Patent Application
20180353748
Non-invasive biosensing technologies have enormous potential for applications ranging from athletics, to neonatology, to pharmacological monitoring, to personal digital health, to name a few applications. The sweat ducts can provide a route of access to many of the same biomarkers, chemicals, or solutes that are carried in blood and can provide significant information enabling one to diagnose ailments, health status, toxins, performance, and other physiological attributes even in advance of any physical sign. Sweat has many of the same analytes and analyte concentrations found in blood and interstitial fluid. Interstitial fluid has even more analytes nearer to blood concentrations than sweat does, especially for larger sized and more hydrophilic analytes (such as proteins).
If biofluid access through the skin has such significant potential as a sensing paradigm, then why has it not emerged beyond decades-old usage in infant chloride sweat assays for Cystic Fibrosis or in illicit drug monitoring patches? Or, why for example did past reverse iontophoresis products for interstitial fluid extraction dominantly through eccrine ducts, such as GlucoWatch, also fail commercially? In part, past challenges and failures have been due to the difficulty of finding ergonomic and acceptable ways to generate the biofluid for sampling (non-invasive, continuous, non-irritating, etc.). In part, past challenges and failures have also been due to the difficulty of obtaining an adequate volume of sample for a measurement of analytes in these types of biofluids. Reducing the sample volume is critical for fast sampling times and/or to allow lower sample generation rates (e.g., less reverse iontophoresis current and related stress on skin). However, simply reducing the sample volume, especially for the case of reverse-iontophoresis, brings about secondary challenges such as pH changes.
A more detailed background description is now provided, beginning here with a discussion of fluid sampling rate. Assume a case where sweat glands dominantly provide the pre-existing pathways. Next, using information from Cunningham in Chapter 7 of the book In Vivo Glucose Sensing, 2010, assume a device having a sampling area of 1 cm2 applied on a person's wrist. Assuming use of a sweat gland density of 150/cm2 for the wrist, a sensor that is 0.55 cm in radius (1.1 cm in diameter) would cover about 1 cm2 area or approximately 150 sweat glands. Now, consider an example sample generation rate by reverse iontophoresis based on 15-150 nL of interstitial fluid being generated with 3 minutes of reverse iontophoresis at 0.3 mA/cm2. Therefore, roughly 5 to 50 nL is generated per minute. Assuming a 1 cm2 area for reverse iontophoresis, then the sample generation rate for the wrist would therefore be roughly 0.03 to 0.3 nL/min/gland. If the fluidic portion of the 1 cm2 device is 127 μm thick (i.e., same as the gel used with the GlucoWatch), then fluidic volume is 12,700 nL. If that volume were to be completely filled with new interstitial fluid, it would require 2822 to 282 minutes (47 to 4.7 hours), which represents a very slow sampling interval. Using another estimate by Pikal and Shah “Transport mechanisms in iontophoresis. I. A theoretical model for the effect of electroosmotic flow on flux enhancement in transdermal iontophoresis” 1990, the sample generation rate of interstitial fluid is 6-19 μL/hr/mA, or about 12.5 μL/hr/mA*1 hr/60 min=0.21 μL/mA/min. For the above case of 0.3 mA/cm2, the sample generation rate would be 63 nL/min/cm2 which for 150 glands/cm2 would be 0.42 nL/min/gland (higher than Cunningham's example, but still requiring greater than 2 hours for the sampling interval). Discrepancies between these numbers could be due to interpretations of volumes based on diluted analyte concentration, or non-ideal factors.
Next, consider the impact of the choice of sensing modality. The actual sampling interval and the time required to sense an analyte varies widely based on the method of sensing the analyte. For example, some sensors consume the sensed analyte (e.g., glucose and enzymatic/amperometric sensing) while others do not consume the analyte and respond by equilibrating to the local concentration of the analyte (e.g., ionselective or electrochemical aptamer-based sensors). An aptamer-based sensor may bind an analyte, but the analyte is not consumed (i.e., once the analyte binds, the same site will not bind further analyte, and the analyte can be released back into solution). Because sensors that consume the analyte do not require a complete refreshing of the sample volume (e.g., new sample replaces or washes away the old sample), sensors that consume the analyte will not apply to how sampling rate and sampling interval are described and calculated herein. Conversely, sensors that do not consume the analyte, therefore, will only respond as quickly as the sample volume is refreshed across the sensor.
Consider an example involving an electrochemical aptamer sensor for vasopressin. Assume the sensor is configured with a linear range of detection centered around vasopressin's normal concentration range in interstitial fluid, where the fluid is extracted by reverse iontophoresis. Unlike an amperometric sensor that consumes the analyte, the aptamer sensor does not consume the vasopressin, nor does the aptamer sensor aggregate detection of vasopressin over time. Therefore, the vasopressin must remain within the detection range if the sensor is to continue to detect vasopressin. The sampling interval for vasopressin, accordingly, would be in the multiple-hour range using a device with slow interstitial fluid refresh rate, such as those described in the above example. For detecting and protecting against dehydration, and other time-sensitive applications, such sampling intervals could be entirely too slow. Cortisol awakening response, for example, occurs within a 30 minute window and requires multiple readings during that window. The multi-hour sampling intervals mentioned above would be entirely too slow for such an application.
Consider instead an interstitial fluid wicking component that is composed of 5 μm deep channels that comprise 5% of the wicking component surface area, resulting in an interstitial fluid volume of at least 5E-4 cm*0.05*1 cm2=2.5E-5 mL or 25 nL. That is a roughly 500× lower sample volume. Such a substantially reduced sample volume can provide one or more significant advances in performance, such as: (1) greatly reduced sampling intervals (e.g., as fast as minutes even for sensors that do not consume the analyte); and (2) greatly reduced current density requirements for reverse iontophoresis. However, reducing the sample volume creates at least one secondary challenge, namely pH changes caused by water electrolysis.
Consider an illustrative example that assumes a 0.3 nL/min/gland interstitial fluid sample generation rate for 0.3 mA/cm2 applied to skin. Assume only needing to fill the dermal duct of the eccrine sweat gland which has a diameter of about 15 μm and a length of about 2000 μm. This ductal volume is therefore 2000*3.14*7.52 μm3=0.35 nL. At 0.3 nL/min/gland it would require approximately 1 minute to get a fresh interstitial fluid sample to the skin surface (and 30 seconds for 0.6 nL/min/gland). This is likely at least one reason why GlucoWatch applies reverse iontophoresis for a period of 3 minutes followed by 7 minutes to allow glucose to diffuse into the gel and be sensed (such that it is not pulled back into skin during the subsequent application of voltage in the opposite polarity).
Further, assume the device covers skin having 100 glands/cm2, where the device area is 1 cm2 and a volume to be filled of 1 μL (the space between the device and skin to be filled is 10 μm thick). It would require 30 minutes to fill this volume at only 0.3 nL/min/gland, and 15 minutes at 0.6 nL/min/gland. If the system is starting at a pH of 7 for the 10 μm thick fluid volume, with reverse iontophoresis durations of 1 min, 10 min, and 30 min, the pH under the negative voltage electrode would drop to extreme levels, i.e., 0.7, 0.3, and −0.7, respectively (first order calculations only). Such pH changes can be impractical from both a skin safety perspective and from a biosensing perspective (e.g., they could degrade numerous types of analytes). Some interesting conclusions can be made. First, using reverse iontophoresis for sample generation of only interstitial fluid, with such small sample volumes, can be impractical, unless significant steps are taken to buffer the pH changes. Therefore in some cases, dilution of the interstitial fluid in sweat could be advantageous as it would also dilute the pH change. Second, for an example like this where the interstitial fluid is transported away and replenished into the sample volume, increasing the sample volume does not reduce the pH change (because doubling the sample volume dilutes the pH change, but also requires 2× more current to fill that sample volume, resulting in the same pH change as before). Third, where the interstitial fluid is transported away and replenished into the sample volume, the pH is theoretically constant for a given sample volume (is irrespective of current density), because both sample generation rate and current density scale linearly.
Next, an erroneous argument could be made that using a scheme where one continually reverses the voltage (like that used by GlucoWatch) could solve all issues with pH. Basically, each time voltage was applied the pH would advantageously start at the opposite end of the pH spectrum (e.g., would be modulated back and forth so the net pH would instead oscillate closer to a neutral pH of 7). However, that assumes the interstitial fluid sample stays in place, which as previously described is accurate for a technology such as GlucoWatch (glucose sensing, thick gel full of fluid), but not for a technology with a greatly reduced sample volume and a net transport of fluid away from the skin to at least one sensor. For example, if it requires 30 minutes to fill the volume and that volume is continually being emptied and taken to a sensor as quickly as it is generated, then the fluid volume is actually even lower (is being depleted) which makes the pH changes even greater. Also, if the fluid were not transported away, then a portion of the fluid could be pulled back into the body by reverse iontophoresis as well. Again, this is not a problem for technology like GlucoWatch (glucose sensing) but is a challenge for other types of devices.
It is also possible, at a cost of reduced reverse iontophoresis current density, to mitigate pH issues by using voltage at the electrodes (i.e., between the electrodes and solution) that is below the electrolysis potential for water: H2O→½ O2(g)+2 H++2 e− for +1.23 V at the anode, and 2 H2O+2 e−→H2(g)+2 OH− for −0.83 V at the cathode. Diamond electrodes can extend this voltage threshold to as much as about 2V. A situation with a lower resistance across the skin (e.g., active sweating stimulated by carbachol iontophoresis) would therefore have reduced total voltage required for reverse iontophoresis. The voltage drop for a system on skin would be in part at the electrodes, in part across the skin, and in part across the tissue/body beneath the skin. For example, consider on the sweating forearm 0.2 mA applied with 5-10 V with a smallest electrode area of 0.95 cm2 for a current density of about 0.2 mA/cm2. Also, consider sweating under a Vitrode-J electrodes of 40 mm diameter placed 3 cm apart also on the forearm which registers a conductance of 100 μS, which for 10 V translates to 1 mA per 12 cm2 or about 0.1 mA/cm2. To be absolutely certain that the voltage across both electrodes was less than 2V (near the point of no electrolysis), based on first-order calculations, the current densities would need to be reduced from about 0.1 mA/cm2 to about 0.05 mA/cm2. Alternately, to be even more accurate and/or safer, the voltage drop at the actual electrodes could be measured by having a second high impedance electrode near the iontophoresis electrode(s). As a result, the total applied voltage could be increased until the point where the electrodes measure voltages associated with generation of electrolysis. At the point of electrolysis, the voltage increase could be halted, or even more desirably, could be slightly decreased to reduce electrolysis. Alternately, the pH at the actual electrodes could be measured with a pH sensitive electrode, and the total applied voltage could be increased until the electrodes begin significantly changing the local pH by electrolysis (at which point the voltage increase could be halted or the voltage decreased). In any or all of these cases, the current densities listed above are lower than the about 0.3 mA/cm2 used by GlucoWatch, which leads us next to further background discussion on what current densities may be required and/or most desirable.
The current densities required for interstitial fluid extraction by reverse iontophoresis can also be compared to other ‘natural’ forms of iontophoresis in the body. A comparison and calculation is made here, with respect to the amount of natural iontophoresis that exists during sweat generation. These calculations are first-order and provide further background information only. Assume that at 1 nL/min/gland the eccrine sweat gland creates a secreted Na+ concentration of around 30 mM (the concentration in secretory coil is likely larger, because some amount of Na+ is reabsorbed by the duct, but such differences will be ignored for present purposes). Next, obtain the amount of Na+ in 1 nL: 1E-9 L*30E-3 Mol/L*6.02E23 Na+/Mol=1.8E13 Na+. Therefore, there is a Na+ generation rate of 1.8E13 Na+/min/gland. The flux of charged Na+ creates an equivalent electrical current (A, or C/s) of 1.8E13 Na+*1.6E-19 C/Na+=3 μC, which is 3 μC/min/gland. Turning that into C/s (for A), you obtain 0.05 μC/s/gland or 50 nA/gland. This Na+ current enters the secretory coil because there is a net negative charge induced (negative voltage) in the secretory coil caused by the injection of Cl− ions which are actively secreted by the cells lining the secretory coil. This Na+ current originates from interstitial fluid and enters the secretory coil through the tight junctions between the 1-2 layers of cells that line the secretory coil. This therefore represents a natural form of reverse-iontophoresis and therefore potentially a natural amount of electro-osmosis created in the secretory coil.
GlucoWatch generated 0.03 to 0.3 nL/min/gland of interstitial fluid using 300 μA/cm2. For sweat, if we assume 100 active glands/cm2, then the 50 nA/gland is equivalent to 5 μA/cm2. Comparatively, GlucoWatch generates 0.03-0.3 nL/min/gland with 300 μA/cm2, while sweat glands naturally generate 1 nL/min/gland with 5 μA/cm2. Therefore, sweat has roughly 3-30× higher fluid flow rate, while using 60× less electrical current than GlucoWatch. This means that, at 1 nL/min/gland, if interstitial fluid were brought into the sweat by this natural form of reverse iontophoresis, the interstitial fluid component would be roughly 200-2000× less in volume than the sweat component. Because blood proteins are estimated to be 1000× or more dilute in sweat, this small interstitial fluid component suggests that even without electroporation, reverse iontophoresis could significantly increase the concentration of certain larger analytes in sweat.
Continuing the discussion from an applied/practical perspective, assume 0.1 nL/min/gland of sweat generation and the device applies only 5 μA/cm2 of reverse iontophoresis current to the skin, then by first order calculation, and assuming the reverse iontophoresis (electro-osmosis) causes protein molecules enter sweat through the tight junctions or other pathways leading to the secretory coil, the device could receive 10× more protein than is found in normal sweat. This would be 60× less current than GlucoWatch. Or assume 0.1 nL/min/gland sweat generation rate, and apply only 50 μA/cm2. The device could theoretically obtain 100× more protein with 6× less current than GlucoWatch. These examples suggest that reverse iontophoresis can enhance analyte concentrations at much lower current densities than those needed by GlucoWatch.
Returning to the previous discussion of current densities without electrolysis of water, about 0.1 mA/cm2 to about 0.05 mA/cm2 are well above the values of current densities shown above for increasing analyte concentrations in sweat. However, if the sweat generation rate were 1 nL/min/gland (not 0.1 nL/min/gland as just described above), then higher current densities would be needed to compensate for additional dilution of analytes in sweat. This reveals yet another need to reduce the sample volume for the sake of allowing sample collection at the lowest possible sample sweat rates, and as stated before, if there is no sweat, allowing interstitial fluid collection at the lowest possible current densities.
One more interesting comparison can be made. If only about 0.1 to 0.05 mA/cm2 or less can be used if electrolysis is to be avoided, and if for GlucoWatch 0.3 mA/cm2 was needed to generate 0.03 to 0.3 nL/min/gland of interstitial fluid, then trying to collect interstitial fluid only without electrolysis is certainly challenging if fast sampling intervals are needed. Clearly reduced volume between sensors and the skin surface is needed. Additionally, a combination of sweat and interstitial fluid can in some cases, allow a shorter sampling rate (faster transport of analytes) while requiring a lower current density.
Also in the prior art such as GlucoWatch, there is not a net flow of analyte to, and across, a sensor. Therefore, the chronological assurance or sampling interval is determined entirely by each time a sample is extracted (the frequency of sample extractions, is the chronological assurance and therefore determines the sampling interval). However, if a device were to have a net flow of analyte to, and across, a sensor, then the chronological assurance is not so simple as it is dependent on the sample volume required in the device and the sample generation and flow rates.
Lastly, in this background discussion of both challenges and opportunities presented by the disclosed invention, two more issues should be raised. First, electrolysis is not the only challenge caused by applying current through the skin. Skin damage, pain, discomfort, or annoyance in some cases could still be experienced by the user even at low current densities, and therefore reducing current densities nearly always improves user acceptance of such a device. Second, if a device transports sampled fluid away (actively, or passively), then electrical contact must still be adequately maintained with the skin, despite the reduced or absent fluid volume. Using a thick gel mitigates this issue completely, but results in a very large sample volume. Therefore, gels will need to be implemented with reduced volumes, electrodes must be kept in close contact with the skin, or the fluid volume between the device and skin must be minimized.
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 intimate proximity with biofluid and analytes as they flow out from the skin surface. With such a new invention, non-invasive and wearable biosensing could become a compelling new paradigm as a biosensing platform.
Embodiments of the disclosed invention provide biofluid sensing devices capable of reduced volume between the sensors and pre-existing pathways such as sweat glands, which decreases the sampling interval and/or reduces the required flow rate of the biofluid that is being generated. Some embodiments of the disclosed invention also mitigate challenges such as pH changes which can occur at an iontophoresis electrode.
In one embodiment, a sensor device for sensing on the skin includes one or more analyte-specific sensors and a volume-reducing component that provides a volume-reduced pathway for biofluid between the one or more sensors and pre-existing pathways in said skin when said device is positioned on said skin. In one embodiment, the biofluid may be more than 50% interstitial fluid. In another embodiment, the biofluid may be more than 50% sweat.
In other embodiments, various methods for integration of volume reducing components, sensors, chemical delivery components, and reverse iontophoresis components are provided. In yet another embodiment, various components and techniques are provided for buffering acid or base generation at an electrode for reverse iontophoresis.
The objects and advantages of the disclosed invention will be further appreciated in light of the following detailed descriptions and drawings in which:
As used herein, “interstitial fluid” or “tissue fluid” is a solution that bathes and surrounds tissue cells. The interstitial fluid is found in the interstices—the spaces between cells (also known as the tissue spaces). Embodiments of the disclosed invention focus on interstitial fluid found in the skin and, particularly, interstitial fluid found in the dermis. In some cases where interstitial fluid is emerging from sweat ducts, the interstitial fluid contains some sweat as well, or alternately, sweat may contain some interstitial fluid. As used herein, “mainly interstitial fluid” means fluid that contains by volume less than 50% sweat (i.e., is primarily interstitial fluid). As used herein, “mainly sweat” means fluid that contains by volume 50% or greater of sweat (i.e., may contain some interstitial fluid, but has equal or greater amount of sweat than interstitial fluid). The percentages of each fluid can be quantified by several methods, such as measuring analyte dilutions in sweat (e.g., some analytes are dilute in sweat but not in interstitial fluid), or such as by measuring and comparing sample generation rates their respective contributions to the total fluid volume quantified (e.g., compare sample generation rates with or without application of reverse iontophoresis; or compare sample generation rates with or without natural or chemically-induced sweat stimulation).
As used herein, “biofluid” is a fluid that is comprised mainly of interstitial fluid or sweat as it emerges from the skin. For example, a fluid that is 45% interstitial fluid, 45% sweat, and 10% blood is a biofluid as used herein. For example, a fluid that is 20% interstitial fluid, 20% sweat, and 60% blood is not a biofluid as used herein. For example, a fluid that is 100% sweat or 100% interstitial fluid is a biofluid. A biofluid may be diluted with water or other solvents inside a device because the term biofluid refers to the state of the fluid as it emerges from the skin. Generally, as compared to blood, sweat is highly dilute of large sized analytes (e.g., greater than 1000× for proteins, etc.) and to a lesser extent, as compared to blood, interstitial fluid is dilute for some larger sized analytes (e.g., 10-100× or more or less depending on the specific analyte, current density, etc.).
As used herein, “pre-existing pathways” refer to pores, pathways, or routes through skin through which interstitial fluid may be extracted. Pre-existing pathways include but are not limited to: eccrine sweat ducts, other types of sweat ducts, hair follicles, inter-cell junctions, tape-stripping of the stratum corneum, skin defects, pathways created by electroporation of skin (e.g., of the stratum corneum), laser poration of skin, mechanical poration of skin (e.g., micro-needle rollers), chemical or solvent based poration of skin, or other methods or techniques. It should be recognized that “pre-existing” does not require that such pathways must be naturally occurring or that such pathways must exist prior to application of the device. Rather, methods of the disclosed invention may be practiced using a pathway that naturally exists or that was created for the particular application. Therefore, any technique to provide pre-existing pathways may be used in conjunction with embodiments of the disclosed invention. For example, a microneedle is a pre-existing pathway if the microneedle uses reverse iontophoresis for analyte extraction. However, generally, non-invasive access is preferred, and naturally occurring pre-existing pathways may be preferred for many applications. As another example, electroporation of the lining of the sweat glands may form or affect a pre-existing pathway. As another example, skin permeability enhancing agents or chemicals may form part or all of a pre-existing pathway. For simplicity of description herein, eccrine sweat glands will be the only pre-existing pathways explicitly discussed, but as noted above, embodiments of the disclosed invention may apply to any pre-existing pathway as defined above.
As used herein, “chronological assurance” means the sampling rate or sampling interval that assures measurement(s) of analytes in a biofluid in terms of the rate at which measurements can be made of new biofluid analytes emerging from the body. Chronological assurance may also include a determination of the effect of sensor function, potential contamination with previously generated analytes, other fluids, or other measurement contamination sources for the measurement(s). Chronological assurance may have an offset for time delays in the body (e.g., a well-known 5-30 minute lag time between analytes in blood emerging in interstitial fluid), but the resulting sampling interval (defined below) is independent of lag time, and furthermore, this lag time is inside the body, and therefore, for chronological assurance as defined above and interpreted herein, this lag time does not apply.
As used herein, “interstitial fluid sampling rate” or “sweat sampling rate” or simply “sampling rate” is the effective rate at which new biofluid sample, originating from the pre-existing pathways, reaches a sensor that measures a property of the fluid or its solutes. Sampling rate is the rate at which new biofluid is refreshed at the one or more sensors and therefore old biofluid is removed as new fluid arrives. In an embodiment, this can be estimated based on volume, flow-rate, and time calculations, although it is recognized that some biofluid or solute mixing can occur. Sampling rate directly determines or is a contributing factor in determining the chronological assurance. Times and rates are inversely proportional (rates having at least partial units of 1/seconds), therefore a short or small time required to refill sample volume can also be said to have a fast or high sampling rate. The inverse of sampling rate (1/s) could also be interpreted as a “sampling interval” (s). Sampling rates or intervals are not necessarily regular, discrete, periodic, discontinuous, or subject to other limitations. Like chronological assurance, sampling rate may also include a determination of the effect of potential contamination with previously generated biofluid, previously generated solutes (analytes), other fluid, or other measurement contamination sources for the measurement(s). Sampling rate can also be in part determined from solute generation, transport, advective transport of fluid, diffusion transport of solutes, or other factors that will impact the rate at which new sample will reach a sensor and/or is altered by older sample or solutes or other contamination sources. During reverse iontophoretic extraction of fluid samples and analytes, some analytes that have a net charge could move faster or slower, with or against, the advective flow of fluid sample. In the event that the analytes are moving faster or slower than the advective flow, the sampling rate is still determined by the advective flow of interstitial fluid and the replenishment of new fluid sample across the sensor as the old sample is replaced. If an embodiment of the disclosed invention does not include a net flow of sample fluid across a sensor, and does include transport of a solute (analyte) to the sensor, then the term sampling rate may be replaced with the term “analyte sampling rate”. As will be described in greater detail below, sampling rate may be interpreted with respect to sensors that do not consume the analyte as part of the process of sensing the analyte, because these sensors are dependent on flow of fresh analyte to the sensors and removal of old analyte away from the sensors.
As used herein, “sweat stimulation” is the direct or indirect causing of sweat generation by any external stimulus. One example of sweat stimulation is the administration of a sweat stimulant such as pilocarpine or carbachol from a sweat stimulating component. Going for a jog, which stimulates sweat, is sweat stimulation, but would not be considered as sweat stimulating component. Sweat stimulation can include sudo-motor axon reflex sweating, passively diffused chemical into skin to stimulate sweat, or any other suitable method for sweat stimulation. As further examples, sweat stimulation can be achieved by simple thermal stimulation, by orally administering a drug, by intradermal injection of drugs such as methylcholine, carbachol, or pilocarpine, and by dermal introduction of such drugs using iontophoresis.
As used herein, “sample generation rate” is the rate at which biofluid is generated by flow through pre-existing pathways. Sample generation rate is typically measured by the flow rate from each pre-existing pathway in nL/min/pathway. In some cases, to obtain total sample flow rate, the sample generation rate is multiplied by the number of pathways from which the sample is being sampled. Similarly, as used herein, “analyte generation rate” is the rate at which solutes move from the body or other sources toward the sensors.
As used herein, “measured” can imply an exact or precise quantitative measurement and can include broader meanings such as, for example, measuring a relative amount of change of something. Measured can also imply a binary measurement, such as ‘yes’ or ‘no’ type qualitative measurements.
As used herein, “sample volume” is the fluidic volume in a space that can be defined multiple ways. Sample volume may be the volume that exists between a sensor and the point of generation of biofluid sample. Sample volume can include the volume that can be occupied by sample fluid between: the sampling site on the skin and a sensor on the skin where the sensor has no intervening layers, materials, or components between it and the skin; or the sampling site on the skin and a sensor on the skin where there are one or more layers, materials, or components between the sensor and the sampling site on the skin.
As used herein, “microfluidic components” are channels or other geometries formed in or by polymers, textiles, paper, or other components known in the art to transport fluid in a deterministic manner.
As used herein, “state void of sample” is where a space or material or surface that can be wetted, filled, or partially filled by a biofluid sample, but which is in a state where it is entirely or substantially (e.g., greater than 50%) dry or void of biofluid sample.
As used herein, “advective transport” is a transport mechanism of a substance or conserved property by a fluid due to the fluid's bulk motion.
As used herein, “diffusion” is the net movement of a substance from a region of high concentration to a region of low concentration. This is also referred to as the movement of a substance down a concentration gradient.
As used herein, a “volume-reduced pathway” or “reduced-volume pathway” is at least a portion of a sample volume that has been reduced by addition of a material, device, layer, or other component, which therefore increases the sampling interval for a given sample generation rate. A volume-reduced pathway can be created by at least one volume reducing component.
As used herein, “volume reducing component” means any component or material that reduces the sample volume and increases the sampling rate and/or the analyte sampling rate. In some cases, the volume reducing component is more than just a volume reducing material, because a volume reducing material by itself may not allow proper device function (e.g., the volume reducing material would need to be isolated from a sensor for which the volume reducing material could damage or degrade, and therefore the volume reducing component may comprise the volume reducing material and at least one additional material or layer to isolate volume reducing material from said sensors).
As used herein, “flux” is the rate of transfer of fluid and/or particles and/or solutes across a given surface. With respect to the eccrine sweat gland, flux can refer to both a fluid (e.g., interstitial fluid, intracellular fluid, etc.) and its contents, or refer to only one or more analytes entering into the sweat gland (e.g., ions, molecules, proteins, etc.). A flux in the sweat gland can occur at all areas, or in subsets of areas (e.g., a part of the dermal duct, or the secretory oil, etc.). A flux can also be referred to as “flux of analyte” or “analyte flux” or other similar uses that refer to a flux of analytes in interstitial fluid, moving along with or against the flow of one or more of these fluids, or moving fully or somewhat independently of flow of these fluids. For example, charges of analytes can be negative or positive, and fluxes can be in the opposite direction of advective flow.
As used herein, “reverse iontophoresis” is a subset or more specific form of “iontophoresis” and is a technique by which electrical current and electrical field cause molecules to be removed from within the body by electroosmosis and/or iontophoresis. Although the description below focuses primarily on electro-osmosis, the term “reverse iontophoresis” as used herein may also apply to flux of analytes brought to or into the devices of the disclosed invention, where the flux is in whole or at least in part due to iontophoresis (e.g., some negatively charged analytes may be transported against the direction of electroosmotic flow and eventually onto a device according to an embodiment of the disclosed invention). Electroosmotic flow (or electro-osmotic flow, synonymous with electroosmosis or electroendosmosis) is the motion of liquid induced by an applied potential across a porous material, capillary tube, membrane, microchannel, or any other fluid conduit. Because electroosmotic velocities are independent of conduit size, as long as the electrical double layer is much smaller than the characteristic length scale of the channel, electroosmotic flow is most significant when in small channels. In biological tissues, the negative surface charge of plasma membranes causes accumulation of positively charged ions such as sodium. Accordingly, fluid flow due to reverse iontophoresis in the skin is typically in the direction of where a negative voltage is applied (i.e., the advective flow of fluid is in the direction of the applied electric field). As used herein, the term “iontophoresis” may be substituted for “reverse iontophoresis” in any embodiment where there is a net advective transport of biofluid to the surface of the skin. For example, if a flow of sweat exists, then negatively charged analytes may be brought into the advectively flowing sweat by iontophoresis. The net advective flow of sweat would typically be needed, because in this case, a net electro-osmotic fluid flow would be in the direction of sweat into interstitial fluid (and without a net advective flow of sweat, the sweat would be lost, and there would be no pathway for transporting the analyte to at least one sensor). Furthermore, because “reverse iontophoresis” is a subset or more specific form of “iontophoresis”, the term “iontophoresis” may refer to both “reverse iontophoresis” and “iontophoresis”. The terms “reverse iontophoresis” and “iontophoresis” are interchangeable in the disclosed invention.
As used herein, the term “analyte-specific sensor” is a sensor specific to an analyte and performs specific chemical recognition of the analytes presence or concentration (e.g., ion-selective electrodes, enzymatic sensors, electrically based aptamer sensors, etc.). For example, sensors that sense impedance or conductance of a fluid, such as biofluid, are excluded from the definition of “analyte-specific sensor” because sensing impedance or conductance merges measurements of all ions in biofluid (i.e., the sensor is not chemically selective; it provides an indirect measurement). Sensors could also be optical, mechanical, or use other physical/chemical methods which are specific to a single analyte. Further, multiple sensors can each be specific to one of multiple analytes.
As used herein, the term “sensor that consumes the analyte” is an analyte-specific sensor that decreases the total amount of analyte present (e.g., glucose and other enzymatic/amperometric sensing).
As used herein, the term “sensor that does not consume the analyte” is an analyte-specific sensor that responds by equilibrating to the local concentration of the analyte (e.g., ionselective or electrochemical aptamer-based sensors) and that does not decrease the total amount of the analyte present. An aptamer-based sensor may bind an analyte, but the analyte is not consumed (i.e., once the analyte binds, the same site will not bind further analyte, and furthermore, the analyte can be released back into solution as well). The definition and calculations for sampling rate and sampling interval described herein apply to cases where the sensors do not consume the analyte.
As used herein, the terms “wicking pressure,” “wicking force,” “capillary pressure,” or “capillary force” means a pressure or force that should be interpreted according to its general scientific meaning. For example, capillary (tube) geometry can be said to have a capillary pressure or a wicking pressure. For example, a wicking textile or gel may have a capillary pressure, even if the material is not geometrically a tube or a channel. Similarly, the (relatively empty) space between a material placed on skin and the skin surface can have an effective wicking pressure. The terms wicking or capillary pressure and wicking or capillary force may be used interchangeably herein to describe the effective pressure provided by any component or material that is capable of capturing biofluid by a negative pressure (i.e., pulling it into or along said component or material). For simplicity, the term “wicking pressure” is used herein to refer to any of the above alternate terms. Wicking pressure also must be considered in its specific context, for example, if a sponge is fully saturated with water, then it has no remaining wicking pressure. Therefore, wicking pressure as used herein describes a device and/or a component during use, and not interpreted in isolation or in contexts other than the disclosed devices or use scenarios.
As used herein, the term “wicking collector” means a component of the disclosed invention that supports the creation of, or sustains, a volume reduced pathway by use of wicking pressure, and/or that is the wicking element adjacent to or on skin that receives biofluid before it reaches a sensor. A wicking collector can be a microfluidic component, a capillary material, a wrinkled surface, a textile, a gel, a coating, a film, or any other suitable component. A single component may serve multiple functions as a wicking collector and, for example, a wicking pump (defined below).
As used herein, the term “wicking pump” refers to a component that supports creation of or sustains a volume reduced pathway by use of wicking pressure, or that receives biofluid after a sensor and has a primary purpose of collecting excess biofluid to allow sustained operation of the device. A wicking pump may also include an evaporative material or surface that is configured to remove excess biofluid by evaporation of water. A wicking pump may be part of the same component or material that serves other purposes (e.g., a wicking collector or a wicking coupler), and in such cases, the portion of said component or material that at least in part receives biofluid after the sensor(s), is also a wicking pump as defined herein.
The term “wicking pump” may also reference alternate configurations, such as a small mechanical pump, or osmotic pressure across a membrane (i.e., the wicking pump would be the membrane and the draw solution or material), so long as the pressure generated satisfies the requirements described herein, and the other materials or components between the wicking pump and skin operate by wicking pressure to maintain their respective sample volumes.
As used herein, the term “wicking coupler” refers to a component that is on or adjacent to a biofluid sensor and that promotes the transport of biofluid or its solutes (e.g., by advective flow, diffusion, or other method of transport) between another wicking component or material and a sensor. In some embodiments, a single component may function as both a wicking coupler and a wicking collector. In other embodiments, a sensor may be configured with a wicking surface or material that functions without a wicking coupler (e.g., an immobilized aptamer layer which is hydrophilic, or polymer ionophore layer which is porous to the analyte). A wicking coupler may be part of the same component or material that serves other purposes (e.g., a wicking collector or a wicking pump), and in such cases, the portion of said component or material that, at least in part, couples biofluid to a sensor(s) and that is on or adjacent to the sensor(s), is also a wicking coupler as defined herein.
As used herein, the term “wicking space” refers to the space between the skin and wicking collector that would be filled by air, skin oil, or other non-biofluid fluids or gases if no biofluid existed. In some embodiments of the disclosed invention, even if biofluid exists, the wicking collector removes some or most of biofluid from the wicking space by action of wicking pressure provided by the wicking collector.
As used herein, “biofluid collector pressed against skin” is a component that at least in part is pressed directly against the skin, and which is at least a part of a volume-reducing component. Further, a biofluid collector includes a plurality of pores or pathways in a material and/or on the surface of a material that is held against skin so that the plasticity of skin allows skin defects, hair, and other sample volume increasing aspects of skin to at least partially conform against the material.
As used herein, “space between skin and a biofluid collector pressed against skin” refers to the space between the skin and a biofluid collector pressed against skin that would be filled by air, skin oil, or other non-sweat fluids or gases if no sweat existed.
As used herein, “pressure element” is any component that at least in part provides pressure to a biofluid collector pressed against skin to create at least in part a reduced sample volume in the space between skin and a biofluid collector pressed against skin.
Embodiments of the disclosed invention apply at least to any type of sensor device that measures at least one analyte in interstitial fluid extracted at least in part by reverse iontophoresis through pre-existing pathways. Further, embodiments of the disclosed invention apply to sensing devices which measure chronological assurance. Further, embodiments of the disclosed invention apply to sensing devices which can take on forms including patches, bands, straps, portions of clothing, wearables, or any suitable mechanism that reliably brings sampling and sensing technology into intimate proximity with biofluid sample as it is transported to the skin surface. While some embodiments of the disclosed invention utilize adhesives to hold the device near the skin, devices could also be held by other mechanisms that hold the device secure against the skin, such as a strap or embedding in a helmet. 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 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. 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 obvious (such as a battery), and for purposes of brevity and of greater focus on inventive aspects, such components are not explicitly shown in the diagrams or described in the embodiments of the disclosed invention.
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The wicking space can change over time due to skin plasticity, for example, skin can swell and become smoother as it hydrates, and skin can flatten if a device applies pressure against the skin surface. Therefore, in an embodiment of the disclosed invention, if the sample volume is to be reduced between the skin 12 and the area of the wicking collector 136 on or adjacent to skin, then at the time of first application of the device 100 to skin, the sample volume of the portion of the wicking collector 136 that interfaces with or is adjacent to skin is less than the sample volume of the wicking space between the wicking collector 136 and skin 12.
In an embodiment, a wicking collector could be constructed of Rayon or other material that has two or more levels of wicking pressures. For example, Rayon has a first and greater wicking pressure when fluid is wicked along grooves in its fibers, and a second and lower wicking pressure when fluid also fills the spaces in between such fibers. Alternately, open-faced rectangular micro-channels could have a higher wicking pressure when they have less biofluid in the channels (i.e., when only wicking along the corners of the channels which have the highest wicking pressure instead of filling the channels). Therefore, an embodiment of the disclosed invention may include a wicking material where the sample volume in said wicking material during use is less than 50% of the total available volume of such said wicking material.
In use, the device 100 may be placed on a person's skin to sense a biofluid. The following exemplary use of the device 100 is described relative to interstitial fluid, although the description applies equally to any biofluid as defined above. The skin adhesive 112 secures the device 100 to the skin 12. The reverse iontophoresis electrode 150 and the counter electrode 152 are used to generate a flow of interstitial fluid. The wicking collector 136 transports interstitial fluid from the skin 12 towards the wicking pump 138. As interstitial fluid moves through the wicking collector 136, the wicking couplers 130, 132, 134 allow the sensors 120, 122, 124, respectively, to sense the interstitial fluid. In an exemplary embodiment, the sensor 120 may comprise an ion-selective electrode for sodium and a reference electrode, the sensor 122 is an amperometric sensor for urea, and the sensor 124 is an electrochemical aptamer sensor for vasopressin.
In an aspect of the disclosed invention, a net advective flow of biofluid from the skin to the sensor(s) in the device is required for the sensor(s) to sense the desired analytes in the biofluid. As previously noted, the terms “iontophoresis” may be substituted for “reverse iontophoresis” in any embodiment for cases where sweat is the primary driver of a net advective transport of biofluid to the surface of the skin. If a flow of sweat exists, then negatively charged analytes, such as acidic analytes or certain proteins or peptides, may be brought into the advectively flowing sweat by iontophoresis. The net advective flow of sweat is a requirement if “iontophoresis” is to be substituted for “reverse iontophoresis” because, in this case, a net electro-osmotic fluid flow would be in the direction of sweat into interstitial fluid (and without a net advective flow of sweat, there would be no pathway for transporting the analyte to at least one sensor as illustrated in embodiments of the disclosed invention). Even if there were a fluid pathway for pure iontophoretic transport of an analyte (i.e., no advective flow) to a sensor, iontophoretic currents are typically dominated by small ions such as Cl−, and few of the other possible negatively charged analytes could be brought to a sensor in meaningful quantities.
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In an aspect of the disclosed invention, the electrode sizes may be designed to mitigate issues with pain or discomfort caused by electrical current passing into the skin 12. Pain or discomfort caused by electrical current in skin does not scale linearly in terms of the relationship of current density to electrode area as taught in P. W. Ledger, Skin biological issues in electrically enhanced transdermal delivery (1992). The smaller the electrode area is, generally the larger the current density that can be used without a perception of the current or perception of pain. For example, an electrode of 24 cm2 area generates a tingle at 0.08 mA/cm2, whereas an electrode of 0.64 cm2 generates a tingle at 0.4 mA/cm2 (varies based on location on skin and from person to person). In an aspect of the disclosed invention, due to the reduced sample volumes, the areas of electrical contact with skin for reverse iontophoresis are reduced. Consider for example, sampling biofluid from pre-existing pathways that are sweat ducts with densities of 100 glands/cm2, then the contact areas needed to cover an average of 5, 10, and 50 glands would be 0.05 cm2, 0.1 cm2, and 0.5 cm2, respectively. With sweat ducts with densities of 200 glands/cm2, then the contact areas needed to cover an average of 5, 10, and 50 glands would be 0.025 cm2, 0.05 cm2, and 0.25 cm2, respectively. Even fewer glands could be covered, so the above areas of contact may represent upper limits for contact areas for one or more embodiments of the disclosed invention. These areas can be of the electrodes themselves or, in the case of intervening materials or layers between the electrodes and skin, can represent the electrical contact area with skin.
In an aspect of the disclosed invention, sample volumes are dramatically reduced compared to the prior devices. However, reduced sample volumes can also cause issues with analyte depletion in the biofluid (e.g., the sensor captures analytes and thereby changes analyte concentration in biofluid, causing the sensor to erroneously measure analyte concentration). For example, consider a sensor with an area of about 0.001 cm2 (about 300 μm×300 μm) with 5E12 aptamer probes/cm2, which is 5E9 probes or about 8E-15 moles of probe. Now, assume for 14.1 nL of solution that flows past the sensors includes 100 nM of cortisol. That is 14.1E-9 L*100 nM/L=1.41E-15 moles of cortisol. There are about 6× fewer available analytes than available probes. Therefore, a sensor with an area of about 0.001 mm2 (about 30 μm×30 μm) might be preferred because it would contain about 8E-17 moles of probe, which is much less than the moles of analyte and therefore the analyte will not be depleted in the sample. For higher concentration analytes (e.g., 100 μM), preferred sensor areas might therefore be about 1 mm2. Furthermore, because embodiments of the disclosed invention work with such small sample volumes, smaller sensor areas are preferred because a larger sensor area would increase the sample volume required for the sensor. Exemplary sensor areas include less than 0.001 mm2, less than 0.01 mm2, less than 0.1 mm2, or less than 1 mm2.
In another aspect of the disclosed invention, the entry of solutes into the secretory coil or sweat duct can be enhanced using non-natural (applied) reverse iontophoresis. As previously described, Na+ enters the secretory coil from the interstitial fluid through the cell-cell junctions or “tight junctions” between cells. When a flux of Na+ is driven by an electric field; the moving Na+ (and other positive ions) drags additional interstitial fluid and possibly other analytes (solutes) into sweat by a process of natural electro-osmosis. An embodiment of the disclosed invention relies on entry primarily through the cell-cell junctions rather than through additional electrically formed pores. Relying entirely on iontophoresis for fluid access with large sample volumes creates large pores and damages the paths through tissue and cells. Whether a majority of the porous pathways are natural or are created may be determined through the measurement of electrical impedance with the skin. Electrical impedance of the skin will increase with decreasing sweat rate as more sodium and chloride is captured by the dermal duct. At a constant sweat rate, it will decrease only if new porous pathways are created through or between plasma membranes of cells by an excess of current or voltage. The same can be true for interstitial fluid extracted without electroporation, because the dermal duct can recapture sodium and chloride at low generation rates for interstitial fluid in the same way it recaptures sodium and chloride for sweat. Exemplary voltages which will not electroporate a single cell plasma membrane are on the order of, but not limited to, 0.15 to 0.3 V, with electroporation typically being rapidly induced at 0.5 to 1 V across a single plasma membrane. The lining of the sweat gland has several cells, with at least two plasma membranes in series for the case of a single cell, such that an exemplary safe upper limit for applied reverse iontophoresis voltage without causing electroporation is 300 to 600 mV or less. For example, the reverse iontophoresis voltage could be ramped slowly or tested at several levels, and the skin impedance could be measured continuously or repeatedly. These measurements may be used to determine a safe level of voltage to avoid new plasma membrane poration to the point where the dominant entry of flux of analytes is due to new pores as opposed to natural pathways that existed before reverse iontophoresis was applied. To that end, an embodiment of the disclosed invention includes an electrode or sensor for measuring skin impedance. For example, with reference to
When new pore formation begins to occur, there is likely a clear non-linear change in skin impedance as the natural pathways will tend to behave more like a classical resistor in response to voltage (albeit not perfectly linear as they are biological structures), and the new pores will create a superlinear response (e.g., they can get bigger and more numerous over time and the impedance will increase above the expected linear line). An applied potential of only 0.5 V to the stratum corneum shows little or no change in electrical conductance over time, and application of 0.75 V and 1 V showed fairly good stability of conductance up to an hour or more. Therefore, in various embodiments, a reverse iontophoresis voltage of less than 3 V and preferably less than 1V may be applied. The predicted current density for an applied voltage of 1 V and is about 0.01 mA/cm2, and, at 0.21 μL/mA/min, the predicted sample generation rate for the interstitial fluid is about 2.1 nL/min/cm2 or 0.02 nL/min/gland for 100 glands/cm2. However, this current density could still increase the analyte concentration coming in from interstitial fluid by 3× for sweat at 0.1 nL/min/gland sweat generation rate. If 3V were used, close to 10× higher concentration might be achieved.
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In another aspect of the disclosed invention, reverse iontophoresis may be applied without causing significant electroporation by allowing adequate time for the skin to recover after voltage is removed or reduced. In that regard, if electroporation occurs, then a non-linear response may exist between the measured skin electrical impedance and increasing applied voltage (primarily electrical resistance), and/or the relationship between voltage and skin electrical resistance may change versus time even at constant voltage. The skin and/or tissue subjected to electroporation tends to heal, and the electrical resistance should recover over time if the voltage is removed. In an embodiment, a device may apply reverse iontophoresis for a period of 10 minutes for a given applied voltage, which if applied continuously would cause significant electroporation, but the device may then allow 50 minutes resting without reverse iontophoresis such that little or no accumulation of electroporation occurs. In another embodiment a device includes a sensor to measure the electrical resistance of the skin, and the application of the reverse iontophoresis could be regulated based on the measured electrical resistance to ensure excessive electroporation of the skin does not occur. For example, if DC voltage were applied for the reverse iontophoresis, then the DC current could also be measured to directly predict the total electrical resistance. For example, the reverse iontophoresis may be regulated to ensure that the electrical resistance of skin does not drop by more than 3× compared to the electrical resistance without reverse iontophoresis. In an embodiment, a first electrical resistance for skin with no iontophoresis and a second electrical resistance for skin with iontophoresis, where said first electrical resistance is no more than 3× greater than said second electrical resistance. This 3× would be in the context of unchanging skin conditions (e.g., start measuring impedance once the skin is fully hydrated or at a constant chemically stimulated sweat rate). One skilled in the art will recognize that embodiments of the disclosed invention may account for variations with electrode distances, changes between users, changes during use for a single user, etc. The absolute voltage applied between electrodes is, at least in part, dependent on electrode distance and physiological factors.
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Exemplary sweat stimulants include acetylcholine, pilocarpine, methacholine, and carbachol, among others. The sweat stimulation mechanism may be used to initiate sweating to establish a reduced volume pathway and/or electrical connection between a reverse iontophoresis electrode and pre-existing pathways. In an embodiment, the sweat stimulant has a sweat stimulation duration of less than 60 minutes and, after sweat stimulation, reverse iontophoresis is applied to extract interstitial fluid. In that respect, acetylcholine is rapidly metabolized by the body, and sweating would stop occurring even within several minutes. This would allow the device 300 to be quickly primed with biofluid, and, by using the reverse iontophoresis electrode 350, then be able to sample interstitial fluid without dilution from sweat (after the sweating ceases). Some embodiments of the disclosed invention may apply reverse iontophoresis periodically rather than continuously (as described above), and a lack of iontophoresis for a period of time (e.g., minutes to hours) could cause the volume reduced pathway to disconnect or terminate fluidically or electrically. Therefore, the temporary stimulation of sweat, or instead stimulation of a constant low flow rate of sweat (e.g., less than 0.1 nL/min/gland) may be helpful as needed to maintain the volume reduced pathway and/or electrical connection for reverse iontophoresis. In other words, an embodiment of the disclosed invention includes the sampling of both stimulated sweat and interstitial fluid generated by reverse iontophoresis.
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In an aspect of the disclosed invention, sweating that occurs during extraction of interstitial fluid could result in unknown dilution of analyte in the interstitial fluid, with exception to analytes that have sweat concentrations similar to those found in interstitial fluid (e.g., unbound concentrations of cortisol). An embodiment may include a sensor (e.g., measuring skin impedance, sodium concentration, or a thermal flow) to measure sweat generation, sweat sampling interval, and/or sweat flow rate in the absence of or during a pause from reverse iontophoresis. This measurement could then be used to determine the amount of dilution of interstitial fluid that occurs during reverse iontophoresis. Similarly, one or more sensors may be used to measure generation rate for interstitial fluid during reverse iontophoresis. For example, the composition of the interstitial fluid may be analyzed to determine if the fluid includes more or less than 50% sweat (i.e., the ratio of sweat to interstitial fluid in the biofluid). Both generation rates and/or flow rates of sweat, interstitial fluid, or the biofluid in general could be measured by at least one sensor. Further, another embodiment of the disclosed invention may include at least one sensor for determining the ratio of sweat to interstitial fluid in the biofluid (e.g., by methods such as measuring analyte dilution caused by sweat). Another embodiment of the disclosed invention may include at least one sensor for measuring at least one of sample generation rate or biofluid flow rate into the device (e.g., using a thermal flow meter).
In an embodiment of the disclosed invention where the sample volume is known and/or can be pre-determined, the above measures of generation rates and flow rates may also be used to provide chronological assurance of the sampling interval. With reference to
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Because sample volumes in embodiments of the disclosed invention are dramatically reduced, which allows for use with very low sample generation rates, a possible confounding factor is therefore transepidermal water loss. This effect is more than just a one-way transfer of water from skin to outside of the body, and depending on osmolality (which has greater osmolality, skin or collected biofluid), the collected sample of biofluid could either be concentrated (water loss) or diluted (water gain).
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The following examples are provided to help illustrate the disclosed invention, and are not comprehensive or limiting in any manner.
Sweating was stimulated using the Wescor Nanoduct iontophoresis protocol with carabachol substituted for pilocarpine (0.5 mA for 1.3 mA-min, area stimulated was on the forearm with a 1.89 cm2 disk). After the forearm stimulation site was left to sweat for 15 minutes, reverse iontophoresis was performed on half of the stimulated area using an ActivaDose controller set to apply 0.2 mA for 10 minutes. The active electrode was half of a 3% agarose disk within a custom holder. After 2 minutes, a 7% suspension of bromophenol blue in cosmetic-grade PDMS oil was applied to the skin to visualize sweating. The voltage applied during reverse iontophoresis was largely constant during most of the test. This experiment was promising as it showed no detectable reduction in sweat rate due to iontophoresis. The sweat stimulation lasted for greater than 24 hours, which is greater than the 1-2 hours expected when using pilocarpine at practical doses (simply increasing the dose cannot provide longer stimulation with pilocarpine because it is rapidly metabolized). Accordingly, with lower doses of carbachol, sweat stimulation may last for more than 3 hours, more than 6 hours, more than 12 hours, or more than 24 hours.
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Also, a faster sampling rate due to a reduced sample volume can be used to reduce the reverse iontophoresis current density. For example, consider a reverse iontophoresis current density of 0.3 mA/cm2 where the sample volume is not reduced, and then an embodiment of the disclosed invention that reduces the sample volume by 500× may use a current density of 0.0006 mA/cm2. This is a first order calculation that assumes advective flow rate of interstitial fluid is proportional to reverse iontophoresis current density. It is recognized that there will be individual variances in current densities used based on the intended application, and there may be a threshold current density that is too low to support a net advective flow toward the sensors. In some cases, sweat generation may provide the needed advective flow toward the sensors. In various embodiments of the disclosed invention, devices may operate with reverse iontophoresis current densities of less than 0.1 mA/cm2, less than 0.05 mA/cm2, less than 0.02 mA/cm2, less than 0.01 mA/cm2, less than 0.005 mA/cm2, or less than 0.002 mA/cm2. Further, because interstitial fluid can have a lag-time compared to blood, applying a device according to an embodiment of the disclosed invention where the dominant pre-existing pathway for analyte extraction is the sweat ducts may have a reduced lag-time compared to another dominant pre-existing pathway because the sweat glands are at least partially closely surrounded in some cases by a capillary bed with blood flow.
Example 3 provides a hypothetical calculation of wicking pressures for elements of the disclosed invention. For purposes of the calculation, the wicking coupler will have the greatest wicking pressure, followed by the wicking pump, and lastly the wicking collector. These relative wicking pressure strengths will ensure that biofluid is continuously removed from the wicking collector so that negligible biofluid remains on the skin surface.
In all of the calculations, the wicking pressures originate from negative Laplace pressure, Δp=γ(1/R1+1/R2), where the surface tension of the biofluid is close to that of pure water (γ of about 70 mN/m). For simplicity, assume the fluid is constant, and principal radii, R1 and R2, are concave (negative). To further simplify the discussion, the effective radii for each sub-component is calculated (i.e., no need to quantify wicking pressure, smaller radii equals larger wicking pressure).
The space between skin and a wicking collector. First is a rough 2D calculation of what is required to reduce the effective biofluid volume to at least 10% of the available biofluid volume, which will also reduce skin surface contamination. Assume 60 μm peak-to-valley skin roughness, which would be greater with hair or skin defects. If the wicking collector is touching the skin, to reduce the effective biofluid volume to 10% of available volume, the biofluid will be wicked into a space that extends only 20 μm out from the skin ridges (assuming triangular ridge shapes), with a meniscus between skin and the collector that spans about 20 μm. Next, assume a biofluid contact angle on skin of θskin=0°, which represents a hydrated and swollen skin surface (typical contact angles are around 90°). Using a wicking collector made of polyamide (nylon, PA46) that is hot-embossed with a network of rectangular channels similar to those shown in
The wicking collector. Assume the wicking collector has square cross-section microchannels with a 1:1 aspect ratio and width w, for which an effective single capillary radius, Rcollector, can be calculated as Rcollector=w/(3 cos(θpoly)−1). The wicking pressure of the collector may sustain the less than 10% biofluid volume between skin and the wicking collector, and therefore Rcollector=Rskin=10 μm. This Rcollector value yields a calculated channel width of about 11-12 μm. A suitable material for the wicking collector is polyamide (nylon), because it is easily microreplicatable, hydrophilic, and relative to many other polymers, exhibits lower non-specific biofluid protein and analyte binding. The wicking collector could be initially be coated with a layer of poly-vinyl-alcohol (PVA) water-dissolvable polymer of 10's of nm thickness, to enable wetting past channel junctions.
The wicking coupler. Next, assume a 10's of μm thick wicking coupler between the wicking collector and a sensor. For device operation, the wicking coupler must keep the sensors continually wetted with a new sample of biofluid. To achieve this, there is at least a 10× decrease in effective capillary radius or Rcoupler about 1 μm (this includes a margin of error to allow for possible variances). There are several materials available from which a wicking material with micrometer-scale capillaries may be fabricated, for example, a hydrophilized nano-cellulose material that is 20 μm thick when hydrated. Nano-cellulose forms a gel-like material which remains cohesive even when hydrated due to microfibril interactions. Nano-cellulose is soft and should promote wetting to sensors. Another attractive possibility is to coat and polymerize a thin film of a hydrogel or super-porous hydrogel, or coating with agar. Hydrated hydrogels can have pore sizes sufficient to allow advective transport of even large proteins. Super-porous hydrogels have a physically open porous network that can be tuned from sizes of 100's of nm to several μm's. A hydrogel wicking coupler has further advantages because hydrogels (1) are pliant when wet and with slight pressure will remain in wetted contact with sensors; and (2) can be coated onto, and in some cases adhered to, the polyamide wicking collector or sensors.
The wicking pump. In this example, the pump serves primarily as a method to collect and dispose of excess biofluid throughout device operation. The wicking pump may have greater wicking pressure than the wicking collector, but its wicking pressure may not exceed that of the wicking coupler or the pump will remove biofluid from the wicking coupler and leave inadequate biofluid on the sensors for accurate measurements. The wicking pump having an effective wicking radius of Rpump=2-3 μm may be fabricated by simple techniques, such as stacking of a plurality of hydrophilic membrane filters (e.g., made of nitrocellulose or other membrane materials) that have well-tuned pore sizes and wicking pressure; or by use of relatively homogeneous beads (e.g., commercial monodisperse Reade Silica powder); a by use of a longer-chain length hydrogel; micro/nano-porous sponges; or other suitable components Again, the effective Rcoupler could be decreased to 10's or 100's of nM to allow a wider selection of materials and effective radius Rpump for the wicking pump. The pump can be designed to store 10's to 100's of μL of biofluid, allowing for continuous use for greater than 24 hours at 0.5 nL/min/gland and 100 glands/cm2 which is greater than 12 hours of continuous use, which is greater than 6 hours of continuous use. Note, the 10% volume between skin and the wicking collector could be further reduced by the wicking pressure of the wicking pump.
Consider a device similar to the device in Example 2 with an effective space between skin and the wicking collector of 50 μm in height. If the wicking collector, wicking pump, and wicking coupler have greater wicking pressure than the wicking space, then the wicking space would be filled with biofluid. The approximate time to refresh this volume with new biofluid can be translated into biofluid sampling interval, and using first order calculations of simply refilling that volume, the sampling interval would be very long. If a wicking collector or other elements, like a wicking pump, are added to reduce or eliminate the sample volume associated with the effective 50 μm of space between skin and the collector, then the wicking collector should have an effective sample volume of less than 50 μm in the area that it is on or adjacent to skin. Otherwise, adding the wicking collector increases the total sample volume, meaning it does not help reduce the sample volume between the device and skin.
This application claims priority to U.S. Provisional Application Nos. 62/196,541 filed Jul. 24, 2015, 62/328,907 filed Apr. 28, 2016, and 62/357,643 filed Jul. 1, 2016, the disclosures of which are hereby incorporated by reference herein in their entirety.
| Filing Document | Filing Date | Country | Kind |
|---|---|---|---|
| PCT/US16/43862 | 7/25/2016 | WO | 00 |
| Number | Date | Country | |
|---|---|---|---|
| 62196541 | Jul 2015 | US | |
| 62328907 | Apr 2016 | US | |
| 62357643 | Jul 2016 | US |
