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DNA-functionalized Nanoceria for Probing Oxidation of Phosphorus Compounds Xiuzhong Wang, Biwu Liu, and Juewen Liu Langmuir, Just Accepted Manuscript • DOI: 10.1021/acs.langmuir.8b03335 • Publication Date (Web): 05 Dec 2018 Downloaded from http://pubs.acs.org on December 6, 2018

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DNA-functionalized Nanoceria for Probing Oxidation of Phosphorus Compounds

Xiuzhong Wang1,2, Biwu Liu2 and Juewen Liu2*

1. College of Chemistry and Pharmaceutical Sciences, Qingdao Agricultural University, Qingdao, 266109, China

2. Department of Chemistry, Waterloo Institute for Nanotechnology, University of Waterloo, Waterloo, Ontario, N2L 3G1, Canada Email: [email protected]

Abstract Chemical reactions without an obvious optical signal change, such as fluorescence or color, are difficult to monitor. Often, more advanced analytical techniques such as HPLC and mass spectroscopy are needed. It would be useful to convert such reactions to changes in optical signals. In this work, we demonstrate that fluorescently labeled DNA oligonucleotides adsorbed on nanomaterials can probe such reactions, and oxidation of phosphorus containing species was used as an example. Various metal oxides were tested and CeO2 nanoparticles were found to be the most efficient for this purpose. Among phosphate, phosphite, and hypophosphite, only phosphate produced a large signal, indicating its strongest adsorption on CeO2 to displace the DNA. This was further used to screen oxidation agents to convert lower oxidation state compounds to phosphate, and bleach was found to be able to oxidize phosphite. Canonical discriminant analysis was performed to discriminate various phosphorus species using a sensor array containing different metal oxides. Based on this, glyphosate was studied for its adsorption and oxidation. Although this method is not specific enough to be selective biosensors, it is useful as a tool to produce sensitive optical signals to follow important chemical transformations.

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Introduction DNA-functionalized nanomaterials have been widely used to develop biosensors, mainly relying on DNA hybridization, aptamer binding, and DNAzyme reactions.1-6 These sensors often have high specificity originated from the DNA probes, while nanomaterials helped for signal transduction, such as fluorescence quenching, generation of color changes, and electrochemical signals.7-11 For many analytes, however, it is quite difficult to obtain highly specific DNA probes. For example, using DNA to detect anions is quite challenging,12-13 since DNA and anions are both negatively charged and they repel each other.14 We reason that instead of using DNA as specific probes, DNA may also be used as a competitor to develop competitive assays.15-16 With programmable structures, high stability, and ease of modification, DNA is ideal competitors with adjustable binding affinities. Such reactions may not be used for sensing due to limited specificity, but under well-controlled conditions, they may be used to study chemical transformations. Phosphorus is an essential nutrient for life.17 Most of phosphorus exists as pentavalent phosphate (PO43−), but anthropological activities have produced large amounts of reduced inorganic phosphorus such as phosphite (PO33−) in the environment, including rivers, lakes and wastewater treatment plants.18 Phosphite is an energetically favorable chemotrophic electron donor due to the extremely low redox potential of the phosphate/phosphite couple (Eϴ′= −650 mV).19 It is commonly used as a reducing agent,20 and many pesticides contain phosphite species.21 With a phosphate backbone, DNA may compete with such phosphorus containing species, and this competition might be useful for studying such pesticides. Glyphosate is one of the most widely used organophosphorus herbicides against weeds,2223

and the phosphorus in glyphosate has the same oxidation state as that in phosphite. Excessive

use of glyphosate increased crop yield, but its environmental contamination has also raised concerns.24 A consensus has been reached that oxidation is the most efficient way to remove glyphosate from polluted environment.25 Various oxidation processes of phosphite have been reported using nitrogen-doped activated carbon as a catalyst,26 photocatalysis with UV/TiO2,27 TiO2 nanotubes doped with cerium,28 H2O2/UV,29-30 and ferrioxalate/UV.31 The reaction process and products were often determined by ion chromatography,32 which is complex, time-consuming, and costly. Therefore, it is useful to find simple methods to oxidize the glyphosate and detect the products. Sodium hypochlorite (NaClO) is the main component of liquid bleach, and its active 2 ACS Paragon Plus Environment

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agent is chlorine.33-34 In this work, chlorine-based bleach was chosen as an oxidant, and DNAfunctionalized metal oxides were used to study related oxidation reactions. For comparison, the oxidation products were also detected by a traditional colorimetric method.

Materials and Methods Chemicals. The sequence of the DNA probe is 5-FAM-ACGCATCTGTGAAGAGAACCTGGG (FAM: carboxyfluorescein), purchased from Integrated DNA Technologies (IDT, Coralville, IA, USA). Sodium chloride and 4-(2-hydroxyethyl)-1-piperazineethanesulfonic acid (HEPES) were purchased from Mandel Scientific (Guelph, Ontario, Canada). Bleach (6% active chlorine) was purchased from GFS Canada Company Inc (Milton, Ontario, Canada), Hydrogen peroxide, sodium perchlorate, nanoceria, malachite green, sodium molybdate dihydrate, glyphosate, sodium pyrophosphate decahydrate (PPi), sodium polyphosphate with an average of 25 phosphate units (Pi25), sodium trimetaphosphate (STMP), sodium triphosphate (STPP), CeO2, and dimethyl phosphate (DMP) were purchased from Sigma-Aldrich. The metal oxide nanoparticles used in this work were described in our previous publication.35 Milli-Q water was used to prepare all the buffers and solutions. DNA-based probing. Desorption of the FAM-labeled DNA from nanoceria was used to evaluate the oxidation reaction. Briefly, to prepare the probe, 50 nM FAM-labeled DNA in HEPES buffer (10 mM, pH 7.4, with 200 mM NaCl) was mixed with nanoceria (a final of 5 μg mL-1) and incubated for 30 min to result in quenched fluorescence. The desorption kinetics were recorded after a quick addition of phosphate to the FAM-DNA/nanoceria mixture and the kinetics of fluorescence increase was followed using a fluorometer (Eclipse, Varian) at room temperature (~23 C). The fluorescence intensity of the free DNA was used to calculate the percentage of desorbed DNA. Colorimetric quantification of phosphate. The malachite green colorimetric assay was used according to the literature with some minor modifications.36 A malachite green reaction solution (R1) was prepared as follows. First, 0.04 g sodium molybdate dihydrate was dissolved in 5 mL of HCl (4 M). This solution was then mixed with 15 mL of 0.045% (w/v) malachite green and stirred for 30 min until it became clear. After centrifugation, the supernatant was stored at 4 C for use. 10 L of various concentrations of phosphate was placed into microcentrifuge tubes and mixed 3 ACS Paragon Plus Environment

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with 8 μL of HCl (0.1 M), 102 μL distilled water and 80 μL R1 for 10 min. Afterwards, the absorbance of the solutions was determined at 670 nm using a UV-vis spectrometer (Agilent 8453A) at room temperature. A mixture of R1 (80 μL), HCl (0.1M, 8 μL), and distilled water (112 μL) was used as blank. Each experiment was run in triplicate. Oxidation of glyphosate. Glyphosate (10 mM) was incubated in different concentrations of bleach for 60 min, after which the solutions were diluted by 1000-fold in HEPES buffer (10 mM, pH 7.4, with 200 mM NaCl) to neutralize the pH. The samples were then used for desorbing the FAMDNA from nanoceria as described above. The final glyphosate (if no reaction occurred) was 10 µM. Sensor array for canonical discriminate analysis. The response of each sensor is plotted by the fluorescence enhancement (F/F0 − 1) from different chemicals. The concentrations of CeO2, ZnO, and Fe3O4 nanoparticles were 5, 20, and 100 g mL-1, respectively. The concentration of different analytes (phosphate, phosphite, hypophosphite, glyphosate, serine and glutamic) was 50 μM each. All the analytes were replicated six times. The fluorescence was recorded after adding the analytes for 60 min. The training data were analyzed using canonical discriminate analysis using the software package from Origin.

Results and Discussion Probing phosphate with DNA adsorbed on CeO2. The probe was prepared simply by adsorbing a carboxyfluorescein (FAM)-labeled 24-mer DNA oligonucleotide with a random sequence on CeO2 nanoparticles, resulting in quenched fluorescence. Adsorption of DNA on CeO2 is quite independent of its base composition, since DNA mainly uses its phosphate backbone to adsorb.37 In general a longer DNA is adsorbed more strongly and thus more difficult to be displaced.15, 35 Our CeO2 nanoparticles were ~5 nm in size from TEM (Figure 1A), while dynamic light scattering showed a quite narrow size distribution of 4.2  2.0 nm (Figure S1). We chose CeO2 for its small size, high DNA adsorption efficiency, and strong fluorescence quenching properties.16, 37-39 In the presence of a competing ligand, such as phosphate, the adsorbed DNA can be displaced, producing fluorescence signals (Figure 1B).39


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Figure 1. (A) A TEM micrograph of the CeO2 nanoparticles used in this work. (B) A scheme of a fluorescently labeled DNA adsorbed by CeO2 and its subsequent displacement by phosphate, recovering the initially quenching signal. (C) Kinetics of fluorescence enhancement of a FAMlabeled DNA (50 nM) adsorbed on CeO2 nanoparticles (100 µg/mL) displaced by the added phosphate in buffer (10 mM HEPES, pH 7.45, 200 mM NaCl). (D) A calibration curve of DNA desorption by phosphate after 60 min reaction in the low phosphate concentration region.

By adding increasing concentrations of inorganic phosphate (orthophosphate), the fluorescence enhancement was faster and stronger (Figure 1C). We plotted the fluorescence increase indicative of desorbed DNA at 60 min (Figure 1D), and the response was initially linear. A detection limit of 0.15 M phosphate was determined based on signal greater than three times of variation of the background. Although quite sensitive, This reaction however was not specific since other molecules and ions, such as arsenate,15 and pyrophosphate (PPi)39 may also displace the DNA. Considering this, we believe this reaction is more useful in a well-defined system free of these interfering species (not as a sensor but as a probe for studying chemical reactions). Responses to hypophosphite, phosphite and phosphate. In this work, we were interested in phosphorus species of different oxidation states. We picked three model compounds shown in Figure 2A: phosphate, phosphite and hypophosphite with the oxidation state of phosphorus being +5, +3, +1, respectively. These species were then added to the DNA/CeO2 probe. Interestingly, we only observed fluorescence with phosphate, while the two lower oxidation state species did not produce any signal (Figure 2B). This indicated that the other two species were poorly adsorbed by CeO2. This large difference in response gave us an opportunity to study related oxidation reactions. The lower oxidation state phosphorus species had fewer number of associated oxygen atoms. Since


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oxygen is the main ligand for interacting with CeO2, the lower oxidation species cannot strongly adsorb on it.

Figure 2. (A) The structures of the phosphate containing species studied in this work. (B) Kinetics of fluorescence enhancement indicative of FAM-labeled DNA displaced by the added different phosphate species (10 µM of phosphorus).

Responses with other metal oxides. In addition to CeO2, we also tested a few other common metal oxide nanoparticles, including ZnO, Fe2O3, Fe3O4, TiO2, ZrO2, and MgO. We wondered if them could work, or even have a better response. First, we followed adsorption of the DNA probe by adding increasing concentrations of each oxide nanoparticle to the FAM-DNA (Figure 3A). In general, stronger quenching was observed with more oxides added, but different oxides have different abilities for DNA adsorption and fluorescence quenching. The strongest quenching was observed with CeO2, likely due to its very small size. Almost no quenching was observed with TiO2, ZrO2 and MgO. TiO2 can adsorb DNA and quench fluorescence,40 but its adsorption is favored at low pH.41 With a positive surface charge, ZnO can effectively adsorb DNA.42 MgO, on the other hand, may not be a good fluorescence quencher since it is an insulator. Therefore, DNA adsorption by metal oxides is affected by particle size, charge, and chemical property of the metal species. Since we observed fluorescence quenching with CeO2, ZnO, Fe2O3, and Fe3O4, we then tested the efficiency of DNA desorption from them by adding 10 M phosphate and the kinetics of fluorescence enhancement was monitored (Figure 3B). It is interesting to notice that CeO2 had 6 ACS Paragon Plus Environment

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both the strongest capacity of DNA adsorption, and the highest desorption efficiency by phosphate. This indicated that CeO2 adsorbed the DNA with an appropriate affinity.

Figure 3. (A) Quenching of the FAM-labeled DNA (50 nM) as a function of concentration of various nanoparticles in buffer (10 mM HEPES, pH 7.45, 200 mM NaCl). (B) Kinetics of fluorescence enhancement indicative of FAM-labeled DNA displaced from different nanoparticles after adding 10 M phosphate.

Colorimetric phosphate determination. Since the DNA-based measurement is new, we also used a traditional colorimetric assay for comparison. Malachite green forms a strongly absorbing blue species (molar absorption coefficient at around 670 nm, 90000 M−1 cm−1) after complexing with phosphomolybdate.39 We first examined the selectivity of the colorimetric assay using a series of phosphorus compounds, including hypophosphite, phosphite, and phosphate. It is interesting to notice that only phosphate changed the color to blue (Figure 4A). Next, different concentrations of phosphate were tested, and the absorbance at 670 nm increase gradually (Figure 4B). The inset of Figure 4B shows the color of these samples, and a nice color gradient was observed. These samples were used to build a calibration curve for subsequent quantification of phosphate (Figure 4C). A good linear correlation between the absorbance at 670 nm and phosphate concentration in the range from 0.25 to 10 M was obtained. The limit of detection was 0.08 M, which was similar to our CeO2/DNA based sensor. We also reacted various polyphosphates with this probe and still only orthophosphate produced a color change (Figure S2).


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Figure 4. (A) A photograph showing the malachite green colorimetric assay of different phosphorus containing compounds: (a) blank, (b) hypophosphite, (c) phosphite and (d) phosphate. (B) The absorbance at 670 nm in the presence of phosphate of different concentrations. Inset: the color of the samples with the concentrations of phosphate marked in M. (C) The linear relationship at low phosphate concentrations.

Screen of oxidation agents. Now that a large difference was observed between different oxidation states of phosphorus, we then used this reaction to study their oxidation. Phosphorus in a low oxidation state may produce phosphate after oxidation, which can be probed by the fluorescence enhancement of the sensor. To test this idea, we first screened a few oxidation methods such as H2O2/UV,29-30 perchlorate acid, H2O2, and a mixture of perchlorate acid and H2O2 to respectively oxidize hypophosphite and phosphite. The products of the above the reactions were then added into the DNA/CeO2 solution. For hypophosphite, almost no fluorescence increased in any of these samples indicating no phosphate was produced (Figure 5A).30 For phosphite (Figure 5B), fluorescence increase was observed with bleach, while the other oxidation conditions still failed. Therefore, cost-effective bleach might oxidize phosphite to orthophosphate but it cannot oxidize hypophosphite. To further understand the oxidation of phosphite by bleach, we the varied the concentration of bleach from 0.15 ~ 14.8%. The efficiency of the oxidation increased roughly linearly below 1.48% bleach, and adding more bleach beyond 7.4% reached saturation (Figure 5C).


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Figure 5. Kinetics of fluorescence enhancement indicative of FAM-labeled DNA displaced from CeO2 nanoparticles by the reaction products of (A) hypophosphite, and (B) phosphite mixed with (a) perchloric acid (100 mM), (b) H2O2 (100 mM), (c) mixture of perchloric acid and H2O2, (d) H2O2 (100 mM)/UV (470 nm for 90 min), and (e) 80% bleach (4.8% active chlorine, about 676 mM). These reaction mixtures were diluted by 1000-fold into the sensor. (C) Phosphite oxidized by different concentrations of the bleach solution.

Adsorption of glyphosate. After understanding the adsorption of various simple phosphorus containing species, we then studied the adsorption of glyphosate also using the FAM-DNA. The structure of glyphosate is shown in Figure 2A. In addition to a phosphite, it also has a carboxyl group, which increased its number of negative charges. We then added glyphosate to the FAMDNA and metal oxide complexes. With a low concentration of glyphosate (e.g. 10 µM), the response was very low for all the oxides (Figure 6A), and almost no DNA desorbed from any of them. With a higher concentration of 50 µM glyphosate, we observed a fast response with CeO2 and ZnO (Figure 6B), while the two iron oxide samples were still silent. We then compared the oxidation products of phosphite and glyphosate, and they had almost the same fluorescence enhancement demonstrating that the products had a similar ability to adsorb on CeO2 (Figure 6C). Using the CeO2 sensor, the response was measured as a function of the concentration of glyphosate (Figure 6D), and a nice concentration-dependent response was observed. As shown Figure 6E, the fluorescence response at 60 min after adding glyphosate was initially linear and then saturated at higher glyphosate concentrations. A detection limit of 0.2 M glyphosate was determined. A careful observation of the data revealed that the shape of the kinetic curves was quite different for phosphate and glyphosate on different nanoparticles (compare Figure 3B and Figure 6B). For example, phosphate had a very slow response for the ZnO sample, but for glyphosate, 9 ACS Paragon Plus Environment

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initially its kinetics was very fast. This could be related to the chelation effect of the phosphate and carboxyl groups in glyphosate. Different metal oxide nanoparticles may have different affinities with the DNA. At the same time, they also adsorb different phosphorus species differently. With such an observation, we may construct a pattern-recognition-based sensor array for discrimination of phosphorous compounds.35, 43-45 We used a set of sensors containing the FAMDNA adsorbed on CeO2, ZnO and Fe3O4, respectively. The response of each sensor was plotted by the fluorescence enhancement from different analytes (50 M each) after 1 h incubation. The training data were analyzed using canonical discriminate analysis from Origin (Figure 6F). Since glyphosate has a structure similar to amino acids, we also included a few amino acids in this study. With this method, glyphosate can be readily discriminated from other phosphorus containing species as well as from the tested amino acids.

Figure 6. (A) Kinetics of fluorescence enhancement indicative of FAM-labeled DNA displaced by 10 M glyphosate on different nanoparticles from (a) Fe3O4, (b) Fe2O3, (c)ZnO and (d) CeO2, respectively. (B) Kinetics of fluorescence enhancement by the added 50 M glyphosate to different FAM-DNA/oxide complexes. (C) Fluorescence enhancement from CeO2 by the added the oxidation products of phosphite and glyphosate with bleach (80%). (D) The CeO2 sensor kinetic 10 ACS Paragon Plus Environment

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response as a function of the concentration of glyphosate from 0.5 to 100 M. (E) A calibration curve corresponding to the percentage of DNA desorption (at 60 min) as a function of glyphosate concentration. (F) A canonical score plot for fluorescence enhancement using three sensors (CeO2, ZnO, and Fe3O4) for the discrimination of phosphate, hypophosphite, phosphite, glyphosate, serine and glutamic. The ellipses represent the 95% confidence level.

Oxidation of glyphosate. Glyphosate is often treated via catalytic oxidation to produce less toxic phosphate products.26 We wanted to test whether the screening method in this work is effective also for glyphosate. We compared the fluorescence enhancement by glyphosate before and after oxidation by bleach (Figure 7A). Glyphosate can partly displace the FAM-labeled DNA as concluded from the moderate fluorescence enhancement. However, a much stronger enhancement was observed after adding the oxidation product of glyphosate. This difference in DNA desorption kinetics may allow us to monitor oxidation of glyphosate. We then tested the effect of bleach concentration from 0.15 to 14.8% (Figure 7B). No DNA desorbed in the presence of bleach alone (curves a-e). In the presence of the glyphosate and bleach, the signal was obvious, and the desorption rate increased with increase of the bleach concentration (curves f-j). Therefore, this system can be used for studying oxidation of glyphosate.

Figure 7. (A) Kinetics of fluorescence enhancement indicative of FAM-labeled DNA displaced by the added the glyphosate (a) before and (b) after oxidation by bleach (80%). (B) The effect of different concentrations of the bleach solution with 0.15, 0.74, 1.48, 7.4 and 14.8%, respectively. (a)-(e) bleach alone showing no response; (f)-(j): bleach mixed with glyphosate (50 M).


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Using DNA to study chemical transformations have been carried out previously mainly by enzyme reactions and aptamers. For example, Li and coworkers studied the hydrolysis of ATP and the adenosine deamination reaction using a DNA aptamer.46-47 Famulok used aptamer for fluorescence polarization based screening of inhibitors.48 In those cases, they relied on specific aptamers. We studied the reduction of Ce4+ to Ce3+ by iodide using a DNAzyme,49 and also the ring opening reaction of sodium trimetaphosphate.39 The present work describes another example to look at redox reactions using DNA/nano conjugates, where the DNA only played a signaling and non-specific competitor role. Organophosphorus pesticides are a group of organic compounds containing C−P, C−O−P, C−S−P or C−N−P bonds. In this paper, glyphosate was used as a model analyte, and its degradation may proceed through one of the two C−N bond positions, results in the formation of aminomethylphosphonic acid or through the cleavage of a C−P bond, forming sarcosine and then glycine.50 Oxidation mechanisms of other organophosphorus pesticides may be related to P–S or P–OH bonds with the use of different oxidants.51 Whether this method can be used for studying oxidation of other pesticides has to be further tested, and likely screening for the adsorbing material has to be carried out in each case.

Conclusions In summary, we have studied adsorption of a few phosphorus containing species of different oxidation states by CeO2 nanoparticles using fluorescent DNA as a probe. The main goal is to demonstrate DNA as a non-specific probe for studying chemical reactions under well-defined conditions. DNA can be adsorbed on many surfaces via different interaction forces.37, 52-53 On metal oxides the role of DNA phosphate backbone is of particular importance.35, 37 Among the tested, only orthophosphate can displace the DNA, with the best performance observed with CeO2 nanoparticles. This allowed us to study the oxidation of lower oxidation state species. A comparison was also made with a classic colorimetric assay for measuring phosphate. If a compound before and after a chemical transformation has different affinities to a surface and DNA might be able to serve as a good probe for this reaction. Thus, the generality of this method can be quite high for monitoring reactions that do not have a change in optical signals.


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Acknowledgement Funding for this work was from The Natural Sciences and Engineering Research Council of Canada (NSERC). Dr. X. Wang was supported by the Shandong Provincial Government Scholarship (China) to visit the University of Waterloo. We also thank the Natural Science Foundation of Shandong Province, China (No. ZR2018MB030).

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