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Microfluidic tool box as technology platform for hand-held diagnostics.

The idea that microfluidic chips can perform all of the various types of point-of-care testing (POCT) [3] in one system is attractive (1). A platform for immunoassay, chemistry, blood gas, electrolyte, nucleic acid, hematology, and coagulation testing is easily envisioned. POCT methods currently use a variety of formats and materials as means of flow propulsion to achieve fluidics that minimize assay steps and improve ease of use (2). Flow propulsion occurs simply by absorbent, flow-through, chromatographic, or capillary actions. Designs are typically single-use disposables made of inexpensive combinations of hydrophilic and hydrophobic materials and are capable of detecting 1 or more analytes. Alternatively, mechanical mixing or centrifugal, pressurized, or electrophoretic forces can be applied, but these systems are generally more costly (3-7).

Absorbing and drying chemical and biological reagents into a matrix such as paper was one of the first technologies to drive point-of-care devices (8,9). Substrates have since been expanded into membranes, films, and polymer surfaces. Adsorbent substrates became useful in formats to move fluid laterally along a strip or cassette base (10, 11). This principle of lateral flow has been particularly useful for chromatographic separation in immunoassay strips. The majority of POCT devices now use combinations of substrates with dry reagents and cassettes. Generally, treatment with additional liquid reagents is needed to reduce sample matrix effects and improve results.

Capillaries and sample wells have also long been used as disposable formats. Cassettes containing molded channels on the millimeter to centimeter scale are formed by injection molding, embossing, printing, and layering (12-18). These devices use capillary passageways to transfer a sample into reagent areas or to draw off excess sample. Precise control of timing and metering microliter volumes is not possible because flow into an absorptive area is difficult to control (13,14). No means of accurately distributing (metering and splitting) small volumes was succesful even after accounting for the impact of venting. Attempts to uniformly distribute the sample over the reagent region led to placing the reagent inside millimeter-sized capillary passages (16-18). Although this lowered sample volumes, the ability to control flow for additional processing (mixing and separating) was not achieved. Centrifugal devices were identified as an alternative format and extended the usage of capillaries, but they require personal computer-sized devices to control fluid flow (19-21).

Recent trends toward miniaturized devices such as micro total analysis systems (TAS), biological microelectromechanical systems (BIO-MEMS), or Lab-on-a-Chip devices are becoming common for applications outside of POCT (22, 23 ). Several attempts to develop POCT devices have produced commercial products (24-27). Such systems have one or more structures of micrometer-sized geometry down to 0.1 [micro]m produced by a variety of microfabrication techniques. Microfabricated surfaces for separation and capture have been described (28, 29). These approaches are currently being to develop microarray systems for high-throughput screening and single-nucleotide polymorphism identification (30, 31), but in general, these systems are too costly for use as disposable hand-held devices.

Our goal was to test a disposable format based on micrometer-sized structures as a POCT device. We wanted a format that uses separable reagent substrates but is capable of the complex processing needed for quantitative analysis of blood and urine specimens. Our approach was to first mold a generic chip with inlets, wells, and vents and then form microstructures such as connecting channels and geometries by micromachining and laser ablation. Surface energies and geometries were varied and characterized by testing with optical reagents. Useful microstructures were then molded into multistep reaction designs and reexamined. Finalized elements were combined into complex protocols directed toward chemistry and immunoassay testing.

Materials and Methods


Because of its tolerance to cracking and chemical inertness, all first models used Makrolon[R] polycarbonate (Bayer AG) as the surface for embedding fluidic geometries by use of injection molding, laser ablation, and diamond milling (microParts). Polycarbonate is hygroscopic, and its surface energy decays when exposed to water, adhesives, and polar molecules. The next round of working models used polystyrene (145D grade; BASF) and were produced by injection molding and laser ablation but without machining. Molds prepared by lithography achieved 0.1 [micro]m tolerances, whereas machined molds achieved 1 [micro]m, as determined with a Dektak surface profiler (Veeco Corp.). In all cases, chip surfaces were made hydrophilic to obtain water contact angles ([[PHI].sub.water]) from 20 to 80 degrees (sessile drop method). Surface treatment was accomplished by use of a high-energy gas of hydrophilic polymer plasma (microParts). Polystyrene exhibited high surface stability ([+ or -] 5 dynes/[cm.sup.2]) as determined by water contact angle measurements over a year (CV = 14%).

Reagents were dried on nitrocellulose membrane, glass, and cellulose substrates (Whatman Ltd) and were punched and placed as 2-[mm.sup.2] areas into chip by use of a robotic arm on an automated assembler (Kuntz Manufacturing Co., Inc.). Reagent pilots were assembled under controlled temperatures (25-35[degrees]C) and humidity (20%-30% relative humidity). At separate stations on a rotary table, liquids were filled directly into chips and pressure-sensitive adhesive lids (Excel Scientific) were applied. Lids used optically clear polypropylene films for optical areas and aluminized Mylar films for areas needing low moisture vapor transmission rates.


Urinalysis reagents for protein, albumin, leukocytes, hemoglobin, creatinine, glucose, nitrite, ketone, and protein were the standard colorimetric chemistry papers (Multistix[R]; Bayer HealthCare LLC). Urinalysis controls (Medical Analysis Systems) were used for test solutions. Chromogenic glucose reagent placed on a nylon membrane (Biodyne, Pall Corp.) was prepared as described elsewhere (32). For testing, whole blood pretreated with heparin was incubated at 25[degrees]C to degrade naturally occurring glucose. The blood was supplemented with 0, 500,1000, 2000, 4000, and 6000 mg/L glucose and assayed on the glucose reference assay instrument (YSI Life Sciences)

Dry and liquid reagents were prepared for a hemoglobin (Hb) [A.sub.1c] immunoassay based on an immunoaffinity approach using conjugated blue latex particle assays (Fig. 1). Assays for both high (8%-13% Hb [A.sub.1c]) and low (3%-8% Hb [A.sub.1c]) concentrations were prepared and shared the same lysis solution (Cellytic-M; Sigma-Aldrich), waste substrate (Drikette paper; Multisorb), and capture phase. The capture phase was made by striping supported nitrocellulose substrate (5.0 [micro]m pore size) by use of a striper (IVEK Corp) to place two 4-mm bands in a 14 x 1.6 mm capture zone. One band contained Hb [A.sub.1c] agglutinator [1 g/L in phosphate-buffered saline (PBS), pH 7.4], and the other contained anti-fluorescein isothiocyanate (FITC) monoclonal antibody (3 g/L in 0.05 mol/L borate, pH 8.5).


For the high-concentration range assay, the conjugate substrate was made by drying blue latex conjugate on glass fiber paper in a 1:4 dilution with casein blocking buffer (Pierce). Blue latex conjugate was made by labeling bovine serum albumin (BSA) with FITC and anti-Hb [A.sub.1c] antibody. The labeled-BSA was attached to blue latex particles (300 run; 67 [micro]eq of COOH/g) at a loading of 30 [micro]g BSA-FITC-anti-Hb [A.sub.1c]/mg of latex. A wash solution of PBS containing 1 g/L BSA was used.

The conjugate pad for the low-concentration range assay was made by drying the blue latex conjugate on a glass fiber membrane in a 1:400 dilution with blocking buffer. The wash solution was a 1:10 dilution of anti-FITC antibody latex conjugate in PBS containing 1 g/L BSA. Anti-FITC antibody conjugate was prepared by loading blue latex particles (1 mg) with 10 [micro]g of antibody.

Calibrators for Hb Alc were prepared by adding synthesized peptide reagent (Glc-Val-His-Leu-Thr-Tyr-Cys) to assayed blood 5%-15% Hb [A.sub.1c]. Blood was collected with potassium oxalate and sodium fluoride as anticoagulants. Calibrator concentrations were assigned by direct comparison with DCA 2000+ results (Bayer).


Use of color generated from the reagents allowed visualization of fluidics and measurement of responses. We used 2 types of optical detectors for reading chips, based on video capture (Coreco Imaging). Both systems were equipped with a spinning table, programmable controller, stepper motor, opto-electronic trigger, and dividing circuitry to allow video capture while the table was spinning or stationary. One system used a color camera with white light-emitting diode (LED) lighting in a ring strobe, and the other used a monochrome camera (CVM-536 and MV 70; Jai Pulnix Corp.) with a 4-wavelength LED illuminator (460, 525, 620, and 660 run). An LED stopwatch and a tachometer were added to track the time and speed during the spin sequence. Capillary flow in the chip was measured in a stationary position. Fluid stops were overcome either by spinning (200, 500, 1500, or 2500 rpm at a 4-cm radius) or in a stationary position by dispensing additional liquid in 0.1-[micro]L increments to increase hydrostatic pressure. Fluidic protocols included timing steps for changes in hydrostatic pressure and were controlled by a programmed sequence. Images taken from the chip experiments were saved by a DVD recorder and included color calibration markers. Collected video files were digitized and reviewed frame by frame to select the proper pictures at the desired time points. The reagent areas in the pictures were then analyzed by appropriate image processing software to calculate the reflectance and convert to an analytical response.


A tool box of microstructures was constructed for fluidic control, metering, liquid application, mixing, and separation. The geometric shapes used included 50-[micro]m diameter posts, 100-[micro]m notches, and connecting capillaries with 10- to 200-[micro]m cross-sectional diameters (Fig. 2). These structures were applied in the interfacial regions between the larger capillaries (1-2 mm depth and width), chambers (0.5-5 mm depth and width), and reagent substrates. Designs were conceptualized by use of computer-aided design (CAD) software and made by micromachining and molding. Ideal arrangements and geometries were arrived at by iterative testing.


Flow propulsion was achieved by use of capillaries 10-100 [micro]m in diameter and with various millimeter lengths, which were embedded in the chip base. Flow through the capillaries occurred only after the chip surface was made hydrophilic at a [[PHI].sub.water] of 20-80 degrees. The adhesive lid with a [[PHI].sub.water] of 73 degrees (CV = 14%) had only a small impact on the propulsion. Flow for viscous fluids such as whole blood required lower contact angles, whereas the mobility in nonviscous fluids such as urine allowed higher contact angles. Capillaries 50-200 [micro]m in diameter and 0.5-5 mm in length were filled with millisecond velocities. Flow was consistent among samples until capillary diameter reached 10 [micro]m; in larger-diameter capillaries, flow became erratic.


Flow stopped at the end of a micrometer-sized capillary when it reached an opening to a wider and deeper passageway, called a "drop stop" (Fig. 2). A substantial increase (>5-fold) in cross-sectional area was needed to bring flow to a stop. Liquid (up to 20 [micro]L) was held back in large chambers when micrometer-sized exit capillaries were used with a drop stop (Figs. 2 and 3). Chambers, capillaries, and small wells millimeters in size all served as deeper passageways. Stops were allowed on a hydrophilic chip base surface ([PHI] of 20-80 degrees) and were not passive but complete stops until additional pressure was applied. The flow was always away from the source and required venting for air inlets and outlets positioned above and below the stops. Vents in contact with liquids required larger capillaries to prevent clogging.

The pressure needed to overcome the fluid stop was directly dependent on the cross-sectional areas of the capillary and the stop. Capillary diameters of 200, 100, and 50 [micro]m had increasing stop pressures at a fixed surface energy and stop diameter. Stop strength also increased with hydrophilicity as did capillary force. Once fluid passed the stop, flow resumed. Absorbent substrates in the reagent well were found to break the stop if brought into contact with liquid in the micrometer-sized capillaries. Stops therefore needed to be distanced from the absorbent substrates to preventing false triggering.


Metering was achieved by use of fixed volume areas with micrometer-sized capillaries at the beginning and end of the metered area (Fig. 3). The exit capillary had flow held back with a stop. The design was capable of measuring volumes down to 50 nL with a CV of 3%. A variety of fixed volume shapes were used as metering areas. Use of micrometer-sized capillaries as inlets and outlets allowed little fluid to be trapped outside of the metering area. A capillary passage 100 [micro]m wide and 50 [micro]m deep had a low dead volume of ~5 nL per mm of length. The small sample losses allowed the liquid splitting needed for multiplexing.

Variations in the stop strength allowed emptying of fixed volume areas at different times. For example, use of [[PHI].sub.water] of 25-35, 35-45, and 45-55 degrees allowed a 200 [micro]m x 150 [micro]m x 1.5 mm capillary opening into a stop to be overcome at decreasing pressures. Although this approach worked, creating separate hydrophilic areas was avoided by holding [[PHI].sub.water], at 25-30 degrees and varying the cross-section area of the capillary from 50 to 300 [micro]m.



The design used for entry of biological samples into the chip consisted of an inlet hole and a capillary feeding a containment chamber from the back or top side (Figs. 2-4). To achieve a uniform amount of specimen leaving the chamber, one or more 100 x 100 [micro]m grooves were placed across the exit path. These acted to force alignment of the fluid front and displace air through the vents. An additional overflow chamber was also useful for lowering operator dependence and allowed overfilling by 1 to 20 [micro]L. Finally, conical inlet holes were added to aid sample transfer from a fingerstick drop or a transfer capillary (Aqua Cap[R]; Drummond Scientific). Conical inlets reduced dependence on alignment and back pressure. Complete filling was observed with urine (specific gravity, 1.005-1.030) and blood (hematocrit, 44%-60%).

Reagent fluids (e.g., dilution or wash buffers) were loaded on the chip either at the time of assay or before, within liquid-holding wells. Liquid wells with micrometer-sized exit capillaries and stops at the exits were used to prevent leakage (Fig. 3). Pressure prompted release of liquids as long as the chamber was vented. Preventing vapor diffusion through an exit capillary into other assay areas was a problem over 30 days unless liquid could be sealed from the capillary. Sealed designs to store several microliters with moisture vapor transmission rates of 1 g/[m.sup.2]-day could be achieved with foils and breakaway designs.


Separation of particles from fluids occurred in chambers 6 mm in length and 1 mm in width at depths from 25 to 500 [micro]m. One end of the chamber was closed and the other open to inlet and outlet capillaries. Chambers allowed separation of erythrocytes (5 [micro]m diameter) and latex particles (10 [micro]m). The time to complete separation was dependent on the depth of the separator, the particle diameter, and the hydrophilicity. In all cases, having a surface energy equal to or less than the surface energy of the particles facilitated separation. Separation of 1 [micro]L of plasma from 10 [micro]L of blood was obtained in 10 s with a capillary depth to particle diameter ratio of 10:1 with no additional pressure. Applied pressure (>2 g) was needed for the complete separations required for hematocrit determinations. Under these conditions, good accuracy (95%) and low imprecision (CV <10%) were achieved for hematocrit measurements (vs HemoCue) when the area of the optical image of blood compared with plasma was used for calculation.

Affinity separation using a membrane inside the chip required microstructures to facilitate homogeneous transfer and lateral flow of liquid. Without the interface, flow was nonuniform, and the liquid flowed around or under the substrate, forming air pockets. Reagents, such as antibodies and other biomolecules, were immobilized on the substrate, which allowed affinity assays to be performed and the separation to be viewed. Imaging of the solvent fronts of colored particles allowed comparison of the flow properties to the ideal (10). The entry structure was the primary determinate, with post structures (50 x 75 [micro]m posts) on a platform just above the bottom of the substrate providing ideal uniform flow distribution. Once flow was initiated, the washing flow was dependent only on the exit structure and capillary strength.


Mixing via fluid passage through one or more parallel micrometer-sized capillaries was sufficient for cell lysis or reagent mixing. Shear forces to mix components when they left the receiving manifold could be varied by adjusting the capillary lengths (2-5 mm) and diameters (50-200 [micro]m) to cause a localized increase in flow velocity. Complete mixing was facilitated by longer and shallower capillaries. Momentary turbulence at the point of confluence was confirmed by high-speed video. Mixing efficiencies were observed optically by the uniform color after mixing of 10 [micro]L of 250 g/L polyethylene glycol ([M.sub.r] 20 000) in 0.5 mol/L NaOH with 10 [micro]L of phenol red dye in 50 mmol/L phosphate buffer (pH 4). Instantaneous lysing of 1 [micro]L of erythrocytes with 100 [micro]L of lithium thiocyanate buffer was observed by examination for intact erythrocytes by blood reagent (MULTISTIX). In cases with incomplete mixing, incomplete lysis appeared as dark green spots on the pad.


Combining of elements produced chips for blood glucose, urinalysis, and immunoassays (Figs. 3 and 4). The glucose chips shown in panels A and B of Fig. 3 were used for 48 whole-blood glucose measurements. Each assay area consists of an inlet port, a lead channel, a fluidic interface area for sample and reagent substrate, and a venting structure through which the air is removed. The array of microstructure posts required for color uniformity is shown in Fig. 3A. Speed grooves and microstructure placement in the prechamber and the reagent chamber were adjusted to allow bubble-free filling and uniform wetting of 300 nL of sample across the glucose reagent substrate (not shown). The glucose chip gave results after 10 s by use of reflectance membranes (CV = 4.2%) with good correlation ([R.sup.2] >0.99). An overfill tolerance >1 [micro]L was observed.

The MULTISTIX urine chemistries were read on a 16-channel chip (Fig. 3, C and D). This chip was capable of multiple steps. After addition of 50 [micro]L of urine to the inlet port, 16 separate 2.0-[micro]L aliquots were split and metered within the chip and, when triggered, transferred into the first chemistry reaction area. The reaction was read optically after 15 s, and then a second pressure triggered the volume into a second chemistry area for another reaction and optical reading. After 60 s, washing of the second chemistry area was initiated, and wash fluid was collected in a waste well. The urine chip produced 16 separate assays with calibration curves comparable to those obtained with dip-and-read reagents. The urine chip provided a sample pretreatment step in the first reaction area. Hydrophobic reagents were retained in paper substrates, whereas water-soluble indicators were washed into later wells. Complete washing of all color from the pads was observed for these with water-soluble dyes. Passing of multiple volumes through the substrate allowed amplification while washing removed interference. The latter was key to providing quantitative results for these reagents by reducing sample matrix bias.

Two examples of immunochips are shown in Fig. 4: one for liquid reagents dispensed at the time of assay (Fig. 4, A and B) and the other with on-board liquid reagents (Fig. 4, C and D). Heterogeneous competitive immunoassays for Hb [A.sub.1c] were tested with both (Fig. 1). The same chips were equally useful for heterogeneous sandwich immunoassays. A homogeneous immunoassay for Hb [A.sub.1c] would not require the separation elements included in these designs but rather a different combination of tool box elements.

The immunochip shown in panels A and B of Fig. 4 consisted of 4 millimeter-sized wells connected by micrometer-sized capillaries and microstructures. A separate liquid dispenser added lysis and wash buffer. The first well served as the specimen inlet with a 300-nL meter volume and as the target for dispensing. The second area contained conjugate pad and was filled with sample when 3.0 [micro]L of lysis buffer was dispensed. Addition of wash fluid at 2 min overcame the stop and started the flow from the conjugate area into the capture substrate in the third well. As 12 [micro]L of wash fluid was added, flow exited the capture area into the waste area containing absorbent in <1 min. At the end of the wash, the bands in the capture area were read.


The immunochip shown in panels C and D of Fig. 4 is an example of a chips with on-board liquids and consists of meter, capture, conjugate, and waste wells connected to 2 liquid holders, a mixer, and 2 liquid overfill chambers. Again, micrometer-sized capillaries and structures were used to interface areas. In practice, a whole blood specimen was added by the user into the inlet (0.5-2 [micro]L), and the 0.3-[micro]L meter area was filled by capillary action. The flow sequence started mixing of the lysis buffer (3.0 [micro]L) from a liquid well with sample, causing the mixture to enter the mixing manifold and pass through a mixing capillary followed by transfer of 2 [micro]L into the conjugate pad well. After incubation for 2 min, the liquid in the conjugate chamber was loaded in the capture chamber, starting wicking into the nitrocellulose capture zone by lateral flow. After antigen binding, the capture zone was washed by 12 [micro]L of additional fluid released from a second liquid well. The wash fluid contained an additional conjugate to increase sensitivity for the low-range assays. Waste containment was added for excess liquids from the specimen port, mixer, and capture phase.

Both chips were tested with the same Hb [A.sub.1c] reagents. The best results produce a response of 3%-13% Hb [A.sub.1c] with an overall SD of 0.18% Hb [A.sub.1c] (Fig. 5). Washing of the nitrocellulose membrane was very complete in both examples, and reaction band color was uniform.


Fluid control, or the pumping and valving of fluid, is a key requirement for bioassays in which fluids must be retained and then dosed into designated areas at desired times (e.g., metering, incubation, reaction, and washing). Fluidic control is achievable in several different ways (23-29, 33, 34), but the use of micrometer-sized structures with separable reagent substrates works well for disposable POCT chips and eliminates electrical, mechanical, or sacrificial valves. The number of times that the flow can be started and stopped is more limited, but complex assay protocols can still be achieved.

Although the use of capillary channels that connect to reagent wells is well established, the use of devices possessing micrometer-sized structures to control flow volumes away from and into reagent substrates is less well known (13-18). Tight control of surface energies and dimensions allows pressure differences for triggers to be better tolerated by low-cost systems. In our chips, the length of time that the pressure needs to be applied is small and once stops are broken, the overall hydrophilicity of the chip continues the flow. Large capillaries (millimeter-sized) require tight control of pressure changes (e.g., centrifugal) and were unsuitable for our chips. There are considerable advantages to the use of micrometer-sized capillaries to feed dead stops because liquids can be kept from contacting dry reagents, and flow rates changes are ON/OFF.


A key distinction is that the designs shown do not require any centrifugal force to operate. Only capillary force is used to move fluid through the chip. Although centrifugal force can be used to overcome the flow stops, stops were also overcome by dispensing of additional liquid upstream from the stop. All other design elements were purely capillary driven (no spinning), including the metering, sample application, separation designs, and mixing designs. Flow stops were used at the end of the on-board liquid containers and again were adjusted for triggering either by various centrifugal forces or by additional dispensing volumes without centrifugal forces. The capillary separation chamber operated with or without centrifugal force.

Fluid metering is critical to performing quantitative assays because it allows measured volumes to be reacted and split for multiplexing. The techniques generally used in capillary systems, such as overflow metering, in which a chamber is filled to an excess that is overflowed, did not have the accuracy needed for nanoliter volumes (13,19, 26). Overflow areas were used for the containment of waste but not metering in this design. Passive stops also could not be used for metering; instead, dead stops were required to prevent contact between liquid and the absorbent reagent substrate (35). The low dead volumes in micrometer-sized capillaries provided the minimal volume loss needed for metering and splitting for multiplexing.

Inlet and outlet ports are critical in POCT chips, because microchips contain air that must be expelled (36). Underfilling resulting from trapped air leads to underestimation. At the same time, sample not completely entering the inlet port and remaining on the surface of the device causes carryover and contamination. Air inside the capillary must be controlled because fluid flows or trapped air will cause flow issues. One solution has been to expel the air before or while liquid is entering the inlet port, such as with a pipette with a plunger (4 5, 7). We found a means of allowing air to escape the chip by placing micrometer-sized structures in the direction of the flow. As the fluid gathers through the structures, air is expelled away from the solvent front. The containment area for samples in the design was important for preventing sample carryover while allowing for overfilling and thus improved the overall ease of use.

Mixing is important in bioaffinity assays because sample dilution and incubations are needed for binding. Centrifugal mixing, in which components are mixed in a chamber by tidal movements or oscillation cycles, is well known (19). Perturbation structures in capillaries have been shown to facilitate mixing (37, 38). In the capillary mixer design tested, momentary turbulence was produced by use of high-velocity micrometer-sized capillaries at the point of confluence exiting the feed chamber. The resulting mixture then quickly hits the bottom chamber, and the fluid is further mixed. Running the mixture through high-velocity micrometer-sized capillaries allow the rapid mixing needed in POCT assays.

Centrifugal separation is a well-understood principle (21). Small chambers with tight wall-to-particle ratios have been less studied. In our design, separation efficiency was controlled by the ratios of particle diameter to chamber depth and surface energies, suggesting that attraction of particles to the wall is the driving force in this separation. Without adjustment of wall surface energy, adhesion was either irreversible or separation did not occur. For POCT applications, separation at exceedingly low forces and times is critical, making this approach attractive.

Separation with affinity-capture substrates allows fluidic exchanges between the detector area and the reacted fluid (10). Approximately 50 exchange volumes are typically seen with nitrocellulose membranes, which helps amplification. The greater volume of nitrocellulose membrane compared with a surface-immobilized capture monolayer allows less fluidic exchange. Both substrate and surface immobilization face problems of uniform solvent flow across the detector area. Although separate reagent substrates are not typically applied to microfluidic chips, the use of microstructures to draw fluids into and away from substrates provided unexpectedly uniform affinity reactions. Posts, tiers, and grooves were key geometric shapes impacting the direction of fluid into the substrate.

In conclusion, disposable chips based on microstructures and a hydrophilic surface energy work well with reagents on substrates and provide a possible POCT platform. Areas for substrate preparation can be separated from chip surfaces, allowing cost savings. No lithography was required for molding. Although capable of complex multistep protocols, the design is fundamentally simple, and the inner workings can be easily kept transparent to the user. A tool box of useful microgeometries allows easy reconfiguring. These chips require no pressurization or electrokinetic or centrifugal forces and provide results typical of those expected in POCT devices with less reagent and smaller samples.


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[1] Diagnostic Division, Bayer Healthcare LLC, Tarrytown, NY.

[2] microParts GmbH, Boehringer Ingelheim, Dortmund, Germany.

[3] Nonstandard abbreviations: POCT, point-of-care testing; Hb, hemoglobin; PBS, phosphate-buffered saline; FITC, fluorescein isothiocyanate; BSA, bovine serum albumin; and LED, light-emitting diode.

* Address correspondence to this author at: Diagnostic Division, Bayer Healthcare LLC, 1884 Miles Ave., Elkhart, IN 46515-0070. E-mail michael.

Received April 28, 2005; accepted June 2, 2005.

Previously published online at DOI: 10.1373/clinchem.2005.052498
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Title Annotation:Oak Ridge Conference
Author:Pugia, Michael J.; Blankenstein, Gert; Peters, Ralf-Peter; Profitt, James A.; Kadel, Klaus; Willms,
Publication:Clinical Chemistry
Date:Oct 1, 2005
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