Selection of Group-Specific PAE-Binding DNA Aptamers via Rational

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Selection of Group-Specific PAE-Binding DNA Aptamers via Rational Designed Target Immobilization and Applications for Ultrasensitive and Highly Selective Detection of PAEs Yu Han, Donglin Diao, Zhangwei Lu, Xiaoning Li, Qian Guo, Yumeng Huo, Qing Xu, Youshan Li, ShengLi Cao, Jianchun Wang, Yuan Wang, Jiaxing Zhao, Zhongfeng Li, Miao He, Zhaofeng Luo, and Xinhui Lou Anal. Chem., Just Accepted Manuscript • DOI: 10.1021/acs.analchem.6b04808 • Publication Date (Web): 17 Apr 2017 Downloaded from http://pubs.acs.org on April 17, 2017

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Analytical Chemistry

Selection of Group-Specific PAE-Binding DNA Aptamers via Rational Designed Target Immobilization and Applications for Ultrasensitive and Highly Selective Detection of PAEs Yu Han1,†, Donglin Diao1,†, Zhangwei Lu1,†, Xiaoning Li1, Qian Guo1, Yumeng Huo1, Qing Xu1,Youshan Li1, Shengli Cao1, Jianchun Wang1, Yuan Wang1, Jiaxing Zhao1, Zhongfeng Li1, Miao He2,*, Zhaofeng Luo3, and Xinhui Lou1,*

1

Department of Chemistry, Capital Normal University, Beijing, 100048, China

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School of Environment, Tsinghua University, Beijing 100084, China.

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University of Science and Technology of China, School of Life Sciences, Hefei 230027, China

* To whom correspondence should be addressed. Tel: +86-10-68902491 ext. 808; Fax: +86-10-68902320; Email: [email protected]; [email protected]

†

Yu Han, Donglin Diao, and Zhangwei Lu contributed equally to this work and should be regarded as

joint First Authors.

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ACS Paragon Plus Environment


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ABSTRACT: Phthalic acid esters（PAEs) are ubiquitous in the environment and some

of them are recognized as endocrine disruptors that cause concerns on ecosystem functioning and public health. Due to the diversity of PAEs in the environment, there is a vital need to detect the total concentration of PAEs in a timely and low-cost way. To fulfill this requirement, it is highly desired to obtain group-specific PAE binders that are specific to the basic PAE skeleton. In this study, for the first time we have identified the group-specific PAE-binding aptamers via rational designed target immobilization. The two target immobilization strategies were adopted to respectively display either the phthalic ester group or the alkyl chain at the surface of the immobilization matrix. The former enabled the rapid enrichment of aptamers after 4 rounds of selection. The top 100 sequences are cytosine-rich (44.7%) and differentiate from each other by only 1−4 nucleotides at limited locations. The top two aptamers all display the nanomolar dissociation constants to both the immobilized target and the free PAEs (DBP, BBP, DEHP). We further demonstrate the applications of the aptamers in the development of high-throughput PAE assays and DEHP electrochemical biosensors with exceptional sensitivity (LOD: 10 pM) and selectivity (> 105-fold). PAE aptamers targeting one of the most sought for targets thus offer the promise of convenient, low-cost detection of total PAEs. Our study also provides insights on the aptamer selection and sensor development of highly hydrophobic small molecules.

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INTRODUCTION Phthalic acid esters（PAEs）are widely used as plasticizers to impart flexibility of plastic products since 1920s.1 To date, PAEs are the most common chemicals that humans are in contact with daily due to their very broad range of applications in building materials, food packaging materials, personal-care products, and so on. PAEs can easily leak into the environment because they are physically, not covalently, mixed with the matrices. Even though PAEs can be eliminated from different environmental matrices via biotic and abiotic pathways, their extensive use and permanent emissions have leaded to their ubiquitous presence in air,2, 3 drinking water, food,2, 4, 5 marine water,6 sediments,7 soil,8 and sludge

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on a worldwide. PAEs come

into human body through breathing, eating, and skin contact via various exposure mechanisms.1, 5, 10, 11 Some PAEs are involved in endocrine disrupting effects that cause reproductive system problems and cancers.10, 12, 13 PAEs present a variety of toxic effects for many other species including terrestrial and aquatic fauna and flora as well. Therefore, their potential and adverse impacts on ecosystem functioning and on public health have aroused considerable and growing attention in recent years. Six PAEs including bis(2-ethylhexyl) phthalate (DEHP), dibutyl phthalate (DBP), and butyl benzyl phthalate (BBP) have been placed on the priority pollutant list of the United States Environmental Protection Agency (U.S. EPA), European Union (EU), and China.1 The WHO recommends the concentration of DEHP in drinking water below 8 µg/L (20.5 nM)1 that is among the lowest concentration limits of pollutants including heavy metal ions, insecticides, and polycyclic aromatic hydrocarbons 3



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(PAHs). The fast and sensitive detection methods are highly desired for PAEs due to their toxicity, pervasiveness, and low concentration limits in the environment and food. The determination of PAEs is largely based on chromatographic-based methods including HPLC,14-17 GC,18 HPLC–MS19 and GC–MS

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. These methods are

sensitive, accurate, and capable for the simultaneous detection of multiple congeners in one run, but have some limitations including high-costs, long detection time, and the need of trained personnel. To solve these problems, the enzyme-linked immunosorbent assays (ELISAs) that use antibodies as recognition elements have recently been developed for PAE detection.22-30 The immunoassays are all highly selective for a specific PAE congener, with low cross-reactivity values with other congeners. However, PAEs consist of a variety of congeners (more than 20) and it is hard to predict the possible presence of certain congeners.1 It is not cost-effective to run multiple ELISAs, let alone ELISAs are only available for several PAE congeners. PAE group-specific binders that can recognize all PAE congeners would certainly be attractive for the efficient and low-cost screening PAEs in environment. Aptamers are single

or double stranded DNAs or RNAs obtained by an in vitro

technique called SELEX (systematic evolution of ligands by exponential enrichment).31, 32 Aptamer can specifically recognize a variety of targets including proteins,33-35 small molecules,36, 37 tissues,38 and cells39, 40. Compared to antibodies, aptamers are advantageous in cost, stability, easy synthesis and modification. Especially, antibodies are generated via the in vivo immuno-reaction of animals, while 4


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aptamers are selected via SELEX, an affinity enrichment-based in vitro technique. SELEX can provide the flexible control on the specificity (compound specific or group specific) of the selected binders by the rational design of the targets used for aptamer selection. In recent years, many aptamers for a variety of environmental pollutants have been identified and used for biosensors.41-44 However, the selection of aptamers for highly hydrophobic small molecule pollutants, such as persistent organic pollutants (POPs), is still quite challenging and rare reported. The most toxic POPs include polychlorinated biphenyls (PCBs), PAHs, polybrominated biphenyl ethers (PBDEs), polychlorinated dibenzo-p-dioxin (PCDDs), polychlorinated dibenzofuran (PCDFs), and PAEs, which all consist of many congeners. So far, only PCB-binding DNA aptamers have been identified.45, 46 Neither the compound- nor group-specific PAE-binding aptamers have been reported. In this study, for the first time, we demonstrate the identification of group-specific PAE-binding DNA aptamers through the integration of the rational design of the initial targets with high-throughput sequencing technology (Figure 1). The aptamers possess the highly conserved cytosine-rich sequences and show the general specificity for the basic PAE skeleton. We further demonstrate the applications of the PAE-binding aptamers in the development of high-throughput assays and electrochemical biosensors for the ultrasensitive and highly specific detection of PAEs.

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Figure 1. Aptamer selection procedure.

EXPERIMENTAL SECTION SELEX. The single-stranded DNA (ssDNA) random library (Pool0, Table S-1) was commercially synthesized. Each ssDNA molecule contains 40-nt randomized nucleotides flanked by two 20-nt primer-binding sequences. Ten microliters of 100 µM Pool0 was diluted in 490 µL of PAE binding buffer (20 mM Tris·HCl, 100 mM NaCl，2 mM MgCl2, 5 mM KCl, 1 mM CaCl2, 1% Tween 20, 0.03% triton X-100, 2% DMSO, pH 7.9), heated at 95°C for 10 min, quickly cooled down to 0°C on ice, and subsequently incubated for 5 min at room temperature. The experimental details on the synthesis of DBP-NH2 and its immobilization onto Epoxy-Activated Sepharose™ 6B are provided in the Supporting Information (SI). The 200 µL of DBP-NH2-coated medium was washed three times by PAE binding buffer and then mixed with Pool0, followed by incubation at room temperature for 1 hr under mild shaking. The DBP-NH2-coated medium bound with aptamers was then separated from the unbound DNAs by ultrafiltration using an ultrafiltration tube with a molecular cut-off of 100 6


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kDa, followed by the three times washes with 500 µL of PAE binding buffer and ultrafiltration at 10000 rpm for 10 min. The 50 µL PAE binding buffer was added to the washed medium and the mixture was heated at 90°C for 9 min under shaking. After that, the supernatant containing the eluted DNAs were collected by ultrafiltration. This elution process was repeated three times in order to recover more bound DNAs. The enriched DNAs were then amplified via PCR using forward and PO4-labeled reverse primers (Table S-1). The total volume of PCR was 2 mL. The PCR cycling conditions consist of an initial 30 s preheating at 95 °C, followed by 20 amplification cycles of denaturation at 95 °C for 30 s, annealing at 56 °C for 30 s, and extension at 72 °C for 30 s. The PCR product was purified by ethanol precipitation. The pool1 was finally generated by performing the λ exonuclease reaction to digest the phosphorylated strand and purification by ethanol precipitation. The second, third, and forth round of SELEX were subsequently conducted following the same procedure described above except that the ～300 pmol library was input to increase the selection stringency. To minimize the nonspecific absorption of DNAs on the medium, the pool was incubated with unmodified medium on a rotary shaker for 30 min in the third and fourth round of selection prior to the incubation with the DBP-NH2-coated medium. High-Throughput Sequencing. To prepare samples for sequencing, the ssDNAs obtained in the fourth round were amplified via PCR using the same PCR cycling conditions and the reverse primer as used in the SELEX process. An indexed forward primer with 6 nt at the 5' end (FP-Sequencing, Table S-1) was used to facilitate the 7



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sequence analysis. The PCR product was separated by a 12% PAGE gel and the band containing the right PCR product (86 bp) was cut, followed by the DNA purification using the UNIQ-10 PAGE-gel DNA Recovery Kit (Sangon Biotech (Shanghai) Co., Ltd. Shanghai, China). The purified PCR product (∼2 µg) was sent to Novogene (Beijing, China) for the high-throughput sequencing using Illumina sequencing technology.47 Imaging of DBP-NH2 Coated Medium Bound with Aptamers Using Inverted System Microscope. The primer-trimmed FAM-labeled aptamers (1 µM each, DBP-1-FAM and DBP-2-FAM, Table S-1) were respectively incubated with DBP-NH2-coated and bare medium at room temperature for 1 hr in 200 µL of PAE binding buffer, and then centrifuged to discard the unbound aptamers in supernatant. The DBP-NH2-coated and bare medium–ssDNA complex sediment were washed with PAE binding buffer three times, resuspended in PAE binding buffer and dropped on a glass slide. Imaging of the samples was then carried out. Determination of Dissociation Constants (Kds). DBP-NH2 was covalently immobilized on DynabeadsTM M-270 Carboxylic Acid magnetic beads and the experimental detail was described in the SI. The Kds of DBP-1 and DBP-2 were then respectively determined using quantitative PCR (qPCR)-based binding assays, where the amount of aptamers bound with DBP-NH2 coated Dynabeads was determined by qPCR. Primer-binding sequences were respectively added at the two terminals of DBP-1 and DBP-2, respectively, to enable the PCR (DBP-1-qPCR and DBP-2-qPCR, Table S-1). DBP-1-qPCR or DBP-2-qPCR at various concentrations (0–300 nM) were 8


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heated at 95°C for 10 min in 100 µL of PAE binding buffer, rapidly cooled down in an ice bath and then incubated for 10 min at room temperature. The 10 µL DBP-NH2-coated beads were washed 3 times in PAE binding buffer and incubated with the aptamers at room temperature for 30 min. After that, the unbound aptamers were removed by washing 4 times with 200 µL PAE binding buffer. The bound aptamers were then eluted from the beads by heating at 90°C for 10 min in 100 µL PAE binding buffer, followed by the magnetic separation. The amount of bead-bound aptamers was then determined by qPCR and the Kd was calculated by nonlinear fitting analysis. Relative Affinity and Selectivity Tests of Aptamers via Competition Assays. DBP-1-qPCR or DBP-2-qPCR (5 µL, 100 µM) and DBP-NH2-coated Dynabeads (10 µL) mixed in 500 µL PAE binding buffer and incubated for 1 hr at room temperature. The bound and unbound aptamers were then separated by magnetic separation and the Dynalbeads were washed three times using 100 µL PAE binding buffer. The aptamer-bound Dynabeads (10 µL) were then added to the 120 µL solution containing each tested molecule at 10 µM. The tested molecules included DBP-NH2, DBP, DEHP, BBP, a mixture of potential interferences (glucose, kanamycin, ampicillin, and ethanol), phthalic acid, benzoic acid, and ethyl acetate. The reference assay contained only PAE binding buffer. Each mixture was incubated with 1 hr and the supernatant was collected. The amount of aptamers in the supernatant was determined via qPCR. DEHP Electrochemical Biosensors. The biosensors were designed and fabricated according to our previously reported signaling probe displaced electrochemical 9



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biosensors (SD-EAB).41 The details on biosensor fabrication were described in the SI. To collect the titration curve of DEHP, the same electrode was incubated with the 100 µL PEA binding buffer containing DEHP at sequentially increased concentrations. After each 30 min’s incubation at room temperature to reach the kinetic equilibrium, the electrode was briefly rinsed and the SWV curve was recorded. The same experimental conditions were used for the selectivity tests. The tested samples include Hg2+, Cr3+, Cd2+, kanamycin, sulfadimethoxine, phthalic acid, benzoic acid, or ethyl acetate. The concentration of each sample was 10 µM. RESULTS AND DISCUSSION Rational Design of the Original Target for Solid Phase Immobilization. PAEs are the ester derivatives of phthalate acid and a typical PAE consists of a phthalate acid ester group and one or two alkyl chains (Figure 2). In order to select group-specific aptamers, it is essential to expose the common group of PAEs for aptamer binding and prevent the rest parts from participating in the aptamer binding. Here we took advantage of the steric hindrance of solid phase to meet this requirement. Theoretically, either phthalate acid ester group or alkyl chains can be exposed via the rational design of the original targets. In the two recently reported works on the aptamer selection of PCB, PCB derivatives with either OH-

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or NH2-46 on one of the benzene rings were used for

target immobilization. Encouraged by their work and also due to the easy synthesis, we started with the synthesis of OH-functionalized DEHP (4-OH-DEHP, Figure 2, E). By this design, 4-OH-DEHP was immobilized on the matrix via the OH-group and the 10


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alkyl chains were exposed at the surface of the matrix. The synthesis of 4-OH-DEHP and its immobilization on Dynabeads® MyOneTM Carboxylic Acid (Figure S-1), 1H NMR (Figure S-2) and EIS-MS (Figure S-3) characterization,

and the SELEX

process (Figure S-4, Table S-1) were all provided in the SI. Unfortunately, we saw very limited affinity improvement of the pool to the target bound beads and the persistently strong affinity to the bare matrix during the 5 cycles of SELEX (Figure S-5, S-6, S-7), suggesting the failure of the enrichment. The immobilization of PAEs via the OH- on the benzene ring should cause the phthalic acid ester groups covered by the alkyl chains. Therefore, the results suggested that the aptamers specific to alkyl chains are hard to be selected.

Figure 2. Chemical structures of (A) PAEs; (B) BBP; (C) DBP; (D) DEHP; (E) 4-OH-DEHP; (F) DBP-NH2.

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Compared to alkyl chains, the phthalic acid ester group may have the higher chance to specifically interact with aptamers by π-π stacking and hydrogen bonding.45 If the phthalic acid ester group is exposed for aptamer binding, the group-specific aptamers might be obtained. For this purpose, the anchor group must be introduced at the end of one alkyl chain. DBP (Figure 2C) was chosen as the parent target for the synthesis of the original target. Considering the mild and quite efficient coupling efficiency of the NHS-mediated carbodiimide reaction, NH2- was chosen to be introduced at the end of one alkyl chain of DBP. Synthesis of DBP-NH2 and Its Immobilization onto Epoxy-Activated SepharoseTM 6B. DBP-NH2 was chemically synthesized via 3 steps as shown in Figure 3A and the success of the synthesis was confirmed by both 1H NMR (Figure S-8, S-9) and EIS-MS (Figure S-10). DBP-NH2 was then coupled onto Epoxy-Activated Sepharose™ 6B.

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The success of the coupling was confirmed by

the elemental analysis (Table S-3). The percentage of carbon element significantly increased from 33.61% (for the bare Sepharose™ 6B) to 50.21% after the coupling. The percentage of nitrogen element also increased from 0.92% to 2.3%, while the percentage of hydrogen element decreased from 9.26% to 7.88%. These changes were in good agreement with the composition change resulted from the coupling of DBP-NH2 (Figure 3B).

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Figure 3. Synthesis of DBP-NH2 (A) and its immobilization onto Epoxy-Activated SepharoseTM 6B (B).

In Vitro Selection of DNA Aptamers using DBP-NH2 as the Original Target. DBP-NH2-binding DNA aptamers were isolated from a random ssDNA library by the traditional SELEX coupled with high-throughput sequencing technology (Figure 1). The SELEX was terminated after the 5th cycle because the serious cross-linking of the enriched pool was observed on the urea gel even after the denaturation treatment (at 95ºC for 10 min in 2 × TBE-urea sample buffer, Bio-rad). This phenomenon suggested the presence of strong interactions among various DNA sequences, which indicated a high possibility of sequence enrichment. The pool4 was then amplified by PCR for the high-throughput sequencing. The high-throughput sequencing results showed that the top 100 most frequently founded sequences (Table S-4) were highly conserved and all came from the same family

(consensus

sequence:

5'-CTTTCTGTCCTTCCGTCACN1-4TCCCACGC

ATTCTCCACAT-3'). Such highly conserved sequences are rarely reported in aptamer selection, especially for small molecule targets. The adenosine (A) was the most 13



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preferred nucleotide for the first N in the consensus sequences and 80 out of 100 sequences have A at this position. Among the top 40 sequences, 32 sequences have the consensus sequence and the other 8 sequences are the single-base mutants of the consensus sequence. Among the top 41-100 sequences, 21 sequences have the consensus sequences while 36 sequences are the single-base mutants and only 3 sequences are with two-base mutations. Quite interestingly, the consensus sequence reveals a strong tendency toward cytosine-richness (44.7%), suggesting these sequences may fold into certain secondary structures such as i-motif.48 The presence of a global consensus in the aptamer sequences suggests that the aptamers may bind with PAEs in a similar secondary structure under the selection condition. The strong tendency toward C-richness has also been reported for PCB-binding aptamers but no consensus

sequence was identified.45 To elucidate the binding mechanism, the

in-deep biophysics study is required and is beyond the scope of this study. The top two aptamers, respectively named as DBP-1 and DBP-2 (Table S-1), were further studied for their binding affinity and specificity to DBP-NH2 and other free PAEs using various characterization techniques as described below. Specific Binding of DBP-1 and DBP-2 with Surface Bound DBP-NH2 Demonstrated by Fluorescent Inverted Microscopy. The specific binding of DBP-1 and DBP-2 with the surface bound DBP-NH2 on SepharoseTM 6B was confirmed by fluorescent inverted microscopy. As shown in Figure 4A, only the DBP-NH2 coated SepharoseTM 6B displayed the bright fluorescence (Figure 4A, the two images on the right side) after respectively incubated with the fluorophore-labeled DBP-1 or 14


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DBP-2 (DBP-1-FAM and DBP-2-FAM, Table S-1). In contrast, no fluorescence emission was observed for the bare SepharoseTM 6B after the 1 hr's incubation with DBP-1-FAM or DBP-2-FAM (Figure 4A, two images on the left side). The results revealed that DBP-1 and DBP-2 had the affinity to the immobilized DBP-NH2, but not to the matrix. Please note that DBP-1 and DBP-2 are the core sequences without the primer-binding sequences. Therefore, the results also suggested that the primer-binding sequences are not necessary for the binding of aptamers with the immobilized DBP-NH2.

Figure 4. Affinity measurements of DBP-1 and DBP-2 for surface bounded DBP-NH2 by fluorescent inverted microscopy (A) and qPCR-based assays (B). The errors of the Kds were calculated from 3 measurements of the same sample. 15



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Determination of Kd via qPCR-based Assays. The affinity measurements for hydrophobic small molecule targets are quite challenging due to their quite limited water solubility. For example, the estimated maximum concentration of DEHP is below 40 µM even in PAE binding buffer that contains the multiple surfactants to enhance the solubility of DEHP. Above this concentration, the mixture is not transparent any more, suggesting the formation of aggregates. Thus, the saturation of the titration curve can’t be achieved and therefore, the Kd can’t be measured in solution or by immobilizing aptamers on surface. Here we immobilized DBP-NH2 on the hydrophilic surface to avoid the solubility problem. In order to facilitate the separation process and minimize the errors, the size-uniform magnetic Dynabeads instead of Sepharose beads were used. The qPCR instead of fluorescence assay was chosen for the quantification of bound aptamers to minimize the amount of DBP-NH2 required to prepare DBP-NH2-coated beads. In addition, we found that the magnetic Dynabeads sometimes caused the fluorescence quench for unknown reasons, which would interfere the determination of Kd as well. Typically, the same amount of DBP-NH2-coated Dynabeads was respectively incubated with aptamers at varied concentrations, followed by a heat elution. The amount of aptamers in the elution buffer was then determined by qPCR. The binding curves were then plotted in Figure 4B. DBP-1 and DBP-2 all displayed the nanomolar Kd (64 ± 10 and 100 ± 1 nM, respectively), while the initial pool showed no affinity to the DBP-NH2-coated Dynabeads. 16


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Relative Affinity and Selectivity Tests of DBP-1 and DBP-2 to Free PAEs via Competition Assays. As shown in Figure 4, the aptamers can specifically bind with the surface immobilized DBP-NH2. Whether they still can bind with free DBP-NH2 and PAEs or not was subsequently tested via competition assays. Three most toxic and widely used PAE congeners, DBP, BBP, and DEHP, were chosen for this study. In addition, the selectivity of the aptamers against the potential interferences commonly existed in the soft drinks and environmental waters such as glucose, ethanol, and antibiotics were also tested. The aptamers were first bound with the DBP-NH2 coated Dynabeads, and then the same amount of the complex was respectively mixed with free DBP-NH2, BBP, DBP, DEHP, and potential interferences (a mixture of glucose, ethanol, kanamycin, and ampicillin). Thus there was a competition between the immobilized DBP-NH2 and free samples. The relative affinity and selectivity of the aptamers for each sample defined as the ratio of the number of aptamers in sample assay to that in the buffer assay. Therefore, the ratios reflect both the relative affinity of aptamers for different PAE congeners and the selectivity of aptamers against the potential interferences. The higher the ratio is, the higher affinity of the aptamer has for the tested sample.

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Figure 5. Relative affinity measurements of DBP-1 (A) and DBP-2 (B) for free DBP-NH2 (10 µM), DBP (10 µM), DEHP (10 µM), and BBP (10 µM) via competition assays. The ratio (the relative affinity) was calculated by dividing the number of aptamers released from the beads in the presence of the sample by the number of aptamers released from the beads in the PAE binding buffer. Other: a mixture of potential interferences (glucose, kanamycin, ampicillin, and ethanol) all at 10 µM. The errors were calculated from 3 individual measurements.

For both DBP-1 and DBP-2, the ratios for free DBP-NH2, DBP, DEHP, and BBP were all greater than 1, suggesting the aptamers all had higher affinity to those free PAEs than to the PAE binding buffer. In addition, the ratios for free DBP, DEHP, and BBP were 1.8-4.1 and 1.4-2.0 times greater than those for free DBP-NH2 for DBP-1 and DBP-2, respectively (Figure 5). The results suggested that both DBP-1 and DBP-2 had the higher affinity for the unmodified PAEs than for DBP-NH2. In contrast, 18


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for both DBP-1 and DBP-2, the ratios for the potential interference (referred as Other in Figure 5) were all smaller than 1, suggesting the aptamers all had lower affinity to the potential interference than to the PAE binding buffer. In another word, the numbers of aptamers released from the beads in the presence of the potential interference were less than those in PAE binding buffer, implying that the potential interferences had no affinity to the aptamers and even inhibited the dissociation of aptamers from the immobilized DBP-NH2 at a certain level. To confirm the results, three more repeated experiments were conducted by another author and the similar results were obtained. Both PAEs and the potential interferences at the tested concentration had no effect on the amplification efficiency of the PCR (data not shown). Thus, both DBP-1 and DBP-2 possessed the excellent selectivity for PAEs against the tested potential interferences. Together, the competition assays demonstrated that both DBP-1 and DBP-2 were universal binders for all the tested free PAEs with high affinity and selectivity. In order to get clues about the binding mechanism, the selectivity of DBP-1 was further investigated by measuring the relative affinity against several small molecules that contain either the ester or the benzoic acid group. As shown in Figure S-11, DBP-1 showed some affinities to phthalic acid, benzoic acid, and ethyl acetate, but they were all much weaker than these to PAEs (Figure 5). The affinity with phthalic acid was slightly higher than that with benzoic acid, further confirmed that the benzoyl group should be the binding site for DBP-1. The similar affinity with ethyl acetate indicated that the ester group was also critical for the binding of DBP-1 with 19



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PAEs. Therefore, DBP-1 showed good group selectivity to PAEs that contain both the ester and the benzoyl groups. Selectivity Tests of DBP-1 via A Gold Nanoparticle-based Fluorescence Resonance Energy Transfer (FRET) Assay. Even though the Kd can’t be measured in solution or by immobilizing aptamers on surface for PAE aptamer due to the limited solubility of PAEs, these measurements are still valuable to further prove the selectivity of the aptamers. For this purpose, we conducted a gold nanoparticle-based fluorescence resonance energy transfer (FRET) assay, in which the fluorophore labeled DBP-1 was pre-absorbed onto gold nanoparticles (AuNPs) and the fluorescence was significantly quenched (Figure S-12). The selectivity of DBP-1 was tested by respectively challenging the aptamer-AuNP complex with heavy metal ions (Hg2+, As3+, Cu2+, Cr3+, Cd2+, Ni2+) and small molecules. The tested concentrations were all 10 µM. The strong fluorescence increase was observed only in the presence of DEHP, suggesting that the aptamers specifically bound with DEHP. The weak fluorescence increases were observed upon respectively challenged with phthalic acid, benzoic acid, or ethyl acetate. The fluorescence increases in the presence of kanamycin and sulfadimethoxine were negligible. The selectivity results were in good agreement with those determined by the competition assay (Figure S-11). Interestingly, the fluorescence decreases were observed for all the tested heavy metal ions (Figure S-12). It was probably due to the limited stability of aptamer-AuNP. The metal ions caused the partially aggregation of AuNPs, leading to the stronger FRET between the dyes and AuNPs. Overall, these results demonstrated the good selectivity of DBP-1. 20


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Applications of PAE-Binding Aptamers in PAE Detection. The PAE-binding aptamers were then conveniently used for the development of high throughput, sensitive, and low-cost methods for the detection of PAEs. The following two examples are intend to briefly demonstrate the potential. A better performance should be achieved after optimizations. The demonstrated methods might also be applicableto other congeners that were not tested in this study. (1) Competition-Based Quantitative-PCR Assay. Similar to the experiments for the determination of relative affinity and selectivity shown in Figure 5, a titration experiment was conducted, where the samples containing DBP at varied concentrations were incubated with the aptamer-bound beads. As the concentration of DBP decreased the ratio decreased. The 100 nM DBP was clearly differentiated from the buffer control for both aptamers (Figure S-13). The competition assays are compatible for the high-throughput detection of PAEs. (2) DEHP Electrochemical Biosensor. The electrochemical biosensors are very attractive for on-site screening of interested targets due to their superior sensitivity compared to their counterpart optical biosensors.41,

49, 50

We constructed an

electrochemical biosensor (Figure 6A) according to our recently reported work, named

signaling-probe

displaced

electrochemical

aptamer-based

biosensor

(SD-EAB).41 A typical SD-EAB is comprised of a gold electrode immobilized with DNA duplexes formed between a thiolated DBP-1 probe (HS-DBP-1, Table S-1) and a ferrocene (Fc) tagged signaling probe (DBP-1-C-Fc, short strand complementary to the middle section of HS-DBP-1, Table S-1). Fc is close to the electrode surface, thus 21



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can efficiently exchange electrons with the underlying electrodes and generates high current. DEHP competitively binds with the aptamer, resulting in the displacement of the signaling probe and a measurable decrease of electrochemical signal. As shown in Figure 6, the PAE sensor was able to detect DEHP with a detection limit of 10 pM (3S/N) and a dynamic range from 10 pM to 100 nM. The linear relationship was observed between the signal decrease and the logarithm of the DEHP concentration. No clear current decrease over the whole titration process (3 hr) was observed when the sensor was incubated in the PAE binding buffer without DEHP (Figure S-14), suggesting that the current decreases observed during the titration of DEHP were generated due to the binding-induced strand displacement instead of the self-dissociation of the signaling probe. In a good agreement with the results collected by the qPCR (Figure 5, Figure S-11) and the FRET assay (Figure S-12), the DEHP sensor has extraordinary selectivity (in excess of 105). The current decreases caused by the typical environment pollutants such as heavy metal ions (Hg2+, Cr3+, and Cd2+) and antibiotics (kanamycin and sulfadimethoxine), phthalic acid, benzoic acid, and ethyl acetate at 10 µM were all smaller than that caused by DEHP at 100 pM. A significant residual current signal was observed when the sensor was challenged with high concentration of DEHP (100 nM). We believe that it was contributed partially to the limited solubility of DEHP and partially to the nonspecifically absorbed signaling probes on the electrode.41,

49

Previously we demonstrated that the use of

AuNP-attached signaling probe to replace the molecular signaling probe dramatically clears up the strong background signal caused by the nonspecific adsorption of the 22


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signaling probe in a split aptamer-based electrochemical sandwich assay.49 The similar strategy should also work for the PAE sensor and the study is under the way. As shown above, the PAE SD-EABs hold a great promise for practical applications due to its compelling sensitive and extraordinary selectivity that have never been achieved before.

Figure 6. PAE SD-EAB for ultrasensitive and ultraselective detection of DEHP: mechanism (A), SWV curves (B), calibration curve (C), and selectivity tests (D).

CONCLUSIONS In summary, we demonstrated the successful in vitro selection of group-specific PAE DNA aptamers via the rational designed target immobilization strategy. The identified PAE aptamers have the highly conserved C-rich sequences and basically come from the same family. The top-2 aptamers all showed nanomolar Kds and high specificity. 23



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We respectively demonstrated the applications of PAE aptamers in both a competition-based qPCR assay and an electrochemical biosensor for the convenient and low-cost detection of PAEs with the ultrahigh sensitivity and exceptional specificity. The PAE aptamers targeting one of the most sought for targets lay foundation for future developing portable, sensitive, and high-throughput assays and biosensors for the detection of PAEs in food and environmental samples. In addition, our study also provides insights on the group-specific aptamer selection of highly hydrophobic small molecules. We investigated the importance of target immobilization strategy in SELEX. Currently, the solid phase-based SELEX technology is the most widely used method for the selection of aptamers for small molecule targets due to the difficulty in the separation of the target bound aptamers and free aptamers by other separation technologies. Compared to proteins, small molecules are small in size and quite often lack the functional groups for surface immobilization. Thus, the way to immobilize small molecules on the surface has a huge impact on the outcome of SELEX and is essentially the most critical step in the whole selection process. In this study, we found serious nonspecific absorption on the beads and quite limited affinity improvement during the SELEX process when the original target (4-OH-DEHP) was coupled to the carboxylic acid-functionalized Dynabead via the OH group on the benzene ring. By this immobilization strategy, the hydrophobic alkyl chains are exposed at the surface of beads and the phthalic acid ester group is covered. In sharp contrast, the clear sequence enrichment was observed after 4 rounds of SELEX when a different target immobilization strategy was used, 24


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where the functional group for surface immobilization was introduced at the end of one alkyl chain. Thus the phthalic acid ester groups (the binding site for aptamers) are on the most outside of beads. This target immobilization way is theoretically beneficial for the specific binding of PAEs with aptamers and for the selection of group-specific aptamers for PAEs. The use of SepharoseTM 6B to replace the Dynabeads should also contribute to the minimization of the nonspecific absorption of DNAs. Finally, the consensus sequences of the PAE aptamers listed in Table S-4 provide a valuable basis for the study of the binding mechanism between PAEs and the DNA aptamers. The mutations of the consensus sequences located at the quite limited positions. In addition, there is a clear negative correlation between the population of the aptamers and the number of mutations occurred in the sequences. More interestingly, these PAE DNA aptamers are C-rich sequences which might form i-motifs that are four-stranded DNA secondary structures. Initially, i-motifs were thought to be stabilized by acidic conditions, where two parallel-stranded DNA duplexes held together in an antiparallel orientation by forming intercalated hemiprotonated C–C base pairs.51 Therefore, the substantial biological investigation of i-motifs was precluded due to their presumed instability at physiological pH. However, recent advances have shown that i-motif stability is highly dependent on factors such as sequence and environmental conditions and i-motifs can also form at neutral pH.

48

In genomic DNA, wherever there are guanine rich sequences, there are

always complementary sequences rich in C. Such sequences can also form i-motifs 25



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but much less is known about their biological function and relatively few examples of i-motif binding ligands which could be used as probes in such investigations. 48 Some PAEs have been recognized as endocrine disrupters and the study of the binding mechanism between PAEs and their aptamers may also provide insight on the biological function of i-motif structures and the endocrine disrupting effect of PAEs. ASSOCIATED CONTENT Supporting Information This material is available free of charge via the Internet at http://pubs.acs.org. ACKNOWLEDGMENTS We are grateful for financial support from the National Natural Science Foundation (21305093,

21675112),

National

key

scientific

instrument and equipment

development plan (2012YQ030111), and Yanjing Young Scholar Program of Capital Normal University.

26


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REFERENCES

1. Net, S.; Sempere, R.; Delmont, A.; Paluselli, A.; Ouddane, B., Environ. Sci. Technol. 2015, 49, 4019-4035. 2. Wormuth, M.; Scheringer, M.; Vollenweider, M.; Hungerbuhler, K., Risk Anal. 2006, 26, 803-824. 3. Wensing, M.; Uhde, E.; Salthammer, T., Sci. Total Environ. 2005, 339, 19-40. 4. Lu, H. X.; Cai, Q. Y.; Jones, K. C.; Zeng, Q. Y.; Katsoyiannis, A., Crit. Rev. Env. Sci. Tec. 2014, 44, 1-33. 5. Cao, X. L., Compr. Rev. Food Sci. F. 2010, 9, 21-43. 6. Vorkamp, K.; Riget, F. F., Chemosphere 2014, 111, 379-395. 7. Daiem, M. M. A.; Rivera-Utrilla, J.; Ocampo-Perez, R.; Mendez-Diaz, J. D.; Sanchez-Polo, M., J. Environ. Manage. 2012, 109, 164-178. 8. Magdouli, S.; Daghrir, R.; Brar, S. K.; Drogui, P.; Tyagi, R. D., J. Environ. Manage. 2013, 127, 36-49. 9. Dargnat, C.; Teil, M.-J.; Chevreuil, M.; Blanchard, M., Sci. Total Environ. 2009, 407, 1235-1244. 10. Matsumoto, M.; Hirata-Koizumi, M.; Ema, M., Regul. Toxicol. Pharm. 2008, 50, 37-49. 11. Fierens, T.; Van Holderbeke, M.; Willems, H.; De Henauw, S.; Sioen, I., Food Chem. Toxicol. 2012, 50, 2945-2953. 12. Gomez-Hens, A.; Aguilar-Caballos, M. P., Trac. Trend. Anal. Chem. 2003, 22, 847-857. 13. Autian, J., Environ. Health Perspect. 1973, 4, 3-26. 14. Mtibe, A.; Msagati, T. A.; Mishra, A. K.; Mamba, B. B., Phys. Chem. Earth 2012, 50, 239-242. 15. Ranjbari, E.; Hadjmohammadi, M. R., Talanta 2012, 100, 447-453. 16. Xie, Q. L.; Liu, S. H.; Fan, Y. Y.; Sun, J. Z.; Zhang, X. K., Anal. Bioanal.Chem. 2014, 406, 4563-4569. 17. Yilmaz, P. K.; Ertas, A.; Kolak, U., J. Sep. Sci. 2014, 37, 2111-2117. 18. Ostrovský, I.; Čabala, R.; Kubinec, R.; Górová, R.; Blaško, J.; Kubincová, J.; Řimnáčová, L.; Lorenz, W., Food Chem. 2011, 124, 392-395. 19. Xu, D.; Deng, X.; Fang, E.; Zheng, X.; Zhou, Y.; Lin, L.; Chen, L.; Wu, M.; Huang, Z., J. Chromatogr. A 2014, 1324, 49-56. 20. Earls, A. O.; Axford, I. P.; Braybrook, J. H., J. Chromatogr. A 2003, 983, 237-46. 21. Ierapetritis, I.; Lioupis, A.; Lampi, E., Food Anal. Methods 2014, 7, 1451-1457. 22. Sun, R. Y.; Zhuang, H. S., Anal. Methods 2014, 6, 9807-9815. 23. Zhou, L.; Lei, Y.; Zhang, D.; Ahmed, S.; Chen, S., Sci. Total Environ. 2016, 541, 570-578. 24. Zhang, M. C.; Hu, Y. R.; Liu, S. H.; Cong, Y.; Liu, B. L.; Wang, L., Food Anal. Methods 2012, 5, 1105–1113. 25. Zhang, M.; Liu, S.; Zhuang, H.; Hu, Y., Appl. Biochem. Biotechnol. 2012, 166, 436-445. 26. Sun, R. Y.; Zhuang, H. S., J. Environ. Sci. Heal. B 2015, 50, 275-284. 27. Cui, X.; Wu, P.; Lai, D.; Zheng, S.; Chen, Y.; Eremin, S. A.; Peng, W.; Zhao, S., J. Agric. Food. Chem. 2015, 63, 9372-9378. 28. Kuang, H.; Xu, L.; Cui, G.; Ma, W.; Xu, C., Food Agr. Immunol. 2010, 21, 265-277. 29. Zhang, M.; Cong, Y.; Sheng, Y.; Liu, B., Anal. Biochem. 2010, 406, 24-28. 27



1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

30. Sun, R.; Zhuang, H., Anal. Biochem. 2015, 480, 49-57. 31. Ellington, A. D.; Szostak, J. W., Nature 1990, 346, 818-822. 32. Tuerk, C.; Gold, L., Science 1990, 249, 505-510. 33. Lou, X. H.; Qian, J. R.; Xiao, Y.; Viel, L.; Gerdon, A. E.; Lagally, E. T.; Atzberger, P.; Tarasow, T. M.; Heeger, A. J.; Soh, H. T., Proc. Natl. Acad. Sci. U. S. A. 2009, 106, 2989-2994. 34. Bock, L. C.; Griffin, L. C.; Latham, J. A.; Vermaas, E. H.; Toole, J. J., Nature 1992, 355, 564-566. 35. Chen, L.; Rashid, F.; Shah, A.; Awan, H. M.; Wu, M.; Liu, A.; Wang, J.; Zhu, T.; Luo, Z.; Shan, G., Proc. Natl. Acad. Sci. U. S. A. 2015, 112, 10002-10007. 36. Yang, K.-A.; Pei, R.; Stojanovic, M. N., Methods 2016, 106, 58-65. 37. Groher, F.; Suess, B., Methods 2016, 106, 42-50. 38. Li, S.; Xu, H.; Ding, H.; Huang, Y.; Cao, X.; Yang, G.; Li, J.; Xie, Z.; Meng, Y.; Li, X.; Zhao, Q.; Shen, B.; Shao, N., J. Pathol. 2009, 218, 327-336. 39. Shangguan, D.; Li, Y.; Tang, Z.; Cao, Z. C.; Chen, H. W.; Mallikaratchy, P.; Sefah, K.; Yang, C. J.; Tan, W., Proc. Nat. Acad. Sci. U.S.A. 2006, 103, 11838-11843. 40. Wang, C. L.; Zhang, M.; Yang, G. A.; Zhang, D. J.; Ding, H. M.; Wang, H. X.; Fan, M.; Shen, B. F.; Shao, N. S., J. Biotechnol. 2003, 102, 15-22. 41. Liu, R.; Yang, Z.; Guo, Q.; Zhao, J.; Ma, J.; Kang, Q.; Tang, Y.; Xue, Y.; Lou, X.; He, M., Electrochim. Acta 2015, 182, 516-523. 42. Mungroo, N. A.; Neethirajan, S., Biosensors 2014, 4, 472-93. 43. Wang, H.; Wang, Y.; Liu, S.; Yu, J.; Xu, W.; Guo, Y.; Huang, J., Chem. Commun. 2015, 51, 8377-8380. 44. Yildirim, N.; Long, F.; Gao, C.; He, M.; Shi, H.-C.; Gu, A. Z. A., Environ. Sci. Technol. 2012, 46, 3288-3294. 45. Mehta, J.; Rouah-Martin, E.; Van Dorst, B.; Maes, B.; Herrebout, W.; Scippo, M.-L.; Dardenne, F.; Blust, R.; Robbens, J. A., Anal. Chem. 2012, 84, 1669-1676. 46. Xu, S.; Yuan, H.; Chen, S.; Xu, A.; Wang, J.; Wu, L. A., Anal. Biochem. 2012, 423, 195-201. 47. Cho, M.; Xiao, Y.; Nie, J.; Stewart, R.; Csordas, A. T.; Oh, S. S.; Thomson, J. A.; Soh, H. T., PNAS 2010, 107, 15373-15378. 48. Day, H. A.; Pavlou, P.; Waller, Z. A. E., Bioorg. Med. Chem. 2014, 22, 4407-4418. 49. Zhao, T.; Liu, R.; Ding, X. F.; Zhao, J. C.; Yu, H. X.; Wang, L.; Xu, Q.; Wang, X.; Lou, X. H.; He, M., Anal. Chem. 2015, 87, 7712-9. 50. Lou, X.; Zhao, T.; Liu, R.; Ma, J.; Xiao, Y., Anal. Chem. 2013, 85, 7574-7580. 51. Gehring, K.; Leroy, J.-L.; Guéron, M., Nature 1993, 363, 561 - 565

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for TOC only

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Selection of Group-Specific PAE-Binding DNA Aptamers via Rational

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