Enhanced Photodegradation of Extracellular Antibiotic Resistance

Article pubs.acs.org/est

Cite This: Environ. Sci. Technol. XXXX, XXX, XXX−XXX

Enhanced Photodegradation of Extracellular Antibiotic Resistance Genes by Dissolved Organic Matter Photosensitization Xin Zhang, Jing Li, Wen-Yuan Fan, Mu-Cen Yao, Li Yuan, and Guo-Ping Sheng* CAS Key Laboratory of Urban Pollutant Conversion, Department of Applied Chemistry, University of Science and Technology of China, Hefei 230026, China

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S Supporting Information *

ABSTRACT: Extracellular antibiotic resistance genes (eARGs) contribute to antibiotic resistance, and as such, they pose a serious threat to human health. eARGs, regarded as an emerging contaminant, have been widely detected in various bodies of water. Degradation greatly weakens their distribution potential and environmental risks. Dissolved organic matter (DOM), mainly consisted of humic substances, carbohydrates, and organic acids, is ubiquitous in diverse waters and significantly affects the degradation of coexisting contaminants. However, the photodegradation of eARGs in natural water, especially regarding the roles of DOM in this process, remains unknown. Herein, we investigated the eARGs photodegradation in waters with and without DOM. Illumination has been found to effectively photodegrade eARGs, and this process was significantly enhanced by DOM. Further experiments revealed that photosensitization of DOM produced hydroxyl radicals (•OH) to enhance plasmid strand breaks and produced singlet oxygen (1O2) to accelerate the guanine oxidation, which in turn promoted the photodegradation of plasmid-carried eARGs. Transformation assays indicated that eARGs transformation efficiencies were reduced after their photodegradation. The presence of DOM accelerated the decreases of eARGs transformation efficiencies under illumination. DOM concentration and some ions (e.g., NO3−, NO2−, HCO3−, Br−, and Fe3+) affected •OH or 1O2 levels, further influencing the photodegradation of eARGs. Overall, eARGs photodegradation in aquatic environments is a crucial process both in the reduction of eARGs concentrations and in transformation efficiencies. This work facilitated us to better understand the fate of eARGs in waters.

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INTRODUCTION Due to the overuse and misuse of antibiotics, antibioticresistant bacteria have become one of the biggest challenges to human health.1 Bacteria can acquire antibiotic-resistance genes (ARGs) from parent or other microorganisms and exhibit resistance to one or multiple antibiotics.2 Diverse ARGs have been detected in wastewater, sludge, and animal waste in high concentrations.3,4 Furthermore, ARGs associated with mobile genetic elements including plasmid, integron, transposons, and bacteriophage can spread between microorganisms through horizontal gene transfer (HGT),5 which takes place through conjugation, transduction, and transformation.6 Intracellular ARGs (iARGs) can transfer by conjugation within microorganisms or transduction mediated by bacteriophage, while extracellular ARGs (eARGs) propagate via transformation.7 These two forms of ARGs have distinct fates in the environment. Previous studies show that natural transformation can occur extensively with high frequency in diverse environments. The efficiencies of plasmid or chromosomal DNA transformation in freshwater, seawater, soil, and sediment were reported to be up to 2.0 × 10−4, 1.7 × 10−6, 6.2 × 10−5, and 5.7 × 10−6, respectively.8−11 Currently, eARGs © XXXX American Chemical Society

have been discovered in various water bodies such as rivers, estuaries, and seawaters.12 Thus, eARGs are considered an emerging contaminant in aquatic environments due to its prevalence and proliferation in water bodies.12 DNA can undergo damage under UV light illumination.13 The UVC (e.g., UV254) can effectively cause DNA damage, which is accepted to be the mechanism of disinfection by UV254.14 Apart from UVC, UVB (280−320 nm) light alone is also capable of causing the DNA damage.13 UVA (320−400 nm) alone is less effective but can cause DNA damage by indirect photosensitization reaction.13 In addition to UV lightinduced DNA damage, various reactive oxygen species (ROS) can also induce DNA damage.15,16 Nevertheless, among the various ROS such as hydroxyl radicals (•OH), singlet oxygen (1O2), superoxide radical (O2·−), and hydrogen peroxide (H2O2), •OH and 1O2 are the only ones that can cause DNA damage.15 Recently, the eARGs encoded on plasmid DNA Received: May 23, 2019 Revised: August 12, 2019 Accepted: August 19, 2019

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DOI: 10.1021/acs.est.9b03096 Environ. Sci. Technol. XXXX, XXX, XXX−XXX

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Environmental Science & Technology

plasmids were used to meet the detection requirements in experiments focusing on plasmid bases damage and strand breaks. At selected intervals, 100 μL of solution was taken from each tube for subsequent tests. To investigate the role of •OH, 0.1% (v/v) isopropyl alcohol (IPA) or 5 mM sodium formate (HCOONa) was added in the DOM solution to quench •OH.21,22 Besides, 5 mM NaN3 was added in the DOM solution to scavenge both •OH and 1O2.21 Considering the longer lifetime of 1O2 in deuteroxide (D2O),23 the photodegradation experiments were also conducted in DOM−D2O solution to validate the role of 1O2. To clarify the role of 3DOM*, the photodegradation of eARGs was further conducted with addition of IPA under N2-saturated condition to exclude both •OH and 1O2.24 Different amounts of chloride, nitrate, nitrite, bromide, and bicarbonate were separately added into DOM solution to investigate the effects of anionic ions. Finally, to investigate the effect of Fe(III) on the photodegradation of eARGs, Fe(III) was spiked into DOM MOPS (3-morpholinopropanesulfonic acid) buffer (10 mM, pH = 7.3) solution to avoid combining of Fe(III) by phosphate.25 Plasmid Purification and qPCR Analysis. The plasmids were purified with a SanPrep Column PCR Product Purification Kit (Sangon Biotech, Co., China) to remove DOM and salts. Plasmids concentration and purity were determined by a microplate reader (SpectraMax M2e, Molecular Devices Co., USA). The eARGs were quantified using both short- and long-amplicon qPCR. The gene sizes of tet A and blaTEM‑1 genes were 1191 and 861 bps, respectively. The short-amplicon qPCR only covered ∼200 bps of genes, and the long-amplicon qPCR was designed to cover the whole genes. The detailed information for short amplicons and long amplicons of tet A and blaTEM‑1 as well as the corresponding primers are shown in Table S1. The short-amplicon qPCR possessed higher amplification efficiency and could reduce the appearance of nonspecific amplification compared to longamplicon qPCR.26 However, the short-amplicon qPCR did not cover whole resistance genes and would underestimate the photodegradation rates of eARGs. Therefore, the shortamplicon qPCR was applied to more accurately determine the relative photodegradation rates of eARGs under different conditions, and the long-amplicon qPCR was used to measure the absolute photodegradation rate constants of eARGs. The qPCR was performed on a LightCycler 96 instrument (Roche, Basel, Switzerland) with each 20 μL of qPCR reaction solution containing 10 μL of SYBR Green II (Takara), 8.2 μL of DNase free water, 0.4 μL of forward or reverse primer (Table S1), and 1 μL DNA template. The protocol of temperature cycling of qPCR was as follows: 95 ◦C for 30 s, followed by 40 cycles: denaturation at 95 ◦C for 5 s, annealing at 54 ◦C (short amplicons) or 52 ◦C (long amplicons) for 30 s, and extension at 72 ◦C for either 30 s (short amplicons) or 60 s (long amplicons) followed by a melting curve for specificity verification. Each sample was analyzed in duplicate, and the standard curve covered 5 orders of magnitude from 102 to 107 copies/μL. The amplification efficiencies ranged from 90% to 110% for short amplicons and from 80% to 120% for long amplicons. The R2 values were above 0.99. eARGs Transformation Experiments. One nanogram of purified plasmid was added into 100 μL of E. coli DH5α competent cells and placed in ice for 30 min, followed by heat shock at 42 ◦C for 45 s. After heat shock, the sample was put back in ice quickly for about 2 min. Then 700 μL of sterile LB medium was added into the bacteria solution and cultured at

have been reported to be effectively degraded under UV254 illumination.7,17 Furthermore, •OH produced from UV/H2O2 can significantly enhance the degradation of eARGs.17 In brief, both UV light and ROS (e.g., •OH and 1O2) are capable of effectively causing DNA or eARGs degradation. In aquatic environments, photochemical degradation is crucial for the attenuation of contaminants. Contaminants in sunlit waters can undergo direct photolysis prevailingly caused by solar UV and indirect photodegradation induced by photochemically produced reactive intermediates.18 The photoproduced reactive intermediates in sunlit waters originate for the most part from ubiquitous dissolved organic matter (DOM).19 The chromophoric components in DOM can be photoexcited to form various reactive intermediates, such as the triplet DOM (3DOM*), •OH, and 1O2,20 which participate in the photodegradation of contaminants. Considering the capabilities of UV and reactive intermediates (e.g., •OH and 1 O2) in the degradation of eARGs, eARGs in sunlit waters are likely to undergo both direct photolysis and reactive intermediates-induced indirect photodagradation. However, little is known about eARGs photodegradation in natural waters, especially concerning the role of reactive intermediates produced from ubiquitous DOM as well as the impacts of eARGs photodegradation on their transformation. In the present study, we aimed to investigate the photodegradation of eARGs in aquatic environments, with special attention to the role of DOM in this process. eARGs transformation efficiencies were examined through transformation experiments to reveal the influences of eARGs photodegradation on their transformation. Moreover, the reactive intermediates and plasmid damage involved in eARGs photodegradation were investigated to clarify the mechanisms of eARGs photodegradation in DOM solution. To better understand the eARGs photodegradation in natural waters, the effects of DOM concentration and common coexisting ions were also evaluated.

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MATERIALS AND METHODS DOM, Plasmid, Enzymes, and Competent Cells. Suwannee River Dissolved Organic Matter (2R101N) purchased from International Humic Substances Society (IHSS) was used as model DOM. The pBR322 plasmid containing tet A- and blaTEM‑1-resistance genes (Sangon Biotech Co., China) was used as model eARGs. Deoxyribonuclease I (DNase I), alkaline phosphatase (ALPase), and E. coli DH5α competent cells were obtained from Sangon Biotech Co., China. Phosphodiesterase I was purchased from NEB Co., USA. Photochemical Experiments. All photochemical experiments were conducted in a merry-go-round photochemical reactor (XPA-7, Nanjing Xujiang Electromechanical Inc., China). A 500 W medium mercury lamp equipped with a filter emitting from 290 to 400 nm was used as the light source. The light intensity (290−400 nm) measured with a UV light photometer (UV-340A, Lutron, China) at the center of quartz tubes was 40 W/m2. The quartz tubes covered with two layers of aluminum foil were used as the dark controls. During all illumination experiments circulating water was used to maintain a reaction temperature of 25 ± 1 °C. The plasmids carried eARGs were irradiated in phosphate buffer (PBS) (20 mM, pH = 7.3) dosing DOM with different concentrations (0−50 mg/L). In the eARGs photodegradation experiments plasmid concentrations were 0.1 mg/L, while 10 mg/L B


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Figure 1. Changes in logarithmic concentrations of (a, c) tet A and (b, d) blaTEM‑1 genes under light illumination or dark controls with or without DOM measured by (a, b) short-amplicon qPCR or (c, d) long-amplicon qPCR. Detailed sequence information for short and long amplicons as well as the primers was found in Table S1. Concentrations of DOM and plasmid are 20 and 0.1 mg/L, respectively. Symbols and lines are the experimental data and corresponding linear regression, respectively. Error bars represent 95% confidence intervals for n = 4.

37 ◦C for 1 h. After serially diluting the cell suspension, 80 μL was placed on selective LB plates with (10 mg/L tetracycline or 100 mg/L ampicillin) or without antibiotics. The plates were cultured at 30 ◦C for 24 h. Transformation efficiencies were calculated using the following equation7 transformation efficiency =

Quantity One, Bio-Rad, and used to represent the contents of bands.

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RESULTS Photodegradation of eARGs. The changes of logarithmic concentrations of eARGs with increasing illumination time (Figure 1) show that both tet A and blaTEM‑1 genes were photodegradable. The photodegradation of eARGs followed pseudo-first-order kinetics with the assumption that the concentrations of reactive intermediates (e.g., ROS and 3 DOM*) kept constant during the steady state illumination in DOM solution according to previous studies.19,23,29 Herein, both short- and long-amplicon qPCR were used to determine the photodegradation rate constants of eARGs. In phosphate buffer, the photodegradation rate constants measured by shortamplicon qPCR were 0.13 h−1 for tet A and 0.18 h−1 for blaTEM‑1 (Table S2). The corresponding photodegradation rate constants for tet A and blaTEM‑1 measured by long-amplicon qPCR were much higher and were 0.37 and 0.41 h−1, respectively (Figure 1 and Table S2). In DOM solution, enhanced photodegradation rates of eARGs were observed (p < 0.05) (Figure 1). The photodegradation rate constants of tet A and blaTEM‑1 determined by short-amplicon qPCR increased to 0.23 and 0.30 h−1, respectively (Table S2). The longamplicon qPCR-determined photodegradation rate constants for tet A and blaTEM‑1 were elevated to 0.72 and 0.78 h−1 (Table S2). No obvious eARGs degradation was observed under dark controls in both PBS buffer and DOM solution (Figure 1), suggesting that the enhanced degradation of eARGs by DOM was due to the photoexcitation of DOM. Decreases of eARGs Transformation Efficiencies. Transformation experiments were carried out to evaluate the effects of eARGs photodegradation on HGT capabilities. The initial transformation efficiencies of tet A and blaTEM‑1 without

transformant CFU total CFU

where the transformant CFU (colony-forming unit) and total CFU represented the colony forming on LB plates with and without antibiotics, respectively. Plasmid Damage Measurements. Purified plasmids (2 μg) were hydrolyzed using 1.0 U of DNase I, 0.1 U phosphodiesterase I, and 1.0 U ALPase in Tris-HCl buffer (pH = 7.6) at 37 °C for 24 h.27 During enzymolysis of plasmid, 1 mM deferoxamine mesylate was added into the solution to avoid the interference of deoxyguanosine oxidation.28 Subsequently, the plasmid hydrolysates were analyzed by highperformance liquid chromatography (HPLC) and highperformance liquid chromatography tandem mass spectrometry (HPLC-MS/MS) to determine the amount of each deoxynuceloside (i.e., deoxyguanosine (dG), deoxyadenosine (dA), deoxythymidine (dT), and deoxycytidine (dC)) and 8hydroxy-2′-deoxyguanosine (8-OHdG), respectively (Text S1 in Supporting Information). Gel electrophoresis was performed to monitor plasmid strand breaks before and after illumination. Eight microliter plasmid samples (10 mg/L) were mixed with 0.8 μL of gel loading buffer; then 5 μL of samples was loaded onto 1% agarose gel with SYBR Green I. Gel electrophoresis was conducted in 1 × TAE solution at 100 V for 50 min. The bands were visualized by using a Tanon 1600 imaging system (Tanon Science and Technology Co., China). The optical density of various bands was analyzed using C


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Environmental Science & Technology any treatment were 4.4 × 10−4 and 4.3 × 10−4, respectively (Figure S1). The tet A and blaTEM‑1 were encoded on the same plasmid, therefore having nearly equal transformation efficiencies.7 On average, 1 out of 2300 E. coli DH5α cells could acquire the resistance with exposure to the plasmids containing eARGs. Under light illumination the transformation efficiencies of eARGs dropped quickly and followed pseudofirst-order kinetics (Figure 2). The decrease rate constants of

Figure 3. Photodegradation rate constants of (a) tet A and (b) blaTEM‑1 genes in DOM (20 mg/L) solution under different conditions (1, DOM; 2, DOM+IPA; 3, DOM+HCOONa; 4, DOM +NaN3; 5, DOM−D2O solution; 6, DOM−N2-saturated; 7, DOM− N2-saturated+IPA; 8, calculated direct photodegradation rate constants by multiplying screening factor of 20 mg/L DOM). Rate constants obtained based on the linear regression of the data measured by short-amplicon qPCR. Error bars indicate 95% confidence intervals for n = 4.

eARGs photodegradation, the DOM solution was spiked with NaN3 to scavenge both •OH and 1O2. Photodegradation rates were found to further decrease with the addition of NaN3 (Figure 3), suggesting that 1O2 might also play a role in the eARGs photodegradation in the presence of DOM. Moreover, photodegradation rates greatly increased (Figure 3) in DOM− D2O solution. Given the higher lifetime of 1O2 in D2O,23 1O2 was confirmed to participate in the eARGs photodegradation. The eARGs photodegradation was performed under anaerobic condition to investigate the role of 3DOM*.23,24 Under N2saturated condition, the photodegradation rate of tet A decreased slightly (p < 0.01) and the photodegradation rate of blaTEM‑1 were nearly unchanged (p > 0.05) compared to the corresponding controls under air-saturated condition (Figure 3). However, under N2-saturated condition with addition of IPA the eARGs photodegradation were inhibited and their photodegradation rates were comparable with their direct photodegradation rates (p > 0.05) (Figure 3). Structure Changes of the Plasmid. Bases damage and strand breaks are regarded to be the two major forms of DNA lesions;15 thus, both were investigated here to clarify the mechanisms of eARGs photodegradation. As shown in Figure 4a, compared to individual light illumination, photodegradation of dG was selectively enhanced (p < 0.05), while the photodegradation rates of the other three deoxynucleosides (i.e., dA, dT, and dC) were nearly unchanged (p > 0.05) in the DOM solution. To confirm the enhanced photodegradation of dG in plasmid, the concentration of 8-OHdG, one of the major oxidative products of dG,16 was determined. Before illumination, the initial background concentration of 8-OH-dG was 1.6 μg/L. After illumination for 12 h, the levels of 8-OH-dG in plasmid were elevated to 14.7 and 23.7 μg/L in the absence and presence of DOM, respectively (Figure 4b). These results proved the promoting role of DOM in the photodegradation of guanine in plasmid, which might partially contribute to the

Figure 2. Changes in the relative transformation efficiencies of (a) tet A and (b) blaTEM‑1 genes under light illumination or dark controls in phosphate buffer with or without DOM (20 mg/L) measured by plasmid transformation experiments. Dark controls refer to the same treatments (stirring at 25 ◦C in the air) with the corresponding experimental groups except for light illumination. Concentration of plasmid is 0.1 mg/L. Symbols are the measured data, and lines are the linear regression of the data. Error bars represent 95% confidence intervals for n = 4.

transformation efficiencies for tet A and blaTEM‑1 were 0.62 and 0.61 h−1, respectively, under individual light illumination (Table S2). In the presence of DOM, the decreases of transformation efficiencies were promoted (p < 0.05) and the rate constants increased to 1.38 h−1 for tet A and 1.41 h−1 for blaTEM‑1 (Figure 2, Table S2). No obvious decreases were observed under dark conditions in both PBS buffer and DOM solution (Figure 2). These results suggest that the photoexcitation of DOM accelerated the decreases of eARGs transformation efficiencies and inhibited their HGT. Involved Reactive Intermediates in the eARGs Photodegradation in DOM Solution. It is widely accepted that DOM can photosensitize to produce reactive intermediates such as •OH, 1O2, and 3DOM* to enhance various contaminants photodegradation in aquatic environments.20 Both the tet A and the blaTEM‑1 genes photodegradation rates decreased (Figure 3) after addition of IPA or HCOONa to quench •OH, reflecting that the photoproduced •OH contributed to the enhanced photodegradation of eARGs in DOM solution. To investigate whether 1O2 participated in the D


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Figure 4. (a) Changes in the concentration of each deoxynuceloside in plasmid under illumination with or without DOM (20 mg/L). (b) Concentration of 8-OHdG in plasmid before or after 12 h light illumination. (* and **) Significance at p < 0.05 and 0.01 (n = 2), respectively. (c) Gel electrophoresis of plasmid and corresponding (d) bands optical density under different conditions (band 1, initial plasmid; band 2, initial plasmid with 20 mg/L DOM; band 3, after 12 h individual light illumination; band 4, after 12 h illumination within 20 mg/L DOM solution). UV dose of 12 h light illumination is 172.8 J/cm2. Concentration of plasmid used here is 10 mg/L.

then further broke to the linear form.30 With spiking of DOM, the bands of untreated plasmids remained nearly unchanged (Figure 4c and 4d, lane2), which indicates that DOM did not interfere with the gel electrophoresis of plasmids. After 12 h individual light illumination, the supercoiled form totally disappeared. The percentage of nicked form increased to about 67% and a linear form appeared, which accounted for 33% (Figure 4c and 4d, lane 3). In DOM solution, all bands nearly disappeared after 12 h illumination (Figure 4c and 4d, lane 4), suggesting that the photoexcitation of DOM further accelerated the strand breaks of plasmids, and smaller DNA fragments might form after 12 h illumination. To investigate the role of ROS in plasmid strand breaks, •OH quenching and 1 O2 enhancing experiments were performed. When dosing IPA to scavenge •OH, the strand breaks were substantially suppressed and the strand breaks rate was almost equal to that in PBS buffer (p > 0.05) (Figure S3), which demonstrates the crucial role of •OH in plasmid strand breaks. Additionally, the strand breaks rate was nearly unchanged in DOM−D2O solution compared to that in DOM−H2O solution (p > 0.05) (Figure S3), suggesting that 1O2 did not participate in plasmid strand breaks. Besides, the concentrations of plasmids were almost unchanged after 12 h illumination in the absence or presence of DOM (p > 0.05) (Figure S4), suggesting no mineralization of plasmids during light illumination.

enhanced eARGs photodegradation. ROS quenching or enhancing experiments were conducted to investigate the roles of various ROS in the plasmid bases damage. With dosing of IPA to quench •OH, the photodegradation rates of all four deoxynucleosides were almost unchanged (p > 0.05) (Figure S2), indicating that •OH did not play a role in the plasmid bases damage. In DOM−D2O solution, only dG damage was promoted (p < 0.05) (Figure S2), which suggests that 1O2 selectively enhanced the damage of guanine. After spiking NaN3 in DOM solution, the dG photodegradation was suppressed and its photodegradation rate was almost equal to that in PBS buffer (p > 0.05) (Figure S2), further demonstrating that 1O2 played a crucial role in enhancing guanine damage. Gel electrophoresis experiments were performed to explore plasmid strand breaks. As shown in Figure 4c, there were two bands in the gel of untreated pBR322 plasmids. The major band between 2.0 kb and 3.0 kb was attributed to supercoiled form (Form I), and the minor band above 5.0 kb was assigned to nicked circle form (Form II) (Figure 4c, lane 1).7 The band generated after illumination between Form I and II was attributed to the linear form (Form III) (Figure 4c, lane 3).7 Compared to the linear form of plasmids, the supercoiled form was usually more compacted and moved faster in gel, while the nicked circle form was looser and moved slower in gel.30 During plasmid strand breaks, the supercoiled form first fractured to the nicked circle form and E


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Environmental Science & Technology Effect of DOM Concentration and Ionic Species on eARGs Photodegradation. The DOM concentration influences the transmission of light as well as the level of ROS in waters, thereby affecting the photodegradation of contaminants.31 Herein, the photodegradation of eARGs was investigated at different levels of DOM. When DOM concentration increased from 0 to 20 mg/L, the photodegradation rate constants of eARGs linearly increased from 0.13 to 0.23 h−1 for tet A and from 0.18 to 0.30 h−1 for blaTEM‑1 (Figure 5). However, the photodegradation rate constants of

Figure 6. Photodegradation rate constants of tet A and blaTEM‑1 genes in DOM (20 mg/L) solution dosing different ions (5 mM Cl−; 1 mM NO3−; 20 μM NO2−; 10 μM Br− and 2.5 mM HCO3−; 5 μM Fe3+). Photodegradation rate constants determined by short-amplicon qPCR. (* and **) Significance at p < 0.05 and 0.01, respectively, compared to the corresponding control group (n = 4).

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DISCUSSION Mechanisms of Enhanced eARGs Photodegradation in DOM Solution. The present study demonstrated that the photodegradation of eARGs could be enhanced with photoexcitation of DOM (Figure 1). DOM is a kind of natural photosensitizer ubiquitously present in waters and can be photoexcited to produce reactive intermediates (e.g., •OH, 1 O2, and 3DOM*) to enhance various contaminants photodegradation.20 Herein, photoproduced ROS from DOM were proven to accelerate eARGs photodegradation (Figure 3). As for bases damage, only the photodegradation of the dG was promoted in the presence of DOM (Figure 4a and 4b). The ROS quenching or enhancing experiments demonstrated the role of 1O2 in enhancing the damage of dG in plasmid, while • OH played negligible role in bases photodamage (Figure S2). According to previous studies •OH can nonselectively oxidize all four bases, while 1O2 selectively oxidizes guanine.15,16 Here, the highly oxidizing •OH failed to induce bases degradation in plasmid, possibly because that the lifetime of •OH was very short, and it was scavenged before reaching the inner bases sites. In addition, the plasmid strand breaks was enhanced in the presence of DOM (Figure 4c and 4d). The •OH quenching and 1O2 enhancing experiments proved that •OH rather than 1O2 resulted in the enhanced plasmid strand breaks (Figure S3). •OH could induce DNA strand breaks by extracting •H from sugar moiety of DNA,15 while 1O2 was unable to induce the strand breaks due to its relatively lower redox potential.33 Regarding to the role of 3DOM*, the photodegradation experiment under N2-saturated condition was usually adopted to investigate the role of 3DOM* in the contaminants photodegradation.23,24,34 In general, the 3DOM* is expected to play an important role in the contaminant photodegradation if the photodegradation of contaminant is enhanced under anaerobic condition, and •OH dose not play a significant role.24,34 However, under N2-saturated condition, the photodegradation rate of tet A decreased slightly and the photodegradation rate of blaTEM‑1 was almost unchanged (Figure 3). To illuminate the role of 3DOM*, the eARGs photodegradation experiments were further conducted with spiking of IPA under N2-saturated condition to exclude both •OH and 1 O2.24 Under this condition, the enhanced photodegradation

Figure 5. Photodegradation rate constants of eARGs within different concentrations of DOM solution (dashed lines represent rate constants after correction by screening factors of DOM; detailed correction procedures are shown in the Supporting Information, Text S4). Rate constants obtained from linear regression of the data measured by short-amplicon qPCR. Error bars indicate the standard deviation of mean for n = 4.

tet A and blaTEM‑1 genes dropped to 0.14 and 0.19 h−1, respectively, with further increasing DOM concentration to 40 mg/L (Figure 5). Various commonly occurring ions in natural waters, such as Cl−, NO3−, NO2−, Br−, HCO3−, and Fe3+, can influence the level of ROS.32 Considering the importance of ROS in the degradation of eARGs, the effects of these coexisting ions at the environmental level on the photodegradation of eARGs in DOM solution were investigated. As in Figure 6, Cl− had a slight inhibiting effect on tet A photodegradation (p < 0.05), while it had no influence on blaTEM‑1 photodegradation (p > 0.05). After addition of nitrate, the eARGs photodegradation rates were greatly enhanced (p < 0.01) and the degradation rate constants of tet A and blaTEM‑1 increased by 82.4% and 75.0%, respectively. The dosage of nitrite also significantly increased the photodegradation of eARGs (p < 0.01), and tet A and blaTEM‑1 photodegradation rate constants increased by 16.8% and 19.6%, respectively (Figure 6). When dosing Br−, the eARGs photodegradation was inhibited (p < 0.01) and the rate constants decreased by 23.2% and 27.2% for tet A and blaTEM‑1, respectively (Figure 6). Addition of HCO3− significantly decelerated the photodegradation of tet A (p < 0.01) and blaTEM‑1 (p < 0.05), the corresponding rate constants decreased by 23.6% and 23.5%, respectively (Figure 6). With the addition of Fe3+, the photodegradation of tet A and blaTEM‑1 were enhanced (p < 0.01), and the corresponding degradation rate constants increased by 59.7% and 44.4%, respectively, compared to that of the controls (Figure 6). F


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also quench 1O2.38 Therefore, at a high concentration, DOM’s contribution to ROS removal probably exceeded its contribution to ROS production. This resulted in slower eARGs photodegradation rates at higher DOM concentrations. In addition to DOM itself, various ions in water can also influence ROS levels. According to previous work, Cl−, Br−, and HCO3− were all able to scavenge •OH and 3DOM* and produced secondary reactive intermediates (e.g., •Br, •Br2−, •Cl, •Cl2−, and CO3·−).39,40 The photodegradation rate of contaminant induced by reactive intermediates was dependent on both the steady state concentration of reactive intermediates and the second-order rate constant between contaminant and reactive intermediates.29 The steady state concentrations of secondary reactive intermediates were usually higher than that of • OH.39,40 However, the reaction activities of plasmid with reactive bromine species or carbonate radicals were possibly much lower than that with •OH, thus resulting in decreased eARGs photodegradation rates in the presence of Br− or HCO3−. In addition, the quench of 3DOM* by Br− or HCO3− could result in the decrease of 1O2 concentration, which would also inhibit the eARGs photodegradation. Nevertheless, Cl− had a negligible effect on the eARGs photodegradation (Figure 6), probably because Cl− could not quench •OH and 3DOM* effectively at a low concentration.39,41 In terms of NO3− and NO2−, these two anions could be photoexcited to produce • OH,42 thereby enhancing the photodegradation of eARGs (p < 0.01). Besides, iron-based compounds such as Fe(OH)2+ and Fe−DOM complex were expected to be important photoactive species in aquatic environments, which could be photoexcited to produce •OH.43 This probably resulted in the accelerated photodegradation of eARGs in DOM solution with the addition of Fe3+. In brief, DOM concentration and ions influenced the eARGs photodegradation in DOM solution by affecting the •OH or 1O2 levels. Other water elements with the power to influence the levels of •OH or 1O2 may also change the photodegradation rates of eARGs in aquatic environments. Environmental Implications. DOM and especially humic substances were usually considered to protect eARGs against degradation and maintain their persistence in the environment.35 However, in sunlit waters, the protective effect of DOM on eARGs may be challenged based on the results of the present study. The photosensitization of DOM can produce ROS to enhance the photodegradation of eARGs. To better understand the fate and environmental risk of eARGs in water, it is necessary to take into consideration both the adsorption protection and the photodegradation effects of DOM on eARGs. In future work, the role of DOM in the degradation of eARGs in sunlit waters should be reevaluated by investigating both enzyme hydrolysis and photodegradation in the presence of DOM. This study revealed that transformation efficiencies of eARGs drastically decreased under illumination. Consequently, sunlight illumination may make a great contribution to weaken the dispersion of eARGs in aquatic environments. Wastewater effluents contain abundant and diverse eARGs,4 which would undergo photodegradation and decreased transformation efficiency upon being discharged in natural waters. Due to the higher ROS quantum yield of effluent organic matter compared to that of natural DOM and the higher content of nitrate and nitrite in effluent,44,45 the eARGs in effluent water may undergo faster photodegradation than those in natural waters. On the basis of this, it is recommendable to store eARGs-rich effluent water in artificial wetlands or open-air

was thoroughly inhibited (p > 0.05) (Figure 3).This suggests that 3DOM* did not play a role in the eARGs photodegradation. In addition, after dosing NaN3 to scavenge 1O2, the enhanced dG damage was totally suppressed (p > 0.05) (Figure S2), reflecting that 3DOM* did not participate in dG damage. Furthermore, with spiking of IPA to quench •OH, the enhanced strand breaks was also totally inhibited (p > 0.05) (Figure S3). This indicates that 3DOM* could not lead to strand breaks. In brief, 3DOM* could not induce plasmid damages, thus resulting in the eARGs photodegradation. After excluding the role of 3DOM*, the contributions of 1O2 and •OH were calculated based on the quenching experiments (Text S2). During the photodegradation of tet A, the contribution of 1O2 (average 25.0%) was nearly equal to that of •OH (average 24.4%) (Table S3). Regarding to blaTEM‑1 gene, the contribution of 1O2 (average 21.2%) was slightly lower than that of •OH (average 27.4%) (Table S3). 1O2- and • OH-mediated indirect photodegradation played crucial role in the eARGs photodegradation, and its contribution was comparable with that of direct photodegradation (Table S3). Overall, DOM was photosensitized to produce 1O2 enhances dG oxidation and •OH to enhance plasmid strand breaks, thereby promoting plasmid-encoded ARGs photodegradation and decreasing their transformation efficiencies. Relationship between eARGs Photodegradation and the Decrease in Transformation Efficiencies. Transformation of eARGs can lead to dissemination of antibiotic resistance and even lead to antibiotic resistance of pathogenic bacteria.35 Therefore, it is crucial to know the transformation potential of eARGs. Substantial decreases of transformation efficiencies of eARGs were observed after photodegradation, and DOM photosensitization could promote this process (Figure 2). However, the photodegradation rates of eARGs measured by long-amplicon qPCR were lower than the decrease rates of transformation efficiencies (Table S2). This implies that the qPCR measurements underestimated the loss of transformation efficiencies of eARGs. Certain sequence or reactive sites outside tet A and blaTEM‑1 genes might be also related to their transformation. The photodegradation of these sequences or reactive sites could also result in the decreases of eARGs transformation efficiencies. For example, the promoter of gene was crucial for gene expression after transformation.36 Damage of promoters of eARGs would reduce eARGs expression, thereby exhibiting a decrease in transformation efficiencies. Factors Affecting the eARGs Photodegradation in DOM Solution. With increasing DOM concentration, photodegradation rates of eARGs first increased and then decreased (Figure 5). The steady state concentration of both •OH and 1 O2 increased with increasing DOM levels (Text S3 and Figure S5), indicating that the decreases of eARGs photodegradation rates under a high DOM concentration (e.g., >20 mg/L) did not result from changes in ROS production. Due to the light screening effect,29 DOM would compete with plasmids for photons, probably inhibiting direct eARGs photodegradation. However, after eliminating the DOM light screening effect on the direct photodegradation of eARGs (Text S4 and Figure S6), the photodegradation of eARGs still decreased when the DOM concentration was above 20 mg/L (Figure 5), which suggests that the decreases of eARGs photodegradation under higher DOM level might be attributed to the quenching effect of DOM on ROS.37,38 DOM exhibited a high reaction rate toward •OH (5.6 × 109 M−1 s−1).37 In addition, DOM could G


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units in order to benefit of sunlight illumination effects on eARGs.

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ASSOCIATED CONTENT

* Supporting Information S

The Supporting Information is available free of charge on the ACS Publications website at DOI: 10.1021/acs.est.9b03096.

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Measurements of the concentrations of deoxynucleosides and 8-OHdG; calculation of the contributions of direct photodegradation and ROS-induced indirect photodegradation; determination of the steady state concentrations of •OH and 1O2; calculation of screening factors and correction of the photodegradation rate constants; initial transformation efficiencies of tet A and blaTEM-1 genes without any treatment; deoxynuceloside (dC, dG, dA, and dT) photodegradation in PBS buffer or in 20 mg/L DOM solution dosing ROS quenchers or in 20 mg/L DOM−D2O solution; lightinduced strand break of plasmid in PBS buffer or DOM solution (20 mg/L) characterized by gel electrophoresis; concentrations of plasmid DNA before or after 12 h illumination with or without DOM; steady state concentrations of •OH and 1O2 under illumination within different concentrations of DOM; screening factors of different concentrations of DOM; sequences information of short and long amplicons for the tet A and blaTEM-1 genes and the corresponding primers; photodegradation rate constants of eARGs measured by short-amplicon qPCR (SA-qPCR) and long-amplicon qPCR (LA-qPCR) and the decrease rate constants of eARGs transformation efficiencies; calculated contributions (%) of direct photodegradation, •OH and 1O2 to the photodegradation of tet A and blaTEM-1 (PDF)

AUTHOR INFORMATION

Corresponding Author

*Fax: +86-551-63601592. E-mail: [email protected]. ORCID

Guo-Ping Sheng: 0000-0003-4579-1654 Notes

The authors declare no competing financial interest.

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ACKNOWLEDGMENTS The authors thank the National Natural Science Foundation of China (51738012, 51825804, and 51821006) for partial support of this study.

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REFERENCES

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Enhanced Photodegradation of Extracellular Antibiotic Resistance

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