Substituted Aromatic-Facilitated Dissemination of Mobile Antibiotic

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Characterization of Natural and Affected Environments

Substituted Aromatic-Facilitated Dissemination of Mobile ARGs via an Anti-Hydrolysis Mechanism across an EPS Permeable Barrier Weijun Shou, Fuxing Kang, Shuhan Huang, Chunyao Yan, Jiaxin Zhou, and Yijin Wang Environ. Sci. Technol., Just Accepted Manuscript • DOI: 10.1021/acs.est.8b05750 • Publication Date (Web): 18 Dec 2018 Downloaded from http://pubs.acs.org on December 19, 2018

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Environmental Science & Technology

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Substituted Aromatic-Facilitated Dissemination of Mobile ARGs via an Anti-Hydrolysis

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Mechanism across an EPS Permeable Barrier

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Weijun Shou, Fuxing Kang*, Shuhan Huang, Chunyao Yan, Jiaxin Zhou, and Yijin Wang

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College of Resources and Environmental Sciences, Nanjing Agricultural University, Jiangsu 210095,

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China

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*Corresponding author. [email protected]/[email protected] (F. Kang)

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Manuscript prepared for Environmental Science& Technology

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December 03, 2018

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1

ACS Paragon Plus Environment


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ABSTRACT: Mobile antibiotic resistance genes (ARGs) in environmental systems may pose a

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threat to public health. The coexisting substituted aromatic pollutants may help the ARGs cross the

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extracellular polymeric substance permeable barrier into the interior of cells, facilitating ARG

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dissemination, but the mechanism is still unknown. Here, we demonstrated that a specific anti-

27

hydrolysis mechanism of mobile plasmid in the extracellular matrix makes a greater contribution to

28

this facilitated dissemination. Specifically, fluorescence microtitration with a Tb3+-labeled pUC19

29

plasmid was used to study the formation of substituted aromatic-plasmid complexes associated with

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ARG dissemination. Manipulations of the endA gene and EPS confirmed that these forming

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complexes antagonize the EPS-mediated hydrolysis of the plasmid. Fourier transform infrared

32

spectroscopy (FTIR), X-ray photoelectron spectroscopy (XPS), and computational chemistry

33

demonstrated that substituents alter the polarity of aromatic molecules, making the carbon at the 6-

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position of 1, 3-dichlorobenzene as well as the labile protons (-NH2/-OH) of m-phenylenediamine,

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aniline, and 2-naphthol interact with the deprotonated hydroxy group of the phosphate (P-O···H-

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C/N/O), mainly via hydrogen bonds. Linear correlations among ARG disseminations, association

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constants, and bonding energies highlight the quantitative dependency of ARG proliferation on a

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combination of functionalities templated by D-ribose-phosphate.

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2


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INTRODUCTION

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Dissemination of mobile genetic elements has attracted much attention in the past decade1 due

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to the potential hazard of antibiotic resistance genes (ARGs) to public health. Like antibiotics,

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metals,2 nanoparticles,3, 4 ionic liquids,5 and organic pollutants6 can facilitate ARG dissemination in

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different environments. There is evidence that ionic compounds/metal species may help ARGs cross

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EPS permeable barriers into the interior of cells,7 facilitating ARG dissemination. Nevertheless, the

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mechanisms of free ARG proliferation facilitated by co-existing substituted aromatic pollutants

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occurring within the microbially extracellular interface remain unknown, warranting further

48

investigation.

49

To ensure survival under hostile environmental conditions, bacterial cells have developed many

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defense mechanisms to deal with exogenous agents, the secretion of extracellular polymeric

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substances (EPS) is the one of such defense mechanisms. The EPS, constituted by the majority of

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chemically active polysaccharides and bioactive proteins, can accommodate the bacterial cells

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themselves to form a stable microhabitat8 for defending against ARG invaders.9, 10 Thus, the EPS

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matrix is extremely important for serving as an environment-microbe interface in which alien

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chemical/biological components encounter and combat with endogenously defensive materials.

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Therein, extracellular endonucleases11 and alien ARGs can interact in the EPS matrix of biofilms,8, 10

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and these ARGs are easily destroyed by a process called hydrolysis. Apparently, in the EPS matrix

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acted as permeable barrier, such self-defense of biological significance is practically consistent with

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the interdiction of ARG dissemination among bacterial cells/populations in the environmental sense.

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It is important to explore how characters and types of typically organic pollutants affect ARG

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dissemination occurring at this interface.

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Mobile ARG dissemination is affected by the structure of aromatic molecules. Because 3



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plasmids containing ARGs consist of neutral hydrophobic (genetic bases) and charged hydrophilic

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moieties (D-ribose-phosphate),10 they can anchor the weak/nonpolar aromatic hydrocarbons and

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polar substituted aromatics. For instance, polyaromatic hydrocarbons (PAHs) (e.g., phenanthrene

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and pyrene) combine with genetic bases via hydrophobic interactions12,

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dissemination.7 Therein, noncovalent interactions disrupting the hydrogen bonds between genetic

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bases13-15 are associated with π electrons in PAHs serving as inhibitors.16 However, substituents in

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aromatic hydrocarbons, including hydroxy, amino, and chlorine moieties, can alter the

70

physicochemical properties of the aromatic nucleus17-19 (molecular polarity, hydrophobicity,

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electron-donating character, etc.) by taking a π electron from the nucleus. Undoubtedly, the change

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in types of bonds between the aromatics and plasmids can be affected by the substituents on the

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aromatic ring, which might cause aromatics to behave as facilitators (rather than inhibitors16, 20 like

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the parent aromatic hydrocarbons) of ARG proliferation. Apparently, it calls for the reconsideration

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of the roles of substituted aromatics in ARG dissemination occurring in the EPS permeable barrier.

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Few reports have been published on the correlative mechanism between the substituted aromatic-

77

plasmid complex and ARG dissemination.

13

to inhibit ARG

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Currently, the careful determination of the key functionalities of the plasmid that is

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responsible for the association with the substituted aromatics and the quantitative elucidation of

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the relationship between a substituted aromatic-plasmid complex and ARG dissemination are both

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of utmost importance. Based on environmental chemistry studies conducted using a variety of

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techniques (e.g., spectroscopy,21 molecular manipulation,22 and computational chemistry23), the

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interactions between aromatic species and DNA mainly depend on the types of functional groups

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present on the substituted aromatic compounds and the binding sites located at the genetic bases

85

(interlamination), groove, and external backbone;17,

24,

4


25

intercalation of the aromatic

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hydrocarbons between genetic bases has been known to inhibit ARG proliferation.7, 16, 20 These

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successful cases will offer more comprehensive insight into the relationship between

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multisubstituted aromatic compounds and the facilitation of ARG proliferation, which could allow

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more in-depth mechanistic studies occurring in the EPS permeable barrier, ultimately improving

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our understanding of environmental processes of freely mobile genetic elements in combined

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pollution systems (e.g., sludge-engineered wastewater and/or stock manure-irrigated soil ).

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It is expected that the combined approaches of fluorescence microtitrimetry, batch

93

experiments, spectroscopy, and computational chemistry can offer deeper and more

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comprehensive insight into coupling mechanisms as well as the association with substituted

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aromatic-facilitated ARG proliferation (flowchart can be found in Figure S1). In the present study,

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four substituted aromatic compounds (1,3-dichlorobenzene, 2-naphthol, aniline, and m-

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phenylenediamine) are used throughout all experiments because they are ubiquitous in the

98

environment and have multiple electrophilic activities dominated by functionalities. Manipulations

99

of the endA gene (responsible for production of extracellular DNase) and EPS were used to explore

100

the anti-hydrolysis mechanism occurring in the EPS matrix. To further characterize the key

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functionalities of the plasmid, X-ray photoelectron spectroscopy (XPS) and Fourier transform

102

infrared spectroscopy (FTIR) are used to investigate the D-ribose-phosphate-oriented coupling

103

mechanisms responsible for ARG proliferation. A group of batch experiments are performed to

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explore the dependency of ARG dissemination efficiency (T/T0) on bonding energy at key bonding

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sites. The results obtained from the computational chemistry are used to further confirm the

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contribution of major functionalities for the facilitated ARG dissemination utilizing an electron

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localization function (ELF). To obtain a group of statistically significant data, in addition to the above

108

four substituted aromatics, 1,3-dinitrobenzene and 1-naphthylamine are also used to investigate 5



109

aromatic-plasmid associations and the associated relative ARG dissemination efficiency (T/T0).

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MATERIALS AND METHODS

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Materials. Four substituted aromatic compounds, 1,3-dichlorobenzene (1,3DCB), 2-naphthol

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(2NAOL), aniline (AN), and m-phenylenediamine (1,3MPD), were used in our experiments (Sigma-

113

Aldrich, USA). Therein, in the fluorescence microtitration with a Tb3+-labeled pUC19 plasmid, ARG

114

transformation, and computation for bonding energy, 1,3-dinitrobenzene (1,3DNB) and 2-

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naphthylamine (2NA) were additionally used to obtain a better statistical result. Physicochemical

116

properties of all substituted aromatic compounds, including the molecular weight (MW), molecular

117

structure, water solubility (SW), and n-octanol-water partition coefficient (KOW), are given in the

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Table S1. A circular plasmid, pUC19, with 2686 base pairs containing an ampicillin resistance gene

119

was used to perform all experiments (Takara, China). Escherichia coli K-12 (E. coli K-12) and DH5α

120

(E. coli DH5α) were used to prepare the competent cells according to a previous method.7 Their major

121

genomic difference is that endA gene of E. coli DH5α is knocked out,26, 27 because DNase I in the

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extracellular matrix, decoded by the endA, serves as a key barrier to hydrolyze exogenous plasmid

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(genetic sequence can be found in the sequence of the endA gene in the Supporting Information).

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Tb(NO3)3 (purity > 99%, Sigma-Aldrich, USA) was used as the fluorescence probe to explore the

125

formation of substituted aromatic-plasmid complexes. Peptone, yeast extract, agar, NaCl, and D-

126

glucose (biotechnology grade, Thermo Fisher Oxoid, UK) were used to prepare the liquid and solid

127

media. Ampicillin sodium (purity > 98%) was also purchased from Sigma-Aldrich, USA. Ultrapure

128

water was used in all experiments (electrical conductivity = 18.2 MΩ × cm, Millipore, USA).

129

Fluorescence Microtitration with the Tb3+-Labeled Plasmid. A combination of fluorescence-

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quenching microtitrimetry with Tb3+-labeled pUC19 was used to quantitatively explore the

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complexes of the substituted aromatic compounds with pUC19. Tb3+ was used to label the genetic 6


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bases and phosphate groups of the plasmid.28 The Tb3+-labeled plasmids could be excited at 280 nm

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to produce a narrow emission signal at 490 nm. Formation of the substituted aromatic-plasmid

134

complexes could quench this fluorescence chromophore, and the magnitude of the quenching was

135

used to quantify both their association constants and the number of bonding sites. Specifically, a 2.5-

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μL aliquot of aqueous Tb(NO3)3 solution (1.0 mmol/L) was added to 20 mL of aqueous plasmid

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solution (2.0 mg/L), and the mixture was incubated for 20 min at 25°C. The mixture was centrifuged

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through an ultrafiltration centrifugal tube with a polyethersulfone (PEF) 3000-Da membrane

139

(Millipore, USA) for 10 min at 4°C and 6000 g. The plasmid pellet was resuspended in Milli-Q water

140

and centrifuged to force the free Tb3+ into the aqueous solution. After this washing process was

141

performed three times, the plasmid pellet was finally dissolved in 20 mL of Milli-Q water. Substituted

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aromatic compounds (as 10 mmol/L methanol stock solutions) were titrated into the Tb3+-labeled

143

plasmid using a chromatographic injector (25-μL scale, Agilent, USA) and then magnetically stirred

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for 20 min at 160 revolutions per minute (rpm), pH 7.0, and 25 °C. Three-dimensional excitation-

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emission matrix (3DEEM) fluorescence spectroscopy was conducted at an excitation (EX)

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wavelength of 200–370 nm (5 nm bandwidth) and an emission (EM) wavelength of 280–550 nm (5

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nm bandwidth) at a scanning rate of 3000 nm/min (F96Pro, Lengguang, China). The microtitration

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and fluorescence detection procedures were repeated until at least six sets of titration data were

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collected for each substituted aromatic compound. The fluorescence peak (EX/EM = 280 nm/490

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nm) was continuously monitored by the microtitration and detection processes.

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The changes in the fluorescence intensity of Tb3+-labeled plasmid caused by substituted aromatic compounds could be described by the Sterne-Volmer equation:29-32 F0  1  K q 0 [Q]  1  KSV [Q] F

(1),

where F0 and F are the relative fluorescence intensities of the chromophore (Tb3+-labeled plasmid) 7



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in the absence and presence of the quencher (substituted aromatic compound), respectively; Kq is the

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bimolecular quenching rate constant; τ0 is the average lifetime of the fluorophore in the absence of

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the quencher; [Q] is the concentration of the quencher; and KSV is the Stern-Volmer quenching

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constant. For the static quenching process, equation 2 was used to determine the association constant

159

(logKA) and the number of bonding sites (n):33

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log[( F0  F ) / F ]  log K A  n log[Q]

(2).

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Propagation of ARG in the Presence of Substituted Aromatic Compounds. An exposure

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experiment was performed to obtain the plasmid pUC19 stained with the substituted aromatic

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compounds. A dilution method was used to prepare an aqueous plasmid solution (0.5 μg/mL), and a

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stock solution of the aromatic compound in methanol as the carrier (0–0.5 µmol/L) was added. The

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methanol carrier-to-water ratio was kept below 0.1%. All samples were gently shaken manually for

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5 seconds and then left to stand for 2 h to facilitate the association of the aromatic compounds with

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pUC19, and then the samples were centrifuged in 1.5-mL tubes with a disposable 3000-Da

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ultrafiltration membrane (Millipore, USA) for 10 min at 4 °C and 3000 g to remove the free

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substituted aromatic compounds from the solution. It is noteworthy that this process could reduce the

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effect of the unbound/free substituted aromatic compounds on the competent cells in the subsequent

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transformational experiment.

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E. coli K-12 and DH5α competent cells were first treated using EPS manipulation: without

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manipulation of EPS (referred as pristine cells) and with removal of EPS from the surface of E. coli

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K-12 and DH5α using the above sonication/centrifugation method (referred to as low-EPS cells).34

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Briefly, E. coli K-12 and DH5α were cultured in LB medium for 48 h at 37°C and 120 rpm, followed

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by a washing operation35 and removal of EPS on the surface of E. coli cells.18, 34 The obtained E. coli

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K-12 and DH5α were used to prepare the respective competent cells.7 8


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A chemical transformation was used to assess the effect of formation of the substituted aromatic-

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plasmid complex on genetic dissemination. As described previously,7, 16 pUC19 (0.25 μg, 200 μL)

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bound to the substituted aromatics was added to competent E. coli cells (K-12 and DH5α, OD560nm =

181

0.40) after the removal of unbound/free substituted aromatic compounds by centrifugation through a

182

3000-Da ultrafiltration membrane (Millipore, USA) for 10 min at 4 °C and 3000 g. The mixture of

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the competent cells and pUC19 was placed in an ice-water bath for 3.0 min and then immediately

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heat shocked at 42 °C for 90 seconds. The bacterial cells absorbing the pUC19 were immediately

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placed in an ice-water bath for 3 min and then mixed with super optimal broth with catabolite

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repression (SOC). After incubation at 120 rpm for 45 min at 37°C, the bacteria-inoculated liquid (100

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μL) was uniformly spread on the surface of LB solid culture media containing 100 mg/L ampicillin

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sodium. The petri dish was left upright for 30 min to ensure that the bacterial solution was fully

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integrated into the LB solid culture media, and then the sample was left inverted for 36 h at 37°C.

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The transformation was defined as the logarithm of the ratio of the number of transformants (unit

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number) to the mass of the added plasmid (0.25 μg), as follows:

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Transformation (T) = log (number of transformants plasmid pUC19)

(3).

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The relative ARG dissemination (T/T0) was defined as the ratio of pUC19 transformation in the

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presence of the substituted aromatic compounds (T, 0.4 μmol/L) to that in the absence of the

195

substituted aromatic compounds (T0).

196

Anti-Hydrolysis of Substituted Aromatic-Plasmid Complexes in the EPS Matrix. An in vivo

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experiment was specifically performed to study the anti-hydrolysis of substituted aromatic-plasmid

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complexes in the EPS matrix. In this experiment manipulating the endA gene and EPS, two competent

199

cells, E. coli K-12 and DH5α, were used to determine the effect of the EPS matrix on the anti-

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hydrolysis of substituted aromatic-plasmid complexes. 9



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Specifically, the extracted EPS from the surface of E. coli K-12 or DH5α30,

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was used to

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hydrolyze the substituted aromatic-plasmid complex. The “substituted aromatic-plasmid complex”

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pellets, obtained by a 3000-Da ultrafiltration membrane (Millipore, USA), were dissolved into the

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extracted EPS solution (23.8 mg Dry Weight/L, DW/L) to obtain the desired 0.5 μg/mL plasmid

205

complex. After incubation at 25 °C and pH 7.0 for 30 min, the absorbance of each sample (I) was

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immediately determined at 260 nm using a microplate reader (Tecan Infinite 200, Switzerland). In

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addition, the absorbance of a reference sample (I0) containing only plasmid and EPS was measured

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under the same conditions. According to the definition from Kunitz,36 the apparent degradation

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efficiency, regarding the substituted aromatic-pUC19 complexes, could be defined as the difference

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in absorbance at 260 nm before and after reaction of the plasmids with extracted EPS (I-I0).

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XPS and FTIR Analyses. X-ray photoelectron spectroscopy (XPS, UlVAC-PHI5000, Japan)

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was used to explore the changes in the surface functionalities of the pUC19 plasmid caused by the

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substituted aromatic compounds. After the microtitrimetry and fluorescence quenching (aromatic

214

compounds/plasmid = 1/100), all samples were freeze-dried under vacuum and stored at -65 °C for

215

one week (Labconco, USA). The surface valence electrons of the atoms in the functional groups on

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pUC19 were analyzed with a multidetection analyzer under 10-8 Pa residual pressure (UlVAC-

217

PHI5000, Japan). The C 1s peak at 284.6 eV was used as the reference to correct for the surface

218

charging effects. Fourier transform infrared spectroscopy (FTIR, Bruker, USA) analyses were

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performed to characterize the functional groups of these dried biofilms using a previously described

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method.

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Computational Chemistry. Based on the changes in the functional groups indicated by the XPS

222

and FTIR analyses, we constructed a D-ribose-phosphate model containing a pair of D-ribose

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moieties connected via a phosphoester. To reduce the computational load, the genetic bases, which 10


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were originally connected to the D-ribose in the pUC19 plasmid, were removed because evidence

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from the XPS and FTIR analyses suggested that the internal bases were not key bonding sites

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responsible for the interactions with the substituents on the substituted aromatic compounds.

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Molecular models including the substituted aromatic compounds, D-ribose-phosphate, and

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“substituted aromatic-to-D-ribose-phosphate” complexes were constructed using Gauss View

229

software.37 Before the computations were carried out, all molecular models were preliminarily

230

optimized at a B3LYP/3-21G level to the lowest energy. Afterwards, all models were formally

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computed at a B3LYP/6-31G(d) level of theory16, 17 with DFT-D3 for long-range corrections.38 All

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atoms in the constructed models could freely move in the geometry optimization computation.

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Solvent (water) effects were implicitly considered in these computations. The bonding energy for

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each “substituted aromatic···D-ribose-phosphate” complex was computed based on frequency

235

analysis. The calculation was performed as follows:

236

△ 𝐺 = 𝐺substituted aromatic···𝐷 ― ribose ― phosphate ―(𝐺substituted aromatic + 𝐺𝐷 ― ribose ― phosphate)

237

(4),

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where 𝐺substituted aromatic···𝐷 ― ribose ― phosphate, 𝐺𝐷 ― ribose ― phosphate, and 𝐺substituted aromatic are the

239

Gibbs free energies (G) of the “substituted aromatic···D-ribose-phosphate” complex (new bond

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formed between the plasmid and the substituted aromatic compound), D-ribose-phosphate, and

241

substituted aromatic compound, respectively.

242

Electron Localization Function (ELF). The ELF was used to quantitatively elucidate the

243

bonding properties based on the electronic interactions.39 Based on the wave function obtained from

244

the computational study, rigorous ELF topological structures considering the bonding properties

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between the aromatic compounds and D-ribose-phosphate were analyzed using Multiwfn software40

246

as follows: 11



247 248 249 250

ELF(𝐫) =

1

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(5),

1 + [𝐷(𝐫)/𝐷0(𝐫)]2

where 1 |∇ρ(𝐫)|2

[

1

𝐷(𝐫) = 2∑𝑖𝜂𝑖|∇𝜑𝑖(𝐫)|2 ― 8

2/3

𝐷0(𝐫) = 3 10(3𝜋2)

]

(6),

53

(7),

𝜌(𝐫)

𝜌(𝐫)

251

where r denotes a point in three-dimensional space; (r) and (r) are the electron density function

252

and orbital wave function, respectively; D(r) is the excess kinetic energy density caused by the Pauli

253

repulsion; and D0(r) is the Thomas-Fermi kinetic energy density. The ELF values were within the

254

range of [0,1]. A large ELF value means that the electrons are highly localized, which typically

255

corresponds to a covalent bond, a lone pair or atomic inner shell regions.

256

RESULTS AND DISCUSSION

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Formation of Aromatic-Plasmid Complexes Responsible for Facilitated ARG Propagation.

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Formation of substituted aromatic-plasmid complexes was explored using fluorescence quenching

259

microtitrimetry with a Tb3+-labeled pUC19 plasmid. Using the Sterne-Volmer equation (equation 1),

260

good linear correlations between the fluorescence intensities and the amount of substituted aromatic

261

compounds were observed (Figure 1a). The fitting parameters can be found in Table S2. Quenching

262

constants (KSV) increased in the order 7.99 × 105 (1,3DCB-plasmid), 2.41 × 106 (2NAOL-plasmid),

263

7.50 × 106 (AN-plasmid), and 4.64 × 106 L/mol (1,3MPD-plasmid), which suggests that both

264

hydroxy- and amino-substituted aromatic compounds have greater capacities for quenching Tb3+-

265

labeled plasmid chromophores than does the electron-poor 1,3DCB. Generally, the fluorescence

266

lifetime (τ0) of terbium is 3.20 × 10-5 s.41 On the basis of equation 1, the bimolecular quenching rate

267

constants (Kq values) for all of the above combinations were estimated to be approximately (2.50–

268

14.50) × 1010 L/mol/s. The value of Kq for a diffusion-controlled quenching process is less than 1.0

269

× 1010 L/mol/s;42 thus, the higher Kq values found in the present study suggest that a static quenching 12


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process related to the formation of the “substituted aromatic-plasmid” complexes was occurring.

271

Furthermore, based on equation 2, we calculated the association constant (logKA) and the number of

272

bonding sites (n) for substituted aromatic compounds on the pUC19 plasmid. Figure 1b shows that

273

four linear fits were parallel with each other, and Table S2 shows that the corresponding n values for

274

substituted aromatic-plasmid complexes were equal to 1.00 ± 0.10, which shows there was only one

275

binding site for aromatic compounds on pUC19. Based on the intercepts, the association constants

276

(logKA) were 5.82, 6.37, 6.99, and 7.07 for the 1,3DCB-, 2NAOL-, AN-, and 1,3MPD-pUC19

277

complexes, respectively. 1,3MPD, AN, and 2NAOL had stronger affinities for pUC19, which

278

suggests that electron-donating groups on the aromatic hydrocarbons made them bind more tightly in

279

pUC19 complexes than electron-poor 1,3DCB species.

280

The formation of substituted aromatic-pUC19 complexes had an important effect on ARG

281

propagation. Figure 1c-j shows the transformation of ARG to E. coli K-12 and E. coli DH5α. With

282

the addition of substituted aromatic compounds at concentrations up to 0.5 µmol/L, the

283

transformation of ARG to E. coliK-12 was enhanced from 1.57 to 2.29 (1,3DCB), 1.93 (2NAOL),

284

1.92 (AN), and 2.23 log transformants/μg pUC19 (1,3MPD), i.e., enhanced by 1.2−1.5 times. It is

285

apparent that hydroxy- and amino-substituted aromatic compounds made a greater contribution to

286

ARG propagation and that all tested substituted aromatic compounds served as facilitators of ARG

287

propagation. Meanwhile, transformations of ARG to E. coli DH5α showed that these substituted

288

aromatic compounds also served as facilitators (Figure 1g-j). As the concentrations of substituted

289

aromatic compounds increased from 0 to 0.5 µmol/L, the transformations changed from 3.02 to 3.30

290

(1,3DCB), 3.27 (2NAOL), 3.29 (AN), and 3.67 log transformants/μg pUC19 (1,3MPD), i.e.,

291

increased by 1.1−1.2 times. The results further confirm that the formation of substituted aromatic-

292

pUC19 complexes facilitated ARG propagation. 13



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Transformation of ARG to E. coli cells is associated with the endA gene, which is associated

294

with plasmid resistance. In Figure 1c-j, it is noteworthy that DNase I, expressed by the endA gene in

295

E. coli K-12, serves as an important extracellular barrier to hydrolyze exogenous plasmids. Thus, the

296

transformation of ARG to E. coli K-12 is lower than that to E. coli DH5α at the same concentration

297

gradient of substituted aromatic facilitators. Overall, these results further confirm that the formation

298

of substituted aromatic-plasmid complexes assists in the entry of these ARGs to the interior of cells

299

via the reinforcement of the anti-hydrolysis of pUC19, and consequently, substituted aromatic

300

compounds may serve as facilitators of ARG propagation.

301

Anti-Hydrolysis of pUC19 Complexes across the EPS Matrix. EPS covering the surface of

302

competent cells serves as an important permeable barrier for hydrolyzing plasmids,7, 43 weakening

303

ARG transformation. Thus, the stability of the pUC19 plasmid, enhanced by the formation of the

304

complexes, is associated with the mechanism by which substituted aromatic compounds facilitate

305

ARG dissemination. An experiment involving EPS manipulation (Figure 2a-b) showed that the

306

transformation of ARG to K-12 (Figure 2a) and DH5α (Figure 2b) was co-affected by the simulated

307

composite aromatic system (0.40 μmol/L of composite substituted aromatics including 25% 1,3DCB,

308

25% 2NAOL, 25% AN, and 25% 1,3MPD) and EPS. As for pristine E. coli K-12 (Figure 2a),

309

transformation of ARG was only 1.57 log transformants/μg pUC19. As we increased composite

310

aromatics to 0.40 μmol/L, the transformation increased to 1.95 log transformants/μg pUC19. After

311

the EPS was removed from the surface of E. coli K-12, transformation continued to move higher, to

312

approximately 2.55 log transformants/μg pUC19. Comparison of data from the three groups suggests

313

that the EPS covering the surface of E. coli K-12 serves as a permeable barrier to ARG proliferation.

314

Figure 2b shows a similar result. Transformation of ARG to E. coli DH5α was also enhanced

315

from 3.02 to 3.35 log transformants/μg pUC19, after pUC19 was exposed to the mixture of substituted 14


Page 15 of 34


316

aromatics (0.4 μmol/L). After the EPS was removed from surface of E. coli DH5α, the transformation

317

continuously increased to approximately 3.85 log transformants/μg pUC19. This result re-confirms

318

that ARG transformation was inhibited by the EPS permeable barrier on the surface of E. coli DH5α.

319

In the combination with transformation of ARG to E. coli K-12, these results underscore that

320

transformation of ARG to E. coli K-12 was co-regulated by EPS and composite aromatics.

321

Here, it is noteworthy that although E. coli DH5α does not contain the endA gene, transformation

322

of ARG was also enhanced after removal of EPS from the surface of cells (Figure 2b). This seems to

323

suggest that besides the DNase barrier in the extracellular matrix of E. coli DH5α, there is another

324

permeable system that obstructs ARG transfer into cells. Actually, in addition to endA and the

325

physical barrier, E. coli contains a suite of multiple-enzyme systems (EcoR I-V) that encode

326

extracellular restrictive endonucleases.8, 9 Therefore, this enhanced transformation in E. coli DH5α

327

by the removal of EPS might be attributed to the resistance of primary endonucleases encoded by

328

EcoR I-V, the second physical barrier of EPS to alien ARGs.22, 23 Additionally, transformation of

329

ARGs to E. coli DH5α was higher than that to E. coli K-12, which suggests that extracellular DNase

330

I, encoded by the endA gene of E. coli K-12, is important for impeding the entry of these ARGs to

331

the interior of cells. Overall, Figure 2a-b also verifies that the formation of substituted aromatic-

332

plasmid complexes is responsible for ARG proliferation, but the EPS permeable barrier covering the

333

surface of host cells inhibits this proliferation.

334

Figure 2c shows how the degradation efficiency of pUC19 was affected by the EPS matrix (I-I0,

335

0.4-μmol/L of each substituted aromatic compound, 25°C). To obtain statistically significant data

336

(shown in Figure 2d), 1,3DNB (1,3-dinitrobenzene) and 2NA (2-naphthylamine) were also used to

337

investigate the apparent degradation efficiency of substituted aromatic-pUC19 complexes mediated

338

by EPS matrix. According to the definition from Kunitz,36 the apparent degradation efficiency (I-I0) 15



Page 16 of 34

339

may be expressed as the difference in absorbance at 260 nm between before and after reaction of

340

plasmids with extracted EPS (30 min). Our results show that the absorbance of plasmids increased

341

by 0.115−0.127 (EPS extracted from E. coli K-12) and 0.107−0.130 (EPS extracted from E. coli

342

DH5α), which shows that secreted EPS hydrolyzes the plasmids to cause the increase in absorbance.

343

Meanwhile, it is noteworthy that in the presence of five substituted aromatic compounds (except

344

1,3DNB), the apparent degradation efficiencies, mediated by EPS extracted from E. coli K-12, were

345

much higher than those from EPS extracted from E. coli DH5α. This suggests that EPS, produced by

346

E. coli K-12 containing the endA gene, has a stronger capability of hydrolysis of exogenous pUC19.

347

Overall, the formation of aromatic-plasmid complexes reinforces the anti-hydrolysis ability of pUC19

348

in the extracellular matrix, and EPS obstructs the passage of exogenic ARGs into cells.

349

Figure 2d shows the correlation between the relative dissemination efficiency (T/T0, on the y

350

coordinate) and the apparent degradation efficiency (I-I0,on the x coordinate); there was a good

351

negative linear correlation between them (slop, K = -22.7; correlation coefficient, R2 = 0.98). As I-I0

352

increased, the T/T0 values for transformation of pUC19 to E. coli K-12 linearly decreased from

353

approximately 1.6 to 1.1. This shows that the formation of substituted aromatic-to-pUC19 complexes

354

went against the degradation of pUC19 mediated by EPS and that less degradation of pUC19 helped

355

the transformation of pUC19 to host cells. In addition, with the addition of I-I0 from approximately

356

0.117 to 0.127, the T/T0 values for transformation of pUC19 to E. coli DH5α also obeyed a linear

357

reduction from approximately 1.15 to 1.0 (slop = -15.1, R2 = 0.94), which re-confirms that

358

extracellular degradation of pUC19 mediated by EPS went against the transformation of exogenous

359

ARGs to host cells. Overall, the results from Figure 2a-d underscore that the association of pUC19

360

with substituted aromatic compounds facilitated the transformation efficiency of pUC19 to host cells

361

via a reinforced anti-hydrolysis mechanism occurring in the extracellular matrix. 16


Page 17 of 34


362

Formational Mechanism of Substituted Aromatic-pUC19 Complexes. The results in Figure

363

3 were used to study the binding mechanisms in the formation of the substituted aromatic compound-

364

pUC19 complexes, which is responsible for the structural stability of the pUC19 plasmid. After

365

reactions with substituted aromatic compounds, the changes in the valence electrons on the oxygen

366

and phosphorus atoms in pUC19 were analyzed using X-ray photoelectron spectroscopy (XPS). For

367

the pUC19 plasmid, the O 1s signal at 532.3 eV can be assigned to the phosphate, and the signal at

368

530.5 eV corresponds to the bases and/or D-ribose.44 After reactions with 1,3DCB, 2NAOL, AN, and

369

MPD, the signal at 532.3 eV redshifted by 0−0.4 eV (to 532.3, 532.6, 532.5, and 532.5 eV,

370

respectively). With the exception of 1,3DCB, the obvious redshifts caused by reactions with the

371

substituted aromatic compounds show that the phosphate group shares its surplus electrons with the

372

substituents on the aromatic compounds, which results in increased bond energies.

373

After the reactions with 1,3DCB, 2NAOL, AN, and 1,3MPD, the O 1s signal at 530.5 eV shifted

374

to 530.6, 530.7, 530.6, and 530.7 eV, respectively. Such shifts toward higher binding energies seem

375

to suggest that all substituents influence the changes in the valence electron structure of O 1s in bases

376

and/or D-ribose. However, considering the O 1s signal at 532.3 eV and the polarity of the molecules

377

and their functional groups, the changes in the 530.5 eV signal following the reactions with 2NAOL,

378

AN, and 1,3MPD can be attributed to their interactions with D-ribose-phosphate rather than their

379

interactions with the bases. The reaction with 1,3DCB induced only a slight shift to 530.6 eV (a

380

change of 0.1 eV), but the reaction did not change the signal at 532.3 eV, which shows that 1,3DCB

381

preferentially reacted with D-ribose rather than the phosphate groups of the plasmid. This result was

382

further supported by the computational data shown in Figure 4. Overall, Figure 3 shows that the

383

synergistic mechanisms involving D-ribose-phosphate are responsible for the facilitated ARG

384

dissemination by substituted aromatic compounds. 17



Page 18 of 34

385

The changes in the valence electrons (P 2p) on phosphorous in the pUC19 plasmid after the

386

reactions with the aromatic compounds were further analyzed (Figure 3b). The signal at 133.0 eV for

387

the pristine plasmid shifted toward lower binding energies of 132.7 eV for 1,3DCB-plasmid, 132.8

388

eV for 2NAOL-plasmid, 132.7 eV for AN-plasmid, and 132.7 eV for 1,3MPD-plasmid. Except for

389

1,3DCB, these shifts show that surplus electron density on the phosphate group shifted to between

390

the oxygen and the aromatic compound because of the formation of a new bond between the oxygen

391

in the phosphate group and the aromatic compound. The reaction with 1,3DCB did not cause an

392

obvious shift in the P 2p signal at 133.0 eV, which further suggests that D-ribose was mainly

393

responsible for the 1,3DCB-plasmid complex. Including the other tested substituted aromatics,

394

changes in functionality of pUC19 after the reaction with substituted aromatics were also observed

395

by FTIR (Figure S2), which further confirms that D-ribose-phosphate was responsible for the

396

formation of these complexes (FTIR analysis of aromatic-plasmid complexes of the Supporting

397

Information). Overall, the data shown in Figure 3 verify that the D-ribose-phosphate moiety of the

398

pUC19 plasmid was crucial to the interaction between the plasmid and all the aromatic compounds

399

and thus facilitated ARG dissemination via a reinforced anti-hydrolysis to DNase I in the EPS matrix.

400

On the basis of XPS (Figure 3) and FTIR analyses (Figure S2), we analyzed the electron

401

localization to further elucidate the binding mechanisms of the substituted aromatic compound-

402

plasmid complex formation at the key D-ribose-phosphate. To better understand the bonding

403

mechanisms, the strongest bond was the focus of further investigation using an electron localization

404

function (ELF) isosurface plot if there were multiple bonds in the aromatic compound-plasmid

405

complex. First, the 1,3DCB-plasmid complex (Figure 4a) showed that the carbon at the 6-position of

406

1,3DCB bound to the oxygen in D-ribose via a C-H···O bond (bond length = 1.8 Å). The high electron

407

density on the oxygen shifted toward the hydrogen atom of D-ribose of the plasmid (Figure 4b), which 18


Page 19 of 34


408

confirms that hydrogen bonding was responsible for the formation of the 1,3DCB-plasmid complex.

409

2NAOL, AN, and 1,3MPD combined with the pUC19 plasmid via at least two hydrogen bonds

410

to their substituents. The H on the carbon at the 2-position of 2NAOL bound to one of the two oxygens

411

of the phosphate group (C-H···OP) to form a 2.0-Å hydrogen bond (Figure 4c). The OH in 2NAOL

412

bound with another O of the phosphate group (C-O···H···OP) via a stronger 1.5-Å hydrogen bond.

413

Two hydrogens from 2NAOL were shared by two oxygens from a phosphate group. The stronger

414

hydrogen bond was further analyzed through an ELF isosurface plot (Figure 4d). The proton of the

415

hydroxy group of 2NAOL was more likely to bind with the phosphate group through a coherent

416

electron cloud; the electron on the surface of the oxygen of 2NAOL was transferred to the hydrogen.

417

These observations suggest that the oxygen of the phosphate group had a stronger electron-donating

418

ability than that of the deprotonated hydroxy moiety in 2NAOL. These results show that although

419

both the deprotonated oxygen of the phosphate group and the hydroxy group of 2NAOL were

420

electron-rich, they attracted each other through an apparent hydrogen bond.

421

Two hydrogens of the amino group of an AN molecule bound to the D-ribose-phosphate

422

backbone (Figure 4e). One of the hydrogens bound to an O atom of the phosphate group via a strong

423

1.1-Å hydrogen bond. Another hydrogen is bound to the O atom of a D-ribose in the plasmid via a

424

2.1-Å hydrogen bond. Figure 4f shows further analysis of the binding mechanism of the strong 1.1-

425

Å hydrogen bond based on the ELF isosurface data. The hydrogen atom shifted from the N atom of

426

the amino group to an O atom of the phosphate. More importantly, the region of the O with the highest

427

electron density nearly overlapped with the proton. These results suggest that a strong hydrogen bond

428

was responsible for the formation of the plasmid-AN complex. Furthermore, because 1,3MPD

429

possessed the same amino substituent, it exhibited a similar bonding pattern (Figure 4g). Like the

430

attraction between AN and D-ribose, two oxygens of the phosphate bound to a pair of hydrogens in 19



Page 20 of 34

431

the amino group of 1,3MPD via 1.9- and 2.1-Å bonds. Figure 4h shows the further analysis of this

432

pair of bonds and shows that there was a slight deformation in the electron cloud toward the protons

433

of 1,3MPD. Amino-substituted aromatic compounds mainly bound to the plasmid through [N-

434

(H···O)2-P] interactions. Overall, results from Figure 4, supported also by Figure 3, verify that the D-

435

ribose-phosphate site in pUC19 was responsible for their combinations with substituted aromatic

436

compounds.

437

Effect of the Bonding Energy on ARG Propagation. Based on the analyses of the major

438

bonding sites shown in Figure 4, we further evaluated the bonding energies of the aromatic

439

compounds to the D-ribose-phosphate. As shown in Figure 5a, the x-axis represents the association

440

constants (logKA), which were determined by fluorescence microtitration with the Tb3+-labeled

441

plasmid (Figure 1a-b); the y-axis is the bonding energy of substituted aromatic compounds toward

442

the D-ribose-phosphate backbone of the plasmid. Except for the 1,3DCB-plasmid complex, there was

443

a good linear correlation between ΔG and logKA. The linear slope (k = -9.0) shows that there was a

444

negative correlation between these parameters, suggesting that the bonds to the phosphate site

445

dominated the substituted aromatic compound-plasmid complexes. However, the 1,3DCB-plasmid

446

complex did not follow this linear relationship. Based on the data shown in Figure 4, the bonds in the

447

1,3DCB-plasmid complex were mainly to the D-ribose rather than to the phosphate (the other

448

aromatic compounds tested in this study were mainly bound to the phosphate). Thus, although

449

1,3DCB also facilitated the genetic transformation (Figure 1), the slight difference in binding site

450

caused its departure from the linear regression. This inconsistency in the bonding sites, which was

451

caused by the differences in the substituents, was supported by the XPS (Figure 3) and computational

452

analyses (Figure 4). In addition, except for that of 1,3DCB, the bonding energies of the substituted

453

aromatic compounds at the phosphate gradually increased from about -100 to -150 kJ/mol (the 20


Page 21 of 34


454

negative sign indicates the process is spontaneous, and the absolute value represents the energy) as

455

the logKA increased from 3.4 to 8.2, which suggests that functionalities in aromatic hydrocarbons had

456

different affinities to phosphate sites in pUC19 when the complexes formed in a reactive system.

457

Figure 5b further shows the relationship between the relative ARG dissemination efficiency

458

(T/T0) and the logKA. Two good linear correlations were observed between the T/T0 and the logKA

459

(EPS of E. coli K-12: slope =0.03; EPS of E. coli DH5α: slope = 0.09), suggesting the logKA to the

460

D-ribose-phosphate moieties was responsible for the ARG propagation seen with substituted aromatic

461

pollutants. Meanwhile, it is noteworthy that the slope of T/T0 vs logKA for E. coli DH5α was obviously

462

greater than that for E. coli K-12. This shows that when the endA gene was removed, the substituted

463

aromatic compounds had greater effects on ARG dissemination to E. coli DH5α. Overall, the results

464

from Figure 5 demonstrate that this combination at D-ribose-phosphate can reinforce the anti-

465

hydrolysis of pUC19 in the extracellular matrix, facilitating ARG dissemination.

466

21



Page 22 of 34

467

ASSOCIATED CONTENT

468

Supporting Information

469

Supporting Information Available: Sequence of endA gene, and FTIR analysis of aromatic-plasmid

470

complexes (Materials and Methods of the Supporting Information); detailed flowchart of substituted

471

aromatic-facilitated dissemination of mobile ARG (Figure S1); FTIR analyses of functional groups

472

in plasmid pUC19 (Figure S2); physicochemical properties of selected organic compounds (Table

473

S1), the fitting parameters of the interactions between substituted aromatic compounds and the Tb3+-

474

labeled plasmids (Table S2).

475

AUTHOR INFORMATION

476

Corresponding Author

477

*Telephone/Fax: +86-25-8439-5860; e-mail: [email protected]/[email protected].

478

Notes

479

The authors declare no competing financial interests.

480

ACKNOWLEDGMENTS

481

We wish to thank Dr. Tian Lu for his discussions of the ELF analysis. This work was supported by

482

the National Science Foundation of China (Grant Nos. 21777071, 41502170, and 21777071), the

483

Fundamental Research Funds for the Central Universities (Grant No. KYZ201870), the National

484

Science Foundation for Postdoctoral Scientists of China (Grant No. 2014M561662), and the National

485

Undergraduate Innovative Test Program (201810307003).

22


Page 23 of 34


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References

488

1.

489

contaminants: studies in northern Colorado. Environ. Sci. Technol. 2006, 40, (23), 7445–7450.

490

2.

491

Tiedje, J. M. Diverse and abundant antibiotic resistance genes in Chinese swine farms. Proc. Natl.

492

Acad. Sci. USA 2013, 110, (9), 3435–3440.

493

3.

494

to ZnO nanoparticles facilitates horizontal transfer of antibiotic resistance genes. NanoImpact 2018,

495

10, 61-67.

496

4.

497

Nanoalumina promotes the horizontal transfer of multiresistance genes mediated by plasmids across

498

genera. Proc. Natl. Acad. Sci. USA 2012, 109, (13), 4944-4949.

499

5.

500

is enhanced by ionic liquid with different structure of varying alkyl chain length. Front. Microbiol.

501

2015, 6.

502

6.

503

relationship detected between soil bioaccessible organic pollutants and antibiotic resistance genes at

504

dairy farms in Nanjing, Eastern China. Environ. Pollut. 2015, 206, 421-428.

505

7.

506

efficiency of plasmid DNA exposed to PAH contaminants. PloS One 2013, 8, (3), 134–143.

507

8.

508

New York and London, 2012.

509

9.

Pruden, A.; Pei, R.; Storteboom, H.; Carlson, K. H. Antibiotic resistance genes as emerging

Zhu, Y. G.; Johnson, T. A.; Su, J. Q.; Qiao, M.; Guo, G. X.; Stedtfeld, R. D.; Hashsham, S. A.;

Wang, X.; Yang, F.; Zhao, J.; Xu, Y.; Mao, D.; Zhu, X.; Luo, Y.; Alvarez, P. Bacterial exposure

Qiu, Z.; Yu, Y.; Chen, Z.; Jin, M.; Yang, D.; Zhao, Z.; Wang, J.; Shen, Z.; Wang, X.; Qian, D.

Wang, Q.; Lu, Q.; Mao, D.; Cui, Y.; Luo, Y. The horizontal transfer of antibiotic resistance genes

Sun, M.; Ye, M.; Wu, J.; Feng, Y.; Wan, J.; Tian, D.; Shen, F.; Liu, K.; Hu, F.; Li, H. Positive

Kang, F.; Wang, H.; Gao, Y.; Long, J.; Wang, Q. Ca2+ promoted the low transformation

Chaloupka, J., Extracellular enzymes of microorganisms. Springer Science & Business Media:

Felmlee, T.; Pellett, S.; Welch, R. A. Nucleotide sequence of an Escherichia coli chromosomal 23



Page 24 of 34

510

hemolysin. J. Bacteriol. 1985, 163, (1), 94–105.

511

10. Choi, J.; Lee, S. Secretory and extracellular production of recombinant proteins using

512

Escherichia coli. Appl. Microbiol. Biot. 2004, 64, (5), 625–635.

513

11. Ming Chung, C.; Shwu Yuh, C.; Shing Li, C.; Show Mei, C. Cloning and expression in

514

Escherichia coli of the gene encoding an extracellular deoxyribonuclease (DNase) from Aeromonas

515

hydrophila. Gene 1992, 122, (1), 175–180.

516

12. Yen, S. F.; Gabbay, E. J.; Wilson, W. D. Interaction of aromatic imides with DNA. 1.

517

Spectrophotometric and viscometric studies. Biochemistry 1982, 21, (9), 2070–2076.

518

13. Nagata, C.; Kodama, M.; Tagashira, Y.; Imamura, A. Interaction of polynuclear aromatic

519

hydrocarbons, 4-nitroquinoline 1-oxides, and various dyes with DNA. Biopolymers 1966, 4, (4), 409-

520

427.

521

14. Palek, E.; Fojta, M. Peer reviewed: detecting DNA hybridization and damage. Anal. Chem. 2001,

522

74A–83A.

523

15. Wolfe, A.; Shimer Jr, G. H.; Meehan, T. Polycyclic aromatic hydrocarbons physically intercalate

524

into duplex regions of denatured DNA. Biochemistry 1987, 26, (20), 6392-6396.

525

16. Kang, F.; Hu, X.; Liu, J.; Gao, Y. Noncovalent binding of polycyclic aromatic hydrocarbons with

526

genetic bases reducing the in-vitro lateral transfer of antibiotic resistant genes. Environ. Sci. Technol.

527

2015, 49, (17), 10340–10348.

528

17. Kumar, C.; Asuncion, E. H. DNA binding studies and site selective fluorescence sensitization of

529

an anthryl probe. J Am. Chem. Soc. 1993, 115, (19), 8547–8553.

530

18. Shou, W.; Kang, F.; Lu, J. Nature and value of freely dissolved EPS ecosystem services: Insight

531

into molecular coupling mechanisms for regulating metal toxicity. Environ. Sci. Technol. 2018, 52,

532

(2), 457–466. 24


Page 25 of 34


533

19. Kunitz, M. A spectrophotometric method for the measurement of ribonuclease activity. J. biol.

534

Chem 1946, 164, 563–568.

535

20. Hu, X.; Kang, F.; Gao, Y.; Zhou, Q. Bacterial diversity losses: A potential extracellular driving

536

mechanism involving the molecular ecological function of hydrophobic polycyclic aromatic

537

hydrocarbons. Biotechnology Reports 2015, 5, 27-30.

538

21. Salt, D. E.; Prince, R. C.; Baker, A. J.; Raskin, I.; Pickering, I. J. Zinc ligands in the metal

539

hyperaccumulator Thlaspi caerulescens as determined using X-ray absorption spectroscopy. Environ.

540

Sci. Technol. 1999, 33, (5), 713–717.

541

22. Smart, R. C.; Hodgson, E., Molecular and biochemical toxicology. John Wiley & Sons: Hoboken:

542

New Jersey, 2018.

543

23. Kuntz, I. D. Structure-based strategies for drug design and discovery. Science 1992, 257, (5073),

544

1078–1082.

545

24. Sirajuddin, M.; Ali, S.; Badshah, A. Drug–DNA interactions and their study by UV–Visible,

546

fluorescence spectroscopies and cyclic voltametry. J Photoch. Photobio. B 2013, 124, 1-19.

547

25. Li, F.; Li, X.; Liu, X.; Zhang, L.; You, L.; Zhao, J.; Wu, H. Noncovalent interactions between

548

hydroxylated polycyclic aromatic hydrocarbon and DNA: Molecular docking and QSAR study.

549

Environ. Toxicol. Pharm. 2011, 32, (3), 373–381.

550

26. Durfee, T.; Nelson, R.; Baldwin, S.; Plunkett, G.; Burland, V.; Mau, B.; Petrosino, J. F.; Qin, X.;

551

Muzny, D. M.; Ayele, M.; Gibbs, R. A.; Csorgo, B.; Posfai, G.; Weinstock, G. M.; Blattner, F. R.

552

The complete genome sequence of Escherichia coli DH10B: insights into the biology of a laboratory

553

workhorse. J. Bacteriol. 2008, 190, (7), 2597–606.

554

27. Surette, M. G.; Miller, M. B.; Bassler, B. L. Quorum sensing in Escherichia coli, Salmonella

555

typhimurium, and Vibrio harveyi: a new family of genes responsible for autoinducer production. Proc. 25



Page 26 of 34

556

Natl. Acad. Sci. USA 1999, 96, (4), 1639–1644.

557

28. Yonuschot, G.; Mushrush, G. W. Terbium as a fluorescent probe for DNA and chromatin.

558

Biochemistry 1975, 14, (8), 1677–1681.

559

29. Boaz, H.; Rollefson, G. The quenching of fluorescence: Deviations from the Stern-Volmer law.

560

J Am. Chem. Soc. 1950, 72, (8), 3435–3443.

561

30. Eftink, M. R. Fluorescence methods for studying equilibrium macromolecule-ligand interactions.

562

Method. Enzymol. 1997, 278, 221–257.

563

31. Lakowicz, J. R.; Weber, G. Quenching of fluorescence by oxygen: Probe for structural

564

fluctuations in macromolecules. Biochemistry 1973, 12, (21), 4161–4170.

565

32. Valeur, B.; Berberan-Santos, M. N., Molecular fluorescence: principles and applications. John

566

Wiley & Sons: 2012.

567

33. Bi, S.; Ding, L.; Tian, Y.; Song, D.; Zhou, X.; Liu, X.; Zhang, H. Investigation of the interaction

568

between flavonoids and human serum albumin. J Mol. Struct. 2004, 703, (1), 37–45.

569

34. Kang, F.; Qu, X.; Alvarez, P. J.; Zhu, D. Extracellular saccharide-mediated reduction of Au3+ to

570

gold nanoparticles: New insights for heavy metals biomineralization on microbial surfaces. Environ.

571

Sci. Technol. 2017, 51, (5), 2776−2785.

572

35. Kang, F.; Alvarez, P. J.; Zhu, D. Microbial extracellular polymeric substances reduce Ag+ to

573

silver nanoparticles and antagonize bactericidal activity. Environ. Sci. Technol. 2013, 48, (1), 316–

574

322.

575

36. Kunitz, M. A spectrophotometric method for the measurement of ribonuclease activity. J Biol.

576

Chem. 1946, 164, 563–568.

577

37. Dennington, R.; Keith, T.; Millam, J.; Eppinnett, K.; Hovell, W. L.; Gilliland, R. GaussView,

578

version 5. Semichem Inc., Shawnee Mission, KS 2009. 26


Page 27 of 34


579

38. Trucks, G.; Schlegel, H.; Scuseria, G.; Robb, M.; Cheeseman, J.; Scalmani, G.; Barone, V.;

580

Mennucci, B.; Petersson, G.; Nakatsuji, H. Gaussian 09, revision E. 01; Gaussian, Inc. Wallingford,

581

CT 2009.

582

39. Savin, A.; Nesper, R.; Wengert, S.; Fässler, T. F. ELF: The electron localization function.

583

Angewandte Chemie International Edition 1997, 36, (17), 1808-1832.

584

40. Lu, T.; Chen, F. Multiwfn: A multifunctional wavefunction analyzer. J Comput. Chem. 2012, 33,

585

(5), 580–592.

586

41. Dieke, G.; Hall, L. Fluorescent lifetimes of rare earth salts and ruby. J Chem. Phys. 1957, 27, (2),

587

465–467.

588

42. Lakowicz, J., Principles of frequency-domain fluorescence spectroscopy and applications to cell

589

membranes. In Fluorescence Studies on Biological Membranes, Springer: 1988; pp 89–126.

590

43. Mah, T.-F.; Pitts, B.; Pellock, B.; Walker, G. C.; Stewart, P. S.; O'toole, G. A. A genetic basis

591

for Pseudomonas aeruginosa biofilm antibiotic resistance. Nature 2003, 426, (6964), 306–310.

592

44. Vilar, M. R.; Botelho do Rego, A. M.; Ferraria, A. M.; Jugnet, Y.; Nogues, C.; Peled, D.; Naaman,

593

R. Interaction of self-assembled monolayers of DNA with electrons: HREELS and XPS studies. J.

594

Phys. Chem. B 2008, 112, (23), 6957–6964.

595 596

27



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597 598

Figure 1. Effect of aromatic-plasmid associations on ARG propagation. Figure 1a-b present the

599

interactions between substituted aromatic compounds and the Tb3+-labeled plasmid. (a) Stern-Volmer

600

plot; (b) Plot of log [(F0- F)/F] vs log[Q]. Figure 1c-j are the transformation of ARG to E. coli K-12

601

and E. coli DH5α on the basis of the quadruplicates. (×): maximum and minimum values; (□): median;

602

(―): mean. The blue box represents the range of the data set. Transformations of ARG to E. coli K-

603

12 and E. coli DH5α in the presence of four substituted aromatics present a statistical significance (p

604

< 0.01), comparing to those in control groups.

28


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605 606

Figure 2. Influence of EPS permeable barrier on ARG propagation. (a) transformation of ARG to E.

607

coli K-12 co-regulated by EPS and composite aromatics; (b) transformation of ARG to E. coli DH5α

608

co-regulated by substituted aromatics and EPS (In the Figure 2a and b, 0.40 μmol/L of composite

609

substituted aromatics including 25% of 1,3DCB, 25% of 2NAOL, 25% of AN, and 25% of 1,3MPD).

610

(c) apparent degradation efficiency (I-I0) of plasmid pUC19 by extracted EPS (23.8 mg DW/L) in the

611

presence of 0.4-μmol/L of each substituted aromatic compound at 25℃; (d) correlation between I-I0

612

and relative ARG dissemination efficiency (T/T0, 0.4-μmol/L of each substituted aromatic compound,

613

25℃). Linear correlation between T/T0 and I-I0 is fitted in Figure 2d (E. coli K12: slop = -22.7, R2 =

614

0.98; E. coli DH5α: slop = -15.1, R2 = 0.94). Increase in the transformation efficiency of ARG to E.

615

coli K-12 and DH5α with the presence of substituted aromatic compounds/with EPS removal was

616

statistically significant (p < 0.01). In the Figure a-b, (×): maximum and minimum values; (□): median; 29



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617

(―): mean. The black box represents the range of the data set. Averages of triplicates are shown with

618

bidirectional error bars to represent standard deviations in Figure 2c and d.

619 620

30


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621

622 623

Figure 3. XPS of O 1s (a) and P 2p (b) of the plasmid pUC19 before and after reaction with 1,3DCB,

624

2NAOL, AN, and 1,3MPD.

625

31



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626 627

Figure 4. Bond lengths and electron localization function (ELF) analyses between substituted

628

aromatic compounds and plasmid pUC19 (green sphere: Cl; white sphere: hydrogen; red sphere:

629

oxygen; blue sphere: nitrogen; orange sphere: phosphorus; gray sphere: carbon). In the ELF

630

isosurface plots, blue lines represent the isosurface of the van der Waals (vdWs) forces defined by

631

the Bader theory (electron density = 0.001 isosurface). a-b: 1,3DCB-plasmid complex; c-d: 2NAOL-

632

plasmid complex; e-f: AN-plasmid complex; g-h: 1,3MPD-plasmid complex.

32


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633

634 635

Figure 5. Association among genetic transfer efficiency (T/T0), logKA, and bonding energy at the key

636

plasmid backbone. (a): ΔG at the D-ribose-phosphate backbone vs logKA; (b): logKA vs T/T0. The

637

Gibbs free energy (G) was computed at a the B3LYP/6-31G* level using DFT-D3 for the dispersion

638

correction.

639

𝐺substituted aromatic‧‧‧‧𝐷 ― ribose ― phosphate ―(𝐺substituted aromatic + 𝐺𝐷 ― ribose ― phosphate).

640

𝐺substituted aromatic‧‧‧‧𝐷 ― ribose ― phosphate, 𝐺𝐷 ― ribose ― phosphate, and 𝐺substituted aromatic are the G

641

values of the “substituted aromatic‧‧‧‧D-ribose-phosphate” complex (new bond formed between

642

plasmid and the substituted aromatic compounds), D-ribose-phosphate, and substituted aromatic

643

compound, respectively. Averages of triplicates are shown with bidirectional error bars to represent

644

standard deviations.

The

change

in

the

ΔG

was

calculated

645 646

33


by

the

equation

△𝐺= Here,


647

Graphic abstract:

648 649 650 651 652 653

34


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Substituted Aromatic-Facilitated Dissemination of Mobile Antibiotic

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