Reversible Electrochemical Sensor for Detection of High-Charge

Subscriber access provided by UNIV OF CAMBRIDGE

Article

Reversible Electrochemical Sensor for Detection of High Charge Density Polyanion Contaminants in Heparin Jacob Lester, Timothy Chandler, and Kebede Lemma Gemene Anal. Chem., Just Accepted Manuscript • DOI: 10.1021/acs.analchem.5b03347 • Publication Date (Web): 20 Oct 2015 Downloaded from http://pubs.acs.org on October 27, 2015

Just Accepted “Just Accepted” manuscripts have been peer-reviewed and accepted for publication. They are posted online prior to technical editing, formatting for publication and author proofing. The American Chemical Society provides “Just Accepted” as a free service to the research community to expedite the dissemination of scientific material as soon as possible after acceptance. “Just Accepted” manuscripts appear in full in PDF format accompanied by an HTML abstract. “Just Accepted” manuscripts have been fully peer reviewed, but should not be considered the official version of record. They are accessible to all readers and citable by the Digital Object Identifier (DOI®). “Just Accepted” is an optional service offered to authors. Therefore, the “Just Accepted” Web site may not include all articles that will be published in the journal. After a manuscript is technically edited and formatted, it will be removed from the “Just Accepted” Web site and published as an ASAP article. Note that technical editing may introduce minor changes to the manuscript text and/or graphics which could affect content, and all legal disclaimers and ethical guidelines that apply to the journal pertain. ACS cannot be held responsible for errors or consequences arising from the use of information contained in these “Just Accepted” manuscripts.

Analytical Chemistry is published by the American Chemical Society. 1155 Sixteenth Street N.W., Washington, DC 20036 Published by American Chemical Society. Copyright © American Chemical Society. However, no copyright claim is made to original U.S. Government works, or works produced by employees of any Commonwealth realm Crown government in the course of their duties.

Page 1 of 14

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

Analytical Chemistry

Reversible Electrochemical Sensor for Detection of High Charge Density Polyanion Contaminants in Heparin Jacob Lester, Timothy Chandler, Kebede L. Gemene* Department of Chemistry, Northern Kentucky University, Nunn Drive, Highland Heights, KY 41099 Abstract: We present a simple, rapid and inexpensive electrochemical sensor based on a reversible pulsed chronopotentiometric polyanion-selective membrane electrode for the detection and quantification of oversulfated chondroitin sulfate (OSCS) and other high charge-density polyanions that could potentially be used to adulterate heparin. The membrane is free of ion exchanger and is formulated with plasticized poly(vinyl chloride) (PVC) and an inert lipophilic salt, tridodecylmethylammonium-dinonylnaphthaline sulfonate (TDMADNNS). The neutral salt is used to reduce membrane resistance and to assure reversibility of the sensor. More importantly, TDMA+ is used as the recognition element for the polyanions. Here an anodic galvanostatic current pulse is applied across the membrane to cause the extraction of the polyanions from the sample into the membrane and potential is measured at the sample-membrane interface. The measured electromotive force (emf) is proportional to the concentration and the charge density of the polyanions. High charge-density polyanion contaminants and impurities in heparin can be detected using this method since the overall equilibrium potential response of polyions increases with increasing charge density of the polyions. Here, first the potential response of pure heparin is measured at a saturation concentration, the concentration beyond which further addition of heparin does not produce a change in potential response. Then the potential response of heparin tainted with different quantities of the high charge-density contaminant is measured at a fixed total polyion concentration (heparin concentration + contaminant concentration). The latter gives a greater negative potential response due to the presence of the high charge-density contaminant. The increase in the negative potential response can be used for detection and quantification of high charge-density contaminants in heparin. We demonstrate here that 0.3% (w/w) OSCS as well as 0.1% (w/w) dextran sulfate can be detected in heparin at 20-mg/ml total polyion concentration. 1% (w/w) of dextran sulfate can readily be detected in heparin at only 2-mg/ml total polyanion concentration with a linear response (R2 = 0.994).

ACS Paragon Plus Environment

1


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

Page 2 of 14

Heparin is a highly sulfated anionic polysaccharide widely used as an anticoagulant/antithrombotic agent during clinical procedures such as cardiac/vascular surgery and kidney dialysis and as a therapeutic agent for the treatment of many clotting disorders such as deep vein thrombosis and acute coronary syndrome.1,2 In addition, it is used in many medical devices and diagnostic products that may be coated with or contain heparin,3 including catheters, extracorporeal blood loops and blood collection tubes. From late 2007 to early 2008 several hundred severe adverse reactions, including over 200 deaths4 were reported worldwide after intravenous administration of certain lots of commercial heparin products. It was later discovered using orthogonal analytical methods that the lots of commercial heparin products associated with the adverse reactions were contaminated with a high negative charge-density hypersulfated glycosaminoglycan (GAG) called oversulfated chondroitin sulfate (OSCS).5,6, OSCS is a semisynthetic highly sulfated polysaccharide synthesized by oversulfation of chondroitin sulfate A (CSA), which is obtained from animal cartilage.6 It has a chemical structure and an anticoagulant activity similar to that of heparin.6,8,9 Perhaps because of the similarities in chemical structures and biological activities between OSCS and heparin, the United States Pharmacopeia (USP) and the European Pharmacopeia (Ph. Eur.) testing monographs of heparin sodium that existed during the 2007-2008 heparin contamination crisis failed to detect the presence of the contaminant OSCS in heparin products. This could be because, historically, pharmaceutical grade heparin was identified based on measurement of activity using plasma-clotting assays rather than based on molecular properties.10 Thus, after the heparin contamination crisis of 2007-2008, USP and Ph. Eur. revised their heparin sodium testing monographs by including analytical techniques that are sensitive to the structural differences and the charges of the GAGs to ensure the purity of heparin active pharmaceutical ingredient (API) before distribution for clinical use. Accordingly, the current revised monographs require testing of heparin API for purity by orthogonal analytical techniques including nuclear magnetic resonance spectroscopy, 1H NMR9,11,12 (sensitive to structure) and strong anion exchange high performance liquid chromatography (SAX-HPLC) with ultraviolet spectroscopy and mass spectrometry detection13-16 (sensitive to charge). These analytical techniques are well established and reliable. Nevertheless, they require highly trained expert operators and very expensive equipment, which may be available only in specialized laboratories. However, heparin is used extensively and is obtained from a variety of manufacturers worldwide, some of them not properly regulated.4,17 In addition, since the OSCS identified as the cause of severe adverse reactions during the 2007-2008 heparin crisis was found to be a non-natural inexpensive semisynthetic GAG, it is believed that it was added deliberately to the heparin supply chain for economic reasons.5,13,18,19 Thus, screening of heparin for purity is warranted not only at the manufacturing facilities but also at many different levels, including locally in hospitals, for quality assurance of the heparin supply chain. Therefore, the development of additional analytical methods, in particular, reliable methods that are inexpensive and can be operated by users with limited expertise is needed. To this end, many analytical techniques have been developed in the last few years for the detection and quantification of OSCS and other contaminants and impurities in heparin products, in addition to 1H NMR and SAX-HPLC. Capillary electrophoresis (CE)20-22 has been extensively explored. In fact, it was one of the methods in the orthogonal analytical methods that were first used to identify OSCS as the contaminant that caused severe


2

Page 3 of 14

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


events during the 2007-2008 heparin crisis and was included in the early revised USP and Ph. Eur. testing monographs.4,5,22 It was later substituted by SAX-HPLC17 due to its limitations to fully resolve OSCS from heparin.12 Colorimetric microplate based assays have been successfully used to detect OSCS in heparin at very low detection limits.18,23,24 Fluorescence assays25,26 have also been used. Recently, an ultrasensitive fluorescence assay using a nanometal surface energy transfer (NSET) based gold-heparin-dye nanosensor that could detect 1.0x10-9 % (w/w) OSCS in heparin was reported.2 Mass spectrometry has been successfully used for the detection of contaminants in heparin after chromatographic separations or enzymatic digestions of the GAGs.3,4,19 Infrared spectroscopy27, enzymatic digestion4,18,28,29 and polyacrylamide gel electrophoresis (PAGE)8 were also used for the detection of GAG contaminants in heparin. Potentiometric polyion-selective electrodes were also shown to be effective to detect and quantify30,31 OSCS in heparin. Recently, a highly sensitive disposable strip-test type potentiometric sensor with detection limit of 0.005% (w/w) OSCS within heparin was reported.32 Potentiometric polyion-selective electrodes were first introduced in the early 1990s by the Meyerhoff group

1,33-36

These devices have been thoroughly developed and

well characterized within the past few decades and demonstrated to have many potential biomedical applications,37,38 including measuring heparin in undiluted whole blood33 and determination of DNA.39 A very sensitive potentiometric aptasensor with DNA nanostructure-based magnetic beads was described recently using polyion sensitive electrodes.40 These devices offer unique sensor qualities; they are simple, inexpensive, rapid, portable, robust and reliable. The main limitation of these sensors is that they are irreversible. This limits them to single-use disposable devices, which makes them inconvenient for continuous monitoring purposes such as screening of infusion heparin, for example. Fully reversible pulsed chronopotentiometric polyion sensors were described recently for polycations41,42 and polyanions.43 The reversibility of these sensors is made possible by using membranes without intrinsic ionexchange properties. This suppresses spontaneous extraction of the polyions, which otherwise renders the sensors irreversible. The membrane contains inert lipophilic salts of the form R+R- (where R+ and R- are a lipophilic cation and a lipophilic anion, respectively). Ideally, this membrane is irresponsive to polycations or polyanions under zero current potentiometry conditions. However, under pulsed chronopotentiometric conditions, the membrane can be made sensitive to polycations or polyanions, on demand, by perturbing the equilibrium lipophilic ions distribution within the membrane phase (polarization of the membrane) by passing an appropriate galvanostatic current pulse through the membrane. This is followed by a zero-current (open circuit) pulse. Finally, full membrane restoration is attained by applying an equilibrium (stripping) potential across the membrane to depolarize the membrane and eject ions that were extracted into the membrane during the galvanostatic current pulse. This results in membrane regeneration and a fully reversible sensor response.41-44 These sensors have been demonstrated to have many potential analytical applications including detection of heparin and protamine,41-46 detection of protease activities,47-49 as a detector in flow injection analysis (FIA) of polyions50 and as a detector of polyions in liquid chromatography (LC).51 We report here the first reversible electrochemical detection and quantification of high charge density polyanion contaminants in heparin products using pulsed chronopotentiometric polyanion-selective electrodes. The membrane

used

is

plasticized

poly(vinyl

chloride)

(PVC)

containing


tridodecylmethylammonium-

3


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

Page 4 of 14

dinonylnaphthaline sulfonate (TDMA-DNNS) as the lipophilic salt. When an anodic current pulse is applied, polyanions are extracted into the membrane via cooperative ion-pair formation at the sample/membrane interface. The resulting phase boundary potential is a function of the concentration of the polyions and the equilibrium constant of the ion pair formation.35 Since ion pair formation is electrostatic in nature, polyanions with higher charge-density form stronger cooperative ion pairs with TDMA+ and have greater equilibrium constant of extraction into the membrane phase. Therefore, they have a greater negative (anionic) phase boundary potential response. Consequently, heparin samples that are tainted with high charge density contaminants give greater negative potential responses than pure heparin. The increase in the negative potential response of pulstrode sensors to tainted heparin compared to the response to pure heparin can be used to detect and quantify high charge-density contaminants within the heparin sample. This method achieves detection limit of 0.3% (w/w) OSCS, which is one order of magnitude lower than the No-Observable-Effect Level (NOEL)23,53 of OSCS in heparin in animal models (swine) that are sensitive to the effects of OSCS in a similar manner as humans6 and 0.1% (w/w) dextran sulfate when the total polyanion concentration is 20 mg/ml. This technique is based on a simple principle of charge density as explained above. There is no need for separation, enzymatic digestion or any major sample pretreatment for detection of contaminant GAGs in heparin. In addition, potential hypersulfated contaminants such as dextran sulfate and pentosan polysulfate25 or impurities such as oversulfated heparin that may be difficult to detect by 1H NMR due to the lack of the characteristic N-acetyl methyl proton signal of OSCS at 2.16 ppm1,14,24 can be easily detected by this method. In general, pulsed chronopotentiometry with polyanion-selective electrodes may be used as a simple and universal detector of all contaminants that have higher charge-density than heparin in heparin API.

Experimental Section Reagents. High molecular weight poly(vinyl chloride) (PVC), o-nitrophenyl octyl ether (o-NPOE), 2-ethylhexyl sebacate

(DOS),

tetradodecylammonium

tetrakis(4-chlorophenyl)

borate

(ETH

500),

tridodecylmethylammonium chloride (TDMAC), tetrahydrofuran (THF), heparin sodium salt (from porcine intestinal mucosa), dextran sulfate and all salts were purchased from Sigma-Aldrich (St. Louis, MO). Dinonylnaphthalene sulfonic acid (as 50% solution in xylene) was a gift from King Industries (Norwalk, CT). OSCS was a gift from Prof. Meyerhoff (University of Michigan, Ann Arbor, MI). Aqueous solutions were prepared by dissolving the appropriate compounds in Nanopure deionized water (18.2 MΩ cm). The neutral inert salt, TDMA-DNNS was prepared in our lab by metathesis of tridodecylmethylammonium chloride and dinonylnaphthalene sulfonic acid in benzene.

Membrane Preparation. Polyion-selective membrane ( 200 µm thick) was prepared by solvent casting with THF as a solvent, a membrane cocktail containing 10 wt % of the inert lipophilic salt TDMA-DNNS, 30 wt% PVC and 60 wt% NPOE.

Electrodes. Membranes (8 mm diameter) were cut with cork borer from the parent membrane and mounted into electrode bodies (Oesch Sensor Technology, Sargans, Switzerland). The actual membrane area was 23 mm2. The


4

Page 5 of 14

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


inner solution was in contact with an internal Ag/AgCl electrode. The external reference electrode was a doublejunction Ag/AgCl electrode with saturated KCl as inner solution and a 1 M LiOAc bridge electrolyte. A high surface area coiled Pt wire was used as a counter electrode. The working electrodes were conditioned for at least 12 h prior to experiments and kept in the conditioning solution when experiments were not underway. The inner filling and conditioning solution was 10 mM NaCl in 10 mM phosphate buffer at pH 7.4. The background solution was 10 mM NaCl in 10 mM phosphate buffer at pH 7.4 Experimental Setup. A conventional three-electrode setup was used for the pulsed chronopotentiometric measurements, where the TDMA-DNNS based polyion-selective membrane with an internal Ag/AgCl electrode acted as the working electrode and the external reference electrode and counter electrode were immersed into the sample solution. The measurements were conducted with an AFCBI bipotentiostat (Pine Instruments, Grove City, PA) controlled by USB-6212 interface board and LabVIEW 2011 data acquisition software (National Instruments, Austin, TX) on a PC computer. An uptake current pulse time of 1 s, a zero-current pulse time of 0.5 s and a stripping potentiostatic pulse time of 15 s were used throughout the experiment. Baseline potential pulse of 0 V versus Ag/AgCl was applied as a stripping potential. The potentials were sampled at 2 ms intervals and recorded in two files, viz., as raw data and averaged data of the last 10% of the anodic current pulse and the zerocurrent pulse. The potential sampled during the last 10% of the anodic current pulse time was used in all measurements. All experiments were conducted at room temperature (22–23 °C).

Results and Discussion In this work, a reversible electrochemical polyanion sensor based on pulsed chronopotentiometry (pulstrode) is employed for the detection and quantification of high charge-density contaminants in heparin. The working mechanism of this sensor is similar to that of the well-established irreversible, single-use, classical potentiometric polyanion sensor. However, unlike classical potentiometric polyanion sensor, this sensor is reversible and reusable. Full reversibility of this sensor can be realized by utilizing a plasticized PVC membrane containing a neutral inert lipophilic salt of the form R+R- (where R+ and R- are TDMA+ and DNNS-, respectively), instead of the dissociated anion exchanger, TDMACl, used in classical potentiometric polyion sensors. The schematic illustration of the membrane polarization/depolarization principle by the application of consecutive current and potential pulses is shown in Figure 1A.50 Figure 1B shows the observed current-time (green) and potential-time (red) response behavior for the measurement of 0.4 mg/ml heparin in a background solution of 10 mM phosphate buffer, pH 7.4 containing 10 mM NaCl. Here, an anodic galvanostatic current pulse (shown by P1 and green line in Figure 1B) is applied across the membrane to perturb the equilibrium ion distribution shown by (i) in Figure 1A within the membrane and cause membrane polarization via ion redistribution. Here, TDMA+ and DNNS- migrate toward the sample side and the inner filling solution side of the membrane surface, respectively, and concomitantly, the polyanion (Pz-) is extracted from the sample into the polymeric membrane via ion-pair formation with TDMA+ at the sample/membrane interface as shown under (ii) in Figure 1A. The phase boundary potential, which is a function of the polyanion concentration, is measured simultaneously at the sample/membrane interface (red line and P1 in Figure 1B). This was followed by a 0.5-s zero-current (opencircuit) pulse (P2). During this time the membrane tends to relax toward its initial ion distribution condition


5


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

Page 6 of 14

under open-circuit and concomitantly the ions extracted from the contacting aqueous phases during the first uptake current pulse tend to leave the membrane to their respective aqueous phases and potential is measured

simultaneously again at the same membrane interface ((iii) in Figure 1A and P2, red line in Figure 1B).

Figure 1. (A) Schematic illustration of membrane polarization/depolarization mechanism under consecutive current and potential pulses: (i) initial uniform equilibrium concentration profile within the membrane; (ii) polarized membrane profile under galvanostatic current pulse; (iii) membrane relaxation under zero-current (open-circuit) pulse; (iv) membrane restoration under potentiostatic pulse. (B) Observed current-time (green) and potential-time (red) response behavior for the measurement of 0.4 mg/ml heparin, as a model system – potentials are sampled at the end of the galvanostatic pulse (a) and at the end of the zero-current pulse (b)

The advantage of this zero-current pulse in the triple-pulse sequence of pulstrode is that an analytically useful potential can be sampled at the end of this pulse. However restoration of the membrane phase equilibrium ion concentration profile under zero-current condition is very slow54 and cannot be attained in a reasonable measuring time. Lindner’s group described a reverse current pulse method, where a galvanostatic current pulse of opposite sign is applied across the membrane following perturbation of the uniform equilibrium membrane ion distribution by the initial current pulse, for rapid membrane restoration.54 However, this method requires permselectivity of the membrane and cannot work with membranes without excess ion exchanger. In pulstrode, full membrane restoration is attained by applying an equilibrium (stripping) potential after the current pulses. Bakker’s group recently started using a potential equal to the open-circuit potential measured before the current excitation pulse as the membrane restoration potential.55,56 In this work, a 15-s potentiostatic pulse of 0 V versus the reference electrode (P3, red line in Figure 1B) was used as the stripping potential for the membrane restoration. Here all the ions extracted from the aqueous phase are stripped out of the membrane as schematically illustrated by (iv) in Figure 1A and the initial uniform ion distribution within the membrane is restored as depicted by (iv) and (i) in Figure 1A. This effective membrane restoration enables a reversible and very reproducible measurement of polyanions under pulstrode. Indeed, the potentials sampled at the end of the 1-s galvanostatic pulse at (a) and at the end of the 0.5-s zero-current pulse at (b) in Figure 1B for the detection of 0.4


6

Page 7 of 14

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


mg/ml heparin were found to be 1.0189±0.0009 V and 0.3573±0.0004 V, respectively (n=11, where n is the number of data points sampled at (a) in Figure 1B). Polyanions are extracted into the sensing membranes by strong cooperative ion-pair formation with TDMA+ at the membrane/sample interface and the resulting phase boundary potential is a function of the concentration and the charge density of the polyanion.35 Thus, it is invisioned that pulstrode polyanion sensors can be reliably used for detection of high charge density contaminants in heparin. As an example of practical applicability of the sensor, first, the detection of dextran sulfate in heparin was explored. Dextran sulfate and other semisynthetic oversulfated GAGs such as pentosan polysulfate (PPS) could be potentially used as economically motivated adulterants (EMAs) in heparin in the future, even more likely than OSCS, which has been studied so extensively and cannot pass heparin purity tests by the existing methods. These GAGs are heparin mimetic, i.e., they have similar structures and biological activities with heparin18,25. Indeed, they have been used as anticoagulants, historically. However, these highly sulfated polysaccharides have been shown to initiate activation of the contact systems and can cause severe adverse reactions like OSCS.5,57,58 They serve as a negatively charged surface that can initiate the contact system; it was found that the contact system recognized negative charges rather than specific chemical structures.55,56 This strongly supports the importance of detecting these highly sulfated high charge density polyanions in heparin to ensure the safety of heparin API. It also shows the convenience and the potential of pulstrode polyanion sensors for the detection of such contaminants in heparin.

Figure 2. Schemes of the structures of the major repeating disaccharide units of heparin, oversulfated chondroitin sulfate (OSCS) and dextran sulfate (DexS). The charges per repeating disaccharide units of Heparin, OSCS and DexS are -4, -5, and -6, respectively.


7


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

Page 8 of 14

Figure 2 illustrates, schematically, the structures of the major repeating disaccharide units of polyanions of different charge densities that were tested in this work, heparin, oversulfated chondroitin sulfate and dextran sulfate. Heparin, OSCS and DexS have charges of -4, -5, and -6, respectively, per repeating disaccharide units.

Figure 3. Observed time-dependent responses of pulstrode polyanion sensor to varying concentrations of heparin (A) and dextran sulfate (B) in a background of 10 mM phosphate buffer containing 10 mM NaCl at pH 7.4. Each step in the time-trace is composed of 11 data points sampled at 16.5 s interval, at the end of the galvanostatic current pulse (average values with standard deviation < 0.2 mV, for each step are shown). Each triple pulse is composed of a 1-s 15 µA galvanostatic pulse, a 0.5-s zero-current pulse and a 15-s 0 V, versus Ag/AgCl reference electrode, potentiostatic pulse.

Figure 4. Observed pulstrode sensor response calibration curves to varying concentrations of heparin (A) and dextran sulfate (B), from the potential-time traces shown in Figure 3.

Figure 3 shows the observed potential-time traces of the pulstrode polyanion sensor to varying concentrations of heparin (A) and dextran sulfate (B) and Figure 4 shows the corresponding pulstrode sensor calibration curves from the potential-time traces in Figure 3. As expected, dextran sulfate gave considerably larger negative potential response than heparin at each given concentration due to its higher charge density. At 0.2-mg/ml concentrations of the polyanions the observed potential response produced by dextran sulfate is 58 mV more negative than that of heparin. The pulstrode sensor response reproducibilities were investigated by alternative


8

Page 9 of 14

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


measurements in a background solution of 10 mM phosphate buffer containing 10 mM NaCl at pH 7.4 and in a 2 mg/ml polyanion solution in the same background and shown in Figure 5 (A) heparin and (B) dextran sulfate. Good stabilities and reproducibilities were observed for measurements in both the background solutions and the solutions of the polyanions. For heparin measurement, the standard deviations were 1 mV both for the background solution (top) and the heparin solution (bottom). For dextran sulfate measurement, the standard deviations were 1 mV for the background solution (top) and 0.8 mV for the dextran sulfate solution (bottom).

Figure 5. Pulstrode response reproducibilities on alternative measurements in a background solution (top) and in a 2 mg/ml polyanion solution (bottom) of (A) heparin and (B) dextran sulfate. The electrodes were rinsed thoroughly between the measurements. All the other measuring conditions, including pulse parameters are the same as Figure 3.

Figure 6. Pulstrode polyanion sensor response to 2 mg/ml heparin and then to serial additions of dextran sulfate (A) and the corresponding linear curve (B). The total concentration of the polyanions was fixed at 2 mg/ml. All the other measuring conditions are as given in Figure 3.

Figure 6 shows the response of pulstrode polyanion sensors for the direct detection of the contaminant dextran sulfate in heparin. First, the response of the sensor was measured in high concentration (2 mg/ml) heparin. Then more heparin was added and the sensor response was measured again. No change in potential response was observed on the second addition of heparin and this proved that the heparin was at a high concentration where it produces the maximum potential response. Then 1% (w/w) dextran sulfate was added while the total concentration of polyanions was kept constant. This produced a potential difference (∆E) of -3 mV from the response to heparin. Given that the observed standard deviations were ≤ 1 mV, it can be safely stated that as low as 1% of dextran sulfate can be detected in a total polyanion concentration of only 2 mg/ml. Serial additions of


9


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

Page 10 of 14

dextran sulfate while the total polyanion concentration was kept constant produced significant corresponding potential responses. The change in potential (∆E) as a function of the % of dextran sulfate in heparin was found to be linear with R2=0.994 up to 10% (w/w) of dextran sulfate as shown in Figure 6B. It is known that the contaminant species can be detected by the sensor only if it is present in the solution at a concentration greater than its detection limit. Thus, measuring in a solution of higher total polyanion concentration is beneficial to lower the detection limit, as % (w/w) of the contaminant, in heparin. Figure 7 depicts the pulstrode polyanion responses to pure heparin (A), heparin tainted by 0.3% and 1.0% (w/w) OSCS (B) and heparin tainted by 0.3% and 1.0% (w/w) dextran sulfate (C) at total polyanion concentration of 20 mg/ml. As can be seen, the contaminants are clearly discernable at a concentration as low as 0.3% (w/w). The response of heparin tainted with 0.3% (w/w) OSCS is 8±1 mV more negative than pure heparin while the response of heparin tainted with 0.3% (w/w) dextran sulfate is 27.0±0.9 mV more negative than pure heparin.

Figure 7. Responses of pulstrode polyanion sensor to pure heparin (A), heparin tainted by 0.3% and 1% (w/w) OSCS (B) and heparin tainted by 0.3% and 1% (w/w) dextran sulfate (C) at total polyanion concentration of 20 mg/ml. All the other measuring conditions are as given in Figure 3.

Addition of 1% (w/w) of the contaminant polyanions, OSCS and dextran sulfate, produced a change in potential responses of -26.3±1.2 mV and -52.4±1.4 mV, respectively, relative to pure heparin (see Figure 7). This is not surprising, in fact, since 1% (w/w) of the contaminant polyanion produces 0.2 mg/ml of the contaminant in the solution and this is in the high sensitivity response region of the pulstrode polyanion sensor (see Figure 3). Further increase of the total polyanion concentration improves the detection limit as% (w/w) of contaminant in heparin, considerably. Obviously, if the measurement is conducted in high polyanion concentrations as used in the other methods32 including 1HNMR, lower level of contaminants can be detected.


10

Page 11 of 14

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


Conclusions

We have presented a simple, inexpensive, robust and rapid electrochemical method for the detection and quantification of high charge density polyanion contaminants in heparin. The membrane is formulated with the polyanion carrier, TDMA+, which is known to form a strong cooperative ion pair with polyanions (i.e. heparin, OSCS, dextran sulfate, PPS). The potential response generated during measurement of these polyanions is proportional to the polyanion’s charge density; that is, larger charge densities are associated with larger overall potential responses. Thus, the sensor is more sensitive to the high charge density polyanions. We have demonstrated the detection of 0.3% (w/w) OSCS and 0.1% (w/w) dextran sulfate in heparin in a total polyanion concentration of 20 mg/ml with our reusable sensor. The No-Observable-Effect Level (NOEL) of OSCS in humans has not been published, to our knowledge. However, in animal models that are sensitive to the effects of OSCS in a similar manner as humans, the NOEL of OSCS was found to be 3%, which is much higher than our detection limit. The detection limit of the sensor can be even decreased further by using higher concentration of the solution of the polyanions, as used in other methods,32 since the polyanions are readily soluble and the performance of the sensor is not significantly affected by concentration of the polyanions. Therefore, it is anticipated that this sensor can be used reliably for the detection of high charge density contaminants in heparin, obviating the need for sophisticated and expensive methods.

Author Information Corresponding author Telephone: 859-572-7543 Email: [email protected] Notes The authors declare no competing financial interest.

Acknowledgement: This work was supported by Institutional Development Award (IDeA) from the National Institute of General Medical Sciences of the National Institute of Health under grant number 5P20GM103436 and Northern Kentucky University Center for Integrative Natural Sciences And Mathematics, CINSAM.


11


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

Page 12 of 14

References 1.

Ma, S.; Yang, V. C.; Meyerhof, M. E. Anal. Chem. 1992, 64, 694-697.

2.

Kalita, M.; Balivada, S.; Swarup, V. P.; Mencio, C.; Raman, K.; Desai, U. R.; Troyer. D.; Kuberan, B. J. Am. Chem. Soc. 2014, 136, 554-557.

3.

Nemes, P.; Hoover, W. J.; Keire, D. A. Anal Chem. 2013, 85, 7405-7412.

4.

Keire, D. A.; Ye, H.; Trehy, M. L.; Ye, W.; Kolinski, R. E.; Westenberger, B. J.; Buhse, L. F.; Nasar, M.; Al-Hakim, A. Anal. Bioanal. Chem. 2011, 399, 581-591.

5.

Guerrini, M.; Beccati, D.; Shriver, Z.; Naggi, A.; Viswanathan, K.; Bisio, A.; Capila, I.; Lansing, J. H.; Guglieri, S.; Fraser, B.; Al-Hakim, A.; Gunay, N. S.; Zhang, Z.; Robinson, L.; Buhse, L.; Nasr. M.; Woodcock, J.; Langer, R.; Venkataraman, G.; Linhardt, R. J.; Casu, B.; Torri, G.; Sasisekharan, R. Nat. Biotechnol. 2008, 26, 669-675.

6.

Kishimoto, T. K.; Viswanathan, K.; Ganguly, T.; Elankumaran, S.; Smith, S.; Pelzer, K.; Lansing, J. C.; Sriranganathan, N.; Zhao, G. L.; Galcheva-Gargova, Z.; Al-Hakim, A.; Bailey, G. S.; Fraser, B.; Roy, S.; RogersCotrone, T.; Buhse, L.; Whary, M.; Fox, J.; Nasr, M.; Dal Pan, G. J.; Shriver, Z.; Langer, R. S.; Venkataraman, G.; Austen, K. F.; Woodcock, J.; Sasisekharan, R. N. Engl. J. Med. 2008, 358, 2457–2467.

7.

Keire, D. A.; Mans, D. J.; Ye, H.; Kolinski, R. E.; Buhse, L, F. J. Pharm. Biomed. Anal. 2010, 52, 656-664.

8.

Li, B.; Suwan, J.; Martin, J. G.; Zhang, F.; Zhang, Z.; Hoppensteadt, D.; Clark, M.; Fareed, J.; Linhardt, R. J. Biochem. Pharmacol. 2009, 78, 292-300.

9.

Zhang, Z.; Li, B.; Suwan, J.; Zhang, F.; Wang, Z.; Liu, H.; Mulloy, B.; Linhardt, R. J. J. Pharm. Sci. 2009, 98, 4017-4026.

10. Guerrini, M.; Zhang, Z.; Shriver, Z.; Naggi, A.; Masuko, S.; Langer, B.; Casu, B.; Linhardt, R. J.; Torri, G.; Sasisekharan, R. P. Natl. Acad. Sci. USA 2009, 106, 16956-16961. 11. Beyer, T.; Diehl, B.; Humpfer, E.; Schafer, H.; Spraul, M.; Schollmayer, C.; Holzgrabe, U. J. Pharm. Biomed. Anal. 2008, 48, 13-19. 12. Bigler, P.; Brenneisen, R. J. Pharm. Biomed. Anal. 2009, 49, 1060-1064. 13. Trehy, M. L.; Reepmeyer, J. C.; Kolinski, R. E.; Westenberger, B. J.; Buhse, L. F. J. Pharm. Biomed. Anal. 2009, 49, 670-673. 14. Keire, D. A.; Mans, D. J.; Ye, H.; Kolinski, R. E.; Buhse, L. F. J. Pharm. Biomed. Anal. 2010, 52, 656-664. 15. Keire, D. A.; Trehy, M. L.; Reepmeyer, J. C.; Kolinski, R. E.; Ye, W.; Dunn, J.; Westenberger, B. J.; Buhse, L. F. J. Pharm. Biomed. Anal. 2010, 51, 921-926. 16. Beyer, T.; Matz, M.; Brinz, D.; Radler, O.; Wolf, B.; Norwig, J.; Baumann, K.; Alban, S.; Holzgrabe, U. Eur. J. Pharm. Sci. 2010, 40, 297-304. 17. Ye, H.; Toby, T. K.; Sommers, C. D.; Ghasriani, H.; Trehy, M. L.; Ye, W.; Kolinski, R. E.; Buhse, L. F.; Al-Hakim, A.; Keire, D. A. J. Pharm. Biomed. Anal. 2013, 85, 99-107. 18. Sommers, C. D.; Keire, D. A. Anal. Chem. 2011, 83, 7102-7108. 19. Li, G.; Cai, C.; Li, L.; Fu, L.; Chang, Y.; Zhang, F.; Toida, T.; Xue, C.; Linhardt, R. J. Anal. Chem. 2014, 86, 326330. 20. Tami, C.; Puig, M.; Reepmeyer, J. C.; Ye, H.; D’Avignon, D.A.; Buhse, L.; Verthelyi, D. Biomaterials 2008, 29, 4808–4814. 21. Volpi, N.; Maccari, F.; Linhardt, R. J. Anal. Biochem. 2009, 388, 140–145. 22. Somsen, G. W.; Tak, Y. H.; Torano, J. S.; Jongen, P. M. J. M.; de Jong, G. J. J. Chromatogr. A 2009, 1216, 41074112. 23. Bairstow, S.; McKee, J.; Nordhaus, M.; Johnson, R. Anal. Biochem. 2009, 388, 317–321. 24. Sommers, C. D.; Mans, D. J.; Mecker, L. C.; Keire, D. A. Anal. Chem. 2011, 83, 3422–3430.


12

Page 13 of 14

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


25. Alban, S.; Luhn, S.; Schiemann, S. Anal. Bioanal. Chem. 2011, 399, 681-690. 26. Jagt, R. B.; Gomez-Biagi, R. F.; Nitz, M. Angew. Chem., Int. Ed. Engl. 2009, 48, 1995–1997. 27. Spencer, J. A.; Kauffman, J. F.; Reepmeyer, J. C.; Gryniewicz, C. M.; Ye, W.; Toler, D. Y.; Buhse, L. F.; Westenberger, B. J. J. Pharm. Sci. 2009, 98, 3540–3547. 28.

Brustkern, A. M.; Buhse, L. F.; Nasr, M.; Al-Hakim, A.; Keire, D. A. Anal. Chem. 2010, 82, 9865–9870.

29. Xiao, Z.; Tappen, B. R.; Ly, M.; Zhao, W.; Canova, L. P.; Guan, H.; Linhardt, R. J. J. Med. Chem. 2011, 54, 603−610. 30. Wang, L.; Buchanan, S.; Meyerhoff, M. E. Anal. Chem. 2008, 80, 9845-9847. 31. Wang, L.; Meyerhoff, M. E. Electroanalysis 2010, 22, 26-30. 32. Kang, Y.; Gwon, K.; Shin, J. H.; Nam, H.; Meyerhoff, M. E.; Cha, G. S. Anal. Chem. 2011, 83, 3957-3962. 33. Ma, S.; Yang, V. C.; Fu, B.; Meyerhoff, M. E. Anal. Chem. 1993, 65, 2078−2084. 34. Fu, B.; Bakker, E.; Yun, J. H.; Yang, V. C.; Meyerhoff, M. E. Anal. Chem. 1994, 66, 2250−2259. 35. Fu, B.; Bakker, E.; Yang, V. C.; Meyerhoff, M. E. Macromolecules 1995, 28, 5834−5840. 36. Fu, B.; Bakker, E.; Yun, J. H.; Wang, E.; Yang, V. C.; Meyerhoff, M. E. Electroanalysis 1995, 28, 5834−5840. 37. Dai, S.; Esson, J. M.; Lutze, O.; Ramamurthy, N.; Yang, V.; Meyerhoff, M. E. J. Pharm. Biomed. Anal. 1999, 51, 1-14. 38. Yun, J. H.; Han, Y. S.; Chang, L. C.; Ramamurthy, N.; Yang, V.; Meyerhoff, M. E. Pharm. Sci. Technol. To. 1999, 2, 102-110. 39. Durust, N.; Meyerhoff, M. E. J. Electroanal. Chem. 2007, 602, 138-141. 40. Ding, J.; Gu. Y.; Li, F.; Zhang, H.; Qin, W. Anal. Chem. 2015, 87, 6465-6469. 41. Shvarev, A.; Bakker, E. J. Am. Chem. Soc. 2003, 125, 11192–11193. 42. Shvarev, A.; Bakker, E. Anal. Chem. 2005, 77, 5221–5228. 43. Gemene, K. L.; Meyerhoff, M. E. Anal. Chem. 2010, 82, 1612−1615. 44. Shvarev, A.; Bakker, E. Anal. Chem. 2003, 75, 4541–4550. 45. Gemene, K. L.; Bakker, E. Anal. Biochem. 2009, 386, 276–281. 46. Crespo, G.A.; Afshar, M. G.; Baker, E. Angew. Chem. Int. Ed. 2012, 51, 12575-12578. 47. Xu, Y.; Shvarev, A.; Makarychev-Mikhailov, S.; Bakker, E. Anal. Biochem. 2008, 374, 366–370. 48. Fordyce, K.; Shvarev, A. Anal. Chem. 2008, 80, 827-833. 49. Gemene, K. L.; Meyerhoff, M. E. Anal. Biochem. 2011, 416, 67–73. 50. Bell-Vlasov, A.; Zajda, J.; Eldourghamy, A.; Malinowska, E.; Meyerhoff, M. E. Anal. Chem. 2014, 86, 4041-4046. 51. Wang, X.; Balijepalli, A. S.; Meyerhoff, M. E. Electroanalysis 2015, 27, 1-7. 52. Gemene, K. L.; Meyerhoff, M. E. Electroanalysis 2012, 24, 643-648. 53. Mckee, J; Bairstow, S; Szabo, C.; Ray, J.; Wielgos, T.; Hu, P.; Chess, E.; Nordhaus, M.; Hai, T.; Campbell, J. J. Clin. Pharmacol. 2010, 50, 1159-1170. 54. Zook, J. M.; Lindner, E. Anal. Chem. 2009, 81, 5146-5154. 55. Jarolimova, Z.; Crespo, G. A.; Xie, X.; Afshar, M. G.; Pawlak, M.; Bakker, E. Anal. Chem. 2014, 86, 6307-6314. 56.

Afshar, M. G.; Crespo, G. A.; Xie, X.; Bakker, E. Anal. Chem. 2014, 86, 6461-6470.

57. Pan. J.; Qian, Y.; Zhou, X.; Lu, H.: Ramacciotti, E. Zhang, L. J. Biol. Chem. 2010, 285, 22966-22975. 58. Corbier, A.; Berre, N. L.; Rampe, D.; Meng, H.; Lorenz, M.; Vicat, P.; Potdevin, S.; Doubovetzky, M. Toxicol. Sci. 2011, 121, 417-427.


13


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

Page 14 of 14

Table Of Contents (TOC) / Graphic Abstract


14

Reversible Electrochemical Sensor for Detection of High-Charge

Recommend Documents