Electrokinetic Properties of Lubricin Antiadhesive Coatings in

Jan 27, 2016 - Australian Centre for Research on Separation Science, and ARC Centre of Excellence for Electromaterials Science, School of Physical Sci...
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Electrokinetic Properties of Lubricin Antiadhesive Coatings in Microfluidic Systems George W. Greene,*,† Emer Duffy,‡ Aliaa Shallan,‡ Alain Wuethrich,‡ and Brett Paull‡ †

Institute for Frontier Materials and ARC Centre of Excellence for Electromaterials Science, Deakin University, Geelong, VIC Australia ‡ Australian Centre for Research on Separation Science, and ARC Centre of Excellence for Electromaterials Science, School of Physical Sciences, University of Tasmania, Hobart, Australia ABSTRACT: Lubricin is a glycoprotein found in articular joints which has long been recognized as being an important biological boundary lubricant molecule and, more recently, an impressive antiadhesive that readily self-assembles into a well ordered, polymer brush layer on virtually any substrate. The lubricin molecule possesses an overabundance of anionic charge, a property that is atypical among antiadhesive molecules, that enables its use as a coating for applications involving electrokinetic processes such as electrophoresis and electroosmosis. Coating the surfaces of silica and polymeric microfluidic devices with self-assembled lubricin coatings affords a unique combination of excellent fouling resistance and high charge density that enables notoriously “sticky” biomolecules such as proteins to be used and controlled electrokinetically in the device without complications arising from nonspecific adsorption. Using capillary electrophoresis, we characterized the stability, uniformity, and electrokinetic properties of lubricin coatings applied to silica and PTFE capillaries over a range of run buffer pHs and when exposed to concentrated solutions of protein. In addition, we demonstrate the effectiveness of lubricin as a coating to minimize nonspecific protein adsorption in an electrokinetically controlled polydimethylsiloxane/silica microfluidic device.

1. INTRODUCTION Current trends in microfluidic and “lab-on-a-chip” technologies promise a new era of highly integrated functionality which could lead to unprecedented advancements in medical diagnostics1−3 and the study of complex cellular processes.4,5 However, despite such promise, the field of microfluidics still struggles with an old problem: how to simply and cost effectively control the unwanted adhesion of proteins, antibodies, and other notoriously “sticky” biomolecules to the walls and surfaces of these devices.6,7 Nonspecifically adsorbed proteins reduce the signal-to-noise ratio (i.e., sensitivity) and alter the surface charge,7 making it difficult to perform electrokinetic processes in the device with consistent, reliable, and highly repeatable performance. There now exists a large body of research focused on investigating and developing a wide variety of technologies for coating the surfaces of microfluidic devices with antiadhesive materials that include chemically and physically grafted polyethylene glycol based polymers;8 biopolymers such as dextran9 or hyaluronic acid;10 poly(vinylpyrrolidone);11 and zwitterionic polymers12,13 and surfactants.14 While many of these coating technologies are highly effective at blocking adhesion, most possess inherent limitations which significantly limit the wider adoption in commercial microfluidic device technologies which is trending toward inexpensive, point-of© 2016 American Chemical Society

care, and disposable systems. For instance, the chemical grafting of antiadhesive polymers often requires an initial surface functionalization or pretreatment processes which adds undesirable steps and significant cost to the manufacturing process and is often difficult to perform reproducibly on common and inexpensive substrate materials (e.g., polymers such as poly(methyl methacrylate), polydimethylsiloxane (PDMS), or polycarbonate).15−21 Using on-chip electrokinetics (e.g., electroosmosis and/or electrophoresis) to control the flow of fluid and the migration of biomolecules is a desirable feature for lab-on-a-chip devices that potentially expands functionality and facilitates miniaturization and portability of systems. Consistent and reliable electrokinetic operation of a microfluidic system is dependent upon surfaces having a strong and unchanging charge. Unfortunately, most of the effective antiadhesive molecules are charge neutral22 (e.g., PEG) meaning that after grafting/ adsorbing the molecule to the surface of the device, the surface charge is significantly reduced leading to suppressed, nonuniform, inconsistent, and/or unstable electrokinetic properties.11,14 Likewise, many coating materials used to improve Received: September 21, 2015 Revised: January 3, 2016 Published: January 27, 2016 1899

DOI: 10.1021/acs.langmuir.5b03535 Langmuir 2016, 32, 1899−1908

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lubricin coated capillaries under conditions of varying background electrolytes and pH and upon exposure to solutions of proteins. Additional experiments performed in a PDMS/silica microfluidic device with a simple cross design demonstrate the ability of lubricin coatings to almost completely block the nonspecific adsorption of protein to the channel surfaces while allowing facile operation using modest electric field strengths to drive the flow of fluid within the device.

the electrokinetics in microfluidic systems, most notable polyelectrolytes23−25 or ionic surfactants,26−28 often have limited pH ranges,28 can potentially lead to significant levels of nonspecific binding of proteins and antibodies even when the charge of the coating is the same sign as the net charge on the protein,29−31 and may prevent the adhesion of either cationic or anionic proteins but not both.28,32 Lubricin (LUB) is a glycoprotein found in the synovial fluid and covering the cartilage surfaces of articular joints whose important role as a major boundary lubricant in articular joints is now well-known.33−42 The structure and chemistry of the LUB molecule has recently become well characterized and is described in detail in previous reports;34,35,37,38,43 however, a brief description of some of the key features of LUB is as follows. The LUB molecule appears as a long, flexible molecule with a fully extended “contour” length of lc ≈ 200 nm and a diameter of a few nanometers.34 Its molecular weight is approximately Mw ≈ 280−320 kDa which is high compared to the number of amino acids in the sequence, (∼800) due to the heavy glycosylation of the central portion of the molecule that has come to be known as the “mucin domain”.34 Within this mucin domain, short glycan oligomers terminated primarily by polar galactose (∼33% of total glycans) and negatively charged sialic acid (∼66% of total glycans) are O-linked to threonine and serine residues.34,43 This central mucin domain is believed to be responsible for both LUB’s excellent lubrication as well as antiadhesive properties.38,44,45 Flanking either end of the mucin domain are the lightly glycosylated “end domains” of the protein which contain sub domains similar to two globular proteins, somatomedin-B and homeopexin, known to play a special role in cell−cell and cell−extracellular matrix interactions, e.g., binding.34,43 These end domains are therefore extremely “sticky” and are able to adhere to nearly all types of surfaces. These end domains have also been shown to associate with each other to form molecular “loops” and also allow the LUB to easily form dimers, trimers, and tetramers where the loops, joined through associated end domains, adopt “figure eight” and larger, loosely twisted aggregate structures.37 The amphiadhesive nature of the LUB molecule enables its selfassembly onto surfaces to form a dense, well ordered telechelic polymer brush layer by which the “sticky” end-domains adsorb to the substrate while the nonadhesive mucin domain fully extends into the solution in a “loop” conformation.38 Recent experiments have demonstrated that lubricin exhibits impressive antiadhesive properties that are as-good-as and, in some cases, better than chemically grafted layers of polyethylene glycol.44−46 The telechelic brush architecture adopted by the adsorbed LUB effectively hides the underlying substrate while exposing the larger, heavily glycosylated, and low adhesion mucin domain to the surrounding solution.45 However, unlike typical antiadhesive molecules which are charge neutral, LUB possesses an overabundance of negative charge carriers; most of which are found in the mucin domain loop.43,46 The unique combination of large charge density, selfassembly behavior, and excellent antiadhesive properties makes lubricin an attractive coating material for microfluidic applications. This manuscript reports the antiadhesive and electrokinetic properties of lubricin coatings applied to silica, polytetrafluoroethylene (PTFE), and PDMS/silica microfluidic systems. Capillary electrophoresis measurements were used to characterize the magnitude, uniformity, and stability of the electroosmotic mobility and surface zeta potential within

2. EXPERIMENTAL SECTION 2.1. Lubricin Purification. LUB protein was purified using the procedure described in Greene et al.45 (a slightly modified method based upon the protocols outlined in ref 34) from ∼500 mL of bovine synovial fluid sourced from MC Herd (Corio; Victoria, Australia). The synovial fluid was collected percutaneously from the radiocarpal joints of freshly slaughter cattle (∼1 year old; male and female) using a sterile 18 gauge hypodermic needle and stored in a polypropylene bottle, on ice, until the time of processing (approximately 2 h after collection). The only modification of the procedure described in ref 34 was the elimination of the initial membrane filtration and subsequent resuspension step (following the centrifugation of the raw synovial fluid) which was determined to be unnecessary. Instead, following centrifugation, the raw synovial fluid was diluted with a solution of 50 mM sodium acetate, 10 mM EDTA, and Roche inhibitor tablets at pH 5.5 until the pH of the dilution reached pH 5.5. After this dilution, the purification proceeded as described in ref 34 with the hyaluronidase digestion step. The extracted and purified LUB was analyzed for purity using a density gradient SDS-PAGE Biorad gel subsequently stained with coomasie blue. The relative purity of the LUB (as a fraction of the total protein content) was assessed using a Biorad imager and spectroscopic analysis and was found to be approximately 89%. The LUB band appeared on the SDS-PAGE gel at approximately the 280 kDa region, consistent with previous reports.34 The concentration of LUB in the extracted solution was determined using the Biorad protein assay, with BSA as the standard. After the concentration of LUB was assayed, the solution was concentrated using a Millipore Amicon Ultra Centrifugal Filter with a 100 NMWL membrane to yield a final concentration of 150 μg/mL of protein in a buffer consisting of 25 mM sodium phosphate, 150 mM NaCl, 0.5 mM CaCl2, and 0.2 mM alpha lactose at pH 7.4. 2.2. Chemicals. This study utilized a number of different run buffer systems, all of which were prepared using reagent grade materials. The run buffers prepared include 2-(N-morpholino)ethanesulfonic acid buffer (MES; pH 5.0), 3-(N-morpholino)propanesulfonic acid buffer (MOPS; pH 7.2), sodium phosphate buffer (sodium phosphate; pH 6.0, 7.2, pH 8, and pH 12), and trisaminomethane buffer (Tris; pH 9). All run buffers were prepared to the same electrolyte concentration of 15 mM and filtered using a 0.22 μm cellulose acetate syringe filter. Experiments investigating the migration and nonspecific adsorption of protein in LUB coated capillary and microfluidic channels were performed using solutions of chromatographically purified bovine serum albumin (BSA; Sigma-Aldrich, ≥ 98%), reagent grade goat Immunoglobulin G (IgG; Sigma-Aldrich, ≥ 95%), and Alexa Fluor 488 labeled Chicken Anti-Rabbit IgG (Life Technologies; whole antibody). All protein solutions were prepared in 15 mM MOPS buffer at pH 7.2 at specific concentrations provided in the text, figures, and figure legends. 2.3. Capillary Electrophoresis Experiments. Capillary electrophoresis (CE) measurements were used to measure the electroosmotic flow and zeta-potential (ζ) in lubricin-coated silica and PTFE capillaries. All CE experiments were performed on an Agilent 7100 CE system with detection using a DAD (Agilent Technologies, Germany) for direct UV detection at a wavelength of 254 nm. OpenLAB CDS ChemStation Edition software for Windows 7 was used for instrument control and all data acquisition. This study investigated two different capillary systems: a 28 cm (22 cm to detector) fused silica capillary (Polymicro Technologies, Phoenix, AZ) having an inner diameter of 1900

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Langmuir 75 μm and a 34 cm long (28 cm to detector) PTFE capillary (Polymicro Technologies, Phoenix, AZ) having a 100 μm inner diameter. Before coating with LUB, both the silica and PTFE capillaries were cleaned by prerinsing with a solution of 0.1 M KOH for 5 min followed by deionized water for 5 min using high pressure in the forward direction. To coat the capillaries with LUB, the silica and PTFE capillaries were filled (using high pressure) with a 150 μg/mL solution of LUB and allowed to sit idle for 10 min to give the lubricin time to self-assemble into an ordered polymer brush layer. The capillaries were then flushed with deionized water for 5 min using high pressure to remove any unbound LUB molecules. Before performing a measurement, the capillaries were equilibrated to the experimental buffer conditions by flushing them with run buffer for 5 min using high pressure. The electroosmotic flow (EOF) velocity, νEOF, within a LUB coated capillary was determined experimentally by measuring the migration time of acetone (an electrically neutral molecule) through the capillary under the action of an applied electric field using the following expression:

vEOF =

Ldetector tm

Figure 1. Schematic of the PDMS cross channel chip used in the microfluidic experiments. In the device, two orthogonal channels having a 50 μm × 30 μm (rectangular) cross-section connect four fluid “reservoirs” into which electrodes are inserted. The applied electrical potentials applied to the four individual electrodes during the initial “loading” phase and subsequent “injection” phase of the experiment are indicated in the figure. The distance from the Loading Channel Inlet, Loading Channel Outlet, and Buffer Channel reservoirs to the injection point (i.e., the intersection of the channels) was 15 mm. The distance between the Detector Channel reservoir and the injection point was 45 mm.

(1)

where Ldetector is the length of the capillary to the detector and tm is the experimentally measured migration time for the injected acetone to reach the detector. The acetone EOF marker was added to a small aliquot of the experimental run buffer to a concentration of 1% wt. and was introduced into the capillary using hydrodynamic injection for 5 s. The electroosmotic mobility, μEOF, is obtained by normalizing vEOF by the electric field strength E = V/Ltotal: μEOF =

vEOF L /t = detector m E V /Ltotal

(2)

where Ltotal is the total length of the capillary and V is the applied voltage. One of the most important properties in any electrokinetic process is the surface zeta-potential, ζ. The zeta potential indirectly describes the electrostatic double layer surrounding the surface and, while it cannot be directly measured, it is routinely calculated using system appropriate models, such as the well-known Helmzoltz-Smoluchowski equation. So long as the inner diameter of the capillary is at least 7 times larger than the electrostatic double layer thickness (as is the case for the capillaries used in these experiments), the EOF creates a flat, “plug flow” velocity profile which allows the zeta potential on the charged inner wall to be calculated from EOF velocity measurements using the following relationship:47 μ η ζ = EOF ε0ε (3)

LUB coated PDMS/silica microfluidic electrophoresis chip was performed. In this experiment, both uncoated and LUB coated devices were filled with the protein solution and left for 5 min. The uncoated and LUB coated microfluidic electrophoresis chips were rinsed with MOPS buffer for 5 min using vacuum at the outlet and the residual fluorescence was imaged at various exposure times using an inverted fluorescence microscope (Ti−U, Nikon, Tokyo, Japan) equipped with a Nikon high-definition color CCD camera head (Digital Sight DS-Fi1c, Nikon, Japan) and operated with NISElements BR 3.10 software (Melville, NY, USA). Excitation (λex at 450−490 nm) and emission (λem at 520 nm) filters (Semrock, Rochester, NY, USA) were used for all experiments. The images were analyzed with ImageJ to measure the fluorescence intensity as an indication of the extent of protein adsorption on the channel surface. In another experiment, a 0.5 mg/mL solution of Alexa Fluor 488 labeled Chicken Anti-Rabbit IgG in MOPS buffer (pH 7.2) was electrokinetically loaded and injected within the LUB coated microfluidic electrophoresis chip. For this experiment, the LUB coated device was filled with MOPS buffer and 5 μL of a 0.5 mg/mL solution of Alexa Fluor 488 labeled Chicken Anti-Rabbit IgG was added to the reservoir of the “loading channel inlet” (see Figure 1). The electrohydrodynamic loading and subsequent injection of the protein was achieved by applying potentials to electrodes submerged in the reservoirs of the four channels of the device as indicated in Figure 1 next to the headings “Loading” and “Injection”. An inverted fluorescence microscope equipped with a CCD camera was used to visualize the pinched electrokinetic injection of the Chicken AntiRabbit IgG while the detection of the fluorescent intensity of the migrating protein zone was carried out using a photomultiplier tube (PMT; Hamamatsu Photonics KK, Hamamatsu, Japan) connected to the microscope located 20 mm from the injection point. Data acquisition was made using an Agilent interface (35900E) connected to a laptop and operated by Agilent ChemStation for LC software (Agilent Technologies, Waldbronn, Germany).

where ε0 and ε are the vacuum permittivity and medium dielectric constant respectively and η is the medium viscosity. 2.4. Microfluidic Measurements. Microfluidic experiments were conducted using a hybrid PDMS/silica electrophoresis chip. The PDMS layer containing a standard cross microfluidic design was reversibly sealed onto a silica slide to form the channels that were all 50 μm wide and 30 μm deep. Four reservoirs, 20 μL capacity, were punched into the PDMS layer at the terminus of each channel thus defining the “Loading Channel Inlet”, “Loading Channel Outlet”, “Buffer Channel”, and “Detector Channel” as illustrated by the schematic in Figure 1. Before running the experiments, the device was coated with LUB by filling the channels with a 150 μg/mL solution of LUB and allowing it to sit idle for approximately 10 min to give the lubricin time to self-assemble into an ordered polymer brush layer. The device was then rinsed with excess 15 mM MOPS buffer (pH 7.2) to remove any unbound lubricin from the microfluidic device. The device remained filled with MOPS buffer until used for experiments. Microfluidic measurements were performed using 2 mg/mL solutions of Alexa Fluor 488 labeled Chicken Anti-Rabbit IgG in MOPS buffer (pH 7.2; see the Chemicals section). An experiment designed to compare the nonspecific adsorption of a noncoated and 1901

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Figure 2. (a,a′) Representative electropherograms obtained for a neutral marker (acetone) in different run buffers of varying pH in a LUB coated silica (a) and PTFE (a′) coated capillary under an applied electric field strength of E = 357 V/cm and E = 294 V/cm, respectively. We note that the instability in the baseline observed in (a′) at pH 6 and 7.25 was determined to be instrumental rather than caused by adsorption or desorption from the capillary walls. (b,b′) The electroosmotic mobility, μEOF, of the neutral solute as a function of run buffer pH as measured in a LUB coated silica (b) and PTFE (b′) capillary. (c,c′) The zeta potential, ζ, of the LUB coating on the silica (c) and PTFE (c′) capillaries as a function of the run buffer pH. In (b,b′,c,c′), the sequential order (i.e., pH changes) of the individual experimental measurements is shown by the red arrows. The green data points shown in these same figures indicate the value obtained for μEOF (b,b′) and ζ (c,c′) upon repeating the measurement at neutral pH 7.25 to ascertain whether the properties of the LUB coating had been altered by changing the run buffer pH. In (b,c), measurements were run consecutively in the order of lowest to highest E before changing the run buffer pH. The error bars indicate one SD above and below the average of three separate measurements.

3. RESULTS AND DISCUSSION

proteins and other large, charged molecules have been especially difficult to separate using CE due to their naturally high affinity for adsorbing to charged surfaces.48 For proteins, this is often true even if the surface and protein share the same net charge.29 CE was used to measure the EOF velocity within LUB coated silica and PTFE capillaries in order to assess the stability and consistency of the LUB coatings, how the charge of the coatings are altered by changes in the run buffer pH, and how the electrokinetic properties of the coatings are affected by the introduction of “sticky” proteins into the capillary. Figure 2a,a′

3.1. Characterization of the Electrokinetic Properties of LUB Coatings. CE is a separation technique based upon differences in electrophoretic mobilities (μep) (i.e., the differences in the size/charge ratio of different analytes). The apparent electrophoretic mobility for a certain analyte is the sum of its μep and the EOF. CE can be a powerful analytical technique; however, it is particularly sensitive to factors such as electrolyte concentration, pH, or the nonspecific adsorption of certain analytes and/or ions that have the potential to change or alter the charge on the capillary walls. For this reason, 1902

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Figure 3. Comparison of the measured electroosmotic mobility, μEOF, as of function of the run buffer pH in (a) uncoated and LUB coated silica capillaries and (b) uncoated and LUB coated PTFE capillaries. In (a) and (b), measurements were performed under applied electric field strengths of E = 357 V/cm and E = 294 V/cm, respectively.

function of pH measured in uncoated and LUB coated silica and PTFE capillaries have been plotted together in Figure 3a,b, respectively, which shows the greater sensitivity of the EOF in the uncoated capillaries to changes in pH relative to the LUB coated capillaries. The lack of any apparent correlation between the changes in μEOF with pH observed in the uncoated and LUB coated silica and PTFE capillaries demonstrates how little the capillary substrate influences the EOF in the LUB coated systems. Although small in magnitude, the observed changes in μEOF with increasing run buffer pH shown in Figure 2b,b′ appear random and unpredictable, sometimes decreasing and sometimes increasing. However, the small variation in the values of μEOF measured at each pH (as indicated by the very small error bars) suggest that these changes are indeed produced by subtle variations in the charge of the LUB layer under these different buffer conditions. One factor contributing to this observed variation in charge arises from the fact that, while the LUB molecule has an overabundance of negative charge, there is still a nontrivial number of positive charge carriers in the molecule which, to some extent, balance a small portion of the negative charge.46 The extent to which these positive charges balance the negative charge will vary with changes in pH and can alter the degree and distribution of negative charge in the LUB coating in unpredictable ways. However, the observed relationships between μEOF (and ζ) and pH in the silica and PTFE capillaries (Figure 2b,b′) do not appear correlated and suggests, not unexpectedly, that the adsorbed LUB layers in the two capillaries are also structurally different. A previous quartz crystal microbalance study showed that the chemical properties of the substrate surface can affect the density of the self-assembled LUB brush layers.45 In the cited study, the density of the LUB brush absorbed onto a negatively charged thiol-SAM (similar, electrostatically, to silica) was approximately 15% higher than that absorbed to hydrophobic polystyrene (similar, electrostatically, to PTFE). The conformational structure of the adsorbed LUB layer, being essentially a telechelic polyelectrolyte brush, is therefore dependent upon such factors as the molecular grafting density and the magnitude of intermolecular (repulsive and attractive) electrostatic forces. A shift in the ratio of negative to positive charges due to a pH change can thus induce a conformational change. As the mucin domain loop in the LUB becomes more negatively charged with increasing pH, stronger intra- and

shows the typical CE electropherograms obtained following the injection of acetone into a 28 cm long LUB coated silica capillary (Figure 2a) and a 34 cm long LUB coated PTFE capillary (Figure 2a′) under an applied electric field of 10 kV (E = 357 V/cm and E = 294 V/cm, respectively) in various 15 mM background electrolytes having a range of pHs. Using eq 1, νEOF was calculated from the measured migration times. The μEOF and ζ were both calculated from the experimentally measured vEOF using eqs 2 and 3, respectively. The relationship between the μEOF and run buffer pH is shown in Figure 2b,b′ for the LUB coated silica and PTFE capillaries, respectively. The relationship between the ζ and run buffer pH is also shown in Figure 2c,c′ for the LUB coated silica and PTFE capillaries, respectively. For the silica capillaries, measurements were performed using three different electric field strengths, E = 179, 357, and 536 V/cm (corresponding to voltages of V = 5, 10, and 15 kV, respectively) and were run consecutively in the order of lowest to highest E before changing the run buffer pH. For the PTFE capillary, measurements were conducted at only a single electric field strength of E = 294 V/cm (V = 10 kV). In the experiments shown in Figure 2b,b′,c,c′, the LUB coating was only applied once at the start of the experiment and the order in which the measurements were performed (i.e., the sequence of run buffer changes) is indicated by the red arrows in Figure 2b,b′,c,c′ In both capillaries, after completing the highest pH measurement at E = 357 and 294 V/cm in the silica and PTFE capillaries, the pH was returned to neutral pH 7.2 and an additional measurement (shown as the green data point in Figure 2b,b′,c,c′) was performed in order to ascertain the stability of the LUB coating. The values of μEOF obtained in the second and first measurements at pH 7.2 were essentially identical in both capillaries, indicating that no significant desorption or degradation of LUB coating occurred as a result of changing the run buffer pH. In both the LUB coated silica (Figure 2b and c) and PTFE capillaries (Figure 2b′ and c′), the measured values of μEOF and ζ across the full range of pHs measured were found to change very little and remained within a narrow range between μEOF = 2.03.0 × 10−4 cm/(V s) and ζ = −25 to −40 mV. The similarity in the values of μEOF (and ζ) in the silica and PTFE capillaries suggests that the EOF is primarily controlled by the charge within the LUB coating and, to a lesser extent by the underlying substrate. For comparison, the values of μEOF as a 1903

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Langmuir interchain repulsion can lead to it becoming more extended. Since the zeta potential of the surface depends upon the density of the charge (i.e., the number of charges per unit volume rather than the absolute number of charges), it is possible that conformational changes (e.g., swelling) triggered by enhanced electrostatic repulsion can paradoxically lead to a reduction in the LUB layer charge density (and a decrease in the magnitude of the zeta potential) as a result of the average distance between charges becoming greater despite an increase in the absolute number of charges in the layer. Unfortunately, CE measurements provide little insight into the conformational structure of adsorbed surface layers and so additional experimental techniques would be required to determine, for certain, exactly how the structure of the LUB layers respond to changes in pH. Interestingly, and for reasons we cannot explain, an apparent instability in the measured value of μEOF (and ζ) was observed in the LUB coated silica capillary at pH 9 (see Figure 2b,c). Although increased variation in the value of μEOF was observed at each specific electric field strength, E (as indicated by the larger error bars associated with each measurement), a much larger and more significant variation is observed between the different electric field strengths that is not observed at other pHs. Typically, such an instability and large variation in μEOF would indicate a change in the surface charge; i.e., due to desorption of lubricin from the capillary walls. However, the value of μEOF returned to the originally measured valued when the measurement was repeated a second time at pH 7.2 (see the green data point in Figure 2b,c) which strongly suggests that the instability cannot be attributed to LUB desorption from the capillary surface. Because the LUB self-assembles into a well-ordered brush layer, there is a consistency in the LUB layer density and molecular packing that leads to consistent and uniform properties from one coating to another. To demonstrate this consistency, the silica capillary used in this experiment was coated three separate times with LUB (as detailed in the methods) to compare how the measured μEOF varied. Before each coating was applied, the silica capillary was cleaned (and any previous LUB coating was removed) by rinsing the capillary with a 1 M NaOH solution for 10 min followed by a 5 min rinse with clean run buffer (15 mM Sodium Phosphate buffer; pH 7.2). Figure 4 shows the μEOF obtained from the three separate coatings which were found to be essentially identical. 3.2. Effect of Proteins on the Electrokinetic Properties of LUB Coatings. The migration times in a CE measurement are notoriously sensitive to subtle changes in the EOF caused by even minor changes in the surface charge of the capillary walls. Often, one of the primary sources of experimental “drift” in observed migration times is due to the adsorption of analyte molecules and/or buffer specific species to the surfaces of the capillary that leads to a change in the surface charge. Proteins, which often possess the gamut of functional groups, are particularly “sticky” molecules that can drastically alter the electrokinetic properties in a microfluidic system by adsorbing to surfaces.48 An experiment was devised to investigate the stability of the electrokinetic properties of the LUB coatings when exposed to concentrated solutions of proteins. In this experiment, a mixed solution of IgG and BSA proteins in a 15 mM MOPS buffer (pH 7.25) was hydrodynamically injected into a LUB coated PTFE capillary and caused to migrate through the capillary under an electric field strength of E = 294 V/cm. Figure 5a

Figure 4. Electroosmotic mobility measured in a silica capillary coated three separate times with an adsorbed LUB layer. Between each coating, the previous coating was stripped away and the capillary cleaned by rinsing for 10 min with a 1 M NaOH solution followed by a 5 min rinse with clean run buffer. The coating of the capillary with LUB was carried out following the procedure described in the “Experimental Section”. The electroosmotic mobility was calculated using eq 1 from the migration times of a neutral marker in a 15 mM sodium phosphate run buffer (pH 7.25) under an electric field strength of E = 357 V/cm. The error bars indicate one SD above and below the average of three separate measurements.

shows the EOF obtained for the clean background electrolyte and single and mixed solutions of BSA and IgG. At pH 7.25, the BSA is negatively charged and the IgG is near its isoelectric point.49 Consequently, as expected, the IgG is observed migrating with the EOF while the BSA lags the EOF peak resulting in effective separation of the two proteins within the capillary. Any person familiar with CE will notice immediately that the peak shapes obtained for both IgG and BSA in these measurements are of poor quality. The reason for these broad and asymmetric peaks is that these CE experiments were not optimized to obtain the typically desired narrow and symmetric peak shapes typical of CE experiments. On the contrary, these measurements were intentionally performed under nonideal conditions in which the capillary was overloaded with a higher than normal level of protein to amplify any effects on the EOF caused by the adsorption of proteins to the capillary walls. The effects of multiple protein injections on the electrokinetic properties of the LUB coatings were probed by performing a series of consecutively run separations of IgG and BSA (see Figure 5b,c). In this series of measurements, a drift to longer migration times with each consecutive separation was observed for the BSA peak. Normally, a drift in the migration time of an analyte in CE indicates an instability in the EOF; that is, that there has been a change in the surface charge. However, we observe no concomitant change in the migration times for either the EOF or IgG peaks over these same series of measurements. Since no significant change in the EOF or IgG peak migration time was observed over the series of measurements, the observed drift in the BSA peak cannot be attributed to changes in the surface charge, which would occur if the BSA and/or IgG was adsorbing to the LUB coating during the experiment. While it remains unclear, the drift of the BSA peak has to be attributed to some unknown attractive (but not adhesive) interaction between the LUB coating and the migrating BSA which inhibits the migration of the BSA through the capillary. In addition, it was observed that if the capillary was allowed to “rest” for a period of time (in this case, overnight) and the measurement was repeated, the BSA peak was found to return to the original migration time measured in the very first measurement (i.e., run 1). Although we do not 1904

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Figure 5. (a) Representative electropherograms obtained following the injection of the neutral marker (acetone) in 15 mM MOPS run buffer (pH 7.25); a 1 mg/mL solution of BSA; a 1 mg/mL solution of IgG; and a mixed solution of 0.5 mg/mL of BSA and 0.5 mg/mL IgG in MOPS buffer in a LUB coated PTFE capillary separated under an electric field strength of E = 294 V/cm. (b) Measured migration times obtained from 11 consecutively run injections of a mixed solution of 0.5 mg/mL of BSA and 0.5 mg/mL of IgG in MOPS buffer performed in the same capillary as the measurements shown in (a). In between runs 1−10, a 1 min rinse of the capillary was performed before immediately initiating the next experimental measurement. In between runs 10−11, there was a latency period of approximately 8 h before rinsing the capillary and initiating run 11. (c) The observed migration times for both the IgG and the BSA plotted as a function of the run cycle number.

Figure 6. Experiments comparing the nonspecific adsorption from a 1 mg/mL solution of Alexa Fluor 488 labeled Chicken antirabbit IgG in MOPS buffer (pH 7.2) in an uncoated and LUB coated PDMS/silica microfluidic electrophoresis chip (see description in Figure 1). (a) The residual fluorescence of nonspecifically adsorbed antirabbit IgG in the uncoated electrophoresis chip at a 2 s exposure time. (b) The residual fluorescence of nonspecifically adsorbed antirabbit IgG in the LUB coated electrophoresis chip at a 2s exposure time. (c) The residual fluorescence of nonspecifically adsorbed antirabbit IgG in the uncoated electrophoresis chip at a 20 ms exposure time. (d) The residual fluorescence of nonspecifically adsorbed antirabbit IgG in the LUB coated electrophoresis chip at a 200 ms exposure time (i.e., at a 10× longer exposure time than the image shown in (c)). (e) Line scans of the fluorescence intensity obtained for the images shown in (a−d) in the positions indicated by the black and white lines in the corresponding images.

background fluorescence observed in the micrograph for the LUB coated microfluidic chip (Figure 6b) is barely discernible over the natural fluorescent “noise” of the PDMS. Indeed, the line trace of the fluorescence intensity for the LUB coated channel (at 2s exposure time) shown in Figure 6e indicates that the residual fluorescence intensity due to nonspecifically bound protein is marginally greater than the level of background noise (see Figure 6e). The 2 s exposure times used to acquire the images in Figure 6a,b however results in overexposure that is atypical of acquisition times utilized for fluorescence detection in microfluidic systems. A 20 ms exposure time is more typical and at this exposure time it is clear that the intensity of the residual background fluorescence due to nonspecifically bound antirabbit IgG in the uncoated PDMS channel is still significant (see Figure 6c) when compared to the background fluorescent “noise” as seen in Figure 6e. In contrast, even at 200 ms (i.e., at a 10 times longer exposure time), no apparent background fluorescence is seen in the micrograph of the LUB coated PDMS microfluidic chip (see Figure 6d) nor the line trace shown in Figure 6e which indicates that the intensity of the background fluorescence due to nonspecifically adsorbed antirabbit IgG has been reduced below the level of noise. Lubricin coatings in microfluidic devices combine excellent antiadhesive properties with a large and stable surface charge

know for certain, the apparent time dependent nature of this interaction suggests that it may be flow or shear dependent. 3.3. Use of LUB Coatings in Microfluidics. One of the areas where multifunctional electrokinetic and antiadhesive LUB coatings have significant potential is in the area of microfluidics. Achieving maximum signal-to-noise ratios and maintaining consistent and reliable electrokinetic control of fluid flows in microfluidic channels requires elimination of the unwanted adsorption of proteins (or other analytes) to the channel surfaces. The ability of LUB to virtually eliminate the nonspecific adsorption of protein in a PDMS/silica microfluidic electrophoretic chip is illustrated in Figure 6. Using a syringe, a 2 mg/ mL solution of fluorescently labeled antirabbit IgG (10 mM MOPS; pH 7.25) was injected into both an uncoated and LUB coated PDMS microfluidic electrophoretic chips. After a period of 5 min, both uncoated and LUB coated chips were rinsed out with clean MOPS buffer and imaged using a fluorescence microscope (see Figures 6a−d). From these fluorescent micrographs, it is apparent that, at a 2s exposure time, the residual background fluorescence signal from nonspecifically adsorbed antirabbit IgG is substantial in the uncoated PDMS chip (Figure 6a). A line scan of the fluorescence intensity across the channel indicates a very strong background fluorescence signal as shown in Figure 6e. At the same 2 s exposure time, the 1905

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Figure 7. Electrokinetic loading, injection, and fluorescence detection of a 0.5 mg/mL solution of Alexa Fluor 488 labeled Chicken antirabbit IgG in MOPS buffer (pH 7.2) performed in a LUB coated microfluidic PDMS/silica electrophoresis chip using the applied electrical potentials described in Figure 1. (a) White field optical image showing the injection point and configuration of the individual channels in the cross-channel chip. (b) Fluorescence image showing the loading of the antirabbit IgG with white arrows indicating the directions of fluid flow in chip channels. (c) Fluorescence image showing the moment of injection (i.e., the change in voltages) of the antirabit IgG with white arrows indicating the directions of fluid flow in chip channels. (d) Fluorescence image of the same region of the chip shown in (a−c) approximately 10 s after the injection shown in (c) indicating no discernible residual fluorescence due to nonspecifically adsorbed antirabbit IgG. (e) Plot of the fluorescence intensity vs the migration time indicating the detection of the injected antirabbit protein as it migrated down the detector channel. The data shown in (e) was acquired by a PMT detector located 2 cm away from the injection point from a different injection to that shown in the sequence (b−d).

that makes it possible to drive the fluid flow in-chip using electroosmosis at relatively low electric field strengths. Figure 7 shows the electrokinetic injection (Figure 7a−d) and fluorescent detection (Figure 7e) of a 0.5 mg/mL solution of fluorescently labeled antirabbit IgG within a PDMS/silica electrophoretic chip. As described in detail in the methods section, through appropriate manipulation of the electric field within the various channels, a steady flow of antirabbit IgG from the loading channel inlet to the loading channel outlet was achieved and confined in the cross-channel region by convergent flows from the buffer and detector channels (Figure 7b). Changing the electric field clears the antirabbit IgG from the channels with flow from the buffer channel resulting in a plug of antirabbit IgG migrating down the detector channel (Figure 7c). Only a few seconds after the electric field change, within the region of the injection zone, all the antirabbit IgG has been removed from the channels with effectively zero residual background fluorescence (Figure 7d). The plug of antirabbit IgG sent down the detector channel, visible in Figure 7c, was observed by a detector located 2 cm away from the injection zone approximately 45 s after the injection. The change in the fluorescence intensity recorded as the plug of IgG passed by the detector is shown in Figure 7e and resulted in a symmetric and reasonably sharp peak having a full width halfmaximum of 0.32 min. We note that the images and data shown in Figure 7 were taken from the fifth consecutive electrostatic injection performed on this particular chip demonstrating that the low protein adhesion and background fluorescence is long lasting. In between different electrostatic injections of IgG, the microfluidic device was flushed out with clean buffer.

were found to be reasonably high and relatively stable over the range of run buffer pHs tested. It was also found that exposing the LUB coatings to high concentrations of proteins did not lead to significant nonspecific adsorption nor a change in the observed electrokinetics that can be attributed to adsorption. Finally we demonstrated that LUB coatings can be utilized effectively in a microfluidic device to prevent unwanted adsorption of proteins enabling their electrophoretic separation. We demonstrate that LUB coatings exhibit the duel functionality of being an excellent antiadhesive for the prevention of protein fouling while also providing a stable, uniform, and well-defined electrostatic charge which enables the flow of fluid and migration of analytes to be controlled effectively using electric fields and electrokinetic phenomena. The electroosmotic flow and related zeta potentials of the LUB coated silica and PTFE capillaries were found to be reasonably high and relatively unchanging over the range of run buffer pHs tested. It was also found that exposing the LUB coatings to high concentrations of proteins did not lead to significant nonspecific adsorption nor a change in the surface charge that can be attributed to adsorption. Finally we demonstrated that LUB coatings can be utilized effectively in a microfluidic system to prevent unwanted adsorption of proteins to the device surfaces while simultaneously permitting the migration and fluid flow within the device to be controlled electrokinetically.

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AUTHOR INFORMATION

Corresponding Author

*E-mail: [email protected]. Notes

The authors declare no competing financial interest.

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4. CONCLUSIONS We demonstrate that LUB coatings exhibit the dual functionality of being an excellent antiadhesive for the prevention of protein fouling while also providing a stable, uniform, and well-defined surface charge that is important for electrophoretic separations. The EOF mobilities and related zeta potentials of the LUB coated silica and PTFE capillaries

ACKNOWLEDGMENTS This work was funded by the Australian Research Council through a Discovery Early Career Research Award (Project No. DE130101458). G.W.G. would like to thank the ARC for their support with this award. G.W.G. would also like to thank Dr. Noelene Quinsey and the Monash Protein Production Unit, 1906

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Clayton, Monash University, Victoria for their assistance with the purification of the native bovine lubricin protein. G.W.G. also thanks M. C. Herd (Corio, VIC Australia), Prof. Richard Fry (University of Melbourne), and Dr. Raymond Rodgers (University of Adelaide) for their assistance in sourcing synovial fluid. Finally, the authors would also like to acknowledge the ARC Center for Excellence in Electromaterials Science for their support.

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