CONTINUOUSLY CHEMICALLY ENHANCED APTAMER SENSORS

Abstract

A device for measuring the concentration of a target analyte in a sample fluid. The device (100) includes a housing (110) defining a plurality of chambers including at least a first chamber (112) and a second chamber (114). The first chamber includes a sensor fluid (132) therein, and the second chamber includes a reservoir fluid (134) therein. A sample fluid area (116) is also defined by the housing. This sample fluid area is capable of receiving a sample fluid (130) to be tested for the presence or concentration of a target analyte. A first element (140) separates the first chamber from the sample fluid area and restricts diffusion of solutes between the sensor fluid and sample fluid. A second element (142) separates the first chamber from the second chamber and restricts diffusion of solutes between the sensor fluid and the reservoir fluid. The device also includes at least one sensing electrode (120) that is positioned in the sensor fluid.

Description

TECHNICAL FIELD OF THE INVENTION

The present invention relates to aptamer sensors, and to devices and methods including aptamer sensors.

BACKGROUND OF THE INVENTION

This section is intended to introduce the reader to various aspects of art that may be related to various aspects of the present invention, which are described and/or claimed below.

This discussion is believed to be helpful in providing the reader with background information to facilitate a better understanding of various aspects of the present invention. Accordingly, it should be understood that these statements are to be read in this light, and not as admissions of prior art.

Electrochemical sensors such as enzymatic and aptamer-based sensors promise single-step and continuous sensing capabilities that are reagent-free and label-free, but inherent to this same promise is susceptibility to interference and degradation due to fouling, uncontrolled pH or salinity, or other solutes in the sample fluid. For at least pH and salinity, some sample fluids are inherently buffered, such as blood, but other emerging biofluids, such as human sweat or environmental fluids, can have widely ranging pH and salinity. A need exists for improved methods of (1) protecting electrochemical sensors from damaging pH and/or salinity levels (or damaging fluctuations in pH and/or salinity), and (2) protecting electrochemical sensors from other degradation mechanisms as well, such as oxidants, nucleases, or other solutes that can hinder or damage a sensor.

SUMMARY OF THE INVENTION

Certain exemplary aspects of the invention are set forth below. It should be understood that these aspects are presented merely to provide the reader with a brief summary of certain forms the invention might take and that these aspects are not intended to limit the scope of the invention. Indeed, the invention may encompass a variety of aspects that may not be explicitly set forth below.

Many of the drawbacks and limitations stated above can be resolved by creating novel and advanced interplays of chemicals, materials, sensors, electronics, microfluidics, algorithms, computing, software, systems, and other features or designs, in a manner that affordably, effectively, conveniently, intelligently, or reliably brings sensing technology into proximity with biofluid and analytes.

Various aspects of the disclosed invention are directed to aptamer sensors that overcome the drawbacks described above—such as by providing a device that protects aptamer sensors from damage or degradation due to harmful solutes in fluid. To that end, in one embodiment, such a device may be used for detecting the presence of, or measuring the concentration of, a target analyte in a sample fluid. The device includes a housing defining a plurality of chambers including at least a first chamber and a second chamber. The first chamber includes a sensor fluid therein, and the second chamber includes a reservoir fluid therein. A sample fluid area is also defined by the housing. This sample fluid area is capable of receiving a sample fluid to be tested for the presence or concentration of a target analyte. A first element separates the first chamber from the sample fluid area and restricts diffusion of solutes between the sensor fluid and sample fluid. A second element separates the first chamber from the second chamber and restricts diffusion of solutes between the sensor fluid and the reservoir fluid. The device also includes at least one sensing electrode that is positioned in the sensor fluid.

In various embodiments, the first and second elements (that restrict diffusion of solutes between the various fluids) may be chosen from membranes, encasements, and combinations thereof. Further, the reservoir fluid may include enhancing solutes. In this and alternate embodiments, the reservoir fluid may remove at least one harmful solute from the fluid sample.

BRIEF DESCRIPTION OF THE DRAWINGS

The objects and advantages of the disclosed invention will be further appreciated in light of the following detailed descriptions and drawings in which:

FIG. 1 is a diagram of a side view of a membrane sealed electrochemical aptamer sensor with a buffer reservoir according to the present invention.

FIG. 2 is a schematic of a device in accordance with principles of the present invention.

FIG. 3A is a schematic showing that an oil-membrane allows the partitioning of hydrophobic analytes such as cortisol, while blocking hydrophilic solutes such as proteins, salts, and acids/bases.

FIG. 3B is a schematic showing a gold working electrode functionalized with DNA aptamers terminated with a redox couple for an electrochemical aptamer sensor in accordance with principles of the present invention.

FIG. 3C is a schematic showing how the aptamers fold when bound to target analyte, which changes the proximity of the redox couple to the electrode compared to aptamers without target analyte, resulting in a change in electrical current that correlates with the change in analyte concentration.

FIG. 4A is a diagram of side view of an oil-membrane sealed electrochemical aptamer cortisol sensor.

FIG. 4B is a diagram of top view of an oil-membrane sealed electrochemical aptamer cortisol sensor.

FIG. 5A is a schematic showing that, during a concentration-on process, the hydrophobic analyte molecule (A) diffuses to the interface between oil membrane and biofluid, (B) rapidly partitions into the oil (decrease in free energy), (C) diffuses to the other end of the oil membrane, (D) more slowly partitions out of oil with a lag time (increase in free energy), (E) diffuses close to sensor surface, and (F) binds with aptamer.

FIG. 5B is a schematic showing the reverse process of that shown in FIG. 5A.

FIG. 5C is a simple circuit representation of the system (it should be noted that this representation is an approximation and not a theoretically equivalent model).

FIG. 6 is a cortisol titration curve of a cortisol aptamer; the titration curve in FIG. 6 was attained from electrochemical aptamer sensor functionalized gold rod disk electrodes [the cortisol solution is prepared using 1×PBS (pH at 7.5)].

FIG. 7 is a drawing of an of experiment setup for the device with buffer reservoir.

FIGS. 8A-8F are schematic representations of transwell setup steps for characterizing cortisol partitioning through different oils.

FIG. 8G is a graph showing ELISA results of cortisol diffusing through selected oils in the upper well solution after 2 hours of incubation at room temperature.

FIGS. 9A is a graph showing changes in chemical pH over time in an experiment for diffusion using different oils.

FIG. 9B is a graph showing changes in electrical resistance over time in an experiment for diffusion using difference oils.

FIG. 10 is a graph showing the response of water saturated membrane sealed sensors in solution at different salinity level on signal gain (solid line 1000 for Device 1 and solid line 1010 for Device 2) and peak position (dashed line 1020 for Device 1 and dashed line 1030 for Device 2 ) [in FIG. 10, two devices are functionalized (for process see the Methods-Sensor Functionalization section of the Examples section, below) using rod disk electrodes; the devices are tested among 6 beakers at different salinity level and different cortisol concentration level; and the devices are scanned once a minute and remained for a total of 10 minutes in each beaker (chemical environment)].

FIG. 11 is a graph showing the response of 1-decanol membrane sealed sensors with lx and 33×PBS buffer capacity at various pH sample environment, plotted as signal gain (solid line 1100 for 1×buffer and solid line 1110 for 33×buffer) and redox peak position (dashed line 1120 for 1×PBS buffer and 1130 for 33×PBS buffer).

FIG. 12 is a graph showing electrochemical aptamer cortisol sensor in solution with/without cortisol from pH 8.5 to 5.5. Sensor signal gain (solid lines) and redox couple peak position (dashed line).

FIG. 13A is a graph showing the response of electrochemical aptamer cortisol sensors (2 tests per data set) with oil-membranes fabricated with castor oil. The plots contain both the sensor signal gain (solid line) and the redox peak position (dashed line) in different testing environments.

FIG. 13B is a graph showing the response of electrochemical aptamer cortisol sensors (2 tests per data set) with oil-membranes fabricated with 1-decanol. The plots contain both the sensor signal gain (solid line) and the redox peak position (dashed line) in different testing environments.

FIG. 14A is a diagram of a side view of an oil membrane sealed electrochemical aptamer cortisol sensor with buffer reservoir.

FIG. 14B is a diagram of a top view of an oil membrane sealed electrochemical aptamer cortisol sensor with buffer reservoir.

FIG. 14C is a graph showing improved performance on signal gain (solid line) and peak position (dashed line) from 1-decanol membrane protected electrochemical aptamer cortisol sensors with addition of a buffer reservoir when diffusing different sample fluids.

FIG. 15 is a graph showing the response of 1-decanol membrane protected sensors on signal gain (solid line 1500 for Device 1 and solid line 1510 for Device 2) and peak position (dashed line 1520 for Device 1 and dashed line 1530 for Device 2) versus time and introduction of different sample fluids. These two devices are fabricated as the same design in FIGS. 14A and 14B, except that the membrane used here is a proprietary Gore membrane G (3 μm thickness) and the inner layer of Kimwipes™ is saturated with 3 μM cortisol solution instead of PBS buffer. The devices in FIG. 15 are fabricated following same process described in Method—Integration for Device with Buffer Reservoir in the Examples section (below), except that here the membrane used is a proprietary Gore membrane G (3 μm thickness) instead of polycarbonate track-etch membrane and the L-shape inner layer is initially saturated with 3 μM cortisol solution instead of buffer solution.

DEFINITIONS

As used herein, “continuous sensing” with a “continuous sensor” means a sensor that changes in response to changing concentration of at least one solute in a solution such as an analyte. Similarly, as used herein, “continuous monitoring” means the capability of a device to provide multiple measurements of an analyte over time.

As used herein, the term “about,” when referring to a value or to an amount of mass, weight, time, volume, pH, size, concentration or percentage is meant to encompass variations of ±20% in some embodiments, ±10% in some embodiments, ±5% in some embodiments, ±1% in some embodiments, ±0.5% in some embodiments, and ±0.1% in some embodiments from the specified amount, as such variations are appropriate to perform the disclosed method.

As used herein, the term “electrode” means any material that is electrically conductive such as gold, platinum, nickel, silicon, conductive liquid infused materials such as ionic liquids, PEDOT:PSS, conductive oxides, carbon, boron-doped diamond, nanotubes or nanowire meshes, or other suitable electrically conducting materials.

As used herein, the term “blocking layer” or “passivating layer” means a homogeneous or heterogeneous layer of molecules on an electrode which alter the electrochemical behavior of the electrode. Examples include a monolayer of mercaptohexanol on a gold electrode or as another example natural small-molecule solutes in serum that form a layer on a carbon electrode.

As used herein, the term “aptamer” means a molecule that undergoes a conformation or binding change as an analyte binds to the molecule, and which satisfies the general operating principles of the sensing method as described herein. Such molecules are, e.g., natural or modified DNA, RNA, or XNA oligonucleotide sequences, spiegelmers, peptide aptamers, and affimers. Modifications may include substituting unnatural nucleic acid bases for natural bases within the aptamer sequence, replacing natural sequences with unnatural sequences, or other suitable modifications that improve sensor function but which behave analogous to traditional aptamers.

As used herein, the term “redox tag” or “redox molecule” means any species such as small or large molecules with a redox active portion that when brought adjacent to an electrode can reversibly transfer at least one electron with the electrode. Redox tag or molecule examples include methylene blue, ferrocene, quinones, or other suitable species that satisfy the definition of a redox tag or molecule. In some cases, a redox tag or molecule is referred to as a redox mediator. Redox tags or molecules may also exchange electrons with other redox tags or molecules.

As used herein, the term “optical tag” or “fluorescent tag” means any species that fluoresces in response to an optical source such as LED and whose fluorescence is detectable by a photodetector such as a photodiode. Example fluorescent tags include fluorescein and may be used in combination with other fluorescent tags or optical quenchers such a black-hole quencher dyes to change the fluorescence of the optical tag.

As used herein, the term “analyte” means any solute in a solution or fluid which can be measured using a sensor. Analytes can be small molecules, proteins, peptides, electrolytes, acids, bases, antibodies, molecules with small molecules bound to them, DNA, RNA, drugs, chemicals, pollutants, or other solutes in a solution or fluid.

As used herein, the term “membrane” means a polymer film, plug of hydrogel, liquid-infused film, tiny pore, or other suitable material which is permiselective to transport of a solute through the membrane by solute parameters such as size, charge state, hydrophobicity, physical structure, or other solute parameters than can enable permiselectivity. For example, a dialysis membrane is permselective by passing small solutes but not large solutes such as proteins. Membranes as understood herein need not be multiporous, for example, a nanotube or nanopore can act as a permiselective filter and is therefore considered part of a membrane as understood for the present invention.

As used herein, the term “sample fluid” means any solution or fluid that contains at least one analyte to be measured, or any solution or fluid that is tested to determine the presence of at least one analyte therein.

As used herein, the term “sensor fluid” means a solution or fluid that differs from a sample fluid by at least one property, and through which the sensor solution and the sample fluid are therefore separated but are in fluidic connection through at least one pathway such as a membrane. The sensor solution may comprise at least one aptamer as a solute.

As used herein, the term “reservoir fluid” means solution or fluid that differs from sample fluid by at least one property, and through which the sensor solution and the reservoir solution are in fluidic connection through at least one pathway such as a membrane or a pin-hole connection. A reservoir fluid may have multiple function in a device, for example, by introducing a solute continuously or as needed by diffusion equilibrium into the sensor fluid, or for example removing unwanted solutes from a sensor fluid and acting as a “waste removal element”.

As used herein, a “device” comprises at least one sensor based on at least one aptamer, at least one sensor solution, and at least one sample fluid. Devices can sense multiple samples and be in multiple configurations such as a device to measure a pin-prick of blood, or a microneedle or in-dwelling sensor needle to measure interstitial fluid, or a device to measure saliva, tears, sweat, or urine sensor, or a device to measure water pollutants or food processing solutes, or other devices which measure at least one analyte found in a sample fluid.

DETAILED DESCRIPTION OF THE INVENTION

One or more specific embodiments of the present invention will be described below. In an effort to provide a concise description of these embodiments, all features of an actual implementation may not be described in the specification. It should be appreciated that in the development of any such actual implementation, as in any engineering or design project, numerous implementation-specific decisions must be made to achieve the developers' specific goals, such as compliance with system-related and business-related constraints, which may vary from one implementation to another. Moreover, it should be appreciated that such a development effort might be complex and time consuming, but would nevertheless be a routine undertaking of design, fabrication, and manufacture for those of ordinary skill having the benefit of this disclosure. All ranges of parameters disclosed herein include the endpoints of the ranges.

Certain embodiments of the disclosed invention show sensors as simple individual elements. It is understood that many sensors require two or more electrodes, reference electrodes, or additional supporting technology or features which are not captured in the description herein. Sensors measure a characteristic of an analyte. Sensors are preferably electrical in nature, but may also include optical, chemical, mechanical, or other known biosensing mechanisms. Sensors can be in duplicate, triplicate, or more, to provide improved data and readings. Sensors may provide continuous or discrete data and/or readings. Certain embodiments of the disclosed invention show sub-components of what would be sensing devices with more sub-components needed for use of the device in various applications, which are known (e.g., a battery, antenna, adhesive), and for purposes of brevity and focus on inventive aspects, such components may not be explicitly shown in the diagrams or described in the embodiments of the disclosed invention.

As described above, various aspects of the disclosed invention are directed to aptamer sensors that overcome the drawbacks described in the Background section—such as by providing a device that protects aptamer sensors from damage or degradation due to harmful solutes in fluid. In one embodiment, such a device may be used for detecting the presence of, or measuring the concentration of, a target analyte in a sample fluid. To that end, and with reference to FIG. 1, aspects of the present invention apply broadly to optical, aptamer, enzymatic, and other sensors, which may be part of a device for detecting or measuring target analyte in a sample fluid. As shown in FIG. 1, this embodiment of a device 100 includes a housing 110 (made of a material such as plastic) defining a plurality of chambers including at least a first chamber 112 and a second chamber 114. The first chamber 112 includes a sensor fluid 132 therein, and the second chamber 114 includes a reservoir fluid 134 therein. A sample fluid area 116 is also defined by the housing 110. This sample fluid area 116 is capable of receiving a sample fluid 130 to be tested for the presence or concentration of a target analyte. A first element 140 separates the first chamber 112 from the sample fluid area 116 and restricts diffusion of solutes between the sensor fluid 132 and sample fluid 130. A second element 142 separates the first chamber 112 from the second chamber 114 and restricts diffusion of solutes between the sensor fluid 132 and the reservoir fluid 134. The device also includes at least one sensing electrode 120 that is positioned in the sensor fluid 132 in the first chamber 112.

Additionally, a plurality of aptamers (not shown in FIG. 1) may be present in the sensor fluid 132 in the first chamber 112. By being present in sensor fluid 132, the plurality of aptamers is positioned to be able to come into contact with any target analyte in the sample fluid. There are multiple embodiments covering how the plurality of aptamers may be present in the sensor fluid 132. In one embodiment, the plurality of aptamers may be bonded to the sensing electrode 120. Thus, for example, when the device 100 utilizes a plurality of aptamers with a plurality of redox tags associated therewith as the detection mechanism, the sensor fluid 132 and electrode 120 can detect an analyte by bonding the aptamers to the electrode 120 via a mechanism such as thiol bonding of aptamers to a gold electrode. In an alternate embodiment, the plurality of aptamers could be in solution (for example, a large aptamer that is redox tagged with methylene blue could be free in the sensor solution). In this example, the availability of the methylene blue to the electrode 120 (for electron transfer with the electrode 120) could increase or decrease as analyte binding to the aptamer changes its geometry or folding configuration. These examples show that the embodiments of the invention apply broadly to multiple types of device configurations.

With further reference to FIG. 1, in its broadest sense, the first element 140 may be a membrane that protects the sensing electrode 120 by reducing the ability of at least one solute in sample fluid 130 that is potentially harmful to the sensor (such as a nuclease, or fouling proteins such as albumin) from reaching the aptamers, the sensing electrode 120, or other potential components of the sensor.

The first element/protecting membrane 140, of this embodiment, may also enhance the operation of the device 100 by at least partially retaining at least one sensor enhancing solute in the sensor solution 132. One example of a sensor enhancing solute is a buffer. The reservoir fluid 134, in certain embodiments, can also assist in this by including at least one sensor enhancing solute in the reservoir fluid 134, which can be introduced into sensor fluid 132 by passing through second element 142. Reservoir fluid 134 may provide additional functions as well, such as (1) removing at least one harmful solute from sensor fluid 132, and/or (2) removing analyte from sensor fluid 132 (e.g., if analyte easily travels through first element 140 (e.g., membrane) toward sensor fluid 132 but not back into the sample fluid 130, which is the case, for example, when the analyte is a hydrophobic analyte such as cortisol and the first element 140 is an oil-infused membrane). Second element 142 can also be a membrane such as polyethersulfone (PES) or hydrogel, such as an agar hydrogel, that slows diffusion between sensor and reservoir fluids 132, 134. Alternatively, second element 142 may be a long and/or narrow channel which achieves the same effect of slowing diffusion.

Generally, a device as described herein may operate such that a solute that is potentially harmful to the sensing electrode 120 will have lower total mass transport through first element (e.g., membrane) 140 than through second element (e.g., membrane) 142 such that if the harmful solute enters the sensor fluid 132 it will be subsequently removed from sensor fluid 132 by moving into the reservoir fluid 134. Further, any harmful solutes that may be generated by sensing electrode 120 itself, such as an oxidative species that is electrochemically created, can also diffuse through second element (e.g., membrane) 142 to be removed from the sensor fluid 132. While harmful solutes in sensor fluid 132 can move via second element into reservoir fluid 134, the device 100 also operates such that analyte present in sample fluid 130 moves (e.g., diffuses) through first element (e.g., membrane) 140 easily enough such that the concentration of analyte in fluid sample 130 and the concentration of analyte in the sensor fluid 132 are comparable (e.g., differing by less than 50%, or by less than 10%, or by less than 5%). In other words, unlike the harmful solute example discussed above, the analyte does not easily move (e.g., diffuse) through second element (e.g., membrane) 142.

Further, enhancing solutes that are introduced from reservoir fluid 134 via second element 142 should have an adequate enhancing effect on the sensing electrode 120; and, in embodiments, this may be accomplished by ensuring that the enhancing solutes not easily transverse through first element 140. In other words, any restriction on the diffusion of at least one enhancing solute from the reservoir fluid 134 to the sensor fluid 132 provided by the second element (e.g., membrane) 142 is less than any restriction on the diffusion of the at least one enhancing solute from the sensor fluid 132 to the sample fluid 130 provided by the first element (e.g., membrane) 140 Enhancing solutes may include but are not limited to salts, buffers, anti-oxidants, nuclease inhibitors, and molecules that help passivate the sensor electrode such as albumin or peptides. All or a subset of the above features may form an embodiment of the present invention. In one embodiment, wherein the first element and/or second element are membranes, each membrane may be chosen from liquid filled membranes (such as oil-membranes), hydrogels, channels or porous networks, filtration or size-selective membranes, and other suitable materials that achieve the above stated functionality.

With reference now to FIG. 2, where like numerals refer to like features, a portion of a device 200 includes a sample fluid 230, a sensor fluid 232, a reservoir fluid 234, a first element (e.g., membrane) 240 with mass flow 291, and a second element (e.g., pinhole, membrane, or other diffusion restrictive feature) 242 with a mass flow 293. Consider an example where a 0.2 kDa dialysis membrane is first element 240, and assume a plurality of aptamers as solutes in sensor fluid 232 that are 10-100× larger than the analyte to be detected (e.g. phenylalanine, cortisol, etc.). Next, assume the system is designed such that the volume of reservoir fluid 234 is one of at least 2×X, at least 10×, at least 50×, or at least 250× greater than volume of sensor fluid 232, and that the mass flow of aptamer (that occurs at reference numeral 291) is one of at least 2×, at least 10×, at least 50×, or at least 250× less than mass flow of aptamer (that occurs at reference numeral 293), while the mass flow of the analyte (that occurs at reference numeral 291) is one of at least 2×, at least 10×, at least 50×, or at least 250× greater than the mass flow of the analyte (that occurs at reference numeral 293). As a result, the concentrations of analyte will be within one of at least 50%, at least 10%, at least 2%, or at least 0.4% of each other in sensor fluid 232 and sample fluid 230. Reservoir fluid 234 is then able to continuously introduce solutes to sensor fluid 232 or to remove solutes from sensor fluid 232 through second element 242. Solutes to be removed could include aptamers that were cleaved by nucleases such that the redox couple is causing background interference, or solutes to be removed could be nucleases themselves, for example. In either case, the solute removal is enabled by diffusion because the sensor fluid 232 has at least 2×, at least 10×, or at least 100× higher concentration of the solute to be removed than the concentration in the reservoir fluid 234. As a geometrical example, consider a membrane 240 with 0.2 cm² area and 10% porosity to the analyte and a element 242 that is a pinhole in materials 210 and 220 that is 0.001 cm² in area and 0.001 cm in length; the mass transport for a small analyte through the membrane 240 will be equivalent to 0.02 cm² area and the mass transport through the feature 242 0.001 cm² which is 20× different, satisfying the above stated criteria for design as shown in FIG. 2.

EXAMPLES

Electrochemical sensors such as enzymatic and aptamer based sensors promise single-step and continuous sensing capabilities that are reagent- and label-free; but, inherent to this same promise is susceptibility to interference and degradation due to fouling and uncontrolled pH or salinity in the sample fluid. With regard to pH and salinity, some sample fluids are inherently buffered, such as blood. But other emerging biofluids, such as human sweat or environmental fluids, can have widely ranging pH and salinity. This Example presents an oil-membrane sensor protection technique which allows for permeation of hydrophobic (lipophilic) analytes into a sealed sensor compartment containing ideal salinity and pH conditions, while simultaneously blocking common hydrophilic interferents (proteins, acids, bases, etc.) Herein, the oil-membrane sensor protection technique is validated by demonstrating continuous cortisol detection via electrochemical aptamer based sensors. The encapsulated electrochemical aptamer cortisol sensor of this Example exhibits a 5-minute concentration-on rise time and maintains measurement signal of at least 7 hours even in the extreme condition of an acidic solution of pH 3.

Introduction

Electrochemical sensor demonstrations are highly prevalent for testing in (1) buffer fluid or (2) blood. This should not be surprising because these two fluids are highly relevant testing standards and because both fluids are well-buffered in their pH and salinity. Being well-buffered is important to electrochemical sensor operation because pH and salinity can significantly confound sensor response. This challenge can be particularly acute for continuous biosensors because unlike a single-use point-of-care sensor, pH and salinity are much more difficult to control over the longer time period associated with continuous sensing.

Enzymatic sensors are most commonly deployed for both real-time and point-of-care assays because of their ability to couple a biochemical reaction to a change in redox state of a co-factor (e.g., NADH and FADH2) that can be measured directly or indirectly. However, enzymatic sensors are particularly sensitive to changes in pH and salinity, as they are composed of amino acids. Enzymes, as well as other proteins, are inherently pH sensitive because of their labile side-chain protonation sites, and are salinity sensitive because of their charged functional groups. Therefore, any change in salinity and/or pH must be accounted for with regards to converting enzymatic activity to analyte concentration. Changes in salinity and/or pH— and/or failing to account for such changes—increases the risk of inaccurate measurement of analyte concentration when using enzymatic sensors.

Other biosensors that utilize biorecognition elements suffer from similar constraints, including electrochemical aptamer sensors invented by Plaxco and colleagues [Dauphin-Ducharme, P.; Yang, K.; Arroyo-Curras, N.; Ploense, K. L.; Zhang, Y.; Gerson, J.; Kurnik, M.; Kippin, T. E.; Stojanovic, M. N.; Plaxco, K. W. Electrochemical Aptamer-Based Sensors for Improved Therapeutic Drug Monitoring and High-Precision, Feedback-Controlled Drug Delivery. ACS sensors 2019, 4 (10), 2832-2837. https://doi.org/10.1021/acssensors.9b01616]. These sensors rely on the high binding affinity of the analyte to the aptamer. Aptamers are known to be sensitive to both salinity and pH, thus impacting sensor performance and analyte response. Additionally, electrochemical aptamer sensors often use redox couples that are pH sensitive, such as an immobilized methylene blue redox couple.

In biofluids, the majority of the most problematic solutes are hydrophilic, including salts, acids, bases, and larger molecules (such as proteins) that must have a hydrophilic shell to maintain their water solubility. Generalizing the problematic solutes as hydrophilic presents a significant opportunity for more robust sensing of hydrophobic solutes since hydrophilic/hydrophobic selective protection could theoretically be added to sensors. Implementing such a hydrophilic/hydrophobic filter would provide protection even beyond the widely deployed size-selective protective membranes such as those used for in-vivo glucose sensors and other electrochemical aptamer sensors. Furthermore, this hydrophilic/hydrophobic membrane could be adapted to filter out redox-active interfering agents such as the negatively charged FAD/FADH2 and NAD+/NADH coenzymes.

And so, in this Example (and in accordance with the principles of the disclosed invention), the present inventors present a novel approach of oil-membrane sensor protection, which allows for permeation of hydrophobic (lipophilic) analytes into a sealed compartment containing the sensor in desirable pH conditions, while the same oil-membrane simultaneously blocks protein foulants and pH interferents (acids/bases). In this Example, the present inventors specifically validate an oil-membrane sensor protection technique by demonstrating continuous cortisol detection via electrochemical aptamer sensors. The oil-membrane encapsulated electrochemical aptamer cortisol sensor described herein exhibits a 5-minute concentration-on rise time and a measurement signal over at least 7 hours, even under extremely acidic conditions of pH 3. Also discussed are novel methods for rapidly optimizing the oil-membrane technique for each new analyte and application.

From an application perspective, the novel oil-membrane approach presented here is important for numerous applications, including those with hydrophobic analytes, such as orally administered drugs (hydrophobic by design for gut permeation), and steroid hormones (cortisol, testosterone, estrogen, melatonin, etc.). Improved robustness to variable pH also satisfies an acute and immediate need for sweat biosensing, where pH can vary significantly Further, the present inventors believe that having sensor protection even more robust than size-selective membranes could be valuable, especially if the sensor is preserved in optimal buffer conditions. In addition, oil-membrane protection may prove important as biosensing moves into areas that require alternative fluids with non-ideal pH and salinity, such as environmental sensing or food processing. Many emerging potent bio-toxins are hydrophobic because of the need to rapidly permeate biological tissue, and therefore are ideal candidates for oil-membrane protection. Lastly, oil-membrane protection may also be useful because many hydrophobic analytes in biofluids tend to bind to transport proteins, such as albumin, thus greatly reducing the unbound drug concentration for detection by electrochemical sensors. Oil-membrane protection could allow for protein denaturation by using salts or pH outside the membrane barrier to release the hydrophobic analytes such that they are increased to measurable concentrations, while still preserving the sensor environment. Simply, oil-membrane sensor protection has the potential to open up numerous biosensing applications that are currently challenged or even unobtainable with electrochemical sensors.

Device Design

1. Electrochemical Aptamer Sensor for Cortisol Sensing

Electrochemical aptamer sensors are affinity-based biosensors which equilibrate their signal to the analyte concentration. Electrochemical aptamer sensors used in this Example were provided by Eccrine Systems, Inc., Cincinnati, Ohio. In these electrochemical aptamer sensors (FIGS. 3B and 3C), aptamers (oligonucleotides) 344 are immobilized on a gold working electrode 346 using thiol-linkers. Opposite to its thiol-end, the aptamer is tagged with a methylene-blue redox reporter 348. FIG. 3B is a representation of the functionalized working electrode in the “off” position, where the redox couple is extended away from the gold surface when the aptamer sits in its unfolded or partially unfolded state. Upon analyte binding, shown in FIG. 3C, the aptamer folds in on itself, changing the redox-couples 348 distance from the electrode surface. This activity alters the electron transfer rate from the redox couple to the working electrode, resulting in a change in electrical current measured by square-wave voltammetry. Comparing the current before and after binding of analyte, a signal gain can be calculated, which is proportional to the change in the target analyte concentration. Previous studies have shown that pH and salinity have a substantial impact on both binding affinity between the analyte and aptamer, and on the current generated by the pH-sensitive redox reporter. For this reason, the electrochemical aptamer sensor is an excellent platform to test the performance of an oil-membrane for sensor protection.

2. Adding Oil-Membrane Protection onto the electrochemical aptamer Sensor

The oil-membrane encapsulation (see FIG. 3A) involves an oil-impregnated hydrophobic membrane 340 which provides a barrier between the sensing compartment and the sample fluids 330, thereby blocking interferents and improving sensor performance. The oil membrane is designed to provide a constant, stable hydrophobic barrier that allows simultaneous permeation of hydrophobic analytes, such as cortisol molecules, while maintaining desirable aqueous conditions at the sensor 320, such as desirable conditions relative to salinity and pH.

FIGS. 4A and 4B are more detailed schematic representations of the oil-membrane protected electrochemical aptamer cortisol sensors used in this Example. This sensor device 400 uses a 3-electrode system. Four working electrodes 446 a share one common counter electrode 446b and one pseudo-reference electrode 446c. Gold is used for the surface of all electrodes; however, only the working electrodes feature the electrochemical aptamer sensor functionalization with the aptamer and redox couple. After the electrochemical aptamer sensors are fabricated, a layer of Kapton® polyimide film is applied as a chemical and electrical insulation layer. Next, the oil membrane 440 is sandwiched along its perimeter between two layers of adhesive 450, 452 and fixed to the Kapton/substrate 454. The placement of the oil-membrane onto the Kapton/substrate is performed while this device is submerged in a bath of buffer solution such that the buffer solution is sealed inside the device. Lastly, the edges of the oil membrane are further sealed against the Kapton/substrate by applying marine epoxy to improve robustness of the seal. A more detailed material list and fabrication process is included in the Methods section (below).

3. Physics Behind Oil-Membrane Operation and Resulting Design

The permeability of the oil membrane affects sensor response time. An oil membrane will reduce the permeability (diffusion) of analytes to the sensor according to Eq. 1:

$\begin{array}{cc}P=\frac{K\times D}{z}\left(\frac{\mathrm{cm}}{s}\right)& \left(\mathrm{Eq}.1\right)\end{array}$

where the partition coefficient (K) is defined as the ratio of the concentrations at equilibrium of the analyte molecules in the oil vs. in the water; D is the diffusion coefficient for the analyte in the oil; and z is the thickness of the oil layer. Equation 1 is informative to oil membrane design.

The permeability equation (Eq. 1) clearly shows that a thinner membrane and, therefore, thinner oil thickness (z) will increase the membrane permeability (P). For this reason, the present inventors chose to use 11 μm thick polycarbonate track-etch membranes instead of thicker hydrophobic membranes such as conventional porous Teflon films (z=10s to 100s of μm). The oil wicks into and remains in the track-etch membrane pores because of its lower interfacial surface tension with the polycarbonate than the water has with polycarbonate. The membrane permeability equation (Eq. 1) also clearly reveals that a high diffusion coefficient (D) matters as well. The diffusion coefficient of an analyte is dependent on the viscosity of the fluid it is diffusing through. Therefore, an important design element for the oil is to have a low viscosity (diffusion coefficient is inversely proportional to the viscosity of the oil [Walter, A.; Gutknecht, J. Permeability of Small Nonelectrolytes through Lipid Bilayer Membranes. J. Membr. Biol. 1986, 90 (3), 207-217. https://doi.org/10.1007/BF01870127]. Consider an informative example related to the materials used here. The optimal track-etch membranes for this work (see Table 1-below) have an 11 μm membrane thickness and 15.7% porosity, which is equivalent to the analyte having to diffuse through a uniform 70 μm of oil (11 μm/0.157). Now consider castor oil, with a viscosity about 730 times greater than that of water (see Table 2-below). An 11 μm thick castor oil membrane would then be equivalent to a 51,170 μm thick water diffusion distance (11 μum/0.157*730). Alternatively, using a low viscosity oil, such as 1-decanol, the 11 μm thick oil-membrane would have an equivalent thickness of only 945 μm diffusion distance through water (see Table 2).

TABLE 1
Available polycarbonate track-etch membranes' properties
Pore	Pore	Open	Nom.	Nom.
Size	Density	Area	Weight	Thickness
(D, μm)	(pores/cm2)	(%)	(mg/cm2)	(μm)
0.1	4*108	3.1	0.7	6
0.2	3*108	9.4	1.1	10
0.4	1*107	12.6	1.0	10
1.0	2*107	15.7	1.1	11
3.0	2*106	14.1	0.9	9
5.0	4*105	7.9	1.1	10
8.0	1*105	5.0	0.8	7

TABLE 2
Equivalent diffusion distance through
oil-membrane compared to water
						Equivalent	Equivalent
		Vis-	Pore	Open	Thick-	oil	water
	Log	cosity	size	area	ness	thickness	thickness
Oil	P	(cP)	(μm)	(%)	(μm)	(μm)	(μm)
Castor	18	650	0.1	3.1	6	194	141,356
oil			0.2	9.4	10	106	77,695
			0.4	12.6	10	79	57,963
			1.0	15.7	11	70	51,170
			3.0	14.1	9	64	46,617
1-	5	12	0.1	3.1	6	194	26,10
Decanol			0.2	9.4	10	106	1,434
			0.4	12.6	10	79	1,070
			1.0	15.7	11	70	945
			3.0	14.1	9	64	861

Now turning to the oil/water partition coefficient K: Although Equation 1 suggests that a high K is advantageous, it is actually misleading. In a static case, where concentration gradients are held constant on both sides of the oil membrane and the analyte is constantly diffusing through a membrane, Eq. 1 states that the highest possible K is beneficial. However, a very high K can be problematic in dynamic cases. An electrochemical aptamer sensor has to dynamically equilibrate to an often-changing analyte concentration in the biofluid; therefore, it is more important to understand the specific impact of a high K with rising or falling analyte concentrations on the biofluid side of the oil membrane. FIGS. 5A-5B illustrate the dynamic case for the concentration-on and -off process, which will now be described in detail; and then the problematic nature of a high K will be explained.

Referring to FIG. 5A, when the cortisol concentration increases on the biofluid side of the membrane 540 (biofluid shown with reference number 530), the analyte 556 first diffuses to the interface between the biofluid and the oil membrane (shown at arrow “A”). It then partitions into the oil 558 (shown at arrow “B”), followed by its diffusion to the other end of the oil pore inside the membrane (shown at arrow “C”). Due to their hydrophobic nature, analytes such as cortisol energetically prefer to remain in the oil (lower free energy) [Menichetti, R.; Kanekal, K. H.; Bereau, T. Drug-Membrane Permeability across Chemical Space. ACS Cent. Sci. 2019, 5 (2), 290-298. https://doi.org/10.1021/acscentsci.8b00718; Bedrov, D.; Smith, G. D.; Davande, H.; Li, L. Passive Transport of C60 Fullerenes through a Lipid Membrane: A Molecular Dynamics Simulation Study. J. Phys. Chem. B 2008, 112 (7), 2078-2084. https://doi.org/10.1021/jp075149c], and additional retention time can occur as the analyte attempts to leave the oil and partition into the water by the sensor (shown at arrow “D”). Afterwards, analytes diffuse closer to the sensor surface (shown at arrow “E”) and bind with the aptamer 544 (shown at arrow “F”) when the analyte concentration nears the binding affinity of the aptamer. As shown in FIG. 5B, the entire process is reversible.

A high K is problematic for two reasons. First, the oil can become a huge sink for the analyte. Consider a K-value of 10,000 using the 11 μm track-etch membrane discussed above with 15.7% porosity. With the concentration of the analyte in the oil being 10,000 times higher than that in the water, the “effective volume” of the oil in terms of analyte capacity is equivalent to 11 μM* 0.157*10,000=17.27 cm of water. Clearly, K cannot be too large for this reason. A second reason why a high K is problematic is the oil retention of the analyte (lower free energy in the oil than in the water). A higher K can cause the oil to retain the analyte and induce an additional lag time as the analyte attempts to leave the oil and partition back into water. For the concentration-on process, this oil-to-water lag-time is not a major concern for the device because the volume of the buffer solution 532 is small, which reduces the total amount of analyte that must partition into the buffer solution (short lag time). For the concentration-off process, however, it is difficult for the analyte concentration in the buffer solution to decrease quickly, because the analyte concentration in the oil must first decrease, which is limited by the high K value of the oil (inherently a very high concentration of the analyte). Therefore, the oil-to-water lag time as the analyte goes from the oil back into the biofluid can create a major bottleneck for the concentration-off process, because the oil has acquired such a high concentration of the analyte. This much slower concentration-off response time will be observable later in the experimental results.

Although discussion herein on the oil-membrane physics remains focused on analyte transport, it also affects device design for blocking foulants and interferents. K will always have some finite value, even for hydrophilic acids, bases, and salts (it is never zero), and K depends on the charge state of the solute, which can further depend on the pH of the biofluid (solute ionization constants pKa and pKb). For example, a solute might be found primarily in its charged state (98%) within a biofluid of a specific pH. The remaining 2% remains in an uncharged state and may diffuse through the protective oil membrane barrier. For this reason, it is important to understand the effects of pH on analyte properties. Therefore, as the experimental data will show, the oil membrane provides limited protection over time.

Lastly, oil viscosity and oil partition coefficient (K) are not the only relevant materials parameters for oil optimization (See Table 3-below). For most sensor applications the oil must remain liquid at room temperature, while also having low vapor pressure, such that it is not rapidly lost during assembly or potential dry storage. Solubility of the oil in water is also critical because eventual oil loss into water could cause failure in the oil-membrane protection of the sensor. A simple example calculation is as follows. Consider a device that is brought into contact with 0.3 μL of biofluid per minute and a sensor and membrane area of 0.1 cm2 with the membrane containing 0.0173 μL of oil. Assuming that all the oil that could partition into the water does so at any given time (an unrealistic but instructive assumption), if the device were to operate continuously for 24 hours without losing all the oil to the biofluid, the water solubility of the oil would need to be less than 36 mg/L.

TABLE 3
Selected oils that were initially screened along with their relevant properties. LogK is the logarithm
of the ratio of molecule concentration of the oil in octanol and water at equilibrium.
		LogK		Vapor				Double
	Melting	(water	Viscosity	pressure	Solubility	Density		bonds
	point	over	at 25° C.	at 25° C.	in water	at 25° C.	Carbon	(in	Structural
	(° C.)	octanol)	(cP)	(mmHg)	(mg/L)	(g/mL)	#	chain)	family
Tetradecane	6	8.19	2.13	0.0369	0.00033	0.764	14	0	Saturated
									hydrocarbon
Castor oil	−10 to −12	17.72	650	<0.097	<0.001	0.961	57	3	Trigyceride
									mixture
1-Decanol	6.67	4.57	12	0.00851	30	0.830	10	0	Fatty alcohol
Dioctyl	−7.6	6.9	0.224	0.005	—	0.820	16	0	Dialkyl ether
ether
Oleic acid	16.3	7.64	27.64	0.0000005	0.01	0.895	18	1	Unsaturated fatty
									acid
Vitamin K1	−20	10.9	—	—	—	0.984	31	1	Polycyclic
									aromatic Ketone
Mineral oil	—	—	—	<0.1	—	0.83	—	—	—
Dodecane	−7 to −9	6.1	—	0.000015	<0.2	0.845	12	0	Alkyl thiol
thiol

Methods

1. Materials

All oils, reagents and cortisol solution were purchased from Sigma-Aldrich (St. Louis, MO). The 96 transwell plate was purchased from Sigma-Aldrich (St. Louis, MO). The polycarbonate track-etch membrane with 1 μm diameter pore size was obtained from Sterlitech Corporation (Kent, WA). The cortisol aptamer solution and Synthetic Sweat Solution were obtained from Eccrine Systems, Inc (Cincinnati, OH). The titration curve for the cortisol aptamer is shown in FIG. 6. The cortisol aptamer, like other small molecule aptamers, does not require regeneration (inherently fast on and off response times). Samples of adhesive materials (Crystal-clear Gorilla Tape, Kapton® polyimide double side tape and marine epoxy) were purchased from Amazon (Seattle, CA). The acrylic was purchased from McMaster-Carr (Princeton, NJ).

2. Sensor Functionalization

The gold planar electrodes were manufactured using a Temescal FC-1800 E-Beam Evaporator from Ferrotec (CA, USA). Ti (40 nm thick) was deposited on the glass slide as an adhesion promoting layer prior to the 200 nm gold layer. A layer of Kapton® polyimide film, laser cut to size (geometric design shown in FIG. 4), was applied on top of the glass substrate.

For cleaning, the electrode array was first sonicated for 5 minutes. Then it was connected to a CHI E600 electrochemical analyzer through 5252 SOIC clips from Pomona Electronics (CA, USA). Each foot was connected to an individual working channel A Pt wire electrode and a Ag/AgCl electrode were wired as a common counter electrode and common reference electrode, respectively, shared by six channels. The electrode array was immersed in 0.5 M H₂SO₄ solution. Cyclic voltammetry was applied to a 1-V/s scan rate from 0 to 1.6 V to electrochemically clean the surface of the electrode. The surface was rinsed with DI water and dried under an air gun. This was followed by O₂ plasma cleaning for 2 minutes.

A 400 nM cortisol aptamer solution was drop casted onto the surface of the working electrodes and allowed to incubate in the dark for an hour. The cortisol aptamer was obtained from Eccrine Systems, Inc (Cincinnati, OH). The remaining solution was shaken off after incubation. Then 5 mM mercaptohexanol (MCH) solution from Sigma-Aldrich (St. Louis, MO) was used to further passivate the surfaces of the working, counter, and reference electrodes. The functionalized electrodes were stored in a dry hood for 2 hours and protected from the light. When this process was completed, the surface was rinsed with DI water to get rid of extra MCH.

The gold rod electrode was purchased from CH Instruments, Inc (Austin, TX). The cleaning and functionalization process were same as those of the planar electrodes.

3. Integration for Device without Buffer Reservoir

The oil-membrane device shown in FIGS. 4A and 4B includes four sub-assemblies: the sensor substrate 456, the top adhesive assembly 450, the bottom adhesive assembly 452 and 454, and the membrane 440. The membrane was cut to fit the 15 mm×2 mm operation window. The bottom adhesive component was composed of a layer of Crystal-clear Gorilla Tape and a layer of Kapton polyimide double-sided tape. The top adhesive was a layer of Kapton® polyimide double-sided tape attached to a 1.25 mm thick acrylic. Both adhesive components were laser cut into a 20×4 mm rectangle with 15 mm×2 mm open window. And the polycarbonate track-etch membrane was placed between the two adhesives with both sides facing the Kapton® polyimide double-sided tape.

For assembly, the functionalized sensor was first placed into a 100 mm diameter petri dish filled with PBS solution. With the oil-membrane and bottom- and top-adhesive assemblies all attached to each other, they were then placed onto the sensor electrode array. Once assembled, the device was removed from the solution, the extra solution on its outside was dried, and the device was quickly sealed with Brampton marine epoxy from Brampton Technology LTD. (CT, USA) along the perimeter of the device. After 30 minutes, and once the epoxy was cured, the sensor was ready for testing.

4. Integration for Device with Buffer Reservoir

In the device, the sensor electrode array 720 was connected to 5252 SOIC clips from Pomona Electronics (CA, USA) and placed flat. A 10 cm diameter petri dish was filled with lx PBS solution and placed right next to the sensor as a buffer reservoir (with reservoir fluid 734). A 5 mm wide L-shape cut-out of a single layer of Kimtech Science™ Kimwipes™ Delicate Task Wipe was fully saturated with PBS solution. One 5 mm long leg 760 of the L-shape Kimwipes™ tissue was applied onto the sensor. The other leg 762 was connected to the buffer reservoir. A layer of polycarbonate track-etch membrane saturated with oil 764 was applied on top of the Kimwipes™ tissue. A 5 mm wide strip was cut out from a single layer of Kimtech Science™ Kimwipes™ Delicate Task Wipe, wetted with buffer solution, and applied on top to pass sample fluid from a syringe pump 766 over the oil membrane to waste pump. The sample fluid 730 was pumped at 4 μL/min. A 5 mm×5 mm×3 mm acrylic block was then placed on top of the strip as a weight to hold all the layers compressed against each other. This device is shown in the drawing of FIG. 7.

5. Data Generation and Analysis

During the test, the sensor was connected to a CHI E600 (CH Instruments, Inc, Austin, TX) through 5252 SOIC clips. Square wave voltammetry was used within a scanning window of 0 to -0.5 V, with amplitude of 0.035 V and frequency of 500 Hz (for rod electrodes) or 120 Hz (for planar electrodes). The raw data was exported from a saved text file from the CH Instrument (Austin, TX) Software. The signals were processed by a customized model in MATLAB (available at https://www.mathworks.com/products/matlab.html). The signal gain was measured by reading the highest current within two manually set potential points and then subtracting the baseline linearly matched by these two set points.

Experiments and Results

1. Initial Oil Screening Test

As noted in the Device Design section (above), oil choice is a consideration for the successful realization of the oil-membrane technique. The nine different oils listed in Table 3 were initially selected and tested. Initial selection parameters are based on low water solubility and low vapor pressure, to ensure long-term oil integrity within the membrane pores. In addition, oils with positive octanol/water K values are favored because of the hydrophobic nature of the test analyte (cortisol) while achieving a low permeability for hydrophilic interferents (pH and salinity).

In order to rapidly screen the oil candidates, a commercially available transwell setup was modified to measure the concentration of diffused cortisol through transwell membranes with different oils. FIGS. 8A-8E show diagrams of the experimental process. First, the bottom well 868 was filled with 235 μL of lx phosphate buffered saline (PBS) solution with a cortisol concentration of 10 μM 830 (FIGS. 8A and 8B). The traditional transwell membrane of the upper well was removed and replaced with a hydrophobic polycarbonate track-etch membrane used throughout this work (11 μm thick, 15.7% porosity, 1 μm diameter pore size). Each oil was applied to individual track-etch membrane, and the upper transwell 870 was placed into the bottom wells 868 (FIG. 8C). Then, as shown in FIG. 8D, the upper transwell was filled with 180 μL of cortisol-free lx PBS solution 832. The 96-well plate was then covered and incubated for 2 hours at room temperature to allow the cortisol to diffuse from the bottom well to the upper well through the oil-soaked membrane 840. The solution in the upper well was collected for analysis (FIG. 8E) using commercially available ELISA kits.

FIG. 8G displays the ELISA cortisol concentration results obtained by different oil membranes. The solution from the 1-decanol membrane sample contains the most cortisol—more than 5× the cortisol concentration of the second-ranking tetradecane membrane. The third-ranking membrane, castor oil, possesses less than half of the cortisol concentration of the tetradecane membrane. Castor oil is known to be an excellent oil for dissolving drugs and steroid hormones [Huber, C. R. R.; Keysser, C. H. Castor Oil as a Vehicle for Parenteral Administration of Steroid Hormones. J. Pharm. Sci. 1964, 53 (8), 891-895. https://doi.org/10.1002/jps.2600530809; Ogunniyi, D. S. Castor Oil: A Vital Industrial Raw Material. Bioresour. Technol. 2006, 97 (9), 1086-1091. https://doi.org/https://doi.org/10.1016/j.biortech.2005.03.028] and therefore likely possesses a very high K for cortisol. Also, the high viscosity of the castor oil should slow the diffusion of the cortisol through the oil (Eq. 1) resulting in lower cortisol concentration results for castor oil than for the lower-viscosity 1-decanol [Hiss, T. G.; Cussler, E. L. Diffusion in High Viscosity Liquids. AIChE J. 1973, 19 (4), 698-703. https://doi.org/10.1002/aic.690190404].

2. Directly Testing Oils Against Salt and pH Permeability

Before testing the integrated devices, the top three-performing oils (1-decanol, tetradecane, castor oil) were tested directly against high salt solution and extreme acidic solution. A U-boat setup was employed to measure salt diffusion from one buffer solution compartment to another separated by an oil membrane. For the salt diffusion test, a 10 mL buffer solution containing 0.15 M NaCl was placed in a right (control) chamber and 10 mL distilled (DI) water was placed in a left (monitored) chamber. The two chambers were separated by an oil-impregnated membrane secured by two rubber washers. The electrical resistance of the solution was measured in 60 seconds increments using a CHI 600E potentiostat, as shown in FIG. 9B. A similar setup was used for the pH diffusion test. For this test, 10 mL solution at pH 3 was placed in the control (right) chamber and 10 mL DI water was placed in the monitor (left) chamber. The pH value of the monitored solution was measured in 60 seconds increments using an Orion Star™ benchtop pH meter (Thermo Fisher Scientific Inc, Waltham, MA), as shown in FIG. 9A. Test results with highly alkaline solution are not presented here, since the initial focus is on the application of oil membrane in biofluids, such as sweat and saliva whose pH ranges from 4.5 to 7.5.

As shown in FIGS. 9A and 9B, castor oil shows the best performance in terms of blocking interfering species, tetradecane the second, and then 1-decanol the least. The castor oil-saturated membrane maintains the pH within half a unit over 12 hours. Within 8 hours, the tetradecane-saturated membrane can maintain pH changes within one unit. However, 1-decanol fails to maintain the pH within two units after 10 hours. Although it is expected that acids and bases might diffuse through an oil, surprisingly, simple salts such as NaCl diffuse through as well. This becomes less surprising when considering that the octanol/water LogK for Cl− is 0.06 and that for Na+ is −0.77 (low, but adequate to permit diffusion through the oil). Fortunately, for the cortisol aptamer sensor demonstrated in this paper, salinity change does not negatively influence the sensor performance. The salinity test (see FIG. 10) shows higher salt concentration increases the signal current magnitude and to some level increases the absolute signal gain on target concentration. Given the above, the present inventors speculate that by choosing and/or blending different oils, one can likely further increase the resistance of the oil membrane to salt diffusion. Furthermore, as the present inventors will discuss in the next sections, devices were demonstrated as stable with very high salt conditions in the sensor buffer solution (33× buffer, FIG. 11), and as stable with continuous salt mitigation (FIG. 11). Lastly, to confirm that the oil-membrane effectively blocks protein interferents, the present inventors tested gold and boron-doped diamond electrodes that were protected by the oil-membrane and exposed to serum for >12 hours and observed no protein fouling (insignificant changes in electrode impedance/electrode transfer). With the results of FIGS. 8G, 9A, and 9 B, tetradecane, 1-decanol, and castor oil were advanced to the next phase of experimentation with integrated aptamer sensors for cortisol.

3. Initial Sensor Tests as Control Experiments

The sensors were first characterized directly for the effects of pH on sensor response in order to aid in the interpretation of electrochemical aptamer sensor data. To initially find the optimal pH environment for the specific cortisol-binding aptamer, a three-electrode system was tested: an aptamer-functionalized gold disk electrode, a Pt wire counter electrode, and an Ag/AgCl reference electrode. The sensor system was tested for 5 minutes in each solution—with and without cortisol—to measure the on/off signal performance The pH of the sensor solution was varied from pH 8.5 to 5.5. The activity of methylene blue was demonstrated by the signal gain as a function of time pre- and post-cortisol exposure (FIG. 12). The highest signal gain was observed at pH 7.5. As the solution becomes more acidic, the signal gain decreases, and the redox peak shifts towards to a more positive potential.

As an additional control experiment to further reveal the negative impact of pH and the positive impact of oil-membrane protection, another U-boat test setup in a similar manner to that described above was implemented but using a cortisol electrochemical aptamer sensor to monitor sensor functionality. The monitor side was filled with 10 mL of synthetic sweat solution (SSS) at pH 7.5 without cortisol, while the control side was filled with SSS containing 10 μM cortisol concentration at pH 3. Castor oil was used as the oil in the oil membrane. The sensor with the oil-membrane protection responded to cortisol by an increase in redox-couple peak current, and the redox-couple peak position (potential) remained stable over time. Conversely, the same experiment but without oil in the membrane (water-filled) showed that the signal gain response decreased, and the redox peak rapidly shifted in potential. With these results in hand, the next set of experiments pursued the testing of fully integrated devices.

4. Fully Integrated Device Tests

For the remainder of experiments, fully integrated devices were tested using components previously illustrated in FIGS. 4A and 4B. SSS at pH 7.5 was again used as the solution on the sensor-side of the oil-membrane. For the “biofluid” side of the oil-membrane, bulk solution was varied in pH (pH 3 or pH 7.5) and cortisol content (0 μM or 10 μM).

Initially, on the basis of the results in the previous sections, the three best-performing oils (1-decanol, castor oil, and tetradecane) were chosen for the fully integrated device tests. However, the tetradecane was not compatible with the epoxy used to assemble the sensor, nor does it remain stably wetted in the membrane pores (the interfacial tension being potentially too large with respect to the polycarbonate). As a result, FIGS. 13A and 13B only presents the data for castor oil and 1-decanol.

And so, the device concentration-on response time shown in FIGS. 13A and 13B will first be discussed. As discussed in Device Design section, the cortisol diffusion time through the membrane and the resulting device on/off lag time should be affected by both K and viscosity. The present inventors do not have established information or experiment data on the K-value of castor and 1-decanol in relation to cortisol. However, the present inventors can proportionally estimate them on the basis of the LogK(octanol/water) value of castor oil, 1-decanol, and cortisol. Castor oil has a higher LogK(octanol/water) value, which makes it more hydrophobic. The LogK(octanol/water) of 1-decanol is lower but closer to the LogK(octanol/water) value of cortisol than that of castor oil, which means that cortisol is more likely to easily diffuse in and out of 1-decanol as opposed to castor oil which may sequester cortisol within the oil membrane. At the same time, castor oil is 50× more viscous than 1-decanol (Table 3). The expected slower response time for castor-oil-membrane sealed sensor is reflected in FIGS. 13A and 13B, where it takes more than 30 minutes for the signal to reach 90% of its plateau to a 10 μM cortisol change for concentration-on process, while it takes less than 5 minutes for the 1-decanol-membrane sealed sensor.

Next, the device concentration-off response time is discussed. As shown in FIGS. 13A and 13B, for both castor oil and 1-decanol, the response to decreasing cortisol concentration was much slower and arguably unmeasurable compared to the declining signal as a result of gradual sensor degradation over time. At this time, the absence of an off response is speculated to be due to retention in the oil, as discussed in the Device Design section. It should be noted that the present inventors also tested a well-stirred approach for the biofluid which should have increased the diffusion-promoting concentration gradient at the oil membrane/biofluid interface; however, no improvements in concentration-off time were seen.

Lastly, to confirm that each test uses an oil membrane with high integrity (no leaks) and to test the ability of the oil-membrane to protect against pH, the final step in the experiments in FIGS. 13A and 13B was to place the device in a solution of pH 3 and no cortisol content. A pH 3 solution is an extreme testing environment—beyond what is needed for pH variability in most biofluids—but has been tested here to quickly reveal any failure of the oil-membrane protection. As shown in FIGS. 13A and 13B, only small changes in the redox peak position were observed for the castor-oil-membrane sealed sensor; in contrast, for the 1-decanol-membrane sealed sensor, the signal gain started to decrease, while the redox peak position simultaneously shifted toward a more positive potential after 40 minutes. According to the results from the disk sensor test, this shift in redox potential indicates that the sensing solution is becoming more acidic. In other words, charged pH-influencing molecules are partitioning through the oil membrane containing 1-decanol, likely due to its lower K-value and viscosity compared to castor oil. One last observation based on the data of FIGS. 13A and 13B is that, for the castor oil test group, the redox peak does not shift towards a more positive potential, but there is a slow gradual negative potential shift. It is speculated that this could be due to castor oil having a slight water solubility and high molecular heterogeneity. As a result, the hydrophobic tails of castor oil could result in fouling the surface of the gold counter electrode. This would result in greater electrical resistance and, therefore, greater potential to generate the redox peak.

5. Stronger Buffer to Improve pH Protection

With 1-decanol as the highest-performing oil in terms of a fast concentration-on time, but also with reduced protection to pH, a simple solution to improve pH protection was next explored. Simply, the solution on the sensor side is more strongly buffered. The present inventors enhanced the buffer capacity of the SSS sensor solution by adding PBS powder to a final concentration of either lx PBS or 33×PBS. Results shown in FIG. 11 are SSS/1×PBS solution and SSS/33×PBS solution on the sensor side of the oil membrane. A higher buffer capacity notably increased the device longevity for both buffer conditions compared to no additional buffer. The redox peak position of the sensor with lx buffer capacity holds for 140 minutes when exposed to an acidic environment and 440 minutes for the sensor with 33× buffer capacity. Importantly, the lag time for cortisol diffusing through the oil membrane remains between 5-7 minutes. A doubled signal gain was observed from the sensor with 33× buffer capacity. However, with the hand-fabricated sensors used in this Example, variations between the sensors in their signal gain were seen. At 33× buffer capacity, the NaCl concentration is about 4.6 M, supporting the earlier claim (above) that the electrochemical aptamer cortisol sensor operates robustly even in high-salt conditions.

6. An Additional Buffer Reservoir Resolves Concentration-off Challenges and Buffering Capacity

The final design of the device pursued in this work is arguably the most compelling from an application perspective, as it resolves both the longevity concerns of the sensor system as well as the concentration-off response-time challenges discussed both theoretically and shown experimentally in previous sections. As shown in FIGS. 14A-14B, a buffer reservoir containing lx PBS is added with a diffusion-limited connection to the sensor solution side of the oil-membrane. In FIGS. 14A-14C, a constant flow of biofluid is also provided for the first time in this Example. The addition of the buffer reservoir has two purposes. First, the buffer reservoir is able to constantly diffuse in additional buffer to mitigate any change in pH due to the biofluid. This simply requires that the buffer reservoir pathway for additional buffer is less resistant to the diffusion of buffer than the diffusion of acids and bases through the oil membrane pathway (this is easily achievable). Second, the buffer reservoir provides another diffusion pathway to continuously remove the analyte, theoretically resulting in a faster concentration-off response. The experimental results shown in FIG. 14C fully support this expected behavior. By adding the buffer reservoir, the concentration-off process is greatly quickened: the concentration-off time is reduced to 10 minutes at 90% of signal off. Furthermore, there appears to be no limitation on buffer capacity, even in the highly challenging test at pH 3 for the biofluid. It should be noted that the concentration-on process is slowed to about 10 minutes for 90% signal on. This is not unexpected, given that a constant flow of biofluid approach is used in FIGS. 14A-14C and because the buffer reservoir is constantly removing cortisol. Although the approach in FIGS. 14A-14B is unoptimized in this work, the addition of a buffer reservoir is an elegant and simple solution to observed challenges in both buffer capacity and concentration-off response time Thinner 3 μm membranes with uniform porosity were also attempted, which increased the oil/water interface by ˜10× (FIG. 15).

Conclusions

In this Example, the efficacy of an oil-membrane sensor protection approach, including its design and operation with cortisol analyte both in the presence and absence of interferents is demonstrated. Although the cortisol aptamer sensors results can be highly variable from sensor-to-sensor in terms of signal-gain, the effects of oil-membrane protection are consistent and clear across all results and all figures. The results demonstrate the basic feasibility of the oil membrane approach for sensor protection and does so over time scales (>3 hours) and response time (1's to 10's of minutes) that are relevant for both point-of-care and continuous biosensing applications. The buffer-reservoir approach of FIGS. 14A-14C may be the preferred approach for continuous biosensing, given its ability to allow the sensor to respond to both increases and decreases in cortisol concentration, as well as to provide buffering of the sensor for prolonged periods. Although only cortisol is demonstrated in this Example, a rapid oil-optimization can be implemented for each new analyte, as was shown in FIGS. 8A-8G.

While the present invention has been disclosed by reference to the details of preferred embodiments of the invention, it is to be understood that the disclosure is intended as an illustrative rather than in a limiting sense, as it is contemplated that modifications will readily occur to those skilled in the art, within the spirit of the invention and the scope of the amended claims.

Claims

1. A device for measuring the concentration of a target analyte in a sample fluid comprising: a. a housing defining a plurality of chambers including at least a first chamber and a second chamber, the first chamber including a sensor fluid therein, and the second chamber including a reservoir fluid therein;b. a sample fluid area defined by the housing, the sample fluid area capable of receiving a sample fluid;c. a first element that separates the first chamber from the sample fluid area and restricts diffusion of solutes between the sensor fluid and sample fluid;d. a second element that separates the first chamber from the second chamber and restricts diffusion of solutes between the sensor fluid and the reservoir fluid; ande. at least one sensing electrode positioned in the sensor fluid.

2. The device of claim 1, wherein the sensor fluid contains a plurality of aptamers.

3. The device of claim 2, further comprising a plurality of tags associated with the plurality of aptamers, the plurality of tags selected from the group consisting of redox tags and optical tags.

4. The device of claim 1, wherein the at least one sensing electrode contains a plurality of aptamers.

5. The device of claim 4, further comprising a plurality of tags associated with the plurality of aptamers, the plurality of tags selected from the group consisting of redox tags and optical tags.

6. The device of claim 1, wherein the reservoir fluid contains at least one enhancing solute.

7. The device of claim 6, wherein the at least one enhancing solute is selected from the group consisting of a buffer, a salt, an antioxidant, an inhibitor, and a molecule that passivates the sensing electrode.

8. The device of claim 6, wherein any restriction on the diffusion of the at least one enhancing solute from the reservoir fluid to the sensor fluid provided by the second element is less than any restriction on the diffusion of the at least one enhancing solute from the sensor fluid to the sample fluid provided by the first element.

9. The device of claim 8, wherein the at least one sensor enhancing solute is partially retained in the sensor solution.

10. The device of claim 1, further comprising a passivating layer of molecules on the at least one sensing electrode.

11. The device of claim 10, wherein the passivating layer includes a monolayer of mercaptohexanol.

12. The device of claim 1, wherein the sensor fluid includes at least one solute harmful to the at least one sensing electrode, wherein the second element is adapted to allow the at least one solute harmful to the at least one sensing electrode to diffuse from the sensor fluid into the reservoir fluid.

13. The device of claim 12, wherein the harmful solute in the reservoir fluid is at a first concentration and the harmful solute in the sensor fluid is at a second concentration, wherein the first concentration is lower than the second concentration in an amount selected from the group consisting of greater than 2×, less than 10×, and less than 100×.

14. The device of claim 1, wherein the sample fluid includes at least one solute harmful to the at least one sensing electrode, wherein the harmful solute has a lower total mass transport through first element than through second element such that if it enters the sensor fluid it will be removed into the reservoir fluid.

15. The device of claim 1, wherein the target analyte in the sample fluid is at a first concentration, and the target analyte in the sensor fluid is at a second concentration, wherein the first concentration and second concentration differ by an amount chosen from less than 5%, less than 10%, and less than 50%.

16. The device of claim 1, wherein the first element that separates the first chamber from the sample fluid area and restricts diffusion of solutes between the sensor fluid and sample fluid is chosen from a channel, a membrane, and a hydrogel.

17. The device of claim 16, wherein the first element is a membrane chosen from a liquid filled membranes, an oil-membrane, a filtration membrane, and a size-selective membranes

18. The device of claim 16, wherein the first element is a membrane, and the membrane at least partially retains at least one sensor enhancing solute in the sensor solution.

19. The device of claim 1, wherein the second element that separates the first chamber from the second chamber and restricts diffusion of solutes between the sensor fluid and the reservoir fluid is chosen from a channel, a membrane, and a hydrogel.

20. The device of claim 19, wherein the second element is a membrane, and the membrane includes polyethersulfone (PES).

21. The device of claim 19, wherein the second element is a hydrogel, and the hydrogel includes agar.

22. The device of claim 1, wherein the volume of the reservoir fluid in the second chamber is greater than the volume of sensor fluid in the first chamber.

23. The device of claim 22, wherein the difference in volume of the reservoir fluid compared to the volume of the sensor fluid is chosen from at least 2× greater, at least 10X greater, at least 50X greater, and at least 250× greater.

24. The device of claim 22, wherein the sensor fluid contains a plurality of aptamers.

25. The device of claim 24, wherein a first mass flow of the plurality of aptamers that occurs at the first element is less than a second mass flow of plurality of aptamers that occurs at the second element.

26. The device of claim 25, wherein the degree by which the first mass flow of the plurality of aptamers is less than the second mass flow of the plurality of aptamers is selected from the group consisting of at least 2× less, at least 10× less, at least 50× less, and at least 250× less.

27. The device of claim 25, wherein a first mass flow of the analyte that occurs at the first element is greater than a second mass flow of the analyte that occurs at the second element.

28. The device of claim 27, wherein the degree by which the first mass flow of analyte is greater than the second mass flow of analyte is selected from the group consisting of at least 2× greater, at least 10× greater, at least 50× greater, and at least 250× greater.

29. The device of claim 27, wherein a concentration of analyte in the sensor fluid will be within at least a percentage of a concentration of analyte in the sample fluid, wherein the percentage is chosen from 50%, 10%, 2%, and 0.4%.