Peptide Nanocarriers for Detection of Heavy Metal Ions Using

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Peptide Nanocarriers for the Detection of Heavy Metal Ions Using Resistive Pulse Sensing Imogen Heaton, and Mark Platt Anal. Chem., Just Accepted Manuscript • DOI: 10.1021/acs.analchem.9b02353 • Publication Date (Web): 02 Aug 2019 Downloaded from pubs.acs.org on August 5, 2019

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Analytical Chemistry

Peptide Nanocarriers for the Detection of Heavy Metal Ions Using Resistive Pulse Sensing Imogen Heaton, Mark Platt1*

1. Department of Chemistry, Loughborough University, Loughborough, Leicestershire. LE11 3TU, UK. *[email protected]

1 ACS Paragon Plus Environment

Analytical Chemistry 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

Abstract The use of nanocarriers within resistive pulse sensing facilitates the detection and quantification of analytes. To date the field has been dominated by polyionic carriers or nanomaterials. Together they combine the recognition elements of a ligand with a stable support facilitating the sample handling, analysis times and multiplex detection. Here we develop the use of peptide functionalised superparamagnetic nanocarriers to extract and quantify metal ions in solution. The interaction between Nickel and the peptide ligand is measured as a change in translocation velocity of the carrier. The magnitude of change is proportional to the concentration of the metal ions in solution. Unlike DNA aptamers where a change in the tertiary structure and the folding of the polyanionic back bone influences the carrier velocity, the peptides here had a lower net charge under the assay conditions. To try and enhanced the signal, we engineered charged groups within the peptide to explore the effects on the signal. In all cases the metal ion binding dominated the velocity of the carrier. The assay was shown to work across three orders of magnitude and can detect Ni2+ in the presence of some other heavy metal ions. We demonstrate this by quantifying Ni2+ in both tap and pond water. The work allows for future multiplexed sensing strategies using both peptides and DNA aptamers in resistive pulse sensors.


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The field of resistive pulse sensing, RPS, has a range of applications in materials characterisation,1–6 quantification of ligand-target interactions,7–9 mimicking biological pathways and biosensing.10–12 This scalable technology allows single analyte resolution from small molecules up to single cells.5 It has been applied to sensing analytes across the omics from metabolites,13 proteins,9,14,15 DNA sequencing,16–18 epigenetics19,20 and cellular vesicles.21,22 The transport of any analyte through the channel is controlled by tuning the applied potential, pore wall charge, pore size, supporting electrolyte concentration and composition, with a further degree of selectivity by modifying the pore walls with selective ligands15,23–26. DNA aptamers are one example of a ligand used in RPS. They are sequence specific single strand nucleic acids, that bind to target analytes via the formation of a specific tertiary structure.27–29 DNA Aptamer based RPS strategies are three-fold. The first is the modification of the pore mouth/ walls with aptamers to induce a change in current flow upon the binding.30 Second, measuring the translocation rates and velocities of the aptamer sequences through the pores, finally the use of nanomaterials as carriers for multiple aptamers.9,14,31–36 The carrier acts as a support for multiple ligands enhancing the signal, facilitating multiplexed assays, or in cases where the carriers are magnetic particles they can facilitate in the extraction and pre-concentration of the target.37,38 Once immobilised onto a carrier, the binding of the analyte to the aptamer can be measured via two mechanisms. The first is a change in size or shape of DNA strand.13,36 The second is a change in charge density around the carrier.9,13 This is measured through a change in translocation velocity of the carrier and can provide quantitative information.9,14,39 Both strategy are tagless, not requiring the analyte to be labelled in any way. In the first strategy the resolution of the RPS must be such that they can resolve small changes in the size of the DNA. Whilst this is possible with RPS, it means the pore must be comparable in size to the carrier/ aptamer, resulting in a decrease in translocation frequency. The second requires a large change in the tertiary structure on the outer edge of the carrier, figure 1a.40 To date RPS peptides are typically the target and not the capture ligand.41 Initial work developing peptide ligands shows that binding to large analytes can causes changes in translocation rates and velocities.42 There has not yet been a strategy to show a peptide giving a quantitative sensor for metal ions. Here we present the first use of peptide carriers binding to metal ions in solution. We demonstrate this simple concept using a Histidine, His, tagged peptides on the surface of a super paramagnetic nanocarrier, SPN. The His tag binds to Ni2+ ions in solution, and the binding of the metal ions causes a change in charge density around the carrier. This is measured via change in translocation velocity through the RPS. The signal is specific to Ni2+ ions even in the presence of other ions and complex sample matrixes. We test this in both pond water sampled from the university campus and tap water.



Figure 1: a) Example of baseline current and blockade events caused by the carriers translocating the pore, insert showing 60-100 seconds. b) Schematic of the nanocarrier surface and the ligands used. The sections inner and outer refer to the part of the DNA or peptide closest and furthest from the particle surface. c) Schematic of a carrier traversing the RPS device, and the signal. The blockade magnitude ip, and Full width half maximum are shown.

Experimental Chemicals and reagents. Nickel sulphate hexahydrate, chromium chloride hexahydrate, iron chloride, cobalt chloride, and TWEEN 20 were purchased from Sigma-Aldrich, UK. Calcium chloride and potassium chloride were purchased from Fisher Scientific, UK. Carboxylated polystyrene particles were purchased from Bangs Laboratories US, denoted as CPC200 (mode diameter 210 nm, measured concentration 1 x 1012 particles mL-1). Nanopores were purchased from Izon Science Ltd. Reagents were prepared in deionised water (Elga PureLab), with a resistance of 15 MΩ cm. The pH of background electrolyte was altered using KCl and KOH, and measured using a Mettler Toledo easy five pH meter, with a Mettler Toledo InLab micro electrode. Custom peptides and DNA oligonucleotides. Custom peptides were purchased from Mimotopes, Australia, in lyophilised form. They were purified and checked by reverse phase HPLC by the manufacturer. The peptide details are; biotinRTRTRTRTR-OH, called P1, biotin-GTGTGTGTG-OH, P2, biotin- DTDTDTDTD-OH, P3, biotin-RTRTRTRTRHHHHHH-OH, P1H, biotin-GTGTGTGTGHHHHHH-OH, P2H and biotin- DTDTDTDTDHHHHHH-OH, P3H. A custom 10mer DNA oligonuclideotide 4 ACS Paragon Plus Environment

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was purchased from Sigma-Aldrich, in lyophilised form, purified using reverse-phase cartridge purification by the manufacturer. The oligonuclideotide ordered was biotinTTTTTTTTTT. Particle functionalization. Streptavidin-modified magnetic particles were purchased from Ademtech, France. The streptavidin-modified particles were diluted to 1 x 1010 particles mL-1, using 50 mM KCl solutions with 0.05% TWEEN 20. They were vortexed to ensure they were monodispersed with no aggregates before the different peptides or DNA were added in excess. Binding coverage was calculated using the manufacturers calculated biding capacity. The samples were placed on a rotary wheel at room temperature for 30 min. Samples were then placed on a MagRack (life science) until a clear cluster of particles was seen, the buffer was removed and replaced with an equal volume to make a stock solution. RPS set up. A qNano (Izon Science Ltd, NZ) was used to complete all the measurements for this study. A qNano uses data capturing software (Izon Control Suite v3.3) to record the particles as they traverse the pore. The lower fluid cell contained 80 µL of KCl solution and the upper fluid cell contained 40 µL sample solution. After each measurement was taken the nanopore was cleaned by first rinsing the upper fluid cell with background buffer before the buffer was removed and replaced multiple times. The qNano was typically operated with a positive bias, i.e. the positive electrode in the lower fluid cell and the ground electrode in the upper fluid cell, so the particles traverse the pore towards the positive electrode unless otherwise stated. For all experiments a NP200 nanopore was used, more than 200 particles where measured for each sample. To account for any manufacturing variation between pores the baseline current was kept constant throughout all experiments, and the stretch was slightly changed to ensure this, with max 10% difference in baseline between sample runs. Each pore was first characterised using the calibration particles so dayto-day differences could also be accounted for. Binding time. P1H, P2H, and P3H stocks were made up as above, from each two 100 µL aliquots were taken out and nickel was added to one to make 600 nM solutions. These samples were then made up to 200 µL with 50 mM KCl buffer. The samples were then placed on a rotary and left for 5, 10, 15 and 30 min. They were then taken off vortexed and sonicated before being analysed. Flipped Voltage. The qNano was set up with either a positive bias, i.e. the positive electrode in the lower fluid cell and the negative electrode in the upper fluid cell, with an applied voltage of 0.82 V or a negative bias with an applied voltage of -0.82 V and left for 30 minutes. Metal ion interferences. Metal ion stocks solutions were prepared from CaCl2, CrCl3, FeCl3 and CoCl2. They were then added to the P2H particles to give a final metal ion concentration of 600 nM. A mixture of metal ions containing calcium, chromium, iron and nickel was also prepared. Samples were placed on a rotary wheel for 15 min and prior to analysis were vortexed and sonicated. Environmental water samples. Water samples were collected from a lab tap, an outdoor pond next to the chemistry department and deionised water was used as a 5 ACS Paragon Plus Environment


blank. A stock of P2H particles were prepared as before. To each environmental sample (180 L) peptide functionalised particles (10µL), and 1M KCl 0.5% TWEEN 20 (10 µL) were added. The samples when required were also spiked with nickel to give a final concentration of 600 nM of nickel. The samples were placed on a rotary wheel for 15 minutes, and vortexed and sonicated prior to analysis. Particle velocity. Particle velocity through the pore was calculated from the pulse width. As particles translocate the nanopore they produce a pulse, the maximum magnitude of this pulse is recorded as T1.0 as this magnitude decreases multiple time points are recorded correlating to T0.9, T0.8, T0.7 etc. here we use the reciprocal of the value at T0.5 (width of the pulse at 50% peak height) to determine relative particle velocity. These values were then normalised to either calibration particles (CPC200s) or blank peptide functionalised particles ran on the same day, using the same nanopore and experimental conditions. This was done to account for any differences in measured velocity between different pore and different days. Where error bars are shown they represent one Stdev from the average of triplicate runs where a minimum of 200 event are recorded in each run. Each run is carried out on a set of particles that have been functionalised prior to the assay. Zeta potential measurement: Zeta potential of the six peptide functionalised particles was asses using a laser dynamic light scattering (DLS) instrument, Zetasizer nano ZSP, Malvern Instruments LtD, UK. The particles were prepared as above in 50 mM KCl buffer to pH 7 before being measured on the DLS. The samples were added to a folded capillary cell (DTS1060) and ensured to completely cover the electrodes. The cell was equilibrated at 25 oC for 5 s prior to analysis, and three replicated were taken.


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Results and discussion Within RPS experiments each translocation of an analyte/ carrier through the nanopore produces a pulse. The magnitude of the pulse, known as the pulse magnitude, ip, and is related to the volume of the carrier. The width or full width half maximum, FWHM, of the pulse relates to its velocity, figure 1b.43 In the absence of convection, the velocity of the carrier can be proportional to the surface charge.43,44 We have shown how the length of DNA, its packing density and structure can be measured on SPNs via RPS, and that the velocity can be converted into a zeta potential.43 If the zeta potential is not needed, then the velocity can be reported as a relative velocity with respect to a blank or control particle. Here we do not convert the velocity to a zeta potential. Thus we report the velocity as 1/T0.5 values, which correspond to the width of the pulse as half height, similar to the FWHM.32 The modification of a carrier surface with peptides, DNA and proteins is detectable via the change in velocity. To illustrate this, we show the relative velocity of a carrier with no peptide on its surface, Blank, a peptide coated particle, Peptide, and as a comparison a 10mer DNA coated particle, DNA, in figure 2a. The peptide here was short amino acid sequence termed (P3) from table 1. It had an isoelectric point of 0 and a charge at pH 7 of -6.0, its velocity through the RPS was greater than that of the blank and slower than the DNA which was expected. The length of the DNA and peptide is comparable, estimated as 6 and 3.8 nm respectively.

Figure 2: a) Blank SPN, vs. different particle functionalisation. Measured in 50 mM KCl buffered to pH 7, 1.06 V. b) Comparison of peptide sequences without, blue, and with, red, his tag on. P1, P2 and P3 (blue) ran in 50 mM KCl buffered to pH 7,. P1H, P2H and P3H (red) ran in 50 mM KCl buffered to pH 7. Each sample was ran in triplicate, and more than 200

particles were measured each time. Errors bars are one standard deviation from the mean. In this setup a positive bias is applied to the electrode on the opposite side of the nanopore to where the sample is placed. Under this configuration the negatively charged carriers are driven through the nanopore at a frequency and speed 7 ACS Paragon Plus Environment


proportional to the surface charge. It should be noted here that within the RPS platform the upper fluid cell is on top of the pore and there is always a contribution of convection from the fluid flowing between the upper and lower fluid cell along with the electrophoresis. This configuration can aid in the detection of particles of low charge.45 A range of peptides of comparable length and varying charge densities were then immobilised onto different SPN’s. The relative velocity of these functionalised carriers through the RPS are shown in figure 2b. We attempted to use three different peptides of varying charges. The hypothesis was to create a neutral, positive or negative peptide and were chosen to see if the experimental setup could distinguish between the different designs. As can be seen in figure 2b (blue), all the peptides translocated at comparable speeds. Complimentary measurements to determine the zeta potential of the particles can be found in figure S1. Whilst the peptides have a specified charge at pH 7, its clear from figure S1 that the underlying surface charge of the carrier, might contribute to the measured velocity. As we wished to use histidine tagged peptides to extract and detect Ni2+, we added 6 x His to these short peptides producing three additional peptides listed in table 1. The velocities of the His variants are shown in figure 2b (red), and zeta potentials in figure S1. The additional of the His tag resulted in all particles increase in their relative velocities, this was expected given the charge in the His section of -0.4.

Table 1: List of peptide names, sequences, isoelectric points and charge at pH7 calculated from pepcalc.com.

The binding of Ni2+ to His tags is well known, a fact that is exploited in protein purification. In the presence of Ni2+ in solution it would be expected that the 6 x His on the SPNs surface would bind to the ions. It was hypothesised that this should cause a change in charge density around the carrier which could be measured through a change in velocity through the pore. P2H modified SPN’s were added to a solution containing Ni2+ and their relative velocities measured. As can be seen in figure 3ai, in the presence of the Ni2+ ions the particles velocity decreases from 1.00 to 0.81 ms-1. This decrease in speed is attributed to the binding of the Ni2+ to the peptide, causing 8 ACS Paragon Plus Environment

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the particle to have a decrease in negative charge. Thus, when a positive bias is applied to the electrode on the underside of the RPS the electrophoretic speed of the carrier decreases. Conversely if a negative potential is applied to the lower fluid cell, the carriers increase in velocity in the presence of Ni2+, as shown in figure 3aii. Velocities of carriers with different functionalities in the presence/ absence of Ni2+ are shown in figure S2. The data in figure S2 illustrates how the change in velocity in the presence of Ni2+ is specific to the His tag. Our long-term goal is to combine peptide and DNA aptamers in one assay developing multiplexed sensors, thus most of the work here continues to use the positive bias electrode configuration. To confirm enough time had been given for the Ni2+ to bind to the peptides a time dependent study was undertaken, the results are shown in figure 3b. The result indicates that after 15 minutes the particle velocity decreased and remained constant for P1H and P2H, but no consistent change in velocity could be measured for peptide P3H, supplementary data figure S3. We acknowledge that whilst more Ni2+ may bind after 15 mins, the RPS is unable to resolve the changes, and we conclude that the majority of the His sites were occupied after this time period. Given the results in figure 3b, we opted to choose P2H for future experiments, simply because it was the easiest peptide to run and showed the greatest speed change when binding with Ni2+.

Figure 3: a) Changes in P2H functionalised particle speed under different applied potentials, with and without Ni2+. 50 mM KCl buffered to pH 7, -0.82 V (green) and +0.82 V (red). b) Binding of different peptides with Ni2+ over time, P1H, black, P2H, red and P3H, blue. Ran in 50 mM KCl buffered to pH 7, 1.26 V

To show the change in velocity was proportional to the concentration of Ni2+ a dose response curve was created. The concentration of the SPN’s and the density of the peptide on their surface remained constant, and the concentration of Ni2+ varied. As can be seen in figure 4, as the concentration of Ni2+ increases, the velocity of the carriers decreases. The decrease in velocity was observed over three orders of magnitude, which covers the range used to measure Ni in drinking water.46 Important to note that the presence of Ni2+ did not cause the SPN’s to aggregate, see figure S4, and that the signal was reproducible on different batches of carriers figure S5.



1.1

Relative speed (1/T0.5)

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1.0 0.9 0.8 0.7 0

200 400 600 800 1000 1200

Nickel concentration (nM)

Figure 4: Relative particle speeds taken as 1/T0.5, normalised to peptide functionalised beads, vs. varying amounts of nickel ion added. Ran in 50 mM KCl. Each sample was ran in

triplicate, and more than 200 particles were measured each time. Errors bars are one standard deviation from the mean. Whilst poly His tags are known to bind to Ni2+, they are also known to bind to Co2+.47 To illustrate the selectivity of the assay is determined by the ligand itself, and that the presence of other ions did not interfere with the RPS measurement, the assay was repeated in a range of other divalent ions, figure 5a. In the presence of Ni or Co, the velocity of the particles decreased. When Ni was present in a mixture (mix) of other divalent ions (excluding Co) this observation was again, figure 5a. Interestingly the presence of Cr and Fe caused an increase in relative velocity, the cause of which is unclear.


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Figure 5: a) Speed of the P2H functionalised carrier when incubated with Ni2+, Ca2+, Co2+, Cr3+, Fe3+ and a mix without Co2+. All metals present at 600 nM, in 50 mM KCl buffered to pH 7. b) environmental water samples with and without 600 nM Ni2+ present. Samples made up to 50 mM using 1 M KCl. Each sample was ran in triplicate, and more than 200 particles were

measured each time. Errors bars are one standard deviation from the mean. As a final demonstration that the assay can be performed in complex sample matrices, we sampled tap water from the laboratory and water from a local pond on the university campus. Peptide modified carriers were added to the fresh samples, the SPN were mixed in the sample and the conductivity of the sample adjusted using 1 M KCl to make them to 50 mM KCl. In each case the blanks traversed the pore as a consistent speed, figure S6, indicating that any Ni2+ present in the tap and pond water was below our limit of detection. We then repeated this process having spiked each sample with Ni2+. The velocities of the carriers in each of the samples is shown in figure 5b. A decrease in velocity was observed, which confirms the assay worked and that any other analyte present in the samples did not affect the peptide or RPS.

Conclusions Here we present a method to use peptide coated nanocarriers to quantify metal ions in solution. The poly histidine peptide binds to Ni2+ in solution. The binding of the Ni2+ to the carrier is measured as a change in carrier velocity as it traverses the pore. The method was shown to work on environmental samples, where the use of magnetic nanocarriers allows for the Ni2+ to be extracted and quantified in under 30 minutes.

Supporting Information Supporting Information Available: The following files are available free of charge. Blockade magnitudes of peptide particles in the presence of different concentration of Ni2+, and differences between the blank environmental samples when normalised to calibration particles. This material is available free of charge via the internet at http://pubs.acs.org

Author Information Corresponding Author *E-mail: [email protected]. Notes The authors declare no competing financial interest.

Acknowledgements



The authors would like to thank Dr B Kralj for his guidance and support and the Nuclear Decommissioning Authority for their funding.

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Peptide Nanocarriers for Detection of Heavy Metal Ions Using

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