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Response of Aspergillus flavus spores to nitric oxide fumigations in atmospheres with different oxygen concentrations
Journal of Stored Products Research 83 (2019) 78e83
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Response of Aspergillus flavus spores to nitric oxide fumigations in atmospheres with different oxygen concentrations Yong-Biao Liu a, *, Sookyung Oh b, Wayne M. Jurick II c a
U.S. Department of Agriculture, Agricultural Research Service, Crop Improvement and Protection Unit, Salinas, CA, 93905, USA University of California at Davis, Salinas, CA, 93905, USA c U.S. Department of Agriculture, Agricultural Research Service, Food Quality Laboratory, Beltsville, MD, 20705, USA b
a r t i c l e i n f o
a b s t r a c t
Article history: Received 17 December 2018 Received in revised form 5 June 2019 Accepted 6 June 2019 Available online 12 June 2019
Nitric oxide (NO) is a newly discovered fumigant for postharvest pest control. NO fumigation must be conducted under ultralow oxygen conditions because NO reacts with O2 to produce nitrogen dioxide (NO2). In this study, NO fumigations under different O2 concentrations were conducted on Aspergillus flavus spores to determine effectiveness of NO and NO2 in inactivating spores. Spores on gridded cellulose filter discs in Petri dishes were subjected to six fumigation treatments including a control with varying levels of NO under different O2 conditions for 3 h at 15 C. The discs with spores were then cultured on Aspergillus Differentiation medium plates after fumigation for four days at 25 C to count A. flavus colonies. Untreated control discs each had over 50 A. flavus colonies. Three fumigation treatments with 0.1% NO2 or 1.0% NO caused complete inactivation of A. flavus spores. The study demonstrated that NO fumigation with certain levels of NO2 can effectively inactivate A. flavus spores. The results suggest that NO fumigation has potential to be an alternative treatment to control both pests and microbes on stored products. Published by Elsevier Ltd. This is an open access article under the CC BY-NC-ND license (http:// creativecommons.org/licenses/by-nc-nd/4.0/).
Keywords: Nitric oxide Nitrogen dioxide Fumigation Aspergillus flavus Spores Stored products
1. Introduction Stored products often face problems of infestations by pests and microbes. Postharvest losses to insect infestation are widely reported but are not well quantified (Boxall, 2001). Losses from all causes were estimated to be about 10% of production worldwide (Rajendran, 2002; Magan and Aldred, 2007). Contamination of cereal commodities by molds and mycotoxins is also common and represents a significant threat to food quality and results in dry matter, quality, safety and nutritional losses (Sauer et al., 1984; Kenkel et al., 1994; Richard and Payne, 2003; Magan and Aldred, 2007). Effective management of both insects and microbes is required for stored products. Stored product pests are commonly controlled by fumigation with chemical fumigants. In the past, methyl bromide was the
Footnote: This article reports the results of research only. Mention of proprietary products, trade names or commercial products in this publication is not for the purpose of providing specific information and does not constitute an endorsement or a recommendation by the U.S. Department of Agriculture for its use. The USDA is an equal opportunity provider and employer. * Corresponding author. E-mail address: [email protected] (Y.-B. Liu).
dominant fumigant for postharvest pest control on stored products. Methyl bromide fumigation not only controls pests but also controls microbes (Fields and White, 2002). As methyl bromide is being phased out globally and is no longer available for stored product pest control, phosphine and sulfuryl fluoride have become two major alternative fumigants for pest control on stored products (Rajendran, 2001, Fields and White, 2002). However, neither phosphine nor sulfuryl fluoride is an effective alternative to methyl bromide for pest management on stored products. Phosphine is not effective against some insects due to tolerance as well as resistance and phosphine fumigation can last over 10 days to control stored product pests (Hole et al., 1976; Benhalima et al., 2004; Nayak et al., 2007). Phosphine fumigation, however, has inhibitory effects on fungal growth (De Castro et al., 2001). Sulfuryl fluoride fumigation is not effective against insect eggs (Bell et al., 1998) and therefore has limited potential in control postharvest pests. Fungal infection is a serious threat to stored products due to loss and quality reduction of food products and potential health risk due to the presence of mycotoxins. Fungal infection is closely related to moisture, temperature, storage time, and insect infestation (Sauer et al., 1984; Navi et al., 2005). Mycotoxigenic molds in partially dried grain include species in genus Aspergillus, Penicillium, and
https://doi.org/10.1016/j.jspr.2019.06.001 0022-474X/Published by Elsevier Ltd. This is an open access article under the CC BY-NC-ND license (http://creativecommons.org/licenses/by-nc-nd/4.0/).
Y.-B. Liu et al. / Journal of Stored Products Research 83 (2019) 78e83
Fusarium (Richard and Payne, 2003; Magan and Aldred, 2007). Among them, A. flavus Link 1809 is well known for producing aflatoxins that are carcinogenic and is often found in stored products including corns and peanuts (Fakruddin et al., 2015). A. flavus spreads mainly through dispersal of airborne asexual spores (conidia) as inocula. Therefore, effective treatment to inactivate spores is important in preventing infections by A. flavus on stored products. Currently, control of microbes on stored products includes engineering control such as drying grains after harvest to reduce moisture content, modified atmosphere storage, fumigation with sulphur dioxide gas, and application of preservatives (Magan and Aldred, 2007). Phosphine fumigation, however, was found to inhibit A. flavus aflatoxin production on stored maize in 8- and 15day treatments (De Castro et al., 2001). There is a great need for more safe and effective treatments for postharvest pathogens on stored products. It will also be more cost effective if a treatment can control both postharvest pests and pathogens on stored products. Nitric oxide (NO) is a small molecule produced naturally from lightning and combustion of fossil fuel and is also manufactured as an intermediate in fertilizer production. It is also formed by almost all organisms as a ubiquitous cell messenger which modulates a wide variety of physiological processes (Culotta and Koshland, 1992; Beckman and Koppenol, 1996; Lamattina et al., 2003; Moncada and Higgs, 2006) and can be used to treat certain medical conditions such as asthma and cardiovascular diseases (Roberts et al., 1993; Rossaint et al., 1993; Ricciardolo et al., 2004). NO also functions as an antagonist of ethylene biosynthesis and can extend shelf-life and enhance postharvest quality of fresh fruit and vegetables (Wills et al. 2000, 2007; Soegiarto and Wills, 2004; Manjunatha et al., 2010, 2012; Zaharah and Singh, 2011; Saadatian et al., 2012). Recently, nitric oxide was discovered to be a potent fumigant in ultralow oxygen (ULO) atmosphere for controlling postharvest pests (Liu, 2013, 2015, 2016; Liu and Yang, 2016; Liu et al., 2016). Effective NO fumigation treatments have been developed against over 10 insect and mite species at various life stages including both external and internal feeding pests (Liu and Yang, 2016; Liu et al., 2016). NO fumigation not only has high efficacy against pests but also does not leave toxic residues on fumigated products (Liu and Yang, 2016; Yang and Liu, 2017). NO fumigation was also expected to be technically feasible and cost effective (Liu, 2015). NO fumigation must be conducted under ULO conditions to preserve NO to control pests because NO reacts with oxygen spontaneously to produce nitrogen dioxide (NO2). NO has been reported to be effective against bacteria including E. coli (Shank et al., 1962; Brunelli et al., 1995; Ghaffari et al., 2006; Bang et al., 2014). NO2 was found to be effective against microbes including fungal spores and has been studied for sterilizing medical devices by industry (Reed et al., 2014; Akutsu, 2015). In NO fumigations for pest control, there will always be some level of NO2 because it is not practical and necessary to remove all O2 from fumigation chambers. It may also be beneficial to establish NO fumigation treatments with desired levels of NO and NO2 by controlling both O2 levels in a fumigation chamber and the amount of NO injected for control of both pests and pathogens. We hypothesize that a NO fumigation with certain levels of NO2 can achieve effective control of both target pests and pathogens on stored products. Such a treatment will be more beneficial to agriculture industry because one treatment can provide effective control of both pests and diseases on stored products. In the present study, we conducted NO fumigations with certain levels of NO2 to determine their efficacy in controlling A. flavus spores.
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2. Materials and methods 2.1. Chemicals and fungus Nitric oxide (>99.5% purity) and commercial grade nitrogen in compressed cylinders were obtained from a commercial source for the study. NO was released and stored in a N2 washed foil bag (7.5 mils MylarFoil, 20 cm by 40 cm, Impak Corp., Los Angeles, CA) to be used in fumigations tests. Spores of A. flavus strain NRRL 3357 which does not produce aflatoxins from cultures maintained in the USDA-ARS Food Quality Laboratory in Beltsville, MD, were propagated on potato dextrose agar (PDA) plates for seven days at 30 C. Fresh spores from the fungal culture on PDA plates were harvested by suspension in 5 ml water containing 0.05% Triton-X 100 surfactant. Concentrated spore solutions were diluted with water to adjust the spore concentration to about 5 102/ml using a Luna™ Dual Fluorescence Cell Counter (Logos Biosystems Inc, Dongan-gu, Korea) for experiments. 2.2. Procedures of NO fumigation against fungal spores Gridded nitrocellulose mixed ester membrane filter discs (47 mm, Sterlitech Corp, Kent, WA) were placed individually in six vented 60 mm Petri dishes (P5237, Sigma, St. Louis, MO) and wetted with 200 ml water in a clean hood. A L-shaped cell spreader (Celltreat Scientific Products, Pepperell, MA) was used to spread water to wet surfaces of the cellulose filter disc. An aliquot of 100 ml spore solution with a spore concentration of about 5 102/ml was then deposited on each wet cellulose filter disc. The spore solution sample was spread with a L-shaped cell spreader to cover most surface of the disc. Once the disc was dry, the Petri dish was covered with the lid. Petri dishes with spore inoculated cellulose discs for a treatment were stacked together and secured with tape, and then loaded into a 1.9 L jar for fumigation. Each jar with Petri dishes was sealed with a lid with two ports. It was then flushed with N2 to establish a desired ULO condition. An oxygen analyzer (Series 800, Illinois Instruments, Inc., Johnsburg, IL) was used to monitor oxygen levels in each jar. Once the desired ULO condition was established, NO was injected into the jars under a fume hood using an airtight syringe to start NO fumigation treatment. The syringe and attached tubing were flushed with nitrogen prior to NO injection. Jars were then placed at 15 C in an environmental chamber for 3 h to complete the treatment. At the end of fumigation, the jars were moved under a fume hood and each jar was flushed with nitrogen at a flowrate of 5 L/min for 5 min to have at least 10 air exchanges with nitrogen to dilute NO in the jars before opening the jars. The Petri dishes were then removed from the jars. Cellulose discs in the Petri dishes from both controls and treatments were individually placed on 15 60 mm Aspergillus Differentiation Agar culture medium plates (17121, Sigma, St. Louis, MO) supplemented with 100 mg/L of Chloramphenicol (C61000, RPI, Mount Prospect, IL) in a clean hood. The medium plates were then covered and sealed with Parafilm and incubated at 25 C in an environmental chamber to allow surviving spores to form colonies on the medium plates. The plates were periodically inspected visually to check the appearance and development of fungal colonies. Photographs were taken and visualized on a computer screen to count fungal colonies for the treatments and the controls. 2.3. Effects of different fumigation treatments on fungal colony developments from spores A total of six treatments and a control were tested on spores of
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the A. flavus strain. Treatments were established by injecting a volume of NO gas equivalent to a specified percentage of the volume of the jars under different O2 levels. They were T1 (0.1% NO under 10 ppm O2), T2 (1.0% NO under 10 ppm O2), T3 (0.1% NO under 0.1% O2), T4 (1.0% NO under 0.1% O2), T5 (0.02% NO in normal atmosphere), and T6 (0.1% NO in normal atmosphere). As NO reacted with O2 spontaneously to produce NO2 after injection in each jar, the treatments were expected to result in different combinations of NO and NO2. Therefore, the actual treatments were: T1 (0.1% NO, 10 ppm NO2), T2 (1.0% NO, 10 ppm NO2), T3 (0.1% NO2), T4 (0.9% NO, 0.1% NO2), T5 (200 ppm NO2 under normal atmospheric O2 level), and T6 (0.1% NO2 under normal atmospheric O2 level). All treatments were at 15 C and lasted for 3 h. The treatments T1e T6 were terminated by flushing with N2 for 5 min at a flow rate of about 5 L/min. This was designed to avoid reaction of NO in the jars with O2 during the termination to produce NO2 which may affect viability of spores and skew the outcomes of the treatments. In each test, each treatment and the control had three replications. The test was repeated four times. 2.4. Data analysis The data on numbers of colonies on each plate for each treatment were compared using one-way ANOVA and Tukey HSD multiple range test at P ¼ 0.05 using JMP Statistical Discovery Software (SAS Institute, 2012). 3. Results Effects of nitric oxide fumigation treatments on spore viability of fungus A. flavus varied with NO concentration and O2 levels in the treatments. There were significant differences among the treatments in colony formation (F ¼ 177.846; df ¼ 6, 77; P < 0.0001) and three treatments resulted in complete inactivation of A. flavus spores (Table 1). On Aspergillus Differentiation Agar plates, colonies showed characteristic reddish color of A. flavus colonies when viewed from the bottom. Effects of different treatments on colony formation of A. flavus spores were obvious with no colony formation for treatments T2, T4, and T6 plates and just one colony on T3 plate (Fig. 1). As described in Table 1, unfumigated spores germinated and formed over 50 colonies on each Aspergillus Differentiation Agar plate after four days of incubation. Colonies expanded and became fuzzy over time. Fumigations with 1.0% NO under 10 ppm and 0.1% O2 and the fumigation with 0.1% NO under the normal atmosphere, however, had complete inhibition of A. flavus spores. The fumigation with 0.1% NO under 0.1% O2 had almost complete inhibition of spores with only 0.5 colony per plate and there was no significant difference among the above four treatments with 0e0.5 colonies
per plate. The fumigation with 0.02% NO under the normal atmosphere had a partial inhibition of spores with 24.6 colonies per plate and the value was significantly higher than the above four treatments and significantly lower than the colonies for the control. The fumigation with 0.1% NO under 10 ppm O2 had a mean colony number of 41 per plate which was not significantly different from the control (Table 1). 4. Discussion Stored products often have multiple problems of pest infestation and microbial infection. Therefore, it will be ideal if NO fumigation can control both pests and microbes. NO fumigation has been demonstrated to be effective against stored product insects (Liu, 2013; Liu and Yang, 2016). Results of complete inactivation of A. flavus spores from the current study and previous results of effective control of stored product pests (Liu, 2013) suggest that NO fumigation has potential to control both pests and microbes. This would make NO fumigation more useful and cost effective. Before NO injection, both treatments T1 and T2 chambers had 10 ppm O2 and, therefore, were expected to produce 10 ppm NO2 in the treatment chambers after NO injection. However, treatment T2 with 1.0% NO resulted in complete inactivation of A. flavus spores as compared with no significant inactivation in treatment T1. NO has been reported to be effective against bacteria (Shank et al., 1962; Brunelli et al., 1995; Ghaffari et al., 2006; Bang et al., 2014) and fungi (Lazar et al., 2008). NO fumigation was reported to inhibit mycelium growth, sporulation, and spore germination of three fungal species. However, there were no complete inactivation of fungal spores (Lazar et al., 2008). In the study of Lazar et al. (2008), NO gas was injected into sealed petri dishes under the normal atmosphere. Because NO reacts with O2, both NO and NO2 were present and it is therefore not clear whether the antifungal effects were due to NO, NO2, or both. Although the initial 10 ppm O2 level in T2 treatment chamber was expected to result in10 ppm NO2 after NO injection, complete airtight seal of the chambers may not be maintained and a slight leak in of air into the chambers could increase NO2 levels considerably. Given that NO2 had strong antimicrobial effects, NO2 in T2 may have contributed significantly to the complete inactivation of A. flavus spores. Therefore, the complete inactivation of A. flavus spores in the treatment T2 could be due to effects of both NO and NO2. Both T3 and T6 treatments were expected to yield 0.1% NO2 and resulted in 0.5 and 0 colony per plate and represented 99 and 100% inactivation of A. flavus spores. There was no significant difference in the number of A. flavus colonies between the two treatments. In comparison with NO fumigations for pest control on stored products, the 3 h treatment duration in the current study is very
Table 1 Mean numbers of Aspergillus flavus colonies and relative colony reduction in response to 3 h fumigation of spores with nitric oxide under different oxygen levels at 15 C on 4day post-treatment colony formation on Aspergillus Differentiation Agar medium plates at 25 C. Treatment
Control T1: 0.1% NO, 10 ppm O2 T2: 1.0% NO, 10 ppm O2 T3: 0.1% NO, 0.1% O2 T4: 1.0% NO, 0.1% O2 T5: 0.02% NO, 20.9% O2 T6: 0.1% NO, 20.9% O2 a
Concentrationa NO (%)
NO2 (ppm)
0.1 1.0 0 0.9 0 0
10 10 1000 1000 200 1000
No. of Coloniesb (mean ± SE)
Relative colony reduction (%)
51.8 ± 1.5 a 41.0 ± 3.0 b 0.0 ± 0.0 d 0.5 ± 0.3 d 0.0 ± 0.0 d 24.6 ± 2.9 c 0.0 ± 0.0 d
0 20.8 100 99.0 100 52.5 100
Calculated concentrations of NO and NO2 after injected NO reacts with O2 in fumigation chambers. Mean ± SE values followed with the same letter were not significantly different based on one-way ANOVA and Tukey HSD multiple range test at P ¼ 0.05 using the JMP Statistical Discovery software (SAS Institute, 2012). b
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Control
Control
T1
T1
T2
T2
T3
T3
81
T4
T5
T6
T4
T5
T6
Fig. 1. Colony formation of Aspergillus flavus spores on gridded nitrocellulose mixed ester membrane filter discs in Aspergillus Differentiation Agar supplemented with 100 mg/L of Chloramphenicol culture medium plates after 4 d of incubation at 25 C in an environmental chamber after fumigation treatments. The six fumigation treatments were: T1 (0.1% NO, 10 ppm NO2), T2 (1.0% NO, 10 ppm NO2), T3 (0.1% NO), T4 (0.9% NO, 0.1% NO2), T5 (200 ppm NO2 under normal atmospheric O2 level), and T6 (0.1% NO2 under normal atmospheric O2 level). Fumigation treatments were conducted for 3 h at 15 C. Upper panel: top view; lower panel: bottom view.
short. In the previously studies, stored product insects such as Indianmeal moth, confused flour beetle, and rice weevil, the minimum treatment time for control of their most susceptible life stage lasts at least 8e24 h (Liu, 2013). This indicates that the current 3 h treatments for control of A. flavus spores is too short for control of stored product insects. The treatment duration will need to be extended significantly to ensure effective control of postharvest pests. Based on results from the current study, longer NO fumigation treatments under 0.1% O2 are expected to achieve complete inactivation of A. flavus spores. A. flavus as well as other fungal pathogens spread mainly through dispersal of airborne spores. Therefore, effective control of spores is expected to be effective in reducing A. flavus inoculation on stored products. Spores are single cells. In comparison,
mycelium of A. flavus is formed by a network of hyphae and may not be easily killed by NO2 at a reasonable concentration. Therefore, NO fumigation is not expected to completely kill active A. flavus flora on infected stored products but inhibit their spreading. Currently, there is a lack of effective and economical treatments to control fungal spores in stored products to reduce fungal infection. The main method for microbial management is to maintain products dry along with application of fungicides. The maximum moisture contents range from 11 to 16% for stored corn, soybeans, wheat, and sorghum (Calvin et al., 1989). Insect infestation also enhance fungal growth through the damage to stored products, temperature increase by metabolism, and dissemination of spores (Lacey, 1989). Therefore, problems of pest infestation are often accompanied with mold development and there is a need to control
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both pests and microbes. NO fumigation with proper control on oxygen levels has potential to control both postharvest pests and microbes. A. flavus is well known for the production of aflatoxins which are carcinogenic and is a major threat to food safety. However, stored products also face infest by other fungi in all classes (Clarke, 1969). It is reasonable to assume that NO fumigation treatments which are effective against A. flavus spores are also effective against spores of other fungi. NO reacts with O2 spontaneously to produce NO2. Therefore, NO fumigation must be conducted under ULO conditions to preserve NO. The ULO conditions are established by flushing fumigation chambers with N2 gas. Because it is not practical to remove oxygen completely from a fumigation chamber with N2 flush, there are always some oxygen at very low levels remaining in the fumigation chamber. The procedures of NO2 fumigation for pest control can also be modified by controlling ULO conditions to achieve proper levels for NO for pest control but also proper levels for NO2 for controlling microbes. Thereby, NO fumigation has potential to be used control both postharvest pests and pathogens. The results of complete inactivation of A. flavus spores in several NO fumigation treatments in the current study demonstrated that NO fumigation has the potential for controlling both postharvest pests and pathogens. However, the current results of effective inactivation of A. flavus spores were from a small-scale laboratory study in the absence of a stored product. Stored products likely to have different adsorption rates for NO and NO2. NO and NO2 are also likely to have different abilities to penetrate stored products. Therefore, more research is needed to evaluate the potential of NO fumigation in large-scale fumigation tests to develop appropriate fumigation procedures to achieve effective control of postharvest pests and inactivation of spores of A. flavus and other fungi. Acknowledgements We thank T. Masuda and R. Singh for technical assistance. We are very grateful for the invaluable contribution by Dr. Jiujiang Yu who was a research geneticist at USDA-ARS Food Quality Laboratory in Beltsville, MD and retired before the completion of this study. This study was supported in part by TASC grant (C2017-04) from U.S. Depatment of Agriculture, Foreign Agricultural Service and by U.S. Department of Agriculture, Agricultural Research Service Project Plan 2038-22430-002-00-D. References Akutsu, T., 2015. Method and Apparatus for Sterilization with Nitrogen Dioxide. U.S. Patent Application No. US2015/0037206. U.S. Patent and Trademark Office. Bang, C.S., Kinnunen, A., Karlsson, M., Onnberg, A., Soderquist, B., 2014. The antibacterial effect of nitric oxide against ESBL-producing uropathogenic E. coli is improved by combination with micronazole and polymyxin B nonapeptide. BMC Microbiol. 14 (1e9), 65. Beckman, J.S., Koppenol, W.H., 1996. Nitric oxide, superoxide, and peroxynitrite: the good, the bad, and the ugly. Am. J. Physiol. 271, C1424eC1437. Cell Physiol. 40. Bell, C.H., Savvidou, N., Wontner Smith, T.J., 1998. The toxicity of sulfuryl fluoride (Vikane) to eggs of insect pests of flour mills. In: Proc. 7th Intl. Working Conference on Stored Product Protection, vol. 1, pp. 345e350. Benhalima, H., Chaudhry, M.Q., Mills, K.A., Price, N.R., 2004. Phosphine resistance in stored-product insects collected from various grain storage facilities in Morocco. J. Stored Prod. Res. 40, 241e249. Boxall, R.A., 2001. Post-harvest losses to insects e a world overview. Int. Biodeterior. Biodegrad. 48, 137e152. Brunelli, L., Crow, J.P., Beckman, J.S., 1995. The comparative toxicity of nitric oxide and peroxynitrite to Escherichia coli. Arch. Biochem. Biophys. 316, 327e334. Calvin, D., Carson, J., Jacobs, S., 1989. Manage Stored Grain on the Farm. Publication SG-14, Dept. of Entomology, College of Agriculture Science, Pennsylvania State University. https://ento.psu.edu/extension/factsheets/pdf/manageStoredGrain. pdf. Clarke, J.H., 1969. Fungi in stored products. PANS Pest Artic. News Summ. 15,
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Decontamination and deployment of an ASEP-TECH blow/fill/seal system. https://www.noxilizer.com/pdf/ publications/scientific_poster_pda_decontamination_and_depyrogenation_ asep_tech_bfs_system_april2014.pdf. Ricciardolo, F.L.M., Sterk, P.J., Gaston, B., Folkerts, G., 2004. Nitric oxide in health and disease of the respiratory system. Physiol. Rev. 84, 731e765. Richard, J.L., Payne, G.A., 2003. Mycotoxins in Plant, Animal, and Human Systems. Task Force Report No. 139. Council for Agricultural Science and Technology. Roberts Jr., J.D., Lang, P., Bigatello, L.M., Vlahakes, G.J., Zapol, W.M., 1993. Inhaled nitric oxide in congenital heart disease. Circulation 87, 447e453. Rossaint, R., Falke, K.J., Lopez, F., Slama, K., Pison, U., Zapol, W.M., 1993. Inhaled nitric oxide for the adult respiratory distress syndrome. N. Engl. J. Med. 328, 399e405. Saadatian, M., Ahmadiyan, S., Akbari, M., Balouchi, Z., 2012. Effects of pretreatment with nitric oxide on kiwifruit storage at low temperature. 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presence of horticultural produce to extend postharvest life. Nitric Oxide 17, 44e49. Wills, R.B.H., Ku, V.V.V., Leshem, Y.Y., 2000. Fumigation with nitric oxide to extend the postharvest life of strawberries. Postharvest Biol. Technol. 18, 75e79. Yang, X., Liu, Y.-B., 2017. Residual analysis of nitric oxide fumigation on fresh fruit and vegetables. Postharvest Biol. Technol. 132, 105e108. Zaharah, S.S., Singh, Z., 2011. Postharvest nitric oxide fumigation alleviates chilling injury, delay fruit ripening and maintains quality in cold-stored ‘Kensington Pride’ mango. Postharvest Biol. Technol. 60, 202e210.
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Response of Aspergillus flavus spores to nitric oxide fumigations in atmospheres with different oxygen concentrations Yong-Biao Liu a, *, Sookyung Oh b, Wayne M. Jurick II c a
U.S. Department of Agriculture, Agricultural Research Service, Crop Improvement and Protection Unit, Salinas, CA, 93905, USA University of California at Davis, Salinas, CA, 93905, USA c U.S. Department of Agriculture, Agricultural Research Service, Food Quality Laboratory, Beltsville, MD, 20705, USA b
a r t i c l e i n f o
a b s t r a c t
Article history: Received 17 December 2018 Received in revised form 5 June 2019 Accepted 6 June 2019 Available online 12 June 2019
Nitric oxide (NO) is a newly discovered fumigant for postharvest pest control. NO fumigation must be conducted under ultralow oxygen conditions because NO reacts with O2 to produce nitrogen dioxide (NO2). In this study, NO fumigations under different O2 concentrations were conducted on Aspergillus flavus spores to determine effectiveness of NO and NO2 in inactivating spores. Spores on gridded cellulose filter discs in Petri dishes were subjected to six fumigation treatments including a control with varying levels of NO under different O2 conditions for 3 h at 15 C. The discs with spores were then cultured on Aspergillus Differentiation medium plates after fumigation for four days at 25 C to count A. flavus colonies. Untreated control discs each had over 50 A. flavus colonies. Three fumigation treatments with 0.1% NO2 or 1.0% NO caused complete inactivation of A. flavus spores. The study demonstrated that NO fumigation with certain levels of NO2 can effectively inactivate A. flavus spores. The results suggest that NO fumigation has potential to be an alternative treatment to control both pests and microbes on stored products. Published by Elsevier Ltd. This is an open access article under the CC BY-NC-ND license (http:// creativecommons.org/licenses/by-nc-nd/4.0/).
Keywords: Nitric oxide Nitrogen dioxide Fumigation Aspergillus flavus Spores Stored products
1. Introduction Stored products often face problems of infestations by pests and microbes. Postharvest losses to insect infestation are widely reported but are not well quantified (Boxall, 2001). Losses from all causes were estimated to be about 10% of production worldwide (Rajendran, 2002; Magan and Aldred, 2007). Contamination of cereal commodities by molds and mycotoxins is also common and represents a significant threat to food quality and results in dry matter, quality, safety and nutritional losses (Sauer et al., 1984; Kenkel et al., 1994; Richard and Payne, 2003; Magan and Aldred, 2007). Effective management of both insects and microbes is required for stored products. Stored product pests are commonly controlled by fumigation with chemical fumigants. In the past, methyl bromide was the
Footnote: This article reports the results of research only. Mention of proprietary products, trade names or commercial products in this publication is not for the purpose of providing specific information and does not constitute an endorsement or a recommendation by the U.S. Department of Agriculture for its use. The USDA is an equal opportunity provider and employer. * Corresponding author. E-mail address: [email protected] (Y.-B. Liu).
dominant fumigant for postharvest pest control on stored products. Methyl bromide fumigation not only controls pests but also controls microbes (Fields and White, 2002). As methyl bromide is being phased out globally and is no longer available for stored product pest control, phosphine and sulfuryl fluoride have become two major alternative fumigants for pest control on stored products (Rajendran, 2001, Fields and White, 2002). However, neither phosphine nor sulfuryl fluoride is an effective alternative to methyl bromide for pest management on stored products. Phosphine is not effective against some insects due to tolerance as well as resistance and phosphine fumigation can last over 10 days to control stored product pests (Hole et al., 1976; Benhalima et al., 2004; Nayak et al., 2007). Phosphine fumigation, however, has inhibitory effects on fungal growth (De Castro et al., 2001). Sulfuryl fluoride fumigation is not effective against insect eggs (Bell et al., 1998) and therefore has limited potential in control postharvest pests. Fungal infection is a serious threat to stored products due to loss and quality reduction of food products and potential health risk due to the presence of mycotoxins. Fungal infection is closely related to moisture, temperature, storage time, and insect infestation (Sauer et al., 1984; Navi et al., 2005). Mycotoxigenic molds in partially dried grain include species in genus Aspergillus, Penicillium, and
https://doi.org/10.1016/j.jspr.2019.06.001 0022-474X/Published by Elsevier Ltd. This is an open access article under the CC BY-NC-ND license (http://creativecommons.org/licenses/by-nc-nd/4.0/).
Y.-B. Liu et al. / Journal of Stored Products Research 83 (2019) 78e83
Fusarium (Richard and Payne, 2003; Magan and Aldred, 2007). Among them, A. flavus Link 1809 is well known for producing aflatoxins that are carcinogenic and is often found in stored products including corns and peanuts (Fakruddin et al., 2015). A. flavus spreads mainly through dispersal of airborne asexual spores (conidia) as inocula. Therefore, effective treatment to inactivate spores is important in preventing infections by A. flavus on stored products. Currently, control of microbes on stored products includes engineering control such as drying grains after harvest to reduce moisture content, modified atmosphere storage, fumigation with sulphur dioxide gas, and application of preservatives (Magan and Aldred, 2007). Phosphine fumigation, however, was found to inhibit A. flavus aflatoxin production on stored maize in 8- and 15day treatments (De Castro et al., 2001). There is a great need for more safe and effective treatments for postharvest pathogens on stored products. It will also be more cost effective if a treatment can control both postharvest pests and pathogens on stored products. Nitric oxide (NO) is a small molecule produced naturally from lightning and combustion of fossil fuel and is also manufactured as an intermediate in fertilizer production. It is also formed by almost all organisms as a ubiquitous cell messenger which modulates a wide variety of physiological processes (Culotta and Koshland, 1992; Beckman and Koppenol, 1996; Lamattina et al., 2003; Moncada and Higgs, 2006) and can be used to treat certain medical conditions such as asthma and cardiovascular diseases (Roberts et al., 1993; Rossaint et al., 1993; Ricciardolo et al., 2004). NO also functions as an antagonist of ethylene biosynthesis and can extend shelf-life and enhance postharvest quality of fresh fruit and vegetables (Wills et al. 2000, 2007; Soegiarto and Wills, 2004; Manjunatha et al., 2010, 2012; Zaharah and Singh, 2011; Saadatian et al., 2012). Recently, nitric oxide was discovered to be a potent fumigant in ultralow oxygen (ULO) atmosphere for controlling postharvest pests (Liu, 2013, 2015, 2016; Liu and Yang, 2016; Liu et al., 2016). Effective NO fumigation treatments have been developed against over 10 insect and mite species at various life stages including both external and internal feeding pests (Liu and Yang, 2016; Liu et al., 2016). NO fumigation not only has high efficacy against pests but also does not leave toxic residues on fumigated products (Liu and Yang, 2016; Yang and Liu, 2017). NO fumigation was also expected to be technically feasible and cost effective (Liu, 2015). NO fumigation must be conducted under ULO conditions to preserve NO to control pests because NO reacts with oxygen spontaneously to produce nitrogen dioxide (NO2). NO has been reported to be effective against bacteria including E. coli (Shank et al., 1962; Brunelli et al., 1995; Ghaffari et al., 2006; Bang et al., 2014). NO2 was found to be effective against microbes including fungal spores and has been studied for sterilizing medical devices by industry (Reed et al., 2014; Akutsu, 2015). In NO fumigations for pest control, there will always be some level of NO2 because it is not practical and necessary to remove all O2 from fumigation chambers. It may also be beneficial to establish NO fumigation treatments with desired levels of NO and NO2 by controlling both O2 levels in a fumigation chamber and the amount of NO injected for control of both pests and pathogens. We hypothesize that a NO fumigation with certain levels of NO2 can achieve effective control of both target pests and pathogens on stored products. Such a treatment will be more beneficial to agriculture industry because one treatment can provide effective control of both pests and diseases on stored products. In the present study, we conducted NO fumigations with certain levels of NO2 to determine their efficacy in controlling A. flavus spores.
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2. Materials and methods 2.1. Chemicals and fungus Nitric oxide (>99.5% purity) and commercial grade nitrogen in compressed cylinders were obtained from a commercial source for the study. NO was released and stored in a N2 washed foil bag (7.5 mils MylarFoil, 20 cm by 40 cm, Impak Corp., Los Angeles, CA) to be used in fumigations tests. Spores of A. flavus strain NRRL 3357 which does not produce aflatoxins from cultures maintained in the USDA-ARS Food Quality Laboratory in Beltsville, MD, were propagated on potato dextrose agar (PDA) plates for seven days at 30 C. Fresh spores from the fungal culture on PDA plates were harvested by suspension in 5 ml water containing 0.05% Triton-X 100 surfactant. Concentrated spore solutions were diluted with water to adjust the spore concentration to about 5 102/ml using a Luna™ Dual Fluorescence Cell Counter (Logos Biosystems Inc, Dongan-gu, Korea) for experiments. 2.2. Procedures of NO fumigation against fungal spores Gridded nitrocellulose mixed ester membrane filter discs (47 mm, Sterlitech Corp, Kent, WA) were placed individually in six vented 60 mm Petri dishes (P5237, Sigma, St. Louis, MO) and wetted with 200 ml water in a clean hood. A L-shaped cell spreader (Celltreat Scientific Products, Pepperell, MA) was used to spread water to wet surfaces of the cellulose filter disc. An aliquot of 100 ml spore solution with a spore concentration of about 5 102/ml was then deposited on each wet cellulose filter disc. The spore solution sample was spread with a L-shaped cell spreader to cover most surface of the disc. Once the disc was dry, the Petri dish was covered with the lid. Petri dishes with spore inoculated cellulose discs for a treatment were stacked together and secured with tape, and then loaded into a 1.9 L jar for fumigation. Each jar with Petri dishes was sealed with a lid with two ports. It was then flushed with N2 to establish a desired ULO condition. An oxygen analyzer (Series 800, Illinois Instruments, Inc., Johnsburg, IL) was used to monitor oxygen levels in each jar. Once the desired ULO condition was established, NO was injected into the jars under a fume hood using an airtight syringe to start NO fumigation treatment. The syringe and attached tubing were flushed with nitrogen prior to NO injection. Jars were then placed at 15 C in an environmental chamber for 3 h to complete the treatment. At the end of fumigation, the jars were moved under a fume hood and each jar was flushed with nitrogen at a flowrate of 5 L/min for 5 min to have at least 10 air exchanges with nitrogen to dilute NO in the jars before opening the jars. The Petri dishes were then removed from the jars. Cellulose discs in the Petri dishes from both controls and treatments were individually placed on 15 60 mm Aspergillus Differentiation Agar culture medium plates (17121, Sigma, St. Louis, MO) supplemented with 100 mg/L of Chloramphenicol (C61000, RPI, Mount Prospect, IL) in a clean hood. The medium plates were then covered and sealed with Parafilm and incubated at 25 C in an environmental chamber to allow surviving spores to form colonies on the medium plates. The plates were periodically inspected visually to check the appearance and development of fungal colonies. Photographs were taken and visualized on a computer screen to count fungal colonies for the treatments and the controls. 2.3. Effects of different fumigation treatments on fungal colony developments from spores A total of six treatments and a control were tested on spores of
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the A. flavus strain. Treatments were established by injecting a volume of NO gas equivalent to a specified percentage of the volume of the jars under different O2 levels. They were T1 (0.1% NO under 10 ppm O2), T2 (1.0% NO under 10 ppm O2), T3 (0.1% NO under 0.1% O2), T4 (1.0% NO under 0.1% O2), T5 (0.02% NO in normal atmosphere), and T6 (0.1% NO in normal atmosphere). As NO reacted with O2 spontaneously to produce NO2 after injection in each jar, the treatments were expected to result in different combinations of NO and NO2. Therefore, the actual treatments were: T1 (0.1% NO, 10 ppm NO2), T2 (1.0% NO, 10 ppm NO2), T3 (0.1% NO2), T4 (0.9% NO, 0.1% NO2), T5 (200 ppm NO2 under normal atmospheric O2 level), and T6 (0.1% NO2 under normal atmospheric O2 level). All treatments were at 15 C and lasted for 3 h. The treatments T1e T6 were terminated by flushing with N2 for 5 min at a flow rate of about 5 L/min. This was designed to avoid reaction of NO in the jars with O2 during the termination to produce NO2 which may affect viability of spores and skew the outcomes of the treatments. In each test, each treatment and the control had three replications. The test was repeated four times. 2.4. Data analysis The data on numbers of colonies on each plate for each treatment were compared using one-way ANOVA and Tukey HSD multiple range test at P ¼ 0.05 using JMP Statistical Discovery Software (SAS Institute, 2012). 3. Results Effects of nitric oxide fumigation treatments on spore viability of fungus A. flavus varied with NO concentration and O2 levels in the treatments. There were significant differences among the treatments in colony formation (F ¼ 177.846; df ¼ 6, 77; P < 0.0001) and three treatments resulted in complete inactivation of A. flavus spores (Table 1). On Aspergillus Differentiation Agar plates, colonies showed characteristic reddish color of A. flavus colonies when viewed from the bottom. Effects of different treatments on colony formation of A. flavus spores were obvious with no colony formation for treatments T2, T4, and T6 plates and just one colony on T3 plate (Fig. 1). As described in Table 1, unfumigated spores germinated and formed over 50 colonies on each Aspergillus Differentiation Agar plate after four days of incubation. Colonies expanded and became fuzzy over time. Fumigations with 1.0% NO under 10 ppm and 0.1% O2 and the fumigation with 0.1% NO under the normal atmosphere, however, had complete inhibition of A. flavus spores. The fumigation with 0.1% NO under 0.1% O2 had almost complete inhibition of spores with only 0.5 colony per plate and there was no significant difference among the above four treatments with 0e0.5 colonies
per plate. The fumigation with 0.02% NO under the normal atmosphere had a partial inhibition of spores with 24.6 colonies per plate and the value was significantly higher than the above four treatments and significantly lower than the colonies for the control. The fumigation with 0.1% NO under 10 ppm O2 had a mean colony number of 41 per plate which was not significantly different from the control (Table 1). 4. Discussion Stored products often have multiple problems of pest infestation and microbial infection. Therefore, it will be ideal if NO fumigation can control both pests and microbes. NO fumigation has been demonstrated to be effective against stored product insects (Liu, 2013; Liu and Yang, 2016). Results of complete inactivation of A. flavus spores from the current study and previous results of effective control of stored product pests (Liu, 2013) suggest that NO fumigation has potential to control both pests and microbes. This would make NO fumigation more useful and cost effective. Before NO injection, both treatments T1 and T2 chambers had 10 ppm O2 and, therefore, were expected to produce 10 ppm NO2 in the treatment chambers after NO injection. However, treatment T2 with 1.0% NO resulted in complete inactivation of A. flavus spores as compared with no significant inactivation in treatment T1. NO has been reported to be effective against bacteria (Shank et al., 1962; Brunelli et al., 1995; Ghaffari et al., 2006; Bang et al., 2014) and fungi (Lazar et al., 2008). NO fumigation was reported to inhibit mycelium growth, sporulation, and spore germination of three fungal species. However, there were no complete inactivation of fungal spores (Lazar et al., 2008). In the study of Lazar et al. (2008), NO gas was injected into sealed petri dishes under the normal atmosphere. Because NO reacts with O2, both NO and NO2 were present and it is therefore not clear whether the antifungal effects were due to NO, NO2, or both. Although the initial 10 ppm O2 level in T2 treatment chamber was expected to result in10 ppm NO2 after NO injection, complete airtight seal of the chambers may not be maintained and a slight leak in of air into the chambers could increase NO2 levels considerably. Given that NO2 had strong antimicrobial effects, NO2 in T2 may have contributed significantly to the complete inactivation of A. flavus spores. Therefore, the complete inactivation of A. flavus spores in the treatment T2 could be due to effects of both NO and NO2. Both T3 and T6 treatments were expected to yield 0.1% NO2 and resulted in 0.5 and 0 colony per plate and represented 99 and 100% inactivation of A. flavus spores. There was no significant difference in the number of A. flavus colonies between the two treatments. In comparison with NO fumigations for pest control on stored products, the 3 h treatment duration in the current study is very
Table 1 Mean numbers of Aspergillus flavus colonies and relative colony reduction in response to 3 h fumigation of spores with nitric oxide under different oxygen levels at 15 C on 4day post-treatment colony formation on Aspergillus Differentiation Agar medium plates at 25 C. Treatment
Control T1: 0.1% NO, 10 ppm O2 T2: 1.0% NO, 10 ppm O2 T3: 0.1% NO, 0.1% O2 T4: 1.0% NO, 0.1% O2 T5: 0.02% NO, 20.9% O2 T6: 0.1% NO, 20.9% O2 a
Concentrationa NO (%)
NO2 (ppm)
0.1 1.0 0 0.9 0 0
10 10 1000 1000 200 1000
No. of Coloniesb (mean ± SE)
Relative colony reduction (%)
51.8 ± 1.5 a 41.0 ± 3.0 b 0.0 ± 0.0 d 0.5 ± 0.3 d 0.0 ± 0.0 d 24.6 ± 2.9 c 0.0 ± 0.0 d
0 20.8 100 99.0 100 52.5 100
Calculated concentrations of NO and NO2 after injected NO reacts with O2 in fumigation chambers. Mean ± SE values followed with the same letter were not significantly different based on one-way ANOVA and Tukey HSD multiple range test at P ¼ 0.05 using the JMP Statistical Discovery software (SAS Institute, 2012). b
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Control
Control
T1
T1
T2
T2
T3
T3
81
T4
T5
T6
T4
T5
T6
Fig. 1. Colony formation of Aspergillus flavus spores on gridded nitrocellulose mixed ester membrane filter discs in Aspergillus Differentiation Agar supplemented with 100 mg/L of Chloramphenicol culture medium plates after 4 d of incubation at 25 C in an environmental chamber after fumigation treatments. The six fumigation treatments were: T1 (0.1% NO, 10 ppm NO2), T2 (1.0% NO, 10 ppm NO2), T3 (0.1% NO), T4 (0.9% NO, 0.1% NO2), T5 (200 ppm NO2 under normal atmospheric O2 level), and T6 (0.1% NO2 under normal atmospheric O2 level). Fumigation treatments were conducted for 3 h at 15 C. Upper panel: top view; lower panel: bottom view.
short. In the previously studies, stored product insects such as Indianmeal moth, confused flour beetle, and rice weevil, the minimum treatment time for control of their most susceptible life stage lasts at least 8e24 h (Liu, 2013). This indicates that the current 3 h treatments for control of A. flavus spores is too short for control of stored product insects. The treatment duration will need to be extended significantly to ensure effective control of postharvest pests. Based on results from the current study, longer NO fumigation treatments under 0.1% O2 are expected to achieve complete inactivation of A. flavus spores. A. flavus as well as other fungal pathogens spread mainly through dispersal of airborne spores. Therefore, effective control of spores is expected to be effective in reducing A. flavus inoculation on stored products. Spores are single cells. In comparison,
mycelium of A. flavus is formed by a network of hyphae and may not be easily killed by NO2 at a reasonable concentration. Therefore, NO fumigation is not expected to completely kill active A. flavus flora on infected stored products but inhibit their spreading. Currently, there is a lack of effective and economical treatments to control fungal spores in stored products to reduce fungal infection. The main method for microbial management is to maintain products dry along with application of fungicides. The maximum moisture contents range from 11 to 16% for stored corn, soybeans, wheat, and sorghum (Calvin et al., 1989). Insect infestation also enhance fungal growth through the damage to stored products, temperature increase by metabolism, and dissemination of spores (Lacey, 1989). Therefore, problems of pest infestation are often accompanied with mold development and there is a need to control
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both pests and microbes. NO fumigation with proper control on oxygen levels has potential to control both postharvest pests and microbes. A. flavus is well known for the production of aflatoxins which are carcinogenic and is a major threat to food safety. However, stored products also face infest by other fungi in all classes (Clarke, 1969). It is reasonable to assume that NO fumigation treatments which are effective against A. flavus spores are also effective against spores of other fungi. NO reacts with O2 spontaneously to produce NO2. Therefore, NO fumigation must be conducted under ULO conditions to preserve NO. The ULO conditions are established by flushing fumigation chambers with N2 gas. Because it is not practical to remove oxygen completely from a fumigation chamber with N2 flush, there are always some oxygen at very low levels remaining in the fumigation chamber. The procedures of NO2 fumigation for pest control can also be modified by controlling ULO conditions to achieve proper levels for NO for pest control but also proper levels for NO2 for controlling microbes. Thereby, NO fumigation has potential to be used control both postharvest pests and pathogens. The results of complete inactivation of A. flavus spores in several NO fumigation treatments in the current study demonstrated that NO fumigation has the potential for controlling both postharvest pests and pathogens. However, the current results of effective inactivation of A. flavus spores were from a small-scale laboratory study in the absence of a stored product. Stored products likely to have different adsorption rates for NO and NO2. NO and NO2 are also likely to have different abilities to penetrate stored products. Therefore, more research is needed to evaluate the potential of NO fumigation in large-scale fumigation tests to develop appropriate fumigation procedures to achieve effective control of postharvest pests and inactivation of spores of A. flavus and other fungi. Acknowledgements We thank T. Masuda and R. Singh for technical assistance. We are very grateful for the invaluable contribution by Dr. Jiujiang Yu who was a research geneticist at USDA-ARS Food Quality Laboratory in Beltsville, MD and retired before the completion of this study. This study was supported in part by TASC grant (C2017-04) from U.S. Depatment of Agriculture, Foreign Agricultural Service and by U.S. Department of Agriculture, Agricultural Research Service Project Plan 2038-22430-002-00-D. References Akutsu, T., 2015. Method and Apparatus for Sterilization with Nitrogen Dioxide. U.S. Patent Application No. US2015/0037206. U.S. Patent and Trademark Office. Bang, C.S., Kinnunen, A., Karlsson, M., Onnberg, A., Soderquist, B., 2014. The antibacterial effect of nitric oxide against ESBL-producing uropathogenic E. coli is improved by combination with micronazole and polymyxin B nonapeptide. BMC Microbiol. 14 (1e9), 65. Beckman, J.S., Koppenol, W.H., 1996. Nitric oxide, superoxide, and peroxynitrite: the good, the bad, and the ugly. Am. J. Physiol. 271, C1424eC1437. Cell Physiol. 40. Bell, C.H., Savvidou, N., Wontner Smith, T.J., 1998. The toxicity of sulfuryl fluoride (Vikane) to eggs of insect pests of flour mills. In: Proc. 7th Intl. Working Conference on Stored Product Protection, vol. 1, pp. 345e350. Benhalima, H., Chaudhry, M.Q., Mills, K.A., Price, N.R., 2004. Phosphine resistance in stored-product insects collected from various grain storage facilities in Morocco. J. Stored Prod. Res. 40, 241e249. Boxall, R.A., 2001. Post-harvest losses to insects e a world overview. Int. Biodeterior. Biodegrad. 48, 137e152. Brunelli, L., Crow, J.P., Beckman, J.S., 1995. The comparative toxicity of nitric oxide and peroxynitrite to Escherichia coli. Arch. Biochem. Biophys. 316, 327e334. Calvin, D., Carson, J., Jacobs, S., 1989. Manage Stored Grain on the Farm. Publication SG-14, Dept. of Entomology, College of Agriculture Science, Pennsylvania State University. https://ento.psu.edu/extension/factsheets/pdf/manageStoredGrain. pdf. Clarke, J.H., 1969. Fungi in stored products. PANS Pest Artic. News Summ. 15,
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