Introduction

Atrazine (6-chloro-N-ethyl-N′-(1-methylethyl)-1,3,5-triazine-2,4-diamine is a synthetic triazine herbicide used to control grassy and broadleaf weeds in sugarcane, wheat, conifers, sorghum, nuts and corn crops (Iriel et al. 2014; Kumar et al. 2016; Zhao et al. 2017). It was first introduced in twentieth century and often used alone or in amalgamation with other herbicides for agricultural applications (Correia et al. 2007; Kadian et al. 2008; Lewis et al. 2009). It is the second most widely consumed pesticide in the world with annual consumption of about 70,000–90,000 tons (Kumar et al. 2013; Cheng et al. 2016). In India, about 340 tonnes of atrazine is consumed annually (Solomon et al. 2013a, b).

Due to its long half-life of 41–231 days (Karlsson et al. 2016), low adsorption in soils and moderate aqueous solubility, it has a sky-scraping potential to contaminate not only agricultural fields, but also ground and surface water with the highest concentration up to 30 μg/L (Table 1) (Cerejeira et al. 2003; Schwab et al. 2006; Kumar et al. 2013). It was banned in several countries like Italy (Huang et al. 2009), Denmark (Glæsner et al. 2014), Finland and Germany (Vonberg et al. 2014) in the year 1991 and European Union banned atrazine in the year 1992 (White 2016) because its metabolites/residues had the potential to persist in fields and surface water for several years (Bethsass and Colangelo 2013; Nousiainen et al. 2015) resulting in the contamination of surface and water bodies. Some studies reveal the surpass levels of atrazine 3 and 0.1 μgL−1 found in drinking water of the USA and Europe (Mahía et al. 2007). The maximum acceptable concentration (MAC) for atrazine in drinking water is 5 µg/L (Cerejeira et al. 2003). The acceptable daily intake (ADI) is derived on the basis of division of a NOAEL by appropriate uncertainty factors. It is one of the most widely used herbicide, and several times it has been reported to be at levels above the limits in water bodies (Graymore et al. 2001).

Table 1 Chemical and physical properties of atrazine and degradation products
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The production of atrazine increased from 0.26 metric tonnes to 0.67 metric tonnes from the year 2009 to 2012 (Ministry of fertilizers and chemicals, Govt. of India). Environment protection agency classifies atrazine in toxicity class III on a scale of I to IV (I being the most toxic). It is registered for only two crops (apple and sugar) by the central insecticides board and registration committee in India (Bhushan et al. 2013). Atrazine has been classified as an endocrine disrupting pesticide by the US Environmental Protection Agency (Morales-Pérez et al. 2016). The International Agency for Research on Cancer (IARC) has categorized atrazine in the list of carcinogenic pesticides (Mahler et al. 2017). This review covers the toxicity, analytical monitoring by using recent techniques and degradation (chemical, photochemical and microbial) aspects of atrazine.

Toxicity of atrazine

Agricultural chemicals have potential to alter species composition, decrease diversity, interfere with normal succession patterns and alter food webs as a whole (Lin et al. 2016a, b, c). The intensification of industrial and agricultural practices chiefly the utilization of pesticides has in almost every way made our natural resources miserable (Dutta et al. 2016). Various toxic effects of atrazine on humans, plants, animals and microorganisms are displayed in Fig. 1.

Fig. 1
figure 1

Toxicity and detrimental effects of atrazine on humans, plants, animals and microorganisms

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Effect on nontarget plants

One of the most unintentional exposures of pesticides is to the nontarget plants. Atrazine has shown to manifest complete death or stunted growth, translocation, root or shoot uptake, phenotype alteration, mutation and resistance (Burken and Schnoor 1997; Szigeti and Lehoczki 2003; Hassan and Nemat 2005; Nemat and Hassan 2006; Su and Zhu 2006). The different results of atrazine accumulation could be due to the use of different plant species, since distinct mechanisms control the accumulation of atrazine in target and nontarget plants. Concentration- and time-dependent effects of atrazine exposure have been noticed. Atrazine is absorbed by shoots and roots and transported solely by xylem (Francisco 2001; Szigeti and Lehoczki 2003). Pre-emergence herbicides are used to stop the germination of plant seeds. Atrazine is absorbed by leaves as well as roots and has postemergence as well as pre-emergence activity on weeds (Ahrens and Newton 2008). The negative effects of postemergence atrazine treatment upon peroxidase, ascorbate peroxidase and lipid peroxidation were determined in a 15-day experiment (Akbulut and Yigit 2010). Target and nontarget plants exposed to atrazine usually suffer oxidative stress caused by the generation of reactive oxygen species such as superoxide anion radical, hydroxyl radical, singlet oxygen and hydrogen peroxide. Generation of H2O2 in roots of maize exposed to atrazine was also assayed after chemical extraction in a previous study. The most important mechanism used by plants to prevent oxidative stress is through reactive oxygen species scavenging (Burken and Schnoor 1997; Hassan and Nemat 2005; Nemat and Hassan 2006; Szigeti and Lehoczki 2003; Su and Zhu 2006). It has been noticed that the exposure and accumulation of atrazine cause oxidative toxicity and antioxidant response in Zea mays (Li et al. 2012). Gao et al. (2011) indicated atrazine to be a potential threat to seagrass seedling function, and the impact is much higher for adult plants. Exposure to 10 μg/L atrazine significantly lowers the plant fresh weight and total chlorophyll concentration, and up to 86.67% mortality was recorded at the 100 μg/L concentration.

Basically plants have detoxification ability against various pesticides. Enzymes detected in plants responsible for detoxification of various pesticides are cytochrome 450, peroxidases, aryl acylamidases, esterases, lipases, proteases, amidases, oxygenases and reductases (Jiang et al. 2016). It has been noticed that the increasing antioxidant enzyme activities enable Pennisetum americanum seedlings to cope with the oxidative stress induced by moderate concentrations (20 mg kg−1 or below) of atrazine. Cytochrome P450 monooxygenase genes are known to be involved in modification and detoxification of herbicide atrazine in Oryza sativa (McGregor et al. 2008; Tana et al. 2015). Exposure to atrazine can trigger specific GT genes and enzyme activities in Oryza sativa (Lua et al. 2013). Another major detoxification mechanism in leaf tissue of maize is through glutathione conjugation (GS-atrazine). It is considered to be an important biotransformation mechanism of a atrzine in plants. The recovery of atrazine-inhibited photosynthesis is accompanied by a rapid conversation of atrazine to GS-atrazine when atrazine was directly introduced into the leaf tissue. This pathway is relatively inactive in roots and shoots (Shimabukuro et al. 1970). Another detoxification pathway for atrazine in corn corresponds to a chemical hydroxylation process. The mechanism was also described under invitro conditions. Benzoxazinones mixture (DIMBOA, DIBOA, 2-monoglucosyl DIMBOA + 2-monoglucosyl DIBOA) extracted from corn plantlets were able to transform 91% of atrazine into 2-hydroxyatrazine in 24 h. The natural concentration of benzoxazinones in the vacuolar sap of corn seedlings and the pH play a major role in high rate of atrazine chemical hydroxylation in vivo (Raveton et al. 1997). Occurrence of atrazine in water bodies can have serious detrimental effects on nontarget living organisms such as freshwater algae (Baxter et al. 2015, 2016; Bai et al. 2015; Andrus et al. 2013). Atrazine inhibited the growth of Chlamydomonas mexicana and also leads to an increase in carbohydrate content and chlorophyll a accumulation (Kabra et al. 2014). The toxicity of atrazine along with its metabolites desethylatrazine and deisopropylatrazine was evaluated on the amphipods Hyalella azteca and Diporeia sp., and the unicellular algae Pseudokirchneriella subcapitata. It was found to be the most toxic followed by desethylatrazine and deisopropylatrazine and algae being the most sensitive of all. In case of chronic exposure, Diporeia sp. was found to be sensitive as compared to H. azteca by a large magnitude (Ralston-Hooper et al. 2009). Photosynthetic process, cell division, lipid synthesis were majorly affected in green alga, Raphidocelis subcapitata on exposure to atrazine (Ma et al. 2006). According to Solomon et al. (2013a, b), atrazine was not found to cause any lethality or permanent cell damage, but it acts to inhibit photo-phosphorylation.

Effects on aquatic fauna

Atrazine exhibited significant rate of micronuclei and nuclear abnormalities in Channa punctatus (Nwani et al. 2011). Atrazine showed acute toxicity to leopard frog (Rana pipiens), American toad (Bufo americanus), rainbow trout (Onchorhynchus mykiss) and channel catfish (Ictalurus punctatus) (Orton et al. 2006). Chironomus tentans was also studied for the effect of atrazine and binary combination of atrazine with chlorpyriphos. The result depicted atrazine not acutely toxic at even higher levels but when used in combination with chlorpyriphos, methyl parathion and malathion decreases the EC50 (effective concentration) values (Belden and Lydy 2001).

Also the toxicity of active ingredients to pesticide formulations with regard to atrazine, chlorpyriphos and permethrin in glochidia and juvenile life stages of a freshwater mussel (Lampsilis siliquoidea) were compared. The atrazine formulation (Aatrez) was more toxic than technical grade atrazine in chronic tests with juvenile L. siliquoidea. For other pesticides, acute and chronic toxicity of technical grade pesticides were similar to the toxicity of pesticide formulations. Atrazine and formulations did not cause any significant acute toxicity in glochidia and juveniles (Bringolf et al. 2007). In case of freshwater fish, Rhamdia quelen, histopathological changes in liver revealed leukocyte infiltration, hepatocyte vacuolization like steatosis and necrosis areas, leading to raised lesion index levels in all tested concentrations; process of osmoregulation was disturbed and gills showed changes in pavement cells and chloride cells (Mela et al. 2013). The effect of atrazine was also evaluated on some immune parameters of red-eared slider (Trachemys scripta). Lowered serum complement and lysozyme activities, reduced leukocyte number as well as their phagocytic activity and increased neutrophil/lymphocyte ratio depicted a positive correlation between atrazine (high dose) concentration and immunosuppressive effects (Soltanian 2016). The expression of carp HSP70 and 70-kDa heat shock cognate protein (HSC70) with atrazine and chloropyriphos treatment alone or in combination was significantly up-regulated in common carp (Cyprinus carpio L.) providing new insights into the mechanisms used by fish to adapt to stressful environment (Xing et al. 2013). Increased lipid peroxidation and decline in cholesterol and total proteins in liver and muscles were observed for atrazine, glyphosate and quinclorac in tadpoles of Lithobates catesbeianus (Dornelles and Oliveira 2014).

Freshwater clam, Corbicula fluminea, was also studied to evaluate the biochemical and genotoxic effects of the herbicides atrazine and Roundup. Atrazine interfered mostly in biotransformation, while Roundup interfered mainly in antioxidant defenses leading to lipid peroxidation. Herbicides mixture caused a significant increase in the occurrence of DNA damage (Dos-Santos and Martinez 2014). Atrazine shows profound influence on the oxidative stress markers and detoxifying enzyme of the exposed zebra fish (Blahova et al. 2013). Atrazine also causes changes in the glutathione S-transferase isoenzymes (GSTs) activity and their transcription varied within each organ and among organs of common carp (Xing et al. 2012). Atrazine behaves as enzyme inhibitor, impairing hepatic metabolism, and produces genotoxic damage to different cell types as studied in Plotosus lineatus (Santos and Martinez 2012). Various concentrations of atrazine also lead to continuous decline in levels of total protein and serum albumin in grass carp, Ctenopharyngodon idella (Khan et al. 2016a, b). Atrazine is also known to show detrimental effects on the digestive gland of Crassostrea gigas, pacific oyster by modulating important molecular and biochemical parameters within relatively short time period (Lee et al. 2017). Exposure to atrazine may be associated with decreased birth weight and preterm delivery. According to Zadeh et al. (2016), hematological parameters like hemoglobin, hematocrit and RBCs were significantly decreased by chronic toxicity of atrazine in fish, Tor grypus exposed to different levels of atrazine. The increase in concentrations of lactate dehydrogenase (LDH) and decrease in concentrations of creatinine phosphokinases (CPK), serum glutamic-pyruvic transaminase (SGPT) and alkaline phosphatase indicate an adverse effect of atrazine on grass carp, Ctenopharyngodon Idella (Khan et al. 2016a, b). In Channa punctatus, the biochemical parameters such as serum total protein, glucose and cholesterol values were found to decrease, while level of urea significantly increased in all treatments suggesting anemia and hepatic damage. In a risk assessment study of atrazine in American surface waters, it was found that phytoplankton were the most sensitive organisms followed by macrophytes, benthic invertebrates, zooplanktons and fish. In estuarine crab, Neohelice granulata significant decrease in glycogen content and significant diminished content of vitellogenin proteins in ovary was detected after atrazine toxicity (Silveyra et al. 2017). In zebrafish, Danio rerio decrease in glutathione S-transferase and catalase and an increase in superoxide dismutase, glutathione peroxidase and reductase were observed indicating profound influence of atrazine on the oxidative stress markers and detoxifying enzymes (Blahova et al. 2013). Atrazine also affects gill respiration and ion regulation function of fingerlings (Caspian kutum) by damaging tissue, pavement cells and ionocytes (Khoshnood et al. 2015).

Effects on other invertebrates and vertebrates

Oluah et al. (2016) studied the toxicity and the histopathological effects of Atrazine on earthworms, Nsukkadrilus mbae, and reported significant adverse effects. Xu et al. (2006) compared atrazine and chlorotoluron toxicity on Eisenia fetida and found atrazine as more toxic to earthworms; combination showed synergistic effect. Superoxide dismutase (SOD) activity showed an increase. The exposure of chlorpyrifos to E. fetida in combination with atrazine or cyanazine was evaluated, in which the resultant toxicity was greater than the additive (Lydy and Linck 2003). Previous studies suggest that the pessimistic effects of atrazine on neuro-endocrine system occur by changing pituitary hormone levels, such as follicle-stimulating hormone (FSH) and luteinizing hormone (LH) (Cox 2001; Yang et al. 2014). Altered LH leads to prolonged prolactin secretion and subsequent stimulation of the mammary gland proliferative changes and increased incidence of mammary adenocarcinomas and fibroadenomas (Jowa and Howd 2011; Simpkins et al. 2011). Atrazine inhibits the release of gonadotropin-releasing hormone (GnRH), which decreases the secretion of LH that may lead to increased abortions in male Wistar rats (Stoker et al. 2002). Exposure to atrazine affects both germ cells as reduced motility and sperm counts in male rats (Victor-Costa et al. 2010: Pogrmic et al. 2009). Effects of atrazine were also observed as a decrease in levels of serum lipids and liver enzymes in adult mammals (Suzawa and Ingraham 2008: Hayes et al. 2002). Atrazine supposedly increases aromatase enzyme activity via inhibition of phosphodiesterase, which increases the aromatization of testosterone to estrogen (Hayes et al. 2006; Cooper et al. 2007). An increased estrogenic environment may favor altered relative sex hormone levels may affect reproduction and/or development and induction of cancers and proliferation of preexisting estrogen-dependent cancers (Oka et al. 2008). Other effects of metabolites of atrazine on the immune system, central nervous system and cardiovascular function have also been reported (Jin et al. 2010; Christin et al. 2003). In adult humans, non-Hodgkin’s lymphoma associates with the exposure of the compound (Schroeder et al. 2001). According to Gely-Pernot et al. (2015), atrazine exposure interferes with normal meiosis, which affects spermatozoa production in male mice. Oxidative stress and disruptions in calcium homeostasis play an important role in the induction of immunotoxicity in mice by atrazine as depicted by Gao et al. (2016). It is also found to elicit immunotoxic effects on murine lymphocytes, and its presence in the environment might compromise immune function in organisms (Chen et al. 2015a, b). Atrazine is known to reduce the mating rate, number of progeny and competitive fertilization ability; it also alters gene expression and production of proteins in Drosophila melanogaster (Vogel et al. 2015). Marcus and Fiumera (2016) observed reduced pupation rate, emergence rate and longevity of adult in D. Melanogaster after atrazine exposure. Atrazine was also reported to be endocrine disrupters and significant decrease in steroid levels (testosterone and 17β-estradiol), and total proteins were also noted; the histology of ovotestis showed degenerative changes including azoospermia and oocytes deformation (Omran and Salama 2016). 57% reduction in testicular volume was marked in atrazine exposed tadpoles. Also, primary spermatogonial cells and nurse cells were reduced by 70 and 74%, respectively (Tavera-Mendoza et al. 2002). Toxicity profiles of atrazine against microorganisms, aquatic lower invertebrates, higher vertebrates and humans are presented in Table 2.

Table 2 Toxicity and adverse effects of atrazine on humans, aquatic lower invertebrates, higher vertebrates and soil microorganisms
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The prime target of chlorinated atrazine on humans and mammals is the disruption of the endocrine system (Jin et al. 2014; Kroon et al. 2014; Weber et al. 2013). Secondly, it also induces oxidative stress by formation of reactive oxygen species resulting in the reduced semen quality sperm dysfunction and infertility of amphibians, rats and pigs (Gely-Pernot et al. 2015; Jestadi et al. 2014; Kniewald et al. 2000). In females, pesticides imbalances sexual hormones intervene androgen or estrogens receptors for improper function, irregularities of ovarian cycles, instinctive abortion, defect in developmental births, etc. (Bohn et al. 2011). Atrazine forms ROS, which cause single- and double-strand breaks in DNA and thus is genotoxic (Yang et al. 2010). The working of cardiovascular system also gets affected by atrazine exposure (Lin et al. 2016a, b, c; Cosselman et al. 2015). Atrazine is known to cause hepatic damage, as liver is the primary organ for atrazine metabolism in mammals (Campos-Pereira et al. 2012; Gojmerac and Kniewald 1989). Atrazine is also responsible for cardiotoxicity and hepatotoxicity in mice by disruption of ionic balance (Lin et al. 2016a, b).

The negative effects of atrazine on soil and aquatic biota are enormous. The above studies decipher a clear picture about the vulnerability of atrazine to terrestrial and aquatic life forms. The detection of atrazine with better accuracy and reliability is thus essential for safety of human health, biota and environment.

Analytical methods for monitoring atrazine in ecosystems

Basically, single environmental matrix can contained multiple pesticides (Kaur et al. 2017; Kumar et al. 2016, 2017; Singh et al. 2016). The extraction, cleanup and pre-treatment procedure depend upon the physiochemical nature of pesticides and environmental matrix (Kaur et al. 2017; Kumar et al. 2016, 2017). There are number of methods described for the analysis of different classes of pesticides in different matrices (AOAC 1993; Kumar et al. 2015a, b, c, d; Prasad et al. 2013; Kumar and Singh 2016; Singh et al. 2017). In day-to-day laboratory analysis, AOAC method is most authentic. Most common steps for the extractions of pesticides including atrazine have been described by Kumar et al. (2015a, b, c, d) as per the guidelines of AOAC (AOAC 1993; Kumar et al. 2015a, b, c, d; Prasad et al. 2013; Singh et al. 2016, 2017).

Highly sensitive and rapid analytical methods are essential for monitoring of residual atrazine and its metabolites in soil and water bodies (Table 3). It is monitored by using spectroscopic, immunogenic and chromatographic methods which include infrared (IR) spectroscopy, high-performance liquid chromatography (HPLC)/HPLC–MS/HPLC–MS/MS, HPLC, enzyme-linked immunosorbent assay (ELISA) and gas chromatography GC/GC–MS/GC–MS/MS (Miensah et al. 2015; Williams et al. 2014; Bonansea et al. 2013). Recently nano-based solid-based extractions ultra-performance liquid chromatography (UPLC) and GC/LC–MS methods have been developed with very good recovery and detection limits up to ppb (Table 3). The detection of atrazine in biological media is quantified using GC coupled with MS including different detectors like flame ionization detector (Haiyan and Center 2015), electron capture detector (ECD) (Miensah et al. 2015), nitrogen-phosphorus detector (NPD) (Bonansea et al. 2013) and GC coupled with mass spectrometer (Williams et al. 2014).

Table 3 Analytical techniques applied for the detection and quantitative estimation of atrazine and derivatives in various ecosystems
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Quantification of atrazine was also done by a diode array (Yang et al. 2014) and a UV detector (Kong et al. 2016). Immunogenic methods are usually based on ELISA using sheep-based antibodies to atrazine (Na et al. 2012). Other immunogenic methods have been developed in which the antibody is bound to a “dipstick,” and this is used to evaluate concentrations of atrazine in water or liquid food samples (Kaur et al. 2007), while other sampling approaches have used immuno-affinity systems to concentrate atrazine prior to analysis by GC (Tran et al. 2013).

In water samples, atrazine was detected with GC equipped with nitrogen phosphate detector (NPD) and the percentage recovery is very high 96–98% (Nsibande et al. 2015). In sub-surface waters, the atrazine was quantified by both the techniques, i.e., HPLC and GC equipped with NPD, ECD and MS detectors (Barchanska et al. 2014). In aquatic plants and sediments, the concentration of atrazine was quantified by GC coupled with ECD detectors and the recovery percentage is near about 90% (Bennett et al. 2000). In water and soil samples, the solid-phase extraction method is used to concentrate the samples prior for analysis, all the three methods, i.e., GC coupled with all three detectors mentioned above, ELISA and HPLC equipped with UV detector (Hernandez et al. 2000; Lioy et al. 2000). The detection limit of atrazine in biological media moves to a greater extent as it was 0.1 µg/L in the year 2000 (Yokley et al. 2000) and it moves to 50 ng/L in the year 2008 (Gervais et al. 2008). It is all due to the new innovations in day-to-day life. In 2000, the detection was only SPE and GC based. Using styrene di-vinyl benzene sorbents, the detection limits increase to 10 ng/L (Bruzzoniti et al. 2006). Tandem mass spectroscopy combined with ultra-performance liquid chromatography detects up to 50 ng/L of the atrazine in biological media (Kuklenyik et al. 2012).

Qie et al. (2013) developed a technique direct competitive ELISA method (dcTELISA) based on thermistor enzyme for faster detection of atrazine in large-scale samples. In this method, ATZ competes for β-lactamase-labeled ATZ (ATZ–E) for the binding sites on anti-ATZ monoclonal immune response (mAb) which is covalently linked to form immunocomplexes from immune reactor with β-lactamase-labeled ATZ and atrazine. The detection limit was very good with high recovery rate (88–107%) in silage and fresh corn stalk samples. Another novel method for detection of atrazine was based on the SPR (Sepia pterin reductase) determination of P450 mRNA levels in Saccharomyces cerevisiae. The selected oligonucleotide probe that exhibits specificity for P450 mRNA was successfully immobilized on the sensor chip. The mRNA was quantified. It is a highly sensitive and rapid method that permits the detection of atrazine within 15 min (Kim et al. 2003). Kaoutit et al. (2004) also gave a simple conducting polymer-based biosensor approach. A glassy carbon electrode was prepared which is operated at open circuit and served for the immobilization of the enzyme polyphenol oxidase (PPO) during the anodic electropolymerization of polypyrrole (PPy). The concentration of atrazine in aqueous solution is attributed to its inhibitory power toward the catalytic activity of PPO. This biosensor helps in easy detection of photosynthetic inhibiting herbicide monitoring because of its analytical performance and simplicity. A protein-based conjugate method was developed for binding of atrazine with anti-atrazine monoclonal antibodies. Here immobilization was done on gold particles based on test strip method assessed with a photometric device to detect atrazine in very low limits (130 ng/mL) (Byzova et al. 2010). An immunochromatography (ICG) strip test for detection of atrazine in water samples was also developed. Monoclonal antibody (MAb) specific to atrazine was produced from the cloned hybridoma cell (AT-1-M3) and used to develop a direct competitive enzyme-linked immunosorbent assay (DCELISA) and ICG strip. The limit of detection was 3 ng/mL, and it requires only 10 min getting the results and that too in a single step. It came out to be a sensitive and accurate technique (Shim et al. 2006). A disposable immunomagnetic electrochemical sensor involving magnetic particles was developed for the detection of atrazine. The sensor was based on a magnetic monolayer of magnetic particles coated with streptavidin, formed on a gold electrode after application of a magnetic field. The atrazine interacts with biotinyl–Fab fragment K47, and the immune reactions were characterized by impedance spectroscopy. A decrease in electron transfer resistance was observed which could be attributed to rearrangements in the magnetic monolayer. The limit of detection is in the range 10–600 ng/ml; it is a sensitive approach for detection of atrazine acting as an antigen (Helali et al. 2006). Another novel immunoassay was formulated that involved direct coating of haptens on microtiter plates for detection of atrazine. The assay allows hapten-coated plates and uses affinity-purified atrazine which showed very high sensitivity. The limit of detection for atrazine is in the range 0.02–0.7 ng/mL (Suri et al. 2008). Detection of atrazine and triazines in water has also been carried out flow injection chemiluminescence analysis. The aliphatic amines in traizines react with tris(2,2′-bipyridyl)ruthenium(III) to produce chemiluminescence. The presence of natural organic matter (NOM) significantly increased the chemiluminescence, masking the signal generated by atrazine. Isolating the target analyte via solid-phase extraction (SPE) prior to analysis removed this interference and concentrated the samples. The detection limit is 14 ± 2 ng/L (Beale et al. 2009). Pardieu et al. (2009) devised an electrochemical sensor based on molecularly imprinted conducting polymer (MICP). The recognition of atrazine can be quantified by the variation of the cyclic voltammogram of MICP. It shows selectivity toward triazine family and shows a large range of detection from 10−9 to 1.5 × 10−2 mol/L. Impedance spectroscopy transduction combined with the immunosensor technology has been used for the determination of atrazine. The immunoreaction of atrazine on the attached anti-atrazine antibody leads to an increase in the charge transfer resistance which is proportional to the concentration of atrazine. Its limit of detection was 10 pg/mL (Ionescu et al. 2010). Another approach based on antibody replicas for atrazine detection was formulated by Schirhagl et al. (2011). Antibodies were used to pattern the nanoparticles for surface imprinting the polymer layer to produce replicas. Liu et al. (2011) constructed a MIP (molecularly imprinted polymers) chemosensor from a core–shell nanostructure. Vinyl-substituted zinc(II) protoporphyrin (ZnPP) was used as both fluorescent reporter and functional monomer to synthesize atrazine-imprinted polymer shell. The limit for detection is 1.8 mM. Bioluminescent reported bacteria are also utilized for detection of atrazine. Increase in bioluminescent signals is recorded for E. coli. For better interaction between insoluble atrazine and bacterial cells, centrifugation of bacterial cells and analyte dilutions can be performed (Jia et al. 2012). An electrochemical immunosensor for atrazine detection was developed by immobilization of gold nanoparticles on the gold electrode surface. The increase in surface area of work electrode leads to more anti-atrazine monoclonal antibodies capture. Ferricyanide was used as an electrochemical redoxindicator; immunosensor was characterized by cyclic voltammetry and electrochemical impedance spectroscopy. The limit for detection is as low as 0.016 ng/mL (Liu et al. 2014). A field effect transistor was developed based on network of single-walled carbon nanotubes which constitute carbon nanotubes field effect transistors and act as conductor channel for the determination of atrazine in various biological samples with detection limit up to 0.001 ng/mL (Belkhamssa et al. 2016).

Photochemical degradation of atrazine

Several chemical methods have been developed for the degradation during time to time. Konstantinou et al. (2001) have studied the photocatalytic degradation of atrazine and other s-triazine herbicides by using particulate TiO2 as photocatalyst under simulated solar light. The degradation process is highly efficient with traces of atrazine (at ppb level) being decomposed in very short times to less than 0.1 ppb. The process has been shown to lead to the formation of 2,4,6-trihydroxy-1,3,5-triazine (cyanuric acid) as the final product of the degradation process for all the investigated herbicides with several intermediates, rather than to the complete mineralization often observed for other classes of substrates. Monitoring of degradation products has been done using liquid, gas and ion chromatography, and the overall degradation process has been monitored through dissolved and particulate organic carbon measurements.

Atrazine degradation by Fenton’s reagent has been determined as a function of reagent’s concentration and ratios and pH in batch treatments (Barbusiński and Filipek 2001). The actual number and nature of oxidation products have been shown to vary with the concentration. The optimum Fenton’s reagent treatment has been achieved with 1:l molar ratios of FeSO4 and H2O2 (2.69 mM), producing two main terminal products viz, 2-chloro-4,6-diamino-s-triazine (23%) and 2-acetamido-4-amino-6-chloro-s-triazine (28%). Chloride release of 55 ± 9% indicated that dehalogenated s-triazines accounted for the balance of 14C. Atrazine degradation has been found to be pH dependent and decreases from 99% at pH 3 to 37% at pH 9.

In a modified approach, the effects of an inorganic ligand tetrapolyphosphate on the molecular oxygen activation and the subsequent aerobic atrazine degradation by Fe@Fe2O3 core–shell nanowires were investigated at a pH range of 6.0–9.0 (Wang et al. 2014a, b). It was observed that the addition of tetrapolyphosphate enhanced the rate of aerobic atrazine degradation dramatically (955 times) which was even 10 times that of the traditional organic ligand ethylenediamine tetraacetate. The rate enhancement has been attributed to enhanced reduction in Fe(III) to Fe(II) and the subsequent activation of molecular oxygen, owing to the suppressed hydrogen evolution, in the presence of tetrapolyphosphate, from the reduction in proton by Fe@Fe2O3 core–shell nanowires, making more electrons available for the reduction in Fe(III). Moreover, the complexation of tetrapolyphosphate with ferrous ions ensures enough soluble Fe(II) for Fenton reaction and also provides another route to produce more hydroxyl (·OH) radicals in the solution via the single-electron molecular oxygen reduction pathway, thus increasing the rate dramatically.

Ozone being a powerful oxidant has been used in the presence of hydrogen peroxide for degradation of atrazine. Tandem solid-phase extraction procedure was used which includes a C18 reverse-phase support and a strong cation exchanger (Nélieu et al. 2000). It was found that ammeline (2, 4-diamino-6-hydroxy-s-triazine) is the major end product (20% at pH 8) and 2-chloro-4,6-diamino-s-triazine as competitor whose ratio was dependent on the hydroxyl radical content. A number of new intermediates identified were aminoaldehydes and a carbinolamine (Nélieu et al. 2000). Triazine has also been degraded by electrochemical advanced oxidation processes such as anodic oxidation, electro-fenton and photoelectro-fenton using a small open and cylindrical cell with a boron-doped diamond anode. Anodic oxidation has been carried out either with a stainless steel cathode or an O2 diffusion cathode able to generate H2O2 formed at the boron-doped diamond surface in all electrochemical advanced oxidation processes. In the bulk form, Fenton’s reaction between added Fe2+ and electrogenerated H2O2 in electro-Fenton and photoelectron-Fenton, Hydroxyl radical (·OH) is the main oxidant (Borras et al. 2010). It has been observed that almost overall mineralization (94%) is achieved. Atrazine decay always follows a pseudo-first-order reaction, being more rapidly destroyed from ·OH in bulk than at the boron-doped diamond surface. The formation of dealkylated aromatic intermediates such as desethylatrazine and desethyldesisopropylatrazine and cyanuric acid has been revealed by reverse-phase HPLC, and short linear carboxylic acids such as formic, oxalic and oxamic have been identified and quantified by ion-exclusion HPLC. It has been observed that all initial nitrogen is transformed into NO3 − and NH4 + ions (followed by ionic chromatography) in electro-fenton and photoelectro-fenton but not in anodic oxidation, where 36% of nitrogen is released from the solution probably as volatile NO x species.

The recent results showed approximately 80% of atrazine was degraded by ozonation in the presence of hydroxylamine, while only 20% was degraded by ozonation alone. The atrazine degradation involved dealkylation, dechlorination–hydroxylation and olefination (Yang et al. 2016a, b). It has been found that at lower pH the degradation efficiency of atrazine was enhanced by UV/chlorine compared to UV or chlorine alone. The oxidation products of atrazine resulting from dealkylation, dechlorination–hydroxylation, alkylic-hydroxylation, alkylic-oxidation, alkylic-hydroxylation dehydration, deamination–hydroxylation and dechlorination–hydrogenation in UV/chlorine process were detected, which were slightly different from those formed in UV/H2O2 (Kong et al. 2016). Similar products were noticed when nitrite was added to enhance atrazine degradation during ozonation (Yang et al. 2016a, b). The experimental results of electrophotocatalytic reduction (hydroxyl radical reduction) of atrazine with an initial concentration of atrazine (100 mg/L) show that more than 99% of atrazine oxidation was obtained after 30 min of treatment and reaction kinetic constant was about 0.146/min. The analysis with liquid chromatography technique permits to identify, quantify and see the evolution of atrazine by-products which are generated by dechlorination, dealkylation and alkylic-oxidation mechanisms (Komtchou et al. 2016). The photodegradation study of atrazine was demonstrated using either Pt–TiO2 or TiO2 as a photocatalyst under 352 nm light irradiation. The Pt–TiO2-catalyzed atrazine degradation reached 76% in 3 h without adding H2O2 solution or aeration, which was more than 10% higher than the TiO2-catalyzed reaction. The decomposition product of Pt–TiO2-catalyzed atrazine degradation was mainly cyanuric acid (Chen et al. 2017). Shamsedini et al. (2017) noticed the maximum atrazine removal rate was at pH = 11 in the presence of Fe3+–TiO2 catalyst = 25 mg/L and the initial concentration of atrazine equal to 10 mg/L.

Microbial degradation of atrazine

Microorganisms are endowed with enormous and remarkable metabolic capabilities to utilize xenobiotics which are their carbon and energy source. A number of microbial species and strains have been shown to exhibit atrazine metabolism as listed in Table 4. Prokaryotic (Gram-positive and Gram-negative bacteria) and eukaryotic microbial species are involved in atrazine biodegradation both in situ and under in vitro conditions. Bacterial and fungal species usually dechlorinate the atrazine molecule leading to the formation of hyroxyatrazine, deisopropylatrazine and deethylatrazine. Xanthomonas sp. ARB2 (Sawangjit 2016); Enterobacter cloacae JS08 (Solomon et al. 2013a, b); Klebsiella sp. KB02 (Sopid 2012); Comamonas sp. A2 (Yang et al. 2010); Stenotrophomonas maltophilia, Rahnella aquatilis (Marecik et al. 2008); Chelatobacter heintzii, Aminobacter aminovorans, Stenotrophomonas maltophilia (Rousseaux et al. 2001); Pseudaminobacter C147 (Topp et al. 2000a); Bacillus subtilis HB-6 (Wang et al. 2014a, b); Arthrobacter sp. (Getenga et al. 2009; Liu et al. 2010; El Sebai et al. 2011; Zhang et al. 2011; Wang and Xie 2012; Wang et al. 2013); Bacillus licheniformis, B. megaterium (Marecik et al. 2008); Arthrobacter nicotinovorans HIM (Aislabie et al. 2005); Nocardioides sp. SP12 (Piutti et al. 2003); Arthrobacter aurescens TC1 (Strong et al. 2002); Arthrobacter crystallopoietes (Rousseaux et al. 2001); and Nocardioides sp. (Topp et al. 2000b). Some fungal species, viz. Umbelopsis isabellina, Volutella ciliate and Botrytis cinerea, were also found to be involved in degradation of atrazine (Marecik et al. 2008). Microbial species perform atrazine biodegradation by three major pathways out of which one is purely hydrolytic, while remaining two others are mixed (hydrolytic–oxidative) (Fig. 2). The first intermediate product hydroxyatrazine was first extensively converted by Pseudomonas spp. ADP consisting of three gene products atzA, atzB and atzC (Martinez et al. 2001). The dechlorination method (hydrolytic) which is catalyzed by enzyme atrazine chlorohydrolase (atzA or trzN gene product) shows dechlorination followed by elimination of N alkyl substituents to yield cyanuric acid (Solomon et al. 2013a, b). These three genes are widespread and almost found in all the atrazine-degrading strains worldwide (Rousseaux et al. 2001; Topp et al. 2000a). Usually in Gram-positive strains, atzA is replaced by trzN which belongs to hydrolase enzyme which removes several functional groups from the parent compound (Wang et al. 2005; Topp et al. 2000b). The second pathway engrosses the N-dealkylation of the atrazine into deethylatrazine or deisopropylatrazine which is further dealkylated into deisopropyldeethylatrazine or further undergoes hydrolytic to yield cyanuric acid. Rhodococcus strains N186 and 21 and SpTE1 show oxidative reactions by producing enzymes AtzA and TriA which actively deaminates the atrazine metabolites (Seffernick et al. 2001). Dechlorination of dealkylated atraizine is commonly shown by Rhodococcus corallinus NRRLB-containing hydrolase AtzB (Seffernick et al. 2002). Further, in the upper degradation pathway it gets converted into two different aminohydrolases which is encoded by atzB and atzC. Then, the final hydrolytic reaction which is encoded by trzF/atzF, trzD/atzD and trzE/atzE converts cyanuric acid into carbon dioxide (Udiković-Kolić et al. 2010; Wackett et al. 2002).

Table 4 Microorganisms involved in biodegradation of atrazine under in situ and experimental conditions
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Fig. 2
figure 2

Physical and biochemical pathways involved in the microbial biodegradation of atrazine. AC atrazine chlorohydrolase, AM atrazine monooxygenase, HDEH hydroxyl dechloroatrazine ethylaminohydrolase, NEC N-ethylammeline chlorohydrolase, DEM deethylatrazine monooxygenase, NIAIA N-isopropyl ammelide Isopropyl aminohydrolase, DIHA deisopropylhydroxyatrazine aminohydrolase, NEAC N-ethylammeline chlorohydrolase, 2, 4 D6 NEA 1, 3, 5 TEH = 2, 4-dihydroxy-6-(N′-ethyl) amino-1, 3, 5-triazine ethylaminohydrolase, HDAEH hydroxydechloro atrazine ethylaminohydrolase, NIIA N-isopropylammelide isopropylaminohydrolase

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Nitrogen released from atrazine metabolism serves as a nitrogen source for atrazine-degrading bacteria (Vaishampayan et al. 2007; Dutta and Singh 2013; Yang et al. 2010). Some bacteria initiate degradation of atrazine involving the enzyme atrazine chlorohydrolase through the mechanism of hydrolytic dechlorination. Aminohydrolases catalyze two hydrolytic deamination reactions that hydroxyatrazine undergoes; N-isopropylammelide (Getenga et al. 2009; Qingyan et al. 2008) or N-ethylammelide (Topp et al. 2000a) is formed as the intermediate metabolites. These ammelides are finally converted to cyanuric acid (Yang et al. 2010). Another route followed for atrazine degradation is N-dealkylation of the lateral ethyl and isopropyl chains to deethylatrazine, deisopropylatrazine and deethyldeisopropylatrazine (Zhang et al. 2011). These dealkylated atrazine metabolites undergo hydroxylation, and cyanuric acid is formed as the ultimate metabolite (Vaishampayan et al. 2007). Cyanuric acid, formed by either of the metabolic routes, is acted upon by cyanuric acid amidohydrolase, biuret amidohydrolase and allophanate hydrolase enzymes leading to the cleavage of the cyanuric acid to carbon dioxide and ammonia (El Sebai et al. 2011).

Conclusion

The demand for pesticides is on rise globally, especially in emerging economies of the world. India is the second most populated country of the world and is at the center of green revolution. It is, however, under ever-increasing demand of fulfilling the food requirements of huge population and thus relying heavily on synthetic herbicides as a weed control measure. As a consequence, environmental pollution, contamination of reservoirs, effect on food chains and life-threatening toxicities are certain to happen. Atrazine and its adverse effects are considered a highlighted threat to the environmental sustainability. Hence, an urgent need is felt to diverted resources and coordinated efforts to minimize its use, and at the same time, it is essential to monitor its impact on vertebrates, invertebrates and, most importantly, on microbial flora. Microbial biodegradation using in situ approach and use of transgenic strains having enhanced enzymatic activities and superior adaptability is considered a valid option for future studies. In future, use of biopesticides is expected to relieve our dependency on atrazine in order to minimize environmental pollution.