practices or IPM strategies.  Many perceive organic farming as a pesticide-free production system and as the only alternative to conventional farming with synthetic chemicals and nutrients.  Organic farming also uses pesticides, fertilizers, and hormones of natural origin.  For example, potassium salts of fatty acids are used against insects, mites, and fungal diseases.  Mined sulfur is used as a miticide and fungicide.  Popular organic insecticides, based on pyrethrins extracted from Chrysanthemum cinerariaefolium flowers, are very toxic to natural enemies, honey bees, and fish although they are less stable in the environment than synthetic pyrethroids.  The bacterium, Bacillus thuringiensis, which is the source of the toxic insecticidal protein in genetically modified corn, cotton, soybean, and other crops, is widely used in organic farming for managing lepidopteran pests.  Organic produce is also perceived to be healthier than conventional produce although several studies showed that there was no such difference.  A thorough understanding of conventional, organic, and IPM-based production could influence consumers' preference and allows them to make informed, practical, and science-based decisions.

IPM encourages the use of all available control options in a manner that maintains productivity without compromising environmental and human safety.  IPM-based food production can be a better alternative than organic production for various reasons (Table 1).  While several growers already adopt IPM practices, an IPM label or seal can authenticate the production system.

Table 1. Comparison of various food production systems

Since pest control efficacy, productivity, and operational costs are optimized for affordable food production without compromising health aspects, an IPM-based/branded food production system, which utilizes both modern and traditional technologies, might offer a better alternative to the organic system.  IPM-based production allows the use of chemical pesticides to address critical pest issues when needed, without losing the focus on environmental safety and sustainability.  Agriculture is a global enterprise and California agriculture leads and influences farming practices around the world.  While food production with an organic seal can continue, shifting to production with an IPM seal might be a practical and sustainable approach. 

Additional reading:

Dara, S. K.  2015.  Producing with the seal of IPM is a practical and sustainable strategy for agriculture.  UCANR eJournal Strawberries and Vegetables.

Gold, M. V.  2007.  Sustainable agriculture: definitions and terms. USDA-NAL, Beltsville, MD.

NPIC. 2014.  Pyrethrins general fact sheet.

Unsworth J.  2010. History of pesticide use.

Posted on Wednesday, July 19, 2017 at 2:17 PM

Entomopathogenic microorganisms: modes of action and role in IPM

Entomopathogens are microorganisms that are pathogenic to arthropods such as insects, mites, and ticks.  Several species of naturally occurring bacteria, fungi, nematodes, and viruses infect a variety of arthropod pests and play an important role in their management.  Some entomopathogens are mass-produced in vitro (bacteria, fungi, and nematodes) or in vivo (nematodes and viruses) and sold commercially.  In some cases, they are also produced on small scale for non-commercial local use.  Using entomopathogens as biopesticides in pest management is called microbial control, which can be a critical part of integrated pest management (IPM) against several pests.

Some entomopathogens have been or are being used in a classical microbial control approach where exotic microorganisms are imported and released for managing invasive pests for long-term control.  The release of exotic microorganisms is highly regulated and is done by government agencies only after extensive and rigorous tests.  In contrast, commercially available entomopathogens are released through inundative application methods as biopesticides and are commonly used by farmers, government agencies, and homeowners.  Understanding the mode of action, ecological adaptations, host range, and dynamics of pathogen-arthropod-plant interactions is essential for successfully utilizing entomopathogen-based biopesticides for pest management in agriculture, horticulture, orchard, landscape, turf grass, and urban environments.

Entomopathogen groups

Important entomopathogen groups and the modes of their infection process are described below.


There are spore-forming bacterial entomopathogens such as Bacillus spp., Paenibacillus spp., and Clostridium spp, and non-spore-forming ones that belong to the genera Pseudomonas, Serratia, Yersinia, Photorhabdus, and Xenorhabdus. Infection occurs when bacteria are ingested by susceptible insect hosts.  Pseudomonas, Serratia and Yersinia are not registered in the USA for insect control.Several species of the soilborne bacteria, Bacillus and Paenibacillus are pathogenic to coleopteran, dipteran, and lepidopteran insects.  Bacillus thuringiensis subsp. aizawai, Bt subsp. kurstaki, Bt subsp. israelensis, Bt subsp. sphaericus, and Bt subsp. tenebrionis are effectively used for controlling different groups of target insects.  For example, Bt subsp. aizawai and Bt subsp. kurstaki are effective against caterpillars, Bt subsp. israelensis and Bt subsp. sphaericus target mosquito larvae, and Bt subsp. tenebrionis is effective against some coleopterans.

When Bt is ingested, alkaline conditions in the insect gut (pH 8-11) activate the toxic protein (delta-endotoxin) that attaches to the receptors sites in the midgut and creates pore in midgut cells.  This leads to the loss of osmoregulation, midgut paralysis, and cell lysis.  Contents of the gut leak into insect's body cavity (hemocoel) and the blood (hemolymph) leaks into the gut disrupting the pH balance.  Bacteria that enter body cavity cause septicemia and eventual death of the host insect.  Insects show different kinds of responses to Bt toxins depending on the crystal proteins (delta-endotoxin), receptor sites, production of other toxins (exotoxins), and requirement of spore.  The type responses below are based on the susceptibility of caterpillars to Bt toxins.

Type I response – Midgut paralysis occurs within a few minutes after delta-endotoxin is ingested.  Symptoms include cessation of feeding, increase in hemolymph pH, vomiting, diarrhea, and sluggishness.  General paralysis and septicemia occur in 24-48 hours resulting in the death of the insect.  Examples of insects that show Type I response include silkworm, tomato hornworm, and tobacco hornworm.

Type II response – Midgut paralysis occurs within a few minutes after the ingestion of delta-endotoxin, but there will be no general paralysis.  Septicemia occurs within 24-72 hours.  Examples include inchworms, alfalfa caterpillar, and cabbage butterfly.

Type III response – Midgut paralysis occurs after delta-endotoxin is ingested followed by cessation of feeding.  Insect may move actively as there will be no general paralysis.  Mortality occurs in 48-96 hours.  Higher mortality occurs if spores are ingested.  Insect examples include Mediterranean flour moth, corn earworm, gypsy moth, spruce budworm.

Type IV responseInsects are naturally resistant to infection and older instars are less susceptible than the younger ones.  Midgut paralysis occurs after delta-endotoxin is ingested followed by cessation of feeding.  Insect may move actively as there will be no general paralysis.  Mortality occurs in 72-96 or more hours.  Higher mortality occurs if spores are ingested.  Cutworms and armyworms are examples for this category.

Unlike caterpillars, the response in mosquitoes is different where upon ingestion of Bt subsp. israelensis delta-endotoxin, the mosquito larva is killed within 20-30 min.

While Bt with its toxic proteins is very effective as a biopesticide against several pests, excessive use can lead to resistance development.  Corn earworm, diamondback moth, and tobacco budworm are some of the insects that developed resistance to Bt toxins.  Genetic engineering allowed genes that express Bt toxins to be inserted into plants such as corn, cotton, eggplant, potato, and soybean and reduced the need to spray pesticides.  However, appropriate management strategies are necessary to reduce insect resistant to Bt toxins in transgenic plants.

Paenibacillus popilliae is commonly used against Japanese beetle larvae and known to cause the milky spore disease.  Although Serratia is not registered for use in the USA, a species is registered for use against a pasture insect in New Zealand.  In the case of Photorhabdus spp. and Xenorhabdus spp., which live in entomopathogenic nematodes symbiotically, bacteria gain entry into the insect host through nematodes.  Biopesticides based on heat-killed Chromobacterium subtsugae and Burkholderia rinojensis are reported to have multiple modes of action and target mite and insect pests of different orders.


Entomopathogenic fungi typically cause infection when spores come in contact with the arthropod host.  Under ideal conditions of moderate temperatures and high relative humidity, fungal spores germinate and breach the insect cuticle through enzymatic degradation and mechanical pressure to gain entry into the insect body.  Once inside the body, the fungi multiply, invade the insect tissues, emerge from the dead insect, and produce more spores.  Natural epizootics of entomophthoralean fungi such as Entomophaga maimaiga (in gypsy moth), Entomophthora muscae (in flies), Neozygites fresenii (in aphids), N. floridana (in mites), and Pandora neoaphidis (in aphids) are known to cause significant reductions in host populations.  Although these fastidious fungi are difficult to culture in artificial media and do not have the potential to be sold as biopesticides they are still important in natural control of some pest species.  Hypoclealean fungi such as Beauveria bassiana, Isaria fumosorosea, Hirsutella thompsonii, Lecanicillium lecanii, Metarhizium acridum, M. anisopliae, and M. brunneum, on the other hand, are commercially sold as biopesticides in multiple formulations around the world.  Fungal pathogens have a broad host range and are especially suitable for controlling pests that have piercing and sucking mouthparts because spores do not have to be ingested.  However, entomopathogenic fungi are also effective against a variety of pests such as wireworms and borers that have chewing mouthparts.

Related to fungi, the spore-forming microsporidium, Paranosema (Nosema) locustae is a pathogen that has been used for controlling locusts, grasshoppers, and some crickets.  When P. locustae is ingested, the midgut tissues become infected, followed by infection in the fat body tissues.  The disease weakens and eventually kills the orthopteran host within a few weeks.

Various insects killed by different species of entomopathogenic fungi


Entomopathogenic nematodes are microscopic, soil-dwelling worms that are parasitic to insects.  Several species of Heterorhabditis and Steinernema are available in multiple commercial formulations, primarily for managing soil insect pests.  Infective juveniles of entomopathogenic nematodes actively seek out their hosts and enter through natural openings such as the mouth, spiracles, and anus or the intersegmental membrane.  Once inside the host body, the nematodes release symbiotic bacteria that kill the host through bacterial septicemia.  Heterorhabditis spp. carry Photorhabdus spp. bacteria and Steinernema spp. carry Xenorhabdus spp. bacteria.  Phasmarhabditis hermaphrodita is also available for controlling slugs in Europe, but not in the USA.

Infective juvenile of Steinernema carpocapsae entering the first instar larva of a leafminer through its anus.

Nematodes in beet armyworm pupa (left) and termite worker (right).


Similar to bacteria, entomopathogenic viruses need to be ingested by the insect host and therefore are ideal for controlling pests that have chewing mouthparts.  Several lepidopteran pests are important hosts of baculoviruses including nucleopolyhedroviruses (NPV) and granuloviruses (GV).  These related viruses have different types of inclusion bodies in which the virus particles (virions) are embedded.  Virus particles invade the nucleus of the midgut, fat body or other tissue cells, compromising the integrity of the tissues and liquefying the cadavers.  Before death, infected larvae climb higher in the plant canopy, which aids in the dissemination of virus particles from the cadavers to the lower parts of the canopy.  This behavior aids in the spread of the virus to cause infection in healthy larvae.  Viruses are very host specific and can cause significant reduction of host populations.  Examples of some commercially available viruses include Helicoverpa zea single-enveloped nucleopolyhedrovirus (HzSNVP), Spodoptera exigua multi-enveloped nucleopolyhedrovirus (SeMNPV), and Cydia pomonella granulovirus (CpGV).

Most entomopathogens typically take 2-3 days to infect or kill their host except for viruses and P. locustae which take longer.  Compared to viruses (highly host specific) and bacteria (moderately host specific), fungi generally have a broader host range and can infect both underground and aboveground pests.  Because of the soil-dwelling nature, nematodes are more suitable for managing soil pests or those that have soil inhabiting life stages.

Biopesticides based on various entomopathogenic microorganisms and their target pests

Microbial control and Integrated Pest Management

There are several examples of entomopathogen-based biopesticides that have played a critical role in pest management.  Significant reduction in tomato leaf miner, Tuta absoluta, numbers and associated yield loss was achieved by Bt formulations in Spain (Gonzalez-Cabrera et al, 2011).  Bt formulations are also recommended for managing a variety of lepidopteran pests on blueberry, grape, and strawberry (Haviland, 2014; Zalom et al, 2014; Bolda and Bettiga, 2014; Varela et al, 2015).

Lecanicellium muscarium-based formulation reducedgreenhouse whitefly (Trialeurodes vaporariorum) populations by 76-96% in Mediterranean greenhouse tomato (Fargues et al, 2005).  In other studies, B. bassiana applications resulted in a 93% control of twospotted spider mite (Tetranychus urticae) populations in greenhouse tomato (Chandler et al, 2005) and 60-86% control on different vegetables (Gatarayiha et al, 2010).  The combination of B. bassiana and azadirachtin reduced rice root aphid (Rhopalosiphum rufiabdominale) and honeysuckle aphid (Hyadaphis foeniculi) populations by 62% in organic celery in California (Dara, 2015a).  Chromobacterium subtsugae and B. rinojensis caused a 29 and 24% reduction, respectively, in the same study.  IPM studies in California strawberries also demonstrated the potential of entomopathogenic fungi for managing the western tarnished plant bug (Lygus hesperus) and other insect pests (Dara, 2015b, 2016).  Entomopathogenic fungi also have a positive effect on promoting drought tolerance or plant growth as seen in cabbage (Dara et al, 2016) and strawberry (Dara, 2013) and antagonizing plant pathogens (Dara et al, 2017)

Application of SeMNPV was as efficacious as methomyl and permithrin in reducing beet armyworms (S. exigua) in head lettuce in California (Gelernter et al, 1986).  Several studies demonstrated PhopGV as an important tool for managing the potato tubermoth (Phthorimaea operculella) (Lacey and Kroschel, 2009).

The entomopathogenic nematode, S. feltiae,reduced raspberry crown borer (Pennisetia marginata) populations by 33-67% (Capinera et al, 1986).  For managing the branch and twig borer (Melagus confertus) in California grapes, S. carpocapsae is one of the recommended options (Valera et al, 2015).

Entomopathogens can be important tools in IPM strategies in both organic and conventional production systems.  Depending on the crop, pest, and environmental conditions, entomopathogens can be used alone or in combination with chemical, botanical pesticides or other entomopathogens. 

Acknowledgements: Thanks to Dr. Harry Kaya for reviewing this article.


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Dara, S. K. 2015b. Integrating chemical and non-chemical solutions for managing lygus bug in California strawberries.  CAPCA Adviser 18 (1) 40-44.

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Dara, S. K., S.S.R. Dara, and S.S. Dara.  2016.  First report of entomopathogenic fungi, Beauveria bassiana, Isaria fumosorosea, and Metarhizium brunneum promoting the growth and health of cabbage plants growing under water stress.  UCANR eJournal Strawberries and Vegetables, 19 September, 2016. (

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Gelernter, W. D., N. C. Toscano, K. Kido, and B. A. Federici.  1986.  Comparison of a nuclear polyhedrosis virus and chemical insecticides for control of the beet armyworm (Lepidopter: Noctuidae) on head lettuce.  J. Econ. Entomol. 79: 714-717.

González-Cabrera, J., J. Mollá, H. Monton, A. Urbaneja. 2011. Efficacy of Bacillus thuringiensis (Berliner) in controlling the tomato borer, Tuta absoluta (Meyrick) (Lepidoptera: Gelechiidae). BioControl 56: 71–80.

Haviland, D. R.  2014. UC IPM Pest Management Guidelines: Blueberry.  UC ANR Pub. 3542.

Lacey, L. A. and J. Kroschel.  2009.  Microbial control of the potato tuber moth (Lepidoptera: Gelechiidae).  Fruit Veg. Cereal Sci. Biotechnol. 3: 46-54.

Varela, L. G., D. R. Haviland, W. J., Bentley, F. G. Zalom, L. J. Bettiga, R. J. Smith, and K. M. Daane.  2015. UC IPM Pest Management Guidelines: Grape.  UC ANR Pub. 3448.

Zalom, F. G., M. P. Bolda, S. K. Dara, and S. Joseph.  2014. UC IPM Pest Management Guidelines: Strawberry.  UC ANR Pub. 3468. 

Posted on Saturday, May 20, 2017 at 7:20 AM

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