Photolysis of Chlorantraniliprole and Cyantraniliprole in Water and

Article pubs.acs.org/JAFC

Photolysis of Chlorantraniliprole and Cyantraniliprole in Water and Soil: Verification of Degradation Pathways via Kinetics Modeling Ashok K. Sharma,*,† William T. Zimmerman,† Suzanne K. Singles,‡ Kalumbu Malekani,§ Scott Swain,† David Ryan,† Gordon Mcquorcodale,∥ and Laura Wardrope∥ †

Stine Haskell Research Center, E.I. DuPont de Nemours and Company, Newark, Delaware 19714, United States DuPont Experimental Station, E.I. DuPont de Nemours and Company, Wilmington, Delaware 19803, United States ∥ Charles River Laboratories, Tranent, Edinburgh, Scotland EH332NE, United Kingdom ‡

ABSTRACT: Photodegradation of [14C]-chlorantraniliprole (CLAP) and [14C]-cyantraniliprole (CNAP) was investigated in sterile buffer solutions, in natural water, and on soil surfaces. Both compounds displayed rapid degradation in aqueous buffers when exposed to light at concentrations which could result from direct overspray to a shallow water body. While the main products observed had analogous structures, a substantial difference was noted in the rate of degradation of the two compounds despite minimal differences in their structures. Transformations observed were primarily intramolecular rearrangements and degradations resulting from addition of hydroxyl radicals leading to molecular cleavage. Some of the degradation products were transient, and several degradates had isomeric molecular compositions. The sequence of transformations was established definitively with the help of kinetics modeling. Utility of kinetics analysis in verification of the proposed pathways is illustrated. KEYWORDS: chlorantraniliprole, cyantraniliprole, photolysis, degradation, pathway kinetics, soil, buffers

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Technology Ltd., Bicester, UK). The irradiation device was fitted with a UV glass filter (Product 5605 2371, Atlas Material Testing Technology Ltd., Bicester, UK) to block light with wavelength < 290 nm and reduce the intensity of light at >800 nm. All aqueous photolysis experiments were conducted with continuous irradiation over the period specified. The individual samples for irradiation were prepared in separate enclosed quartz vessels equipped with an air inlet and outlet for collection of volatiles. Each sample was maintained beneath the irradiation source in a purpose-built flooding table. Water from a temperature-regulated water bath was continuously circulated through the flooding table throughout the irradiation period to maintain the temperature at 20 ± 2 °C. Individual nonirradiated samples were prepared in separate amber glass jars. These vessels were also placed in a temperature-regulated water bath to maintain the temperature. CNAP aqueous photolysis was carried out in 0.01 M pH 4 buffer, which was prepared by addition of 360 mL of 0.01 M sodium acetate solution to 1640 mL of 0.01 M acetic acid solution in a 2 L volumetric flask. Final pH adjustments were made with 5 M NaOH to pH 4. The buffer solution was filter sterilized by passing through a 0.2 μm filter, and the pH of the buffer solutions was verified to be 3.97. The test substance was added as a water−acetonitrile solution to achieve a concentration of 1 ± 0.02 μg/mL. CLAP photolysis was carried out in 0.01 M pH 7 sterile buffer prepared with maleic acid and at a test substance concentration of 0.6 ± 0.02 μg/mL due to lower water solubility for this compound. Natural water used for photolysis experiments was collected from the Crosswood Reservoir (UK Ordinance Survey map reference: Map 65, Grid Reference NT058570). A subsample of the water was characterized for measurements of pH (7.4), total hardness as CaCO3 (74 mg/L), dissolved organic carbon (2.9 mg/L), total organic carbon (10.2 mg/L), total nitrogen (1.2 mg/L), nitrate (85% conversion to CNAP_M2 as measured by HPLC-UV. The solution containing CNAP_M2 was acidified to pH 3 with acetic acid to prevent its degradation. The acidified solution was applied to a 2 g, 12 cc C18 Mega Bond Elut SPE cartridge (Varian Inc., Lake Forest, CA) and washed with 2 mL of 0.1% formic acid. The column was flushed dry with hexane and vacuum dried. The product was eluted with 10 mL of ethyl acetate with 0.1% formic acid. LC/MS analysis of the sample, in ESI positive mode, gave a molecular weight of 437 and molecular formula C19H13BrN6O2, and the isotope cluster was consistent with a molecule containing one bromine. LC-IR analysis of the sample showed absorption bands for CN at 2228, benzamide CO at 1695, and CN at 1651 cm−1, which were characteristic of functional groups in the structure. Additional heterocycle and benzene double bonds were also present at 1591, 1518, and 1453 and C−O at 1195 cm−1. A portion of the sample was loaded on to SPE cartridge, and the degradate was recovered in acetonitrile-d3 to acquire 1H NMR data. Identification of CNAP_M3. A 300 mL aliquot of the solution containing CNAP_M2 was transferred to a 1 L Erlenmeyer flask, an 6578

dx.doi.org/10.1021/jf5017175 | J. Agric. Food Chem. 2014, 62, 6577−6584

Journal of Agricultural and Food Chemistry

Article

LC/1H NMR-MS. The sample generated contained approximately 93% of M4 by HPLC analysis. LC/MS analysis of the sample, in ESI positive mode, gave a molecular weight consistent with that of CNAP_M4 (molecular weight 344, with acetate adduct at mass 387). The isotope cluster pattern is consistent with a molecule containing one bromine atom. LC-IR of the sample gave an IR spectrum which showed cyano functionality at 2227 cm−1, cyclic amide CO at 1678, and ring double bonds at 1581, 1471, and 1476 cm−1.

equal volume of 0.01 M pH 9 borate buffer was added, and the mixture was stored under ambient conditions of temperature and light for 2 days, until there was approximately 89% conversion to CNAP_M3 as measured by HPLC-UV analysis. The solution containing M3 was concentrated using a 2 g, 12 cc C18 Mega Bond Elut SPE cartridge using pH 5 acetate buffer for preconditioning instead of 0.01% formic acid. The photoproduct was eluted with acetonitrile and reduced to dryness by nitrogen evaporation. The dry solid containing 87% M3 was analyzed by LC-MS/MS, LC-IR, and LC/1H NMR-MS. The solid was dissolved in deuterated acetonitrile prior to LC/1H NMR analysis. LC/MS analysis of the sample in ESI positive mode gave a molecular weight consistent with structure of M3 (molecular weight 437) and isotope cluster pattern consistent with a molecule containing one bromine. The IR spectrum showed a cyano stretch at 2229 cm−1, cyclic amide CO at 1683, C−O stretch at 1250, and ring double bonds at 1461, 1592, and 1528 cm−1. 1H NMR further verified the assigned structure.

Identification of CNAP_M5. The solution containing CNAP_M2 was acidified to a pH of approximately 2−3 with formic acid and the container placed in a water bath at 70 °C for 2 days. The container was then stored under ambient conditions of temperature and light for 5 days, until there was approximately 30% conversion to M5 as measured by HPLC-UV analysis. The solution containing M5 was concentrated using a 2 g, 12 cc C18 Mega Bond Elut SPE cartridge preconditioned with one column volume of methanol followed by one column volume of water containing 0.1% formic acid. The solution was applied to the column and washed with 15−20 mL of 0.1% formic acid. The photoproduct was eluted with a 40/60 acetonitrile/water− 0.1% formic acid. The eluate was reduced to a small volume by nitrogen evaporation and quick frozen using the shelling technique in a dry ice−acetone bath. Remaining solvents were removed on a lyophilizer overnight. The lyophilized sample was stored in a freezer. Final purification was conducted by HPLC prior to recording the spectra. An aliquot of the sample was analyzed by LC-MS/MS, LC-IR, and LC/1H NMR. LC/MS analysis of the sample, in ESI positive mode, gave a molecular ion at 455/457 (1:1) consistent with the structure of M5. Prominent fragments at 424/426 and 266/268 (both showing peak cluster ratios of 1:1 indicative of Br atom in structure) resulted from acylium ion formed from loss of CH3NH and the pyrazole acylium ion moiety after cleavage of the benzamide side. The IR spectrum showed CN at 2232, two amide CO peaks at 1670 and 1639 cm−1, and C− O stretch at 1266 cm−1.

Identification of CNAP_M4. The 600 mL solution containing CNAP_M3 was transferred to a water-jacketed beaker covered with a quartz lid and placed in a Heraeus Suntest unit in a water bath set at 20 °C. The sample was irradiated for 2 days until there was approximately 88% conversion to M4 as measured by HPLC-UV. The solution containing CNAP_M4 was concentrated using a 2 g, 12 cc C18 Mega Bond Elut SPE cartridge. The cartridge was preconditioned with one column volume of acetonitrile followed by one column volume of 1:1 pH 5 acetate buffer/pH 9 borate buffers. The solution was applied to the column and washed with 5 mL of H2O. The column was then flushed dry using house vacuum. The photoproduct was eluted with acetonitrile. The eluate was reduced to dryness by nitrogen evaporation. The dry solid was analyzed by LC-MS/MS, LC-IR, and 6579



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Identification of CNAP_M6. LC/MS analysis of the sample, in ESI positive mode, gave a molecular weight consistent with that of CNAP_M6 (molecular weight 436). The exact mass suggested a molecular formula of C19H13BrN6O2, and the isotope cluster pattern was consistent with a molecule containing one bromine atom. The molecule was clearly an isomer of CNAP_M2 and M3 but quite different in HPLC retention time, and it was stable to base. Stability to base ruled out the possibility of E/Z isomers of M2. Structure assignment was based on the MS fragmentation; NMR of an isolated sample was obtained by aqueous photolysis of CNAP_M1 (ca. 20% yield). NMR spectra for this sample verified the structure.

Determination of Rate Constants. Optimization of kinetics and rate constant calculations was conducted by building a first-order kinetics model in ModelMaker (software purchased from apbenson.com). The models represented by the sequence of transformations are discussed in the Results and Discussion section. In each case alternative models with a different order of transformations, which appeared to be logical based on chemistry, were also tested. The models presented gave the best optimization fit for the observed data. The rate constants generated for optimized kinetics of all experiments are summarized in Table 1, and the data fit is illustrated in various figures.

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RESULTS AND DISCUSSION CLAP and CNAP have a water solubility of 1.2 and 14.6 μg/ mL, respectively. It has been shown in our previous work11 that both compounds are vulnerable to hydrolytic degradation at alkaline pH, therefore, photodegradations were conducted in buffers of either pH 4 (CNAP) or 7 (CLAP) because the contribution from hydrolysis reactions was expected to be minimal and photodegradation reactions were anticipated to dominate. A generalized photodegradation pathway applicable to both compounds is displayed in Figure 3. All products displayed in the figure were not necessarily observed for both compounds under various conditions, and the degradations observed were consistent with those reported earlier.4,6 The similarities and differences between the experiments for both test compounds are discussed. Photodegradation of Chlorantraniliprole. Photolysis of CLAP in pH 7 buffer generated three products which appeared to have been formed sequentially. Degradation initially led to formation of M2, which reached a peak quickly by day 1. As M2 declined, the concentration of M3 increased, which also started to decline during the study period, leading to progressively Table 1. Rate Constants and DT50 Values for CNAP and CLAPa chlorantraniliprole media buffer

nat-water

soil

cyantraniliprole

K (day )

DT50 (day )

K (day )

DT50 (day−1)

P to M1(c) P to M2 M2 to M3 M3 to M4 M4 to sink M2 to M5

NA 1.696 ± 0.069 0.659 ± 0.034 0.313 ± 0.016 0.004 ± 0.002 NA

0.41 1.00 2.25 165.0

0.003 ± 0.001 4.01 ± 0.094 0.011 ± 0.002 0.234 ± 0.051 0.357 ± 0.125 0.007 ± 0.002

231.0 0.17 64.8 3.0 1.9

P to M1(c) P to M3 P to M5 M3 to M4 M4 to sink M1 to sink M5 to sink

0.003 ± 2.311 ± NA 1.166 ± 0.010 ± NA NA

0.379 2.946 0.090 1.003 0.030 0.33 0.001

± ± ± ± ± ± ±

0.010 0.136 0.025 0.038 0.002 E E

1.8 0.24 7.7 0.7 23.3 2.1 693.1

0.043 0.024 0.064 0.034 0.027 0.457 0.091 0.065

± ± ± ± ± ± ± ±

0.006 0.012 0.006 0.007 0.007 0.249 0.036 0.046

16.0 28.5 7.1b

reaction

−1

−1

0.001 0.096

218.0 0.31

0.055 0.003

0.59 66.65

P to M1(c) M1 to sink(c) P to M1 P to M3 M1 to M6 M3 to M4 M4 to sink M6 to sink

−1

b

25.8 1.5 7.7 10.7

a

P = parent compound (CLAP or CNAP), NA = not observed, E = rate constant not optimized, (c) = dark control. bDT50 of parent calculated using sum of rate constants for P to M1 and P to M3 6580



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Figure 3. Transformations of chlorantraniliprole and cyanantraniliprole under photolysis conditions.

higher amounts of M4 (Figure 4). A similar incubation in the absence of light showed no degradation of CLAP.

Figure 4. Chlorantraniliprole photolysis, pH 7 buffer.

The observed data for degradation in buffer were fitted to a kinetics model, illustrated below, in which each successive transformation product is generated as a result of first-order reaction from the preceding component and the last degradate leading to minor unknown products. Optimized kinetics parameters resulting from such a degradation scheme showed a good fit for all significant components (Figure 4), which verified that the products were indeed generated in succession. Even though degradates M2 and M3 had identical molecular weights and mechanistically it was possible to rationalize transformations leading to formation of both compounds directly from the parent, kinetics modeling verified that these two products were indeed formed in succession. Lack of stability of M2 and its hydrolytic transformation to M3 further supported the sequential transformations. Optimized rate constants for each step are listed in Table 1. Photolysis of CLAP in natural water (pH 7.4) also produced the same degradates as the buffer, except degradate M2, which

was not observed. Instead, degradation proceeded directly to M3, which again declined rapidly, leading to increasing amounts of M4. There was a gradual decline in the amount of M4 as the photolysis period progressed. A parallel incubation in the absence of light showed slow degradation of CLAP and formation of a small amount of M1. However, conversion of CLAP to M1 was too slow to make a meaningful contribution to the degradation observed in the presence of light. Therefore, inclusion of degradation to M1 in natural water as a parallel hydrolytic degradation did not alter the kinetics optimization of the photodegradation process. For model optimization in natural water, degradate M2 was ignored, since it was not observed. Instead, direct transformation of CLAP to M3 followed by M4 was optimized to derive the rate constants. The optimized model provided a good fit for the observed data. Figure 5 illustrates the result of inclusion of a simultaneous 6581



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fold faster in natural water as compared to buffer. Even degradation of M4 to minor unknown products (sink) was faster by a factor of 2.5, indicating that degradation of CLAP as well as all photodegradates accelerated in natural water. If pH alone was the primary factor, enhanced degradation of M3 and M4 would not be expected. Degradation of CLAP in soil was not significant over the 30 day duration of the experiment. Photodegradation of Cyantraniliprole. Degradation of CNAP showed many similarities with CLAP photodegradations, but there were substantial differences. A lower pH buffer was chosen for CNAP degradation because only at pH 4 was photodegradation with the least contribution from hydrolytic degradations. Control solutions of CNAP in pH 4 buffer kept in the dark showed minimal conversion to M1. However, unlike CLAP, substantial degradation to cyclodehydration product, CNAP_M1, was noted in natural water dark control (Figure 6).

Figure 5. Chlorantraniliprole photolysis, nat-water.

degradation to M1 using a CLAP to M1 rate constant of 0.0032 day−1 (calculated from dark control experiment). A good data fitting (Figure 5) verified that partial hydrolysis due to increased pH played no significant role in the photodegradation of CLAP. Nevertheless, constituents of natural water did alter the overall photodegradation in two ways. First, the degradation rate increased, and second, a key degradate M2 was no longer detected. Even though M2 was not observed in natural water photodegradation, it was presumed that M2 was a transient intermediate. Apparently, M2 underwent hydrolytic degradation in natural water too rapidly (possibly due to higher pH) to have been detected. Degradation of M2 to M3 under alkaline conditions was noted while attempting to isolate samples of M2, and indeed, it showed rapid conversion to M3 under slightly alkaline conditions. Some hydrolytic conversion of M2 to M3 even in pH 7 buffer cannot be ruled out, since stable isolation of M2 required acidic pH of 5 or less. However, the increase in photolysis rate in natural water probably was not merely due to the higher pH of natural water, because all subsequent transformations were also expedited. The transient behavior of M2 became even more obvious when the analogous product CNAP was investigated, as discussed later. It appeared more likely that faster photochemical reactions were due to dissolved ions and/or organic matter, a phenomenon noted by others as well.7,8 Conversion of parent compound (P) to M3, which was controlled by P to M2 rate constant in buffer, increased by a factor of about 1.5 in natural water (2.3 vs 1.7 day−1, data in Table 1). A faster degradation rate of CLAP was observed in natural water not only for the parent compound but each subsequent transformation M3 to M4 as well as M4 to unknown products. The presence of carbonate, nitrates, sulfates, and dissolved organic carbon has been reported to enhance generation of OH radicals or hydroperoxide radicals.9 Other investigators have also reported quenching of radicals by the same species;10 therefore, it cannot be stated with certainty as to which constituents of natural water facilitated the phototransformation rates. While the transformation mechanism for M2 to M3 (displayed in figure below) is believed to occur with OH anion (pH effect), the same reaction could just as well be triggered by OH radical (photodegradation). In reality, both mechanisms may have been operative to expedite this transformation so much that its degradation in natural water was faster than its formation; hence, M2 was not observed. A comparison of the rate constants for transformation of M3 to M4 (data in Table 1) shows that this reaction was nearly 4-

Figure 6. Cyantraniliprole, dark controls.

This clearly was due to the higher pH (7.4) of natural water. A faster hydrolytic degradation of CNAP in alkaline water, especially when compared with CLAP, has been discussed in detail previously;11 therefore, hydrolytic degradation was anticipated to compete with photodegradation for CNAP in alkaline natural water. Since cyantraniliprole is hydrolytically stable in pH 4 buffer, photodegradation was expected to occur with little contribution from hydrolytic reactions. As noted in Figure 7, degradation was faster than degradation of CLAP, resulting in formation of one main product M2. This product M2 degraded slowly, but it did degrade to three other products. Two of these, M3 and M4, were analogous to those found in CLAP degradation, while M5 was new to CNAP photolysis. Optimization of data using kinetics modeling suggested that M5 was derived from M2 and not from CNAP. It was surprising that CNAP_M2 was slower to degrade photochemically, as compared to CLAP_M2. In part, slower degradation of M2 may have been due to lower pH 6582



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Figure 7. Cyantraniliprole photolysis in pH 4 buffer.

Figure 8. Cyantraniliprole photolysis, nat-water.

of the buffer used, which provided stability toward hydrolytic degradation of M2 to M3. Additional stability toward photodegradation of M2 may also have been caused by the “CN” group due to the extended conjugation of the phenyl and pyrazole rings. Such stabilization possibly made CNAP_M2 somewhat reluctant toward further degradation at pH 4 and allowed it the opportunity to generate an additional degradate M5 in small amounts. Conversion of CNAP_M2 to M3 and M5 is probably due to hydrolytic reactions, even though a photolytic reaction mechanism due to OH radicals can be envisioned equally well. During degradate purification work it was noted that both M2 and M5 were easily converted to M3 in the presence of base or even at pH 7. Therefore, it was not surprising that photodegradation in natural water showed no CNAP_M2 and only a small amount, seldom exceeding 3−4%, for CNAP_M5. Initial formation of M2 undoubtedly did occur in natural water, but its further degradation to M3 was too rapid to allow detection of M2, just as it was noted for CLAP. A small portion did, however, manage to transform to M5 even in natural water. Similar to CLAP, further degradations, such as M3 to M4 and M4 to other unknown products, were observed in buffer as well as natural water, except that these transformations were again accelerated in natural water even for CNAP. In the case of CNAP it was anticipated that formation of M1 via hydrolysis should compete with photolysis in natural water. Since CNAP_M1 was not actually observed during photolysis, we decided to explore this issue via kinetic modeling. While all observed degradates in natural water were optimized for their formation/degradation rates, the rate constant for hydrolysis in natural water was added to the kinetic model (figure below) using a rate constant computed from the dark control experiment. Such modeling for CNAP suggested that nearly 5−10% of total residue should have been observed as M1 in early stages of the study (Figure 8). After all, the presence of light would not be expected to shut down the hydrolytic degradation. Given that this hydrolytic pathway to CNAP_M1 occurs without light, not finding it means that M1 must be photodegrading just as it is formed via hydrolysis. This indeed was found to be the case when a new photodegradate CNAP_M6 was detected and identified. M6 was only observed at a few sampling intervals and accounted for ∼3−4% of the total mixture. Formation of M6 from M1 was verified by direct photolysis of CNAP_M1 in a separate experiment, albeit yielding a maximum of ∼20% only. The small amount of M6 being observed was attributed to its instability under photo-

degradation conditions. Due to the small amount observed, optimization of its formation rates via modeling was not feasible. Hence, it was not included in kinetics and treated as part of a “sink”, a normal practice in kinetics modeling to deal with all minor and unknown products.

Degradation of cyantraniliprole in air-dried soil was slow, and the degradation rates in the presence as well as in the absence of light showed no meaningful differences. However, its degradation in moist soil, which was maintained at approximately 50% of maximum water-holding capacity, did display significant degradation. CNAP degradation in soil kept in the dark led primarily to formation of CNAP_M1 (Figure 9) along with numerous minor products. Photolysis in moist soil 6583



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One can conclude from this investigation that both CLAP and CNAP are degraded by light essentially along the same pathway in water, leading primarily to formation of degradates M3 and M4. M2 may be encountered in water under acidic pH, but it is unlikely to be observed in significant quantity in natural water or soil. The degradation reactions observed were consistent with those reported previously, except for minor additional products observed for CNAP. Kinetics modeling of the data not only served to compute the degradation rates but also undoubtedly helped clarify the degradation pathway as well.

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AUTHOR INFORMATION

Corresponding Author

Figure 9. Cyantraniliprole soil photolysis, dark control.

*E-mail: [email protected].

enhanced the overall degradation rate of CNAP, and formation of most aqueous photolysis products was noted. Degradation occurred via two distinct pathways. One pathway proceeded via initial formation of CNAP_M1, the hydrolysis product, and its subsequent degradation to M6, while the other followed the natural water photolysis sequence. A kinetics assessment using a model similar to that employed for natural water degradation showed a good fit for all observed components, and the kinetics fits are displayed in Figure 10. Photodegradation in a moist soil

Present Address §

(K.M.) Smithers Viscient, 790 Main Street, Wareham, MA 02571, USA. Notes

The authors declare no competing financial interest.

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ACKNOWLEDGMENTS Contributions of Steve Cheatham and Boiana Budevska for spectral data are sincerely acknowledged. REFERENCES

(1) Cordova, D.; Benner, E. A.; Sacher, M. D.; Rauh, J. J.; Sopa, J. S.; Lahm, G. P.; Selby, T. P.; Stevenson, T. M.; Flexner, L.; Gutteridge, S.; Rhoades, D. F.; Wu, L.; Smith, R. M.; Tao, Y. Pestic. Biochem. Physiol. 2006, 84, 196−214. (2) Selby, T. P.; Lahm, G. P.; Stevenson, T. M.; Hughes, K. A.; Cordova, D.; Annan, B.; Barry, J. D.; Benner, E. A.; Curries, M. J.; Pahutski, T. F. Bioorg. Med. Chem. Lett. 2013, 23, 6341−6345. (3) Caboni, P.; Sarais, G.; Angioni, A.; Vargiu, S.; Pagnozzi, D.; Cabras, P.; Casida, J. E. J. Agric. Food Chem. 2008, 56, 7679−7699. (4) Zhang, J. M.; Hai, W. G.; Wu, Y. L. Chemosphere 2012, 87, 132− 136. (5) Malhat, F.; Abdallah, H.; Hegazy, I. Bull. Environ. Contam. Toxicol. 2012, 88, 349−351. (6) Lavtizar, V.; van Gestel, C. A. M.; Dolenc, D.; Trebse, P. Chemosphere 2014, 95, 408−414. (7) Wallace, D. F.; Hand, L. H.; Oliver, R. G. Environ. Toxicol. Chem. 2010, 29, 575−581. (8) Wols, B. A.; Hofman-Caris, C. H. M. Water Res. 2012, 46, 2815− 2827. (9) Dong, M. M.; Rosaio-Ortiz, F. L. Environ. Sci. Technol. 2012, 46, 3788−3794. (10) Wenger, J. C. Environ. Sci. Technol. 2004, 38, 831−837. (11) Sharma, A. K.; Zimmerman, W. T.; Lowrie, C.; Chapleo, S. J. Agric. Food Chem. 2014, 62, 3531−3536.

Figure 10. (a) Cyantraniliprole soil photolysis. (b) Cyantraniliprole soil photolysis.

surface was clearly faster than degradation in moist soil maintained in the dark due to participation of photolysis reactions. However, the rate of photodegradation even in moist soil was much slower than photodegradation in water itself. This experiment implies that photodegradation of CNAP on the soil surface may not be a significant degradation pathway in the environment because moist soils tend to dry out quickly in sunlight. 6584


Photolysis of Chlorantraniliprole and Cyantraniliprole in Water and

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