Mechanochemical Sensing of Single and Few Hg(II) Ions Using

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Mechanochemical Sensing of Single-and Few-Hg (II) Ions Using Polyvalent Principles Shankar Mandal, Sangeetha Selvam, Prakash Shrestha, and Hanbin Mao Anal. Chem., Just Accepted Manuscript • Publication Date (Web): 12 Sep 2016 Downloaded from http://pubs.acs.org on September 12, 2016

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Analytical Chemistry

Mechanochemical Sensing of Single-and Few-Hg (II) Ions Using Polyvalent Principles Shankar Mandal, Sangeetha Selvam, Prakash Shrestha, and Hanbin Mao* Department of Chemistry and Biochemistry, Kent State University, Kent, OH 44242, USA ABSTRACT: Sensitivity of biosensors is set by the dissociation constant (Kd) between analytes and probes. Although potent amplification steps can be accommodated between analyte recognition and signal transduction in a sensor to improve the sensitivity 46 orders of magnitude below Kd, they compromise temporal resolution. Here, we demonstrated mechanochemical sensing that broke the Kd limit by 9 orders of magnitude for Hg detection without amplifications. Analogous to trawl fishing, we introduced multiple Hg binding units (thymine (T)-T pairs) in a molecular trawl made of two poly-T strands. Inspired by dipsticks to gauge content levels, mechanical information (force/extension) of a DNA hairpin dipstick was used to measure the single or few Hg2+ ions bound to the molecular trawl, which was levitated by two optically trapped particles. The multivalent binding and single-molecule sensitivity allowed us to detect unprecedented 1 fM Hg ions in 20 mins in field samples treated by simple filtrations.

Conventional biosensors consist of two components, a molecular recognition unit and a signal transduction unit. In the first unit, molecular recognition occurs, which sets the specificity for a biosensor. In the second unit, signals from the recognition unit are measured, which fine tunes the sensitivity of the sensor 1-3 given that dissociation constant (Kd) between the analyte and its recognizer is fixed. In the majority of cases, such as ELISA (Enzyme Linked ImmunoSorbent Assay),4 these two units require different sets of infrastructure. A separate signal transduction often needs extra steps, such as various amplifications, to generate measurable signatures. Although physical separation of the two units prevents cross-talks between individual components, it considerably increases complexity in the traditional sensing, which leads to additional errors that propagate through multiple stages and generate false positive or false negative results. To address this issue, we have pioneered single-molecule mechanochemical sensing (SMMS)5,6 in which mechanochemical coupling has been exploited to directly connect molecular recognition and signal transduction units. Mechanochemical coupling occurs when individual macromolecules change their conformation upon ligand binding. This is often accompanied by variation in mechanical properties, such as tension or extension, in the macromolecule. By following the mechanical coupling using instruments such as laser tweezers7 in real time, the single-molecule mechanochemical sensing can be accomplished without extra infrastructure employed in a separate signal transduction unit. The single-molecule templates offer the utmost sensitivity for individual molecules, which, in theory, can break the detection limit set by the Kd in ensembleaveraged sensors. However, such platforms have limited space to accommodate multiple functionalities, which, for example, are necessary to reduce the detection time for extremely low analyte concentrations. To transform singlemolecular sensing into unprecedented devices with expanded capabilities, here, we conceptualized and demonstrated singlemolecule mechano-analytical real time sensing (or SMARTS) devices that amalgamate two new molecular functionalities,

trawling and dipsticking, for detection of single or few Hg2+ ions. Mercury contamination is a prevalent environmental concern as Hg2+ can mutate genomic DNA through binding with a T-T mismatch pair in dsDNA8,9. According to the EPA the maximum level of Hg2+ of 10 nM (2 ppb) in drinking water is considered to be safe10. Most commonly, Hg2+ is determined by atomic absorption spectrometry (AAS) techniques11,12 and other closely related Hg2+ detection methods, such as inductively coupled plasma - mass spectrometry (ICP-MS)13, inductively coupled plasma – optical emission spectrometry (ICPOES)14, cold vapor – atomic absorption spectrometry (CVAAS)15, and cold vapor - atomic fluorescence spectroscopy (CV-AFS)16 with a detection limit at sub nM to pM levels (20 ppt to 0.02 ppt)11,12. Currently, Hg2+ has also been detected using electrochemical17-19, optical20, fluorescence21,22, or colorimetric22 detections at ultrasensitive levels. By exploiting single-molecular sensitivity of newly developed mechanochemical sensing23,24, we expect the detection limit of Hg2+ can be significantly reduced. As detection limit is ultimately determined by dissociation constant between the analyte and the sensing probes, we also employ polyvalent binding principles to further improve the detection limit of mechanochemical sensing. These approaches indeed allowed us to detect Hg2+ concentration at the unprecedented level of 1fM (2×10-4 ppt) EXPERIMENTAL SECTION Materials and Reagents. All chemicals including Tris-HCl, KCl, EDTA, Zn(NO3)2.6H2O, MnCl2.4H2O, CoCl2.6H2O, CuCl2.2H2O, Hg(NO3)2.H2O, 3CdSO4.8H2O, FeSO4.7H2O, and Ni(NO3)2.6H2O were purchased from either Fisher Scientific or Sigma with purities > 99.0 %. Pb(NO3)2 was purchased from MP Biomedicals LLC. DNA oligomers were purchased from Integrated DNA Technologies (www.idtdna.com) and purified with denaturing polyacrylamide gel electrophoresis (PAGE). The DNA sequences are listed in Table S1. Unless specified otherwise, all enzymes were obtained from New England Biolabs (www.neb.com). The streptavidin or anti-

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digoxigenin coated polystyrene beads for laser tweezers experiments were purchased from Spherotech (Lake Forest, Illinois, USA). Preparation of the DNA Constructs Used in the SMARTS. The single-molecule mechanoanalytical real time sensing (SMARTS) device contains a single-molecule DNA template, which was prepared using a protocol described elsewhere25. Briefly, a DNA hairpin which acts as a mechanochemical dipstick reporter was sandwiched between two dsDNA handles (2,028 and 2,690 bp). Opposite to the hairpin, poly-thymine anti-parallel DNA oligomers were introduced to serve as recognition units for Hg2+ (Figure S1). The 2,028 bp handle was prepared via PCR amplification of a pBR322 plasmid (New England Biolab, NEB) using a biotinylated primer 5'-GCA TTA GGA AGC AGC CCA GTA GTA GG3´(IDT, Coralville, IA) and subsequently digested with XbaI restriction enzyme (NEB). The 2,690 bp DNA handle was synthesized by the SacI (NEB) and EagI (NEB) digestions of a pEGFP plasmid (Clontech, Mountain View, CA). This handle was subsequently labeled at the 3′ end by digoxigenin (Dig) using 18 µM Dig-dUTP (Roche) and terminal transferase (Fermentas). The final DNA construct was synthesized using T4 DNA ligase (NEB) through three-piece ligation of the 2028 and 2690 dsDNA handles and an oligonucleotide which can form the DNA hairpin. To optimize the efficacy of the SMARTS device, DNA constructs with different sensing probes and reporters were synthesized (see Figure S1 and Table S1 for details). Design of the Microfluidic Chamber. The singlemolecule mechanochemical sensing experiments were performed in a four-channel microfluidic chamber25 in laser tweezers instrument (Figure S2). The microfluidic chamber was prepared by sandwiching a Para-film (BEMIS, Neenah, WI) between two #1 glass coverslips (VWR). The microfluidic pattern was designed by CorelDraw software (Corel Corporation) and imprinted into the Para-film directly by a laser cutter (VL-200, Universal Laser Systems, Scottsdale, AZ). The patterned Para-film and the two coverslips were thermally sealed at 85 °C. The channel thickness (100 ± 5 µm) was defined by that of the film25. Two microcapillary tubes (ID 20 µm, OD 90 µm) were used to transport the anti-digoxigenin beads attached with the DNA construct in the buffer channel and streptavidin coated beads in the target channel. The same capillary tube was used as a separation marker in the conduit (~500 µm) to switch the DNA construct between the target and the buffer channels (Figure S2 inset). Laser Tweezers Instrumentation. Detailed description of the laser tweezers instrument has been reported elsewhere26. In brief, a diode pumped solid-state laser source (DPSS, 1,064 nm in continuous-wave mode, BL-106C, Spectra-physics) was used to generate P- and S-polarized beams for two laser traps. The position of each trap was detected separately using two position-sensitive photodetectors (DL100, Pacific Silicon Sensor). A steerable mirror (Nano-MTA, Mad City Laboratories) was used to control the S-polarized light at the plane conjugate to the back focal plane of a focusing objective (Nikon CFIPlan-Apochromat ×60, NA = 1.2, water immersion, working distance ~320 µm). During experiments, the laser power in IR mirror and IR static were maintained as 233 and 210 mW, respectively, so that the temperature of the polystyrene beads

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increased less than 1˚C27 which may not affect the diffusion of Hg2+ ions in sample. The tension inside the DNA construct was varied by the movement of the steerable mirror using a LabView program (National Instruments Corporation). Single-molecule Mechanochemical Sensing Experiments. To start the sensing experiments, the DNA construct was first immobilized onto the surface of the digoxigenin antibody coated polystyrene beads (diameter: 2.10 µm) through the antibody-antigen complex. The incubated sample was further diluted to 1 mL in a 10 mM Tris buffer with 100 mM KCl (pH 7.4). The streptavidin coated polystyrene beads (diameter: 1.87 µm) were also dispersed into the same buffer and injected into the microfluidic chamber. A 10 mM Tris buffer containing 100 mM KCl and 5 mM EDTA without Hg2+ was flowed in the top (buffer) channel. The same buffer containing Hg2+ without EDTA was injected in the bottom (target) channel. Two separate laser beams were used to trap two different types of beads (see below). By escorting one of the trapped beads close to another using the steerable mirror, the DNA was tethered between the two trapped beads in ~ 5s in the buffer channel. After the tethering, one of the trapped beads was moved away from another with a loading speed of ~5.5 pN/s and the single tether was confirmed by DNA overstretching plateau at 65 pN or a single breakage event in the force-extension (F-X) curves. This rate allowed us to clearly observe the binding of the Hg2+ by unfolding of the hairpin dipsticks (Figure 1). An unfolding event was identified as a sudden change in the endto-end distance during the force ramping. The unfolding force was measured directly from the F-X curves while the changein-contour-length (∆L) due to the unfolding was calculated by the data points flanking the rupture event using a modified extensible worm-like chain (WLC) model (see Supporting Equation S1)28,29 . RESULTS AND DISCUSSION Single-Molecule Mechanoanalytical Real Time Sensing (SMARTS) Device. The SMARTS device is inspired by the trawling mechanism in fishing and the dipstick used to gauge the content level in a container (Figure 1A). Using a trawl net anchored to two moving boats, trawling has been frequently employed in the fishing industry and scientific surveys. The large surface of a trawl ensures efficient netting of desired products in ocean. In the SMARTS device, we will borrow the trawling mechanism for molecular sensing. We will use individual DNA molecules anchored to two optically trapped beads as templates (Figure 1B). A molecular trawl is made of multiple recognition elements in each of the two ssDNA strands serving as two pincers, which facilitate analyte binding. Instead of using boats to move a trawl, here we keep the molecular trawl standstill while forcing the solution to flow by. Noticing dipsticks can check the content level in a container (Figure 1A), our SMARTS device employs molecular dipsticks made of DNA stem-loop structures (hairpins) to determine whether analytes are caught in the molecular trawl (Figure 1B). When the hairpin is subject to ~15 pN exerted by optical tweezers, the stemloop is unraveled30, which is accompanied by the unzipping of the two pincers in the molecular trawl. The unzipping is halted when the dipstick and the trawl encounter mechanical blocks composed of the analyte-recognizer complexes in the

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SMARTS, which signifies the catch of analyte molecules (Figure 2D). The design of molecular trawl is rather versatile. Either protein or nucleic acids can be used in the SMARTS device to specifically recognize chemical analytes. Since DNA is more stable and cost-effective than proteins, we chose individual DNA molecules as templates for the SMARTS. Although specific aptamers31 can be used as recognition elements for various targets ranging from small molecules, macromolecules, to whole cells1,32, for its simplicity and environmental significance, we used T-T mismatch pairs to recognize Hg2+ ions through the T-Hg2+-T complex9.

Figure 1: A single-molecule mechanoanalytical real time sensing (SMARTS) device using the concepts of a fishing trawl and a dipstick. A) A trawl net anchored to two moving boats used to catch fish. Analogous to a dipstick to gauge the content level in a container, a dipstick can report the amount of fish caught in the net. B) Molecular trawling in a SMARTS device that is anchored to two optically trapped beads. The device contains a molecular trawl with multiple recognition elements in each of the two ssDNA strands, which serve as two pincers to catch analytes (green hexagons). A DNA hairpin acts as a molecular dipstick to report the amount of the bound analytes via mechanochemical unfolding events. Target binding in the SMARTS leads to a reduced signal in change-in-contour-length (∆L, see Figure 2D for calculation).

Molecular Trawling in the SMARTS. To evaluate the efficacy of the molecular trawls for Hg2+ detection, we varied the number of thymines in each trawl while using the same molecular dipstick (hairpin) reporter with a 20-bp stem and a tetrathymine (T4) loop (Figures 2A and S1, see Table S1 for sequence). To determine whether Hg2+ binds to the T4 hairpin loop, we first prepared a control construct without any T-T mismatch in the molecular trawl (Figure S1 and Table S1). We obtained similar hairpin folding and unfolding transitions in a microfluidic chamber that contains a 10 mM Tris buffer (pH 7.4) and a 1 mM Hg2+ solution (Figure S2). The identical change-in-contour-length (∆L) and similar rupture force values

of the hairpin reporter in the two solutions (Figure S3) indicated that either Hg2+ does not bind to the T4 loop, or the binding is too weak to interfere with the mechanochemical signal used for sensing. Therefore, in the following SMARTS designs, we keep the T4 loop in the hairpin reporter.

Figure 2: Molecular trawling for Hg2+ binding. A) Molecular trawls made of 1T, 2T, 4T, 10T, or 50T (Thymine) in each of the two anti-parallel DNA strands. Hg2+ binds to the probe via the T-Hg2+-T complex. The same hairpin dipstick (a 20-bp stem with a 4-nt loop, Figure S1) was used for each trawl. B) Typical force-extension (F-X) curves of the 10T probe in a 10 mM Tris buffer with 100 mM KCl and 5 mM EDTA (pH 7.4, green), and in a 1 nM Hg2+ channel (the ~17 pN features in the red and blue traces show the Hg2+ binding while the 30 pN feature in the blue trace depicts Hg2+ ejection). Colored and black traces represent stretching and relaxing curves, respectively. Curves are offset in x-axis for clarity. C) ∆L histograms for the 10T probe with (red) and without (green) 1 nM Hg2+ in the Tris buffer (N = 332). D) Mechanism of Hg2+ binding to the 10T SMARTS device. E) Recycling of the 10T probe between the Tris buffer and the 1 nM Hg2+ channel. F) Limit of detection (LOD) of Hg2+ for different probes. Error bars were calculated from the predicted LOD for 50% binding of the sensing probe.

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We started with a molecular trawl that contains one thymine in each ssDNA pincer. However, binding of Hg2+ as high as 1 mM was not observed probably due to the weak affinity of the Hg2+ to the single T-T pair. Using a trawl with two or four adjacent thymines (the 2T and 4T probes in Figure 2A) in each pincer revealed binding events in force-ramping experiments (100 µM and 100 nM Hg2+ for the 2T and 4T, respectively, see FigureS4), but not in force-clamp experiments in which a constant tension was maintained in the backbone of each SMARTS device (see FigureS5). Without Hg2+, the hairpin dipstick transits rapidly between folded and unfolded states at constant force (or mechanoescence24, see the 10T probe as an example in Figure S5A). With 1 µM Hg2+, the mechanoescence became significantly slower (Figures S5B and S6), which is understandable as the T-Hg2+-T complexes must dissociate prior to the full unfolding of the hairpin. However, before the formation of the T-Hg2+-T complexes during refolding, the loosely associated Hg2+ ions may attract the two pincers together, which reduces transition kinetics due to the entropic penalty (Figure S7). In addition, the F-X curves showed transitions with reduced change-in-contour length (∆L) or without any feature (Figure 2B&C). When we reduced Hg2+ concentration, it became more difficult to observe the change in mechanoescence due to the fact that during the folding-unfolding transitions, two pincers in the molecular trawl came closer only momentarily, decreasing the chance to catch Hg2+ ions. Therefore, we used F-X detection mode for low analyte concentrations. In this mode, the ∆L was measured from the F-X curve each time a SMARTS device was mechanically stretched. It is clear from Figure 2C that ∆L values for 1 nM Hg2+ are significantly smaller than the buffer, confirming the catch of the Hg2+ by the SMARTS in the target channel. At even lower analyte concentrations, however, not every FX curve shows evidence of catching Hg2+ ions. This observation increases the uncertainty in the Hg2+ sensing if ∆L histograms, which reflect average results, are compared between the buffer and the Hg2+ channels. We therefore opted to determine the binding of Hg2+ to the molecular trawl from individual F-X traces. First, we obtained more than 15 ∆L measurements in the buffer channel to calculate average ∆Laverage with a standard deviation (σ). After the SMARTS was transported to the Hg2+ channel (Figure S2), the binding of the Hg2+ was depicted by the ∆L values less than (∆Laverage-3σ) (Figures 2E and S8A) at the 99.7% confidence level. The detection limit of Hg2+ was set when there was no sign of Hg2+ binding in 20 minutes. It is noteworthy that the (∆Laverage-3σ) does not reveal binding of the Hg2+ to the distal end of a T-T probe, therefore, such a determination gives an upper estimation of the LOD. In quantification of Hg2+ concentrations, this bias constitutes a systematic error which can be rectified by a calibration curve (see Figure 6B below). As shown in Figure 2F, when we increased the number of thymines in molecular trawls, the detection limit reduced. For a 10T SMARTS device, the detection limit was 100 pM. The detection limit reduced to 10 pM when a 50T device was used (Figure 2F). It is noteworthy that the SMARTS device can be transported back to the buffer channel that contains 5 mM

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EDTA to remove the Hg2+ bound to the probe (Figures 2E and S8). During experiments, more than 12 recycles were performed without deteriorating sensing capability of the 10T SMARTS device. Alternatively, the bound Hg2+ can be removed by increasing the tension (blue traces in Figures 2B and S4B) for recycling. Molecular Dipsticks in the SMARTS. The hairpin dipstick we used for the 50T SMARTS device contained only 44 nucleotides (a 20-bp stem and a 4-nt loop, see Table S1). This dipstick may not be able to probe the Hg2+ binding in the pincers longer than the 22T used in the molecular trawl, which limits the sensing area of the SARMTS device (Figure S9). In the next step, we increased the length of the hairpin dipstick to 108 nucleotides (a 52-bp stem and a 4-nt loop, see Table S1) so that it can reach to the bottom of the 50T* molecular trawl (Figure 3A). To our disappointment, almost identical ∆L histograms were obtained with or without Hg2+ ions (Figure 3B&C, see Figure S10 for ∆L measurement), indicating the binding of Hg2+ was not observed in 1 µM Hg2+. The unfolding transition (Figures 3B & S10) revealed a ~ 19 pN plateau during which hairpins started to unzip. Such a plateau provided an opportunity for Hg to dissociate from the T-Hg2+-T complex if the time window is long enough (Figure 3A). To verify this, we performed force-ramping experiments on the 10T probe by reducing the loading rate from 5.5 pN/s to 0.55 pN/s in 100 nM Hg2+. As expected, the binding of Hg2+ was less obvious from the ∆L measurements when the loading rate was reduced (compare blue and green traces in Figure 3D&E).

Figure 3: Molecular dipsticks to report Hg2+ binding. A) Sensing mechanism of the 50T* probe that contains 50 T-T pairs for Hg2+ recognition and a 108-nt dipstick hairpin (see Table S1 for sequence) for target reporting. B) Typical F-X curves for the 50T* probe in a 10 mM Tris buffer with 100 mM KCl and 5 mM EDTA (pH 7.4, green) or in 1 µM Hg2+ (red). Colored and black traces represent stretching and returning curves, respectively. C) ∆L histograms for the 50T* probe in the Tris buffer (green) or 1 µM Hg2+ (red) (N = 220). D) Typical F-X curves for the 10T SMARTS in the Tris buffer (green, loading rate 5.5 pN/s) or in 100 nM Hg2+ with 5.5 pN/s (red) and 0.55 pN/s (blue) loading rates. E) Corresponding ∆L histograms for the 10T device in D) (N = 156). Notice curves in (B) and (D) are offset in x-axis for clarity.

It has been reported that when a hairpin is long, the unfolding transition of the hairpin is no longer cooperative30 . In-

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stead, the saw-tooth unfolding features at ~15 pN could appear, leading to spontaneous dissociation of the Hg2+ ions as observed above. To serve as an effective reporter for analyte binding in the molecular trawling, long hairpin dipsticks should be avoided. We therefore designed two short hairpin dipsticks to gauge the binding of Hg2+ (Figure 4A inset). To effectively probe towards the bottom of the 50T trawl, the two hairpins were designed to have a total of 88 nucleotides with a 10-nt spacer in between (Table S1). Binding of Hg2+ anywhere in the 50T trawl is expected to reduce the change-incontour-length (∆L) of the unfolding events as the T-Hg2+-T complex has been shown to block the unraveling of the dipstick hairpins (see above). The mechanical unfolding of the probe thus prepared (50T**) showed two unzipping features for the two hairpin dipsticks (Figure 4A). To determine the binding of Hg2+, we summed up ∆L values of these two unfolding events with or without Hg2+ (Figure 4B). In the 100 fM Hg2+ channel, ∆Laverage is smaller than the buffer channel. Interestingly, the distribution of ∆L values in Hg2+ is similar to that in buffer. This suggests that Hg2+ is more likely to bind in the middle of the trawl, in which thymines in one pincer can still survey a large area for Hg2+ while maintaining a close distance to the other pincer for T-Hg2+-T formation.

Figure 4: Mechanoanalytical device with two hairpin dipsticks (50T**). A) Representative F-X curves show two folding/unfolding events for the two hairpins at 15-20 pN in the Tris buffer (green) and in 100 fM Hg2+ (red and blue). The ejection of bound Hg2+ is depicted at ~30 pN in the blue trace. Colored and black traces represent stretching and returning curves, respectively. Inset shows the diagram for the 50T** device. Curves are offset in x-axis for clarity. B) Cumulative ∆L histograms for the unfolding events of both hairpins in the Tris buffer (green) and in 100 fM Hg2+ (red) (N=308). C) Probe recycling by switching the 50T** device between a 10 mM Tris buffer (pH 7.4, 100 mM KCl and 5 mM EDTA) and the 100 fM Hg2+ channel.

As discussed previously, at lower Hg2+ concentrations, it is less likely Hg2+ is bound to the SMARTS device each time the force ramping detection was performed. In fact, higher ∆Laverage values with wider ∆L distributions were observed at lower Hg2+ concentrations (Figure S11), suggesting it is less likely for Hg2+ to bind in the trawl closer to the hairpin dipstick. Thus, comparison of ∆L histograms is not effective to deter-

mine the binding of Hg2+ unambiguously. Similar to the approach described above, we followed the ∆L for individual traces. Hg2+ is considered to be detected with 99.7% confidence level if ∆L measured in the Hg2+ channel is less than (∆Laverage - 3σ), where ∆Laverage is the average ∆L in the buffer channel and σ is the standard deviation (Figures 4C and S8B). This calculation allowed us to determine an upper detection limit of 1 fM Hg2+ in 20 minutes using the 50T** probe (see discussion above). Similar to the 10T device, by transporting the 50T** between an EDTA-containing buffer and a Hg2+ channel (Figures 4C and Figure S8B) or using ejection force (blue trace in Figures 4A and S12) to remove Hg2+ ions, the 50T** sensor can be recycled for subsequent tasks without deterioration in the sensing capability. Compared to the most sensitive Hg2+ detection (~ 100 fM)12, our approach is at least 2 orders of magnitude lower in detection limit with simpler sample preparation. It is noteworthy that the detection limit can be further improved by using a longer detection window, a longer molecular trawling, or more trawls in the SMARTS. Given that affinity (Kd) of Hg2+ binding to a single T-T pair is in the µM ranges,33 it is surprising that femtomolar Hg2+ concentrations (9 orders of magnitude lower than the Kd) were detected here. The superior sensitivity can be ascribed to four factors. First, the T-T pairs in the SMARTS device are arranged in a polyvalent manner, which has a well-known entropic effect to increase the binding affinity34. Second, the multiple T-T pairs raise the effective local concentrations, increasing the association rate for Hg2+ binding. Third, the 50T** device can reach deeper into a solution, increasing the interaction area for Hg2+ binding within fixed time. Finally, the single-molecule detection coupled with realtime response of the analyte binding provides extraordinary signal-to-noise level both spatially and temporally, which lays the foundation for all above factors. Specificity of Hg2+ Detection. Before testing field samples using these SMARTS devices, we first evaluated its specificity. Using the approach described in Figure 2, we measured the detection limits with the 10T probe for the Co2+, Ni2+, Fe2+, Cu2+, Hg2+, Cd2+, Mn2+, Pb2+, and Zn2+ ions. Except for the Pb2+ and Ni2+, which showed 1 µM detection limits in 20 minutes, all other ions did not bind to the probe at this concentration. The selectivity for Hg2+ (100 pM for the 10T probe) is therefore at least 4 orders of magnitude higher than the closest competitors, Pb2+ and Ni2+ (Figure 5).

Figure 5. Specificity test of Hg2+ versus other cations in the 10T SMARTS device (n ≥ 12). Error bars were calculated from the predicted LOD for 50% binding of the sensing probe.

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Quantification of Hg2+ in the Field Samples. To quantify the concentration of Hg2+ using the SMARTS devices, we constructed calibration curves in which the observation time for the first binding event was plotted against known Hg2+ concentrations. Immediately after the sensing probe was switched from the buffer channel to the target channel within 10s, F-X curves were recorded at 20s interval. The time for Hg2+ binding was determined when ∆L of the hairpin reporter was smaller than (∆Laverage-3σ) as discussed above. Such an approach allowed a temporal resolution of 20s, which can be improved by using a short time interval to record F-X curves.

Figure 6. Detection limit of 50T** SMARTS device. A) Dynamic range (100 pM - 1 fM) and B) linear range (100 pM100 fM, with r2 = 0.9665) for Hg2+ detection in the 50T** device. The cyan points depict Hg2+concentration measured from pristine water (A) and 40 times concentrated pristine water sample from Lake Erie. For the 50T** device with two hairpin dipsticks, we found the observation time increased inversely with concentration in the dynamic range of 1 fM-100 pM (Figure 6A), and linearly with the concentration range of 100 pM – 100 fM. Above 100 pM, the observation time for the first binding was too short to accurately determine the concentration. However, using the 10T SMARTS device, the dynamic range for high Hg2+ concentrations (100 pM – 10 µM) can be well established (Figure S13). With the specificity and the quantification procedures firmly established, we then measured Hg2+ concentration in Lake Erie. We first filtered pristine Lake Erie water (Huntington Beach, 08.20.2015) with a 200 nm-pore polystyrene filter, which was followed by direct injection into the microfluidic chamber (Figure S2, please see supporting information for details). By recording the time for the first binding events, we found 1.2 ± 0.4 fM Hg2+ (2.4×10- 4 ± 0.8×10- 4 ppt) in the field sample using exponential fitting equation (Figure 6A). Concentrating the same sample 40 times, the Hg2+ in Lake Erie was determined to be 92 ± 20 fM (equivalent to 2.3 ± 0.5 fM Hg2+(4.6×10- 4 ± 1.0×10- 4 ppt) in the pristine sample) using linear fitting equation (Figure 6B). The small deviation between the two measured values probably reflects that facts that measurement in the linear range (Figure 6B) is more accurate with respect to that close to the boundaries of the dynamic range (Figure 6A). The close values also depict the reliability of the quantification method for the mechanoanalytical device. We expect the linear range of the SMARTS device to quantify Hg2+ ions can increase by increasing the length of the dipstick and the molecular trawls. This will facilitate the accurate quantification of the trace level of Hg2+ ions in the field water sample without pre-concentration steps.

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CONCLUSIONS In summary, we have successfully constructed first-in-class single-molecule mechanoanalytical real time sensing (SMARTS) devices. With the incorporation of molecular trawls for Hg2+ recognition and molecular dipsticks to report Hg2+ binding, an unprecedented detection limit of 1 fM (2 × 10- 4 ppt) Hg2+ was achieved in 20 minutes, which is two orders of magnitude lower than the best Hg2+ detection method reported.12 This detection limit is nine orders of magnitude lower than the Kd, which represents an improvement of three orders of magnitude compared to the amplification based biosensing methods35,36. The SMARTS can be versatile to detect a range of different analytes, which include small molecules as well as proteins and nucleic acids. The novel concept of mechanoanalytical devices described here provides a key contribution to the mechanoanalytical chemistry23, a new field that uses mechanochemical principles for chemical analyses. However, other force based techniques such as Atomic Force Microscopy (AFM)37 can be employed for this type of sensing as well. Due to the single-molecule nature, the throughput of the sensing is low. This can be significantly increased by incorporating high-throughput magnetic tweezers38. On the other hand, the complex instrumental setup can be addressed by using mini optical tweezers that are readily automated39.

ASSOCIATED CONTENT Supporting Information Synthesis of the DNA construct, Experimental setup, Single molecule mechanochemical sensing, Development of molecular trawling in the SMARTS, Method of pre-concentration and quantification of Hg2+ ions in the field samples. Table S1 and Figures S1-S13.

AUTHOR INFORMATION Corresponding Author * E-mail: [email protected]

Author Contributions The manuscript was written through contributions of all authors.

Notes The authors declare no competing financial interest.

ACKNOWLEDGMENT We would like to thank the NSF CHE-1609514 for financial support.

REFERENCES (1) Cammann, K. Z. Anal. Chem. 1977, 287, 1-9. (2) Thevenot, D. R.; Toth, K.; Durst, R. A.; Wilson, G. S.; Biosens. Bioelectron. 2001,16(1-2),121-31. (3) Anthony, T.; George, W.; Isao, K. Biosensors: Fundamentals and Applications; Oxford University Press: Oxford, UK, 1987. (4) Lequin, R. M. Clin. Chem. 2005, 51, 2415-2418. (5) Shrestha, P.; Mandal, S.; Mao, H. Mechanochemical Sensing, RNA and DNA Diagnostics, Springer International Publishing. 2015, 241-258. (6) Shrestha, P.; Mandal, S.; Mao, H. ChemPhysChem 2015, 16, 1829-1837.

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(7) Ashkin, A.; Dziedzic, J. M.; Bjorkholm, J. E.; Chu, S. Opt. Lett. 1986, 11, 288-290. (8) Tchounwou, P. B.; Ayensu, W. K.; Ninashvili, N.; Sutton, D. Environ. Toxicol. 2003, 18, 149-175. (9) Miyake, Y.; Togashi, H.; Tashiro, M.; Yamaguchi, H.; Oda, S.; Kudo, M.; Tanaka, Y.; Kondo, Y.; Sawa, R.; Fujimoto, T. J. Am. Chem. Soc. 2006, 128, 2172-2173. (10) U.S. EPA. National Primary Drinking Water Regulations. US EPA, 2009. http://water.epa.gov/drink/contaminants/index.cfm#List. (11) Cizdziel, J. V.; Gerstenberger, S. Talanta 2004, 64, 918-921. (12) Suvarapu, L. N.; Baek, S. O. J Anal Methods Chem 2015, 372459, 5. (13) Jenner, G. A.; Longerich, H. P.; Jackson, S. E.; Fryer, B. J. Chem Geol 1990, 83, 133-148. (14) Li, Y.; Hu, B. Spectrochim. Acta B 2007, 62, 1153-1160. (15) Ghaedi, M.; Reza Fathi, M.; Shokrollahi, A.; Shajarat, F. Anal. Lett. 2006, 39, 1171-1185. (16) R. P. Mason, W. F. F., H. Hurley, A. K. P. L. Donaghay,; A. K. Hanson Jr., J. M. S. Oceanography 1993, 38, 1227. (17) Kong, R.-M.; Zhang, X.-B.; Zhang, L.-L.; Jin, X.-Y.; Huan, S.Y.; Shen, G.-L.; Yu, R.-Q. Chem. Commun. 2009, 5633-5635. (18) Yang, H.; Zhou, Z.; Huang, K.; Yu, M.; Li, F.; Yi, T.; Huang, C. Org. Lett. 2007, 9, 4729-4732. (19) Gao, C.; Huang, X.-J. Trends Anal. Chem. 2013, 51, 1-12. (20) Jiang, Z.; Fan, Y.; Chen, M.; Liang, A.; Liao, X.; Wen, G.; Shen, X.; He, X.; Pan, H.; Jiang, H. Anal. Chem. 2009, 81, 54395445. (21) Nolan, E. M.; Racine, M. E.; Lippard, S. J. Inorg. Chem. 2006, 45, 2742-2749. (22) Yang, Y.-K.; Yook, K.-J.; Tae, J. J. Am. Chem. Soc. 2005, 127, 16760-16761. (23) Koirala, D.; Shrestha, P.; Emura, T.; Hidaka, K.; Mandal, S.; Endo, M.; Sugiyama, H.; Mao, H. Angew. Chem. Int. Ed. 2014, 53, 8137-8141. (24) Mandal, S.; Koirala, D.; Selvam, S.; Ghimire, C.; Mao, H. Angew. Chem. Int. Ed. 2015, 54, 7607-7611. (25) Koirala, D.; Yu, Z.; Dhakal, S.; Mao, H. J. Am. Chem. Soc. 2011, 133, 9988-9991. (26) Mao, H.; Luchette, P. Sens. Actuators B 2008, 129, 764-771. (27) Selvam, S.; Koirala, D.; Yu, Z.; Mao, H. J. Am. Chem. Soc. 2014, 136, 13967-13970. (28) Baumann, C. G.; Smith, S. B.; Bloomfield, V. A.; Bustamante, C. Proc. Natl. Acad. Sci. USA 1997, 94, 6185-6190. (29) Yu, Z.; Mao, H. Chem. Rec. 2013, 13, 102-116. (30) Woodside, M. T.; Behnke-Parks, W. M.; Larizadeh, K.; Travers, K.; Herschlag, D.; Block, S. M. Proc. Natl. Acad. Sci. USA. 2006, 103, 6190-6195. (31) Yangyuoru, P. M.; Dhakal, S.; Yu, Z.; Koirala, D.; Mwongela, S. M.; Mao, H. Anal. Chem. 2012, 84, 5298-5303. (32) Sefah, K.; Shangguan, D.; Xiong, X.; O'Donoghue, M. B.; Tan, W. Nat. Protocols 2010, 5, 1169-1185. (33) Torigoe, H.; Ono, A.; Kozasa, T. Chemistry 2010, 16, 1321813225. (34) Mammen, M.; Choi, S.-K.; Whitesides, G. M. Angew. Chem. Int. Ed. 1998, 37, 2754-2794. (35) Liu, M.; Zhang, W.; Zhang, Q.; Brennan, J. D.; Li, Y. Angew. Chem. Int. Ed. 2015, 54, 9637-9641. (36) Zhang, S.; Garcia-D'Angeli, A.; Brennan, J. P.; Huo, Q. Analyst 2014, 139, 439-445. (37) Binnig, G.; Quate, C. F.; Gerber, C. Phys Rev Lett 1986, 56, 930-933. (38) De Vlaminck, I.; Henighan, T.; van Loenhout, M. T. J.; Pfeiffer, I.; Huijts, J.; Kerssemakers, J. W. J.; Katan, A. J.; van Langen-Suurling, A.; van der Drift, E.; Wyman, C.; Dekker, C. Nano Lett 2011, 11, 5489-5493. (39) Stephenson, W.; Wan, G.; Tenenbaum, S. A.; Li, P. T. J. Vis. Exp. 2014, 90, 51542.

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TOC

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Mechanochemical Sensing of Single and Few Hg(II) Ions Using

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