- Author: Surendra K. Dara
Use of biopesticides or non-chemical pesticides is encouraged as a part of integrated pest management (IPM) for environmental and human safety and to reduce the risk of insecticide resistance. With the increase in biopesticide use in both organic and conventional cropping systems, it is a good time to review the potential of insect resistance to botanical and microbial pesticides.
Insects and mites develop resistance to chemical pesticides through genetic, metabolic, or behavioral changes resulting in reduced penetration of toxin, increased sequestration or excretion, reduced binding to the target site, altered target site that prevents binding of the toxin, or reduced exposure to the toxin through modified behavior. When the active ingredient is a toxic molecule and has the mode of action similar to that of a chemical compound, regardless of the plant or microbial origin, arthropods are more likely to develop resistance through one or more of the abovementioned mechanisms. When the mode of action is infection by a microorganism, rather than a toxin, arthropods are less likely to develop resistance. Under natural circumstances, plants, insects, natural enemies, and beneficial or harmful microbes continuously co-evolve and adapt to changing environment. When there is a higher selection pressure, such as indiscriminate use of chemical pesticides, increased mutagenesis can lead to resistance issues. A good understanding of insect resistance to biopesticides will help minimize potential risks and improve their efficient use in IPM.
Resistance to botanical pesticides
Nicotine, an alkaloid from Nicotiana spp., is one of the earlier botanical pesticides known. Although nicotine is not currently used as an insecticide, its synthetic alternatives – neonecotinoids – are commonly used against several pests. Botanical insecticide pyrethrum, extracted from the flowers of Chrysanthemum cinerariaefolium, contains insecticidal pyrethrins (synthetic pyrethrins are referred to as pyrethroids). Although insect resistance to pyrethrum or pyrethroid compounds has been known (Whitehead, 1959; Immaraju et al., 1992; Glenn et al., 1994), they have been effectively used against a number of pests through careful placement in IPM, organic, or conventional management strategies. Additionally, pyrethrin products have been effectively used along with piperonyl butoxide, which acts as a synergist and resistance breaker (Gunning et al. 2015).
Another botanical insecticidal compound, azadirachtin, is a tetranortriterpenoid limonoid from neem (Azadirachta indica) seeds, which acts as an insecticide, antifeedant, repellent and insect growth regulator. While neem oil, which has a lower concentration of azadirachtin, has been used in the United States as a fungicide, acaricide, and insecticide for a long time, several azadirachtin formulations in powder and liquid forms have become popular in recent years and were found effective in managing important pests (Dara 2015a and 2016). Feng and Isman (1995) reported that the green peach aphid, Myzus persicae developed resistance to pure azadirachtin under artificially induced selection pressure after 40 generations, but did not develop resistance to a refined neem seed extract. They suggested that natural blend of azadirachtin compounds in a biopesticide would not exert selection pressure that could lead to resistance. Additionally, Mordue and Nisbet (2000) discussed that azadirachtin can play a role in insecticide resistance management because it reduces the detoxification enzyme production as a protein synthesis inhibitor. Azadirachtin also improved the efficacy of other biopesticides in multiple studies (Trisyono and Whalon, 2000; Dara, 2013 and 2015b).
Insects feeding on plant allelochemicals can develop cross-resistance to insecticides (Després et al., 2007). For example, overproduction of detoxification enzymes such as glutathione S-transferases and monooxygenases in the fall armyworm, Spodoptera frugiperda,when it fed on corn and cowpea, respectively, imparted cross-resistance to various chemical pesticides. It is important to keep this in mind when botanical pesticides are used to detect potential resistance issues.
Resistance to bacterial biopesticides
Bacillus thuringiensis (Bt)is a gram-positive soil bacterium, which contains crystalline toxic protein that is activated upon ingestion by an insect host, binds to the receptor sites in the midgut, and eventually causes insect death. Since the mode of action involves a toxin rather than bacterial infection, several insects developed resistance to Bt pesticides or transgenic crops that contain Bt toxins (Tabashnik et al., 1990; McGaughey and Whalon, 1992; Tabashnik, 1994; Iqbal et al., 1996). However, Bt pesticides are still very popular and used against a variety of lepidopteran (Bt subsp. aizawai and Bt subsp. kurstaki), dipteran (Bt subsp. israelensis and Bt subsp. sphaericus), and coleopteran (Bt subsp. tenebrionis) pests.
Spinosad is a mixture of macrocyclic lactones, spinosyns A and spinosyns D, derived from Saccharopolyspora spinosa, an actinomycete gram-positive bacterium, and is used against dipteran, hymenopteran, lepidopteran, thysanopteran, and other pests. Spinosad products, while naturally derived are registered as chemical pesticides, not as biopesticides. Insect resistance to spinosad later led to the development of spinetoram, which is a mixture of chemically modified spinosyns J and L. Both spinosad and spinetoram are contact and stomach poisons and act on insect nervous system by continuous activation of nicotinic acetylcholine receptors. However, insect resistance to both spinosad (Sayyed et al., 2004; Bielza et al., 2007) and spinetoram (Ahmad and Gull, 2017) has been reported due to extensive use of these pesticides. Cross-resistance between spinosad and some chemical insecticides has also occurred in some insects (Mota-Sanchez et al., 2006; Afzal and Shad, 2017).
Resistance to viral biopesticides
Baculovirus infections in lepidoptera have been known for centuries, especially in silkworms. Currently, there are several commercial formulations of nucleopolyhedroviruses (NPV) and granuloviruses (GV). When virus particles are ingested by the insect host, usually lepidoptera, they invade the nucleii of midgut, fatbody, or other tissue cells and kill the host. Baculoviruses are generally very specific to their host insect species and can be very effective in bringing down the pest populations. However, variations in the susceptibility of certain insect populations and development of resistant to viruses has occurred in several host species (Siegwart et al., 2015). Resistance to different isolates of Cydia pomonella granulovirus (CpGV-M, CpGV-S) in codling moth (Cydia pomonella) populations is well known in Germany and other parts of Europe (Sauer et al., 2017a & b).
Resistance to fungal biopesticides
There are several fungi that infect insects and mites. Fungal infection starts when fungal spores come in contact with an arthropod host. First, they germinate and gain entry into the body by breaching through the cuticle. Fungus later multiplies, invades the host tissues, kills the host, and emerges from the cadaver to produce more spores. Entomophthoralean fungi such as Entomophthora spp., Pandora spp., and Neozygites spp. can be very effective in pest management through natural epizootics, but cannot be cultured in vitro for commercial scale production. Hypocrealean fungi such as Beauveria bassiana, Isarea fumosorosea, Metarhizium brunneum, and Verticillium lecanii,on the other hand, can be mass-produced in vitro and are commercially available. These fungi are comparable to broad-spectrum insecticides and are pathogenic to a variety of soil, foliar, and fruit pests of several major orders. Since botanical, bacterial, and viral biopesticides have insecticidal metabolites, proteins, or viral particles that have specific target sites and mode of action, insects have a higher chance of developing resistance through one or more mechanisms. Although fungi also have insecticidal proteins such as beauvericin in B. bassiana and I. fumosorosea and dextruxin in M. anisopliae and M. brunneum, their mode of action is more through fungal infection and multiplication and arthropods are less prone to developing resistance to entomopathogenic fungi. However, insects can develop resistance to entomopathogenic fungi through increased melanism, phenoloxidase activity, protease inhibitor production, and antimicrobial and antifungal peptide production (Wilson et al., 2001; Zhao et al., 2012; Dubovskiy et al., 2013). It appears that production of detoxification enzymes in insects against fungal infections can also impart resistance to chemical pesticides. Infection of M. anisopliae in the larvae of greater wax moth, Galleria mellonella, increased dexotification enzyme activity and thus resistance to malathion (Serebrov et al., 2006).
These examples show that insects can develop resistance to biopesticides in a manner somewhat similar to chemical pesticides, but due to the typically more complex and multiple modes of action, at a significantly lesser rate depending on the kind of botanical compound or microorganism involved. Resistance to entomopathogenic fungi is less common than with other entomopathogens. Since biopesticide use is not as widespread as chemical pesticides, the risk of resistance development is less for the former. However, excessive use of any single tool has the potential for resistance or other issues and IPM, which uses a variety of management options, is always a good strategy.
Acknowledgements: Thanks to Pam Marrone for reviewing the manuscript.
Afzal, M.B.S. and S. A. Shad. 2017. Spinosad resistance in an invasive cotton mealybug, Phenacoccus solenopsis: cross-resistance, stability and relative fitness. J. Asia-Pacific Entomol. 20: 457-462.
Ahmad M. and S. Gull. 2017. Susceptibility of armyworm Spodoptera litura (Lepidoptera: Noctuidae) to novel insecticides in Pakistan. The Can. Entomol. 149: 649-661.
Bielza, P., V. Quinto, E. Fernández, C. Grávalos, and J. Contreras. 2007. Genetics of spinosad resistance in Frankliniella occidentalis (Thysanoptera: Thripidae). J. Econ. Entomol. 100: 916-920.
Dara, S. K. 2013. Strawberry IPM study 2012: managing insect pests with chemical, botanical, and microbial pesticides. UCANR eJournal Strawberries and Vegetables March 13, 2013 (http://ucanr.edu/blogs/blogcore/postdetail.cfm?postnum=9595).
Dara, S. K. 2015a. Root aphids and their management in organic celery. CAPCA Adviser 18(5): 65-70.
Dara, S. K. 2015b. Strawberry IPM study 2015: managing insect pests with chemical, botanical, microbial, and mechanical control options. UCANR eJournal Strawberries and Vegetables November 30, 2015 (http://ucanr.edu/blogs/blogcore/postdetail.cfm?postnum=19641).
Dara, S. K. 2016. Managing strawberry pests with chemical pesticides and non-chemical alternatives. Intl. J. Fruit Sci. https://doi.org/10.1080/15538362.2016.1195311
Després, L., J.-L. David, and C. Gallet. 2007. The evolutionary ecology of insect resistance to plant chemicals. Trends in Ecol. Evol. 22: 298-307.
Dubovskiy, I. M., M.M.A. Whitten, O. N. Yaroslavtseva, C. Greig, V. Y. Kryukov, E. V. Grizanova, K. Mukherjee, A. Vilcinskas, V. V. Glupov, and T. M. Butt. 2013. Can insects develop resistance to insect pathogenic fungi? PloS one 8: e60248.
Feng, R. and M. B. Isman. 1995. Selection for resistance to azadirachtin in the green peach aphid, Myzus persicae. Experientia 51: 831-833.
Glenn, D. C., A. A. Hoffmann, and G. McDonald. 1994. Resistance to pyrethroids in Helicoverpa armigera (Lepidoptera: Noctuidae) from corn: adult resistance, larval resistance, and fitness effects. J. Econ. Entomol. 87: 1165-1171.
Gunning, R., G. Moores, and M. Balfe. 2015. Novel use of pyrethrum to control resistant insect pests on cotton. Acta Hort. 1073: 113-118. https://doi.org/10.17660/ActaHortic.2015.1073.16
Immaraju, J. A., T. D. Paine, J. A. Bethke, K. L. Robb, and J. P Newman. 1992. Western flower thrips (Thysanoptera: Thripidae) resistance to insecticides in coastal California greenhouses. J. Econ. Entomol. 85: 9-14.
Iqbal, K., R.H.J. Vekerk, M. J. Furlong, P. C. Ong, S. A. Rahman, and D. J. Wright. 1996. Evidence for resistance to Bacillus thuringiensis (Bt) subsp. kurstaki HD-1, Bt subsp. aizawai and Abamectin in field populations of Plutella xylostella from Malaysia. Pest Manag. Sci. 48: 89-97.
McGaughey, W. H. and M. E. Whalon. 1992. Managing insect resistance to Bacillus thuringiensis toxins. Science 258: 1451-1455.
Mordue, A. J. and A. J. Nisbet. 2000. Azadirachtin from the neem tree Azadirachta indica: its action against insects. An. Soc. Entomol. Bras. 29: 615-632.
Mota-Sanchez, D., R. M. Hollingworth, E. J. Grafius, and D. D. Moyer. 2006. Resistance and cross-resistance to neonicotinoid insecticides and spinosad in the Colorado potato beetle, Leptinotarsa decemlineata (Say) (Coleoptera: Chrysomelidae). Pest Manag. Sci. 62: 30-37.
Sauer, A. J., E. Fritsch, K. Undorf-Spahn, P. Nguyen, F. Marec, D. G. Heckel, and J. A. Jehle. 2017a. Novel resistance to Cydia pomonella granulovirus (CpGV) in codling moth shows autosomal and dominant inheritance and confers cross-resistance to different CpGV genome groups. PLoS ONE 12(6): e0179157.
Sauer, A. J., S. Schulze-Bopp, E. Fritsch, K. Undorf-Spahn, and J. A. Jehle. 2017b. A third type of resistance of codling moth against Cydia pomonella granulovirus (CpGV) shows a mixture of a Z-linked and autosomal inheritance pattern. Appl. Environ. Microbiol. AEM-01036.
Sayyed, A. H., D. Omar, and D. J. Wright. 2004. Genetics of spinosad resistance in a multi-resistant field-selected population of Plutella xylostella. Pest Manag. Sci. 60: 827-832.
Serebrov, V. V., O. N. Gerber, A. A. Malyarchuk, V. V. Martemyanov, A. A. Alekseev, and V. V. Glupov. Effect of entomopathogenic fungi on detoxification enzyme activity in greater wax moth Galleria mellonella L. (Lepidoptera, Pyralidae) and role of detoxification enzymes in development of insect resistance to entomopathogenic fungi. Anim. Human Physiol. 33: 581-586.
Siegwart, M., B. Graillot, C. B. Lopez, S. Besse, M. Bardin, P. C. Nicot, and M. Lopez-Ferber. 2015. Resistance to bio-insecticides or how to enhance their sustainability: a review. Front. Plant Sci. 6:381. https://doi.org/10.3389/fpls.2015.00381
Tabashnik, B. 1994. Evolution of resistance to Bacillus thuringiensis. Annu. Rev. Entomol. 39: 47-79.
Tabashnik, B. E., N. L. Cushing, N. Finson, and M. W. Johnson. 1990. Field development of resistance to Bacillus thuringiensis in diamondback moth (Lepidoptera: Plutellidae). J. Econ. Entomol. 83: 1671-1676.
Trisyono, A. and M. E. Whalon. 2000. Toxicity of neem applied alone and in combination with Bacillus thuringiensis to Colorado potato beetle (Coleoptera: Chrysomelidae). J. Econ. Entomol. 92: 1281-1288.
Whitehead, G. B. 1959. Pyrethrum resistance conferred by resistance to DDT in the blue tick. Nature 184: 378-379.
Wilson K., S. C. Cotter, A. F. Reeson, and J. K. Pell. 2003. Melanism and disease resistance in insects. Ecology Letters 4: 637-649.
Zhao, P., Z. Dong, J. Duan, G. Wang, L. Wang, Y. Li, Z. Xiang, and Q. Xia. 2012. Genome-wide identification and immune response analysis of serine protease inhibitor genes in the silkworm, Bombyx mori. PloS one 7: e31168.
- Author: Surendra K. Dara
- Author: David Peck, Manzanita Berry Farms
Under the soil is a complex and dynamic world of moisture, pH, salinity, nutrients, microorganisms, and plant roots along with pests, pathogens, weeds and more. A good balance of essential nutrients, moisture, and beneficial microorganisms provides optimal plant growth and yield. These factors also influence natural plant defenses and help withstand stress caused by biotic and abiotic factors.
Several beneficial microbe-based products are commercially available to promote plant growth and improve health, yield potential and quality. Some of them improve nutrient and water absorption while others provide protection against plant pathogens or improve plant defense mechanism. In addition to the macronutrients such as nitrogen, phosphorus, and potassium, several micronutrients are critical for optimal growth and yield potential. Some of the micronutrient products are also useful in promoting beneficial microbes. Understanding the plant-microbe-nutrient interactions and how different products help crop production are helpful for making appropriate decisions.
Mycorrhizae (fungi of roots) establish a symbiotic relationship with plants and serve as an extended network of the root system. They facilitate improved uptake of moisture and nutrients resulting in better plant growth and yield (Amerian and Stewart, 2001; Wu and Zou, 2009; Bolandnazar et al., 2007; Nedorost et al., 2014). Mycorrhizae can also help absorb certain nutrients more efficiently than plants can and make them more readily available for the plant. With increased moisture and nutrient absorption, plants can become more drought-tolerant. Mycorrhizae also help plants to withstand saline conditions and protect from plant pathogens. A healthy root system can fight soil diseases and weed invasion. Additionally, mycorrhizae increase organic matter content and improve soil structure.
Considering an increasing need for fumigation alternatives to address soilborne pathogens in strawberry, mycorrhizae and other beneficial microbes could be potential tools in maintaining plant health. Additionally, recent studies suggest that entomopathogenic fungi such as Beauveria bassiana, Metarhizium brunneum, and Isaria fumosorosea form mycorrhiza-like and endophytic relationships with various species of plants and could help with plant growth and health (Behie and Bidochka, 2014; Dara et al., 2016). These fungi are currently used for pest management, but their interaction with plants is a new area of research. Understanding this interaction will potentially expand the use of the biopesticides based on these fungi for improving plant growth and health. A study was conducted at Manzanita Berry Farms, Santa Maria in fall-planted strawberry crop during the 2014-2015 production season to evaluate the impact of beneficial microbes on strawberry growth, health, mite infestations, powdery mildew, botrytis fruit rot, and yield.
List of treatments, their application rates and frequencies:
- Untreated control: Received no supplemental treatments other than standard grower practices.
- HealthySoil: NPK (0.1-0.1-0.1).
- BotaniGard ES: Entomopathogenic fungus Beauveria bassiana strain GHA. Rate - 1 qrt in 50 gal for a 30 min transplant dip and 1 qrt/ac every 15 days until January and once a month thereafter until April, 2015.
- Met52: Entomopathogenic fungus Metarhizium brunneum strain F52. Rate – 16 fl oz in 50 gal for a 30 min transplant dip and 16 fl oz/ac every 15 days until January and once a month thereafter until April, 2015.
- NoFly: Entomopathogenic fungus Isaria fumosorosea strain FE9901. Rate – 11.55 oz in 50 gal for a 30 min transplant dip and 11.55 oz/ac every 15 days until January and once a month thereafter until April, 2015.
- Actinovate AG: Beneficial soilborne bacterium Streptomyces lydicus WYEC 108. Rate – 6 oz in 50 gal for a 30 mintransplant dip and 6 oz/ac every month.
- TerraClean 5.0: Hydrogen dioxide and peroxyacetic acid. Rate – 1:256 dilution for a 1 min root dip followed by 2 gal/ac 10 days after planting and then 2 and 1 gal/ac alternated every 15 days until April, 2015.
- TerraGrow: Humic acids, amino acids, sea kelp, glucose based carriers, bacteria – Bacillus licheniformis, B. subtilis, B. pumilus, B. amyloliquefaciens, and B. magaterium, and mycorrhizae – Trichoderma harzianum and T. reesei. Rate – 1.13 g in 10 gal for a 1 min root dip followed by 1.5 lb/ac 10 days after planting and once every month until April, 2015.
- TerraCelan and TerraGrow: Same as individual treatments at the time of planting, but TerraClean at 2 gal/ac and TerraGrow at 1.5 lb/ac 10 days after planting followed by monthly treatments until April, 2015.
- O-MEGA: NPK (0.2-1.0-0.5), bacteria – Azotobacter chroococcum, Azospirillum lipoferum, Lactobacillus acidophilus, Pseudomonas fluorescens, Cellulomonas cellulans and the fungus Aspergillus niger. Rate – 20 ml in 1 gal sprinkled on transplants 30 min before planting followed by 1 qrt/ac every week rest of the season.
Strawberry transplants (variety BG-6.3024) were treated at the time of planting on 6 November, 2014 and treatments are also administered periodically through the drip irrigation system following the abovementioned schedule. Each treatment had two 330' long beds each with four rows of plants. Treatments were randomly arranged in two blocks and two sampling plots (20' long) were established within each bed in a block. The impact of the treatments on plant growth (canopy size), health, spider mite populations, botrytis and powdery mildew severity, and yield were monitored periodically. Plant growth was determined by measuring the canopy size. Plant health was rated on a scale of 0 to 5 where 0=dead, 1=weak, 2=moderate low, 3=moderate high, 4=good, and 5=very good. Powdery mildew severity was determined by observing leaf samples under microscope and rating the severity on a scale of 0 to 4 where 0=no infection, 1=1-25%, 2=26-50%, 3=51-75%, and 4=76-100% of leaf area with powdery mildew. Twenty plants or leaf samples per plot were used for these observations. To monitor botrytis fruit rot, a box of fruits from each plot were held at room temperature and disease was rated 3 and 5 days after harvest on a scale of 0 to 4 where 0=no infection, 1=1-25%, 2=26-50%, 3=51-75%, and 4=76-100% of fruit with botrytis. Yield data were also collected from the plots throughout the production season using grower's harvesting schedule. Mite counts were also taken periodically.
Data were analyzed using analysis of variance and significant means were separated using Tukey's HSD means separation test.
Treating the transplants with different treatment materials and planting in respective beds
Newsly transplanted experimental plots.
Chris Martinez (center, front row) and rest of the field crew at Manzanita Berry Farms
Canopy size: Significant differences (P = 0.002) among treatments were seen only on the first observation date on 26 January, 2015 where TerraClean-treated plants were smaller than some of the treatments. There were no significant differences (P > 0.05) in treatments on the following observations in February and March, however TerraClean-treated plants recovered and plants were larger in some of the treatments.
Size of the plant canopy on three observation dates.
Plant health: Treatments did not have a significant (P > 0.05) impact on plant health. Health ratings varied from 4.2 for TerraClean to 4.6 for untreated, BotaniGard, Actinovate, and O-Mega treatments in January. In February, TerraGrow-treated plants had 4.5 rating and BotaniGard and O-Mega treatments had 4.8. March ratings varied between 4.8 and 4.9 in all the treatments. As there were no soilborne diseases during the study period, the impact of the treatments could not be determined, which was the main objective of the study.
Plant health ratings on three observation dates.
Powdery mildew: Disease severity did not differ among treatments (P > 0.05) on 16 April and 16 June, but significant (P = 0.008) differences were observed on 26 June where BotaniGard-treated plants had the lowest. When data were compared for the three observation dates, severity rating varied from 1.8 for BotaniGard to 2.24 for TerraClean.
Powdery mildew severity on individual observation dates (top) and combined for three observations (bottom)
Botrytis fruit rot: There were no significant (P > 0.05) differences among treatments on any of the four observation dates or when data were combined for all observations. In general, fruit rot was less severe 3 days after harvest than 5 days after during the first three observation dates. When data were combined for the observation dates, HealthySoil treatment had a rating of 1 followed by Met52, NoFly, Actinovate, and TerraClean+TerraGrow with a 1.3 rating for 3 days after harvest.
Severity of botrytis fruit rot 3 and 5 days after harvest on individual observation dates (above) and when data were combined (below).
Spider mites: Mite populations were very low in all the plots during observation period and data were not included.
Fruit yield: While the seasonal yield of total, marketable, or unmarketable berries was not significantly (P > 0.05) different for any of the treatments marketable yields had a wider range than unmarketable yields among treatments. The lowest marketable fruit yield was seen in TerraClean (35.6 kg or 79.4 lb) and HealthySoil (35.8 kg or 79.8 lb) while the highest yield was seen in Actinovate (40.1 kg or 89.4 lb) followed by untreated control (39.4 kg or 87.9 lb), O-Mega (39.3 kg or 87.6 lb), Met52 (39.2 kg or 87.4 lb), and NoFly (38.7 kg or 86.3 lb) treatments.
Seasonal yields of total, marketable, and unmarketable strawberries per plot.
This is the first field study evaluating the impact of three popular entomopathogenic fungi along with multiple beneficial microbes on strawberry plant growth, foliar and fruit diseases, and yield. While differences among treatments were not pronounced, it appeared that some had a positive impact on some of the parameters measured. It is interesting to note that yields were higher (although not statistically significant) than the grower standard, HealthySoil. Compared to the grower standard, marketable yield was higher in many other treatments. Since an untreated situation is not common in a commercial field, using beneficial microbes can be useful. Although previous field studies evaluated the impact of with the entomopathogenic fungus B. bassiana in strawberries (Dara, 2013; Dara, 2016), a positive impact on plant growth or yield by I. fumosorosea and M. brunneum in commercial strawberries has never been reported earlier.
Additional studies with different application rates would be useful to understand how beneficial microbes could be exploited more.
Acknowledgments: Thanks to Dave Peck, Manzanita Berry Farms for the collaboration and industry partners for the financial support. Thanks to Chris Martinez and rest of the field crew at Manzanita Berry Farms and Fritz Light and Tamas Zold for the technical assistance.
Amerian, M.R., and W.S. Stewart. 2001. Effect of two species of arbuscular mycorrhizal fungi on growth, assimilation and leaf water relations in maize (Zea mays). Aspects of Appl. Biol. 63: 1-6.
Behie, S.W., and M.J. Bidochka. 2014. Nutrient transfer in plant-fungal symbioses. Trends in Plant Sci. 19: 734-740.
Bolandnazar, S., N. Aliasgarzad, M.R. Neishabury, and N. Chaparzadeh. 2007. Mycorrhizal colonization improves onion (Allium cepa L.) yield and water use efficiency under water deficit condition. Sci. Horticulturae 114: 11-15.
Dara, S. K. 2013. Entomopathogenic fungus Beauveria bassiana promotes strawberry plant growth and health. UCANR eJournal Strawberries and Vegetables, 30 September, 2013. (http://ucanr.edu/blogs/blogcore/postdetail.cfm?postnum=11624)
Dara, S. K. 2016. First field study evaluating the impact of the entomopathogenic fungus Beauveria bassiana on strawberry plant growth and yield. UCANR eJournal Strawberrries and Vegetables, 7 November, 2016. (http://ucanr.edu/blogs/blogcore/postdetail.cfm?postnum=22546)
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, 16 September, 2016.(http://ucanr.edu/blogs/blogcore/postdetail.cfm?postnum=22131)
Nedorost, L., J. Vojtiskova, and R. Pokluda. 2014. Influence of watering regime and mycorrhizal inoculation on growth and nutrient uptake of pepper (Capsicum annuum L.). VII International symposium on irrigation of horticultural crops, Braun P., M. Stoll, and J. Zinkernagel (eds). Acta Horticulturae 1038:559-564.
Wu, Q.S., and Y. Zou. 2009. Mycorrhizal influence on nutrient uptake of citrus exposed to drought stress. Philippine Agri. Scientist 92: 33-38.