A pot experiment was conducted to evaluate the effect of urea on nitrogen metabolism and membrane lipid peroxidation in Azolla pinnata. Compared to controls, the application of urea to A. pinnata resulted in a 44% decrease in nitrogenase activity, no significant change in glutamine synthetase activity, 660% higher glutamic-pyruvic transaminase, 39% increase in free amino acid levels, 22% increase in malondialdehyde levels, 21% increase in Na+/K+- levels, 16% increase in Ca2+/Mg2+-ATPase levels, and 11% decrease in superoxide dismutase activity. In terms of H2O2 detoxifying enzymes, peroxidase activity did not change and catalase activity increased by 64% in urea-treated A. pinnata. These findings suggest that urea application promotes amino acid metabolism and membrane lipid peroxidation in A. pinnata.
Citation: Chen J, Huang M, Cao F, Pardha-Saradhi P, Zou Y (2017) Urea application promotes amino acid metabolism and membrane lipid peroxidation in Azolla. PLoS ONE 12(9): e0185230. https://doi.org/10.1371/journal.pone.0185230
Editor: Alexander Valentine, Stellenbosch University, SOUTH AFRICA
Received: May 14, 2017; Accepted: September 9, 2017; Published: September 25, 2017
Copyright: © 2017 Chen et al. This is an open access article distributed under the terms of the Creative Commons Attribution License, which permits unrestricted use, distribution, and reproduction in any medium, provided the original author and source are credited.
Data Availability: All data are contained within the paper.
Funding: This work was supported by the Earmarked Fund for China Agriculture Research System (CARS-01). The funder had no role in study design, data collection and analysis, decision to publish, or preparation of the manuscript.
Competing interests: The authors have declared that no competing interests exist.
Azolla is a floating aquatic fern that is widely distributed throughout temperate and tropical freshwater ecosystems. It establishes symbiotic associations with nitrogen (N)-fixing blue-green algae, and this association is mainly responsible for its high productivity and its ability to fix N at high rates . For centuries, Azolla has been employed as green manure in rice in China and Vietnam and more recently in other Asian and African countries . The basal application of Azolla at 10–12 t∙ha–1 increases soil N by 50–60 kg∙ha–1 and reduces the requirement for chemical N fertilizers in rice production by 30–35 kg∙ha–1 . Besides its utilization as a green manure, Azolla can be used as an animal feed, a human food, a medicine, and a water purifier. It may also be used for the production of hydrogen fuel, the production of biogas, the control of weeds, the control of mosquitoes, and the reduction of ammonia volatilization which accompanies the application of chemical N fertilizer .
The growth of Azolla is influenced by various environmental factors, and it is well documented that Azolla prefers sites that are rich in all essential plant nutrients, except for N [5, 6]. Similarly, in 2016, we observed a considerable amount of A. pinnata in zero-N plots but not in urea-applied plots previously used in a long-term field experiment with double rice cropping that was initiated in 2008 (Fig 1). This suggests that urea is a stressor to Azolla and a decline in Azolla biodiversity in rice field ecosystems. Furthermore, urea application synchronously inhibits the growth of Azolla and the amount of N fixed [2, 8], thereby indicating that there may be a link between the effect of urea application on the growth and N metabolism in Azolla. Currently, information on the effect of urea application on N metabolism in Azolla is limited.
The picture and data were obtained from a long-term field experiment with double rice cropping from 2008 onwards in Liuyang County, Hunan Province, China in early rice-growing season in 2016. See Qin et al.  for the details of the experiment. Vertical bars represent SE (n = 4).
Plants subjected to adverse environments usually undergo alterations in cell membrane structure and function [9, 10]. Membrane lipid peroxidation is a major contributor to these alterations, which presumably can lead to further deleterious consequences such as inhibition of ATPases [11, 12]. Malondialdehyde (MDA) is one of the major products of membrane lipid peroxidation and has been used as an indicator of membrane lipid peroxide levels [13, 14]. Additionally, in response to adverse conditions (e.g., dry, high, or low temperature, waterlogging), antioxidant enzymes such as superoxide dismutase (SOD), peroxidase (POD), and catalase (CAT) are produced to remove active oxygen radicals . However, whether these physiological processes are induced by urea application in Azolla remain unclear.
In the present study, a pot experiment was conducted to determine several parameters associated with N metabolism and membrane lipid peroxidation in Azolla with and without urea application. The objective of this study was to determine the effect of urea application on the metabolism and membrane lipid peroxidation in Azolla.
Materials and methods
No specific permissions were required for the activities conducted in this study. The experiments did not involve endangered or protected species.
A. pinnata plants used in this study were collected from a zero-N plot used in a long-term field experiment with double rice cropping that was initiated in 2008 in Liuyang County, Hunan Province, China during the early rice-growing season of 2016 (Fig 1). Twelve plastic pots (22 cm × 16 cm × 7 cm; length × width × height) were filled with distilled water to a depth of 5 cm. The pots were separated into two groups: one group (n = 6) received urea (46% N) at a rate of 1.15 g∙pot–1 (equivalent to the recommended field rate of 150 kg N∙ha–1) and the other group (n = 6) did not receive N fertilizers (control). Uniformly sized Azolla plants were selected and cultured in the pots at a rate of 10 g fresh weight (FW)∙pot–1 under laboratory conditions.
Each pot was treated as one replicate. All plants in each pot were sampled at 48 h after culture. Approximately 0.1 g of fresh plant tissues was randomly collected and used in measuring nitrogenase activity. The rest of the plants were frozen in liquid N2 and stored at -80°C for the determination of glutamine synthetase (GS), glutamic-pyruvic transaminase (GPT), Na+/K+-ATPase, Ca2+/Mg2+-ATPase, SOD, POD, and CAT activity and MDA and free amino acid levels. The determination methods of various enzymes were as follows:
Nitrogenase activity was determined by an acetylene reduction assay  using a gas chromatograph. Fresh plant tissues (0.1 g) were placed in 20 ml bottles. Two layers of cheesecloth wetted with 0.2 ml of water were placed in each bottle to supply moisture. Bottles were capped with rubber serum stoppers, and reactions were incubated at 25°C for 30 min under 0.25 atm O2, 0.65 atm argon, and 0.1 atm acetylene. The reactions were initiated by the injection of acetylene and were terminated by adding 0.5 ml of 50% trichloroacetic acid with a syringe. Ethylene was assayed by the procedure utilized by Koch and Evans . Appropriate control reactions were included.
GS activity was measured by a γ-glutamylhydroxamate assay , and one unit (U) of activity was defined as an absorbance change of 0.01 per min at a wavelength of 540 nm. One unit of GS activity is the amount of enzyme catalyzing the formation of 1 μmol γ-glutamylhydroxamate per min at 25°C.
GPT activity was determined by a modified method of Cabaud et al. , which was originally used for the determination of transaminase activity in serum samples. Approximately 0.1 g of a frozen sample was homogenized at 4°C in 0.05 M Tris-HCl extraction buffer (pH 7.2). The crude extract was centrifuged at 8,000g for 10 min. The enzyme activity in the supernatant was measured at 37°C in a reaction buffer containing 200 mM DL-alanine and 2 mM α-ketoglutarate. Around 30 min later, the reaction was stopped by adding 0.5 mL of 2, 4-dinitrophenylhydrazine. The absorbance of the reaction solution (pyruvic acid) was measured at a wavelength 505 nm. Enzyme activity was calculated from a standard calibration curve, and 1 U of the activity corresponded to the formation of 1 μmol pyruvic acid per 30 min.
Na+/K+- and Ca2+/Mg2+-ATPase activity was determined by measuring the amount of inorganic phosphate (Pi) released from ATP , and 1 U of the activity corresponded to 1 μmol Pi released per hour. The enzymatic reactions were performed using assay kits (Jiancheng Biological Engineering Research Institute, Nanjing, China).
SOD activity was determined by monitoring the rate of reduction of nitroblue tetrazolium (NBT) , and 1 U of enzyme activity was defined as the amount of enzyme causing a 50% inhibition in the NBT reduction rate. Frozen sample (0.5 g) were homogenized in an ice cold 100 mM EDTA–phosphate buffer (pH 7.8). The homogenate was centrifuged (10,000×g) for 20 min at 4°C and supernatant was used as the enzyme extract. The activity was determined by using the photochemical p–nitrobluetetrazolium chloride (NBT) reduction method.
POD activity was measured using the guaiacol oxidation method . Frozen sample (0.1 g) was homogenised in 2 mL 50 mM phosphate buffer (pH 6.1). The homogenate was centrifuged at 10,000×g and the supernatant was used as enzyme extract. Peroxidaseactivity was determined with guaiacol as the substrate in a total volume of 3 mL. Increase in the absorbance due to oxidation of guaiacol (ε = 25.5 mM−1 cm−1) was measured at 470nm. 1 U of enzyme activity was defined as an absorbance change of 0.01 per min at a wavelength of 470 nm.
CAT activity was determined by monitoring the rate of decomposition of H2O2 . Frozen sample (0.1 g) was homogenised with 3 mL 50 mM potassium phosphate buffer containing 1 mM EDTA (pH 7.0) and centrifuged at 6000×g for 15 min. The supernatant obtained was used as enzyme extract. The concentration of H2O2 in each sample was calculated using the extinction coefficient (ε = 39.4 mM−1 cm−1) and and one unit (U) of CAT activity is the amount of enzyme dissociating 1 nmol H2O2 min−1 at a wavelength of 240 nm.
The concentration of MDA was assayed by measuring the amount of thiobarbituric acid reactive substances . Frozen sample (0.5 g) were crushed in 10 mL of 80:20 (v/v) ethanol and water solution, followed by centrifugation at 3000×g for 10 min. Absorbance was read at 440, 532 and 600 nm with the help of appropriate equations the TBARS (thiobarbituric acid reactive substances) as MDA equivalents were determined.
The concentration of free amino acids was determined by the ninhydrin colorimetric method . Frozen sample (0.5 g) was crushed in 5 ml of 10% acetic acid and diluted to 100 ml with distilled water. Then the extract was filtered. The total free amino acids were determined by taking 1 ml of filtered liquid 3 ml of hydrogenated ninhydrin reagent in 25 ml volumetric flask. The mixture was heated for 15 min in boiling water bath and cooled to room temperature. It was then diluted to 25 ml with distilled water and laid aside for 10–15 min. The optical density of the solution was checked at 570 nm by spectrophotometer, using distilled water as a blank. Using leucine as a standard, standard curve was prepared by plotting the absorbance of a series of working standards against their respective concentrations.
Results and discussion
Effect of urea application on N metabolism in Azolla
All results described hereafter are presented relative to that in the control. Urea-treated Azolla showed a 44% decrease in nitrogenase activity (Fig 2A), which is in agreement with the findings of previous studies [2, 8]. Prior to this study, information on the effect of urea application on N metabolism in Azolla was limited. GS is a key enzyme in the assimilation and control of N metabolism; it is the first enzyme involved in the conversion of inorganic to organic N in most organisms . Our study showed no significant difference in GS activity between the urea-treated and control Azolla ferns (Fig 2B), indicating that N assimilation is not affected by urea application. However, further investigations are needed to comfirm this speculation, because glutamate dehydrogenase (GDH) is can also perform this function in the aminating direction, especially under high levels of NH4 . In addition, 660% increase in GPT activity after urea treatment was observed (Fig 2C). GPT is a key regulatory enzyme that catalyzes amino from glutamic acid to transfer pyruvic acid to form alanine and α-ketoglutaric acid . This enzyme plays an important role in plant adaptation to adverse conditions, including changes in amino acid metabolism to maintain a supply of amino groups to ensure immediate protein synthesis upon cessation of the adverse conditions . Moreover, free amino acids are known to play significant roles in osmotic adjustment in plants subjected to adverse conditions . Increases in the free amino acid pool (11–52%) have been reported in various plants exposed to salt and temperature stress [31, 32]. Similarly, in this study, concentration of free amino acids was 39% higher with urea treatment (Fig 2D). These results also reveal that urea application induced an increase in amino acid metabolism in Azolla. Amino acid metabolism is a common process in N assimilation, carbon fixation, and secondary metabolism . Fahnenstich et al.  reported that amino acids derived from tricarboxylic acid (TCA) cycle acids increase whereas those from glycolysis intermediates decrease in glycolate oxidase-overexpressing Arabidopsis. Ishikawa et al.  observed that increased amino acid metabolism depletes glycolysis and TCA cycle intermediates in rice cells under oxidative stress. In this study, we did not determine the effect of urea application on the amino acid composition in Azolla. Therefore, further studies are needed to determine the effect of urea application on the physiological functions and regulatory mechanisms governing amino acid metabolism in Azolla.
Effect of urea application on membrane lipid peroxidation in Azolla
The urea treatment resulted in a 22% increase in MDA concentration in Azolla (Fig 3A). Membrane lipid peroxidation can cause further deleterious consequences such as inhibition of ATPase activity [11, 12]. This is supported by the observations in the present study that Na+/K+-ATPase and Ca2+/Mg2+-ATPase activity after urea treatment was respectively 21% and 16% lower (Fig 3B and 3C). Although antioxidative enzymes are produced to protect plant cells against oxidative injury , free radicals can not be scavenged thoroughly in various plants due to the excessive amount of free radicals generated or the severe weakening of its scavenging capacity . A lower scavenging activity in plants indirectly reflects a decrease in the activity of antioxidative enzymes . In this study, urea treatment resulted in an 11% decrease in SOD activity (Fig 3D). The difference in POD activity was not significant between the urea-treated and control ferns (Fig 3E). Azolla subjected to urea treatment showed a 64% higher CAT activity (Fig 3F). These results indicate that the decrease in SOD activity was partially responsible for membrane lipid peroxidation in Azolla after urea application. Furthermore, our results also demonstrate that SOD and CAT activities were more sensitive to urea application than POD activity (Fig 3D–3F). These findings suggest that the responses of various antioxidative enzymes to adverse conditions are plant-specific [38–40]. More importantly, the present study showed that the SOD activity response was inversely correlated to that of CAT activity after urea application (Fig 3D and 3F). In plant antioxidative systems, SOD removes O2–, whereas CAT decomposes H2O2 . Therefore, the reduction in SOD activity might have occurred prior to the disruption of the H2O2-scavenging system, and membrane lipid peroxidation in urea-treated Azolla was induced mainly by O2–. However, further investigations should be conducted to confirm these assumptions.
Urea treatment of Azolla induced an increase in GPT and CAT activity and free amino acid and MDA concentrations and a decrease in nitrogenase and SOD activity. These findings indicate that urea application promotes amino acid metabolism and membrane lipid peroxidation in A. pinnata.
- 1. Arora A, Singh PK. Comparison of biomass productivity and nitrogen fixing potential of Azolla SPP. Biomass Bioenerg. 2003; 24: 175–178.
- 2. Cissé M, Vlek PLG. Influence of urea on biological N2 fixation and N transfer from Azolla intercropped with rice. Plant Soil 2003; 250: 105–112.
- 3. Raja W, Rathaur P, John SA, Ramteke P. Azolla: An aquatic pteridophyte with great potential. Int. J. Res. Biol. Sci. 2012; 2: 68–72.
- 4. Wagner GM. Azolla: A review of its biology and utilization. Bot. Rev. 1997; 63: 1–26.
- 5. Lumpkin TA. Environmental requirements for successful Azolla growth. In: Azolla utilization: Proceedings of the workshop on Azolla use, Fuzhou, Fujian, China. Los Baños: International Rice Research Institute; 1987. pp. 89–97.
- 6. Sadeghi R, Zarkami R, Sabetraftar K, Van Damme P. A review of some ecological factors affecting the growth of Azolla spp. Caspian J. Environ. Sci. 2013; 11: 65–76.
- 7. Qin J, Impa SM, Tang Q, Yang S, Yang J, Tao Y, et al. Integrated nutrient, water and other agronomic options to enhance rice grain yield and N use efficiency in double-season rice crop. Field Crops Res. 2013; 148: 15–23.
- 8. Vlek PLG, Diakite MY, Mueller H. The role of Azolla in curbing ammonia volatilization from flooded rice systems. Fertil. Res. 1995; 42: 165–174.
- 9. Surjus A, Durand M. Lipid changes in soybean root membrances in response to salt treatment. J. Exp. Bot. 1996; 47: 17–23.
- 10. Liljenberg CS. The effects of water deficit stress on plant membrane lipids. Prog. Lipid Res. 1992; 31: 335–343. pmid:1287669
- 11. Kuiper PJC. Functioning of plant cell membranes under saline conditions: membrane lipid composition and ATPases. In: Staples RC, Toenniessen GH, editors. Salinity tolerance in plants: strategies for crop improvement. New York: John Wiley & Sons; 1984. pp. 77–91.
- 12. Crawford RMM, Braendle R. Oxygen deprivation stress in a changing environment. J. Exp. Bot. 1996; 47: 145–159.
- 13. Xu J, Li C, Yang F, Dong Z, Zhang J, Zhao Y, et al. Typha angustifoloa stress tolerance to wastewater with different levels of chemical oxygen demand. Desalination 2011; 280: 58–62.
- 14. Yin X, Zhang J, Guo Y, Fan J, Hu Z. Physiological responses of Potamogeton crispus to different levels of ammonia nitrogen in constructed wetland. Water Air Soil Pollut. 2016; 227: 65.
- 15. Zhang J, Kong Y, Wang S, Yao Y. Activities of some enzymes associated with oxygen metablolism, lipid peroxidation and cell permeability in dehydrated Malus micromalus seedlings. Afr. J. Biotechnol. 2010; 9: 2521–2526.
- 16. Turner GL, Gibson AH. Measurement of nitrogen fixation by indirect methods. In: Bergersen FJ, editor. Methods for evaluating biological nitrogen fixation. New York: John Wiley & Sons; 1980. pp. 111–138.
- 17. Koch B, Evans HJ. Reduction of acetylene to ethylene by soybean root nodules. Plant Physiol. 1966; 41: 1748–1750. pmid:16656468
- 18. Chien HF, Lin CC, Wang JW, Chen CT, Kao CH. Changes in ammonium ion content and glutamine synthetase activity in rice leaves caused by excess cadmium are a consequence of oxidative damage. Plant Growth Reg. 2000; 36: 1–7.
- 19. Cabaud P, Leeper R, Wróblewski F. Colorimetric measurement in serum: glutamate-oxalacetate, glutamate-pyruvate transaminase. Amer. J. Clin. Path. 1956; 26: 1101–1105.
- 20. Ohnishi T, Gall RS, Mayer LM. An improved assay of inorganic phosphate in the presence of extractable phosphate compounds: application to the ATPase assay in the presence of phosphocreatine. Anal. Biochem. 1975; 69: 261–267. pmid:129016
- 21. Giannopolitis CN, Ries SK. Superoxide dismutase. I. Occurrence in higher plants. Plant Physiol. 1977; 59: 309–314. pmid:16659839
- 22. Maehly AC, Chance B. The assay of catalases and peroxidases. In: Glick D, editor. Methods of biochemical analysis. New York: Interscience Publishers; 1954. pp. 357–425.
- 23. Aebi H. Catalase. In: Bergmeyer HU, editor. Methods of enzymatic analysis. New York: Academic Press; 1974. pp. 673–677.
- 24. Heath RL, Packer L. Photoperoxidation in isolated chloroplasts. I. Kinetics and stoichiometry of fatty acid peroxidation. Arch. Biochem. Biophys. 1968; 125: 189–198. pmid:5655425
- 25. Rosen H. A modified ninhydrin colorimetric analysis for amino acids. Arch. Biochem. Biophys. 1957; 67: 10–15. pmid:13412116
- 26. Lea PJ, Blackwell RD, Joy KW. Ammonia assimilation in higher plants. In: Mengel K, Pilbeam DJ, editors. Nitrogen metabolism of plants. Oxford: Clarendon Press; 1992. pp. 153–186.
- 27. Miflin BJ, Habash DZ. The role of glutamine synthetase and glutamate dehydrogenase in nitrogen assimilation and possibilities for improvement in the nitrogen utilization of crops. J. Exp. Bot. 2002; 53: 979–987. pmid:11912240
- 28. Chao L, Baofu P, Weiqian C, Yun L, Hao H, Liang C, et al. Influences of calcium deficiency and cerium on growth of spinach plants. Biol. Trace Elem. Res. 2008; 121: 266–275. pmid:17960330
- 29. Huber W, Sankhla N. Eco-physiological studies on Indian arid zone plants. II. Effects of salinity and gibberellin on the activity of the enzymes of amino-acid metabolism in leaves of Pennisetum typhoides. Oecologia 1973; 13: 271–277. pmid:28308582
- 30. Ashraf M, Tufail M. Variation in salinity tolerance in sunflower (Helianthus annuus L.). J. Agron. Crop Sci. 1995; 174: 351–362.
- 31. Fougère F, Rudulier DL, Streeter JG. Effects of salt stress on amino acid, organic acid, and carbohydrate composition of roots, bacteroids, and cytosol of alfalfa (Medicago sativa L.). Plant Physiol. 1991; 96: 1228–1236. pmid:16668324
- 32. Yoon Y, Kuppusamy S, Cho KM, Kim PJ, Kwack Y, Lee YB. Influence of cold stress on contents of soluble sugars, vitamin C and free amino acids including gamma-aminobutyric acid (GABA) in spinach (Spinacia oleracea). Food Chem. 2017; 215: 185–192. pmid:27542466
- 33. Pratelli R, Pilot G. Altered amino acid metabolism in glutamine dumper1 plants. Plant Signal. Behav. 2007; 2: 182–184. pmid:19704691
- 34. Fahnenstich H, Flügge U, Maurino VG. Arabidopsis thaliana overexpressing glycolate oxidase in chloroplasts: H2O2-induced changes in primary metabolic pathways. Plant Signal. Behav. 2008; 3: 1122–1125. pmid:19704454
- 35. Ishikawa T, Takahara K, Hirabayashi T, Matsumura H, Fujisawa S, Terauchi R, et al. Metabolome analysis of response to oxidative stress in rice suspension cells overexpressing cell death suppressor bax inhibitor-1. Plant Cell Physiol. 2010; 51: 9–20. pmid:19919949
- 36. Ge T, Sui F, Bai L, Lu Y, Zhou G. Effects of water stress on the protective enzyme activities and lipid peroxidation in roots and leaves of summer maize. Agric. Sci. China 2006; 5: 101–105.
- 37. Yan B, Dai Q, Liu X, Huang S, Wang Z. Flooding-induced membrane damage, lipid oxidation and actived oxygen generation in corn leaves. Plant Soil 1996; 179: 261–268.
- 38. Xue T, Hartikainen H. Association of antioxidative enzymes with the synergistic effect of selenium and UV irradiation in enhancing plant growth. Agric. Food Sci. Finland 2000; 9: 177–186.
- 39. Tewari RK, Kumar P, Sharma PN. Oxidative stress and antioxidant responses in young leaves of mulberry plants under nitrogen, phosphorus or potassium deficiency. J. Integr. Plant Biol. 2007; 49: 313–322.
- 40. Hafsi C, Romero-Puertas MC, del Río LA, Abdelly C, Sandalio LM. Antioxidative response of Hordeum maritium L. to potassium deficiency. Acta Physiol. Plant 2011; 33: 193–202.