Device fabrication, optimization, and automation

BY :Rui Yang, Jayson V. Pagaduan, Ming Yu & Adam T. Woolley

Abstract Microfluidic systems with monolithic columns have been developed for preconcentration and on-chip labeling of model proteins. Monoliths were prepared in microchannels by photopolymerization, and their properties were optimized by varying the composition and concentration of the monomers to improve flow and extraction. On-chip labeling of proteins was achieved by driving solutions through the monolith by use of voltage then incubating fluorescent dye with protein retained on the monolith. Subsequently, the labeled proteins were eluted, by applying voltages to reservoirs on the microdevice, and then detected, by monitoring laserinduced fluorescence. Monoliths prepared from octyl methacrylate combine the best protein retention with the possibility of separate elution of unattached fluorescent label with 50 % acetonitrile. Finally, automated on-chip extraction and fluorescence labeling of a model protein were successfully demonstrated. This method involves facile sample pretreatment, and therefore has potential for production of integrated bioanalysis microchips. Keywords Biomarkers . Chromatography . Microfluidics . Monoliths . Solid-phase extraction Introduction Detection of biomarkers is of great importance for diagnosis, monitoring and treatment of diseases, for example different types of cancer [1–6] and pregnancy complications [7, 8]. Significant research effort has been devoted to developing efficient and effective methods for detection of diseasespecific biomarkers. Despite impressive progress achieved to date, development of effective and scalable analytical techniques for protein biomarkers, pathogenic bacteria, and viruses remains a significant challenge [9]. Modern bioanalytical techniques, for example liquid chromatography coupled with mass spectrometry, can be used to identify biomarkers, but cost and scalability are two drawbacks [10]. Enzyme-linked immunosorbent assay (ELISA) is another powerful means of measurement of biomarkers, but ELISA is most effective for batches of similar analyses in multiwell plates [11]. Microfluidics, especially integrated devices, have emerged as promising techniques because they consume small volumes of fluid and are rapid, fabrication cost is low, and they are portable [12–15]. Furthermore, miniaturization of traditional analyses can enable automation and parallelization of tests with reduced amounts of sample in less time [16, 17]. Finally, human error and contamination can potentially be reduced by integration of sample preparation, separation, detection, and data processing on a single microfluidic device [18]. One of the most difficult steps in microfluidic integration is sample preparation [19]. Among the different sample-preparation techniques, solid-phase extraction (SPE) is widely used for preconcentration and purification [20]. Affinity and reversed-phase are two common column types in SPE. The former has been used to extract or enrich bio-recognizable substances, for example cancer biomarkers or PCR products [21–23], whereas the latter is more suitable for purification of non-polar to moderately polar compounds [24]. In conventional packed particle reversed-phase columns, the supports can be fabricated in a variety of ways from a variety of materials with different useful functionality. As a Published in the topical collection celebrating ABCs 13th Anniversary. R. Yang : J. V. Pagaduan : M. Yu : A. T. Woolley (*) Department of Chemistry and Biochemistry, Brigham Young University, Provo, UT 84602, USA e-mail: atw@byu.edu Anal Bioanal Chem (2015) 407:737–747 DOI 10.1007/s00216-014-7988-0 Author's personal copy result, they are widely used in microfluidics, as summarized in recent reviews [25, 26]. Several methods have been used to trap particles within microfluidic devices, including frits [27], weirs [28], pillars [29], and column height constraints [30]. In addition, fritless designs have been developed for packing particles [31, 32]. However, packed columns have limitations associated with packing difficulties and complicated design; this increases complexity when they are integrated into microchips. Monolithic columns are increasingly used in microfluidics because of their ease of preparation, lack of retaining structures, and adjustable porosity and surface area [33]. The first use of a monolith in a microfluidic system for SPE was reported by Svec et al. [34]; enrichment of Phe–Gly–Phe–Gly up to 1000-fold was reported. Similarly, Tan et al. [35] developed a device with multiple hydrophobic monoliths fabricated within channels in a cyclic olefin copolymer (COC) chip, in which imipramine was extracted from human urine. Shediac et al. [36] made an acrylate-based porous polymer monolith as a stationary phase for microchip electrochromatography of amino acids and peptides. Rohr et al. [37] used a monolith to assist with the mixing of two fluids, and Yu et al. [38] formed a monolith from a thermally responsive monomer, which then acted as a valve when the temperature was varied. In many of these applications, the monoliths were used for a single function rather than to create a fully integrated analytical system. Importantly, there is a need for integrated microfluidic systems with monoliths for sample preparation. Recently, Nge et al. [39] reported use of a monolith prepared from butyl methacrylate for SPE and on-chip labeling. However, pretreatment of the monolith by rinsing with 30 % acetonitrile was necessary to obtain the best retention. In addition, monolith formulation was not fully optimized for flow and retention characteristics. In this paper we report the fabrication and optimization of microfluidic columns for SPE and on-chip labeling. Monoliths were prepared by in-situ photopolymerization in microchannels. Different types and concentrations of monomers were evaluated, and retention of model proteins was observed without the need for column preconditioning. On-chip labeling of model proteins was achieved by driving solutions through the monolith by use of voltage, and incubating fluorescent dye with the protein retained in the monolith. The labeled protein was subsequently eluted, by applying voltages to reservoirs on the microdevice to drive eluent through the monolith, and detected by monitoring laser-induced fluorescence. Monoliths prepared from octyl methacrylate resulted in the best combination of protein retention while still enabling unattached fluorescent label to be eluted in a separate fraction by use of 50 % acetonitrile. Finally, we demonstrated automation of on-chip capture, fluorescence labeling, and elution of proteins. Experimental Materials and reagents Cyclic olefin copolymer plates (either 6″×6″, 1 mm thick or 4″×6″, 2 mm thick) were obtained from Zeon Chemicals (Zeonor 1020R, Louisville, KY, USA). Methyl methacrylate (MMA), butyl methacrylate (BMA), octyl methacrylate (OMA), lauryl methacrylate (LMA), 2,2-dimethoxy-2- phenylacetophenone (DMPA), 1-dodecanol, ethylene dimethacrylate (EDMA), and isopropyl alcohol were purchased from Sigma–Aldrich (St Louis, MO, USA). Cyclohexanol and dimethyl sulfoxide (DMSO) were from J. T. Baker (Phillipsburg, NJ, USA). Tween 20 was purchased f r om M alli n c k r o dt B a k e r ( P a ri s, K Y, U S A ). Hydroxypropylcellulose (HPC, 100 kDa average molecular weight) was from Aldrich (Milwaukee, WI, USA). Sodium dodecyl sulfate (SDS) was obtained from Fisher Scientific (Pittsburgh, PA, USA). Bovine serum albumin (BSA) and heat shock protein 90 (HSP90) were purchased from New England Biolabs (Ipswich, MA, USA). BSA was labeled with fluorescein isothiocyanate (FITC), and HSP90 was labeled with Alexa Fluor 488 TFP ester. Both fluorophores were obtained from Invitrogen (Carlsbad, CA, USA). Anhydrous sodium carbonate, sodium bicarbonate, and acetonitrile (ACN) were obtained from EMD Chemicals (Gibbstown, NJ, USA). Bicarbonate buffer solution was prepared by mixing sodium carbonate and sodium bicarbonate with deionized water and diluting to 10 mmol L−1 carbonate, resulting in pH 9.3. Off-chip labeling of HSP90 with Alexa Fluor 488 TFP ester was achieved by use of a process similar to that described by Nge et al. [40]. Briefly, HSP90 solution was prepared in bicarbonate buffer at a concentration of 220 μg mL−1 . Alexa Fluor 488 TFP ester solution (5 μL), concentration of 10 mg mL−1 in DMSO, was added to 250 μL protein solution and incubated in the dark overnight at room temperature. Unconjugated dye was removed from the protein by filtration through an Eppendorf 5418 centrifugal filter. Labeled protein samples were collected and stored in the dark at 4 °C until use. Device fabrication Individual COC plates were obtained by cutting a COC sheet into pieces, each of length 5 cm and width 2.5 cm, with an electric motor saw. Reservoirs were produced by drilling holes in the cover plate before device bonding. The microdevices were fabricated by use of a combination of photolithographic patterning, etching, hot embossing, and thermal bonding, as described by Kelly et al. [41]. Bonding of COC was performed at 110 °C for 24 min. A simple, two-reservoir layout (Fig. 1a) was used for preliminary testing, and a six-reservoir layout was used for automated and integrated SPE and onchip labeling (Fig. 1b). The channels in the design were 738 R. Yang et al. Author's personal copy approximately 50 μm wide and 20 μm deep. Channels were rinsed with isopropyl alcohol before polymerization of the monolith. Monoliths were fabricated by a modification of a method reported elsewhere [39]. Porogens, photoinitiator, and Tween 20 were weighed in accordance with the values listed in Table 1 and mixed with each different monomer (i.e., MMA, BMA, OMA, or LMA). The solution was sonicated until the photoinitiator was completely dissolved, then degassed for 5 min. It was next loaded into the device, and black tape was used as a mask to expose only the desired region of the chip to UV irradiation. Exposure was conducted with a SunRay 400 lamp (Intelligent Dispensing Systems, Encino, CA, USA) at 200 W for 12–15 min. A 2 mm long monolith was formed in each microdevice at the location indicated in Fig. 1. After polymerization, devices were rinsed with isopropyl alcohol. Each device was then washed with deionized water several times and air-dried before characterization and testing. Scanning electron microscopy (SEM) was performed with a Philips XL30 ESEM FEG instrument in low-vacuum mode. A potential of 10–12 V was applied to the surface, depending on the extent to which the monolith charged. The edge that contained the monolith was cut manually by use of a microtome with a glass knife. Once the monolith was exposed, the surface was cleaned by use of adhesive tape to remove debris. The sample was then mounted on aluminium stubs with carbon tape and coated with silver by use of a Polaron sputterer to reduce charging during SEM imaging. The samples were coated under an applied potential of 2.5 kVand a current of 18–20 mA for 3 min. Device operation Before sample loading, monolithic columns were rinsed with 2-propanol several times, to clean the surface, and then bicarbonate buffer was directed into the channel. Next, the stability of the current was examined by applying +600 V to reservoir 2 and grounding reservoir 1 for 1 min; simultaneously, the microdevice was observed by use of an optical microscope to make sure no bubbles were trapped in the microchannel. Retention and elution on monoliths To evaluate the extent to which different samples were retained on the monoliths, fluorescent dyes (FITC and Alexa Fluor 488 TFP ester, each 100 nmol L−1 ) and two labeled proteins (BSA and HSP90, 200 ng mL−1 ) were transferred to reservoir 1 and loaded by applying +400 V to reservoir 2 for 5 min and grounding reservoir 1, as shown in Fig. 1a. Rinsing was conducted by replacing the sample in reservoir 1 with buffers of different ACN concentration (30 % or 50 %) and applying +400 or +600 V to reservoir 2 for 2 min. For elution, the rinse buffer in reservoir 1 was replaced with eluent Fig. 1 Schematic diagrams of designs and photographs of microfluidic devices. (a) A single-channel device with two reservoirs for SPE and on-chip labeling. (b) A six-reservoir device for integrated and automated experiments in which the reservoirs are: 1 and 2, loading buffer; 3, elution buffer; 4, dye; 5, protein; and 6, rinsing buffer. All the channels are of width 50 μm and depth 20 μm Table 1 Monolith composition Monomer Cross-linker Porogen Photoinitiator Surfactant EDMA Cyclohexanol 1-Dodecanol DMPA Tween 20 29.7 % 14.9 % 18.8 % 17.8 % 1.0 % 17.8 % On chip preconcentration and fluorescence labeling of model proteins 739 Author's personal copy consisting of 85 % ACN, 15 % bicarbonate buffer, 0.05 % HPC, and 0.05 % SDS; reservoir 1 was then grounded and + 600 V or +1000 V was applied to reservoir 2. On-chip labeling For on-chip labeling experiments (Fig. 1a), unlabeled protein samples were loaded in the same way as in the retention and elution experiments. Next, reservoir 1 was rinsed and filled with fluorescent dye solution (10 mg mL−1 ) in DMSO. This solution was driven through the column by applying the same voltages as in loading for 10 min, followed by incubation for 10–15 min with the voltage off. Rinsing was performed by replacing the labeling solution in reservoir 1 with buffers of different ACN concentration (30 % or 50 %) and applying the same voltages as in the previous step for 10 min. For elution, the rinse solution in reservoir 1 was replaced with eluent consisting of 85 % ACN and 15 % bicarbonate buffer. During elution, reservoir 1 was grounded while +600 V was applied to reservoir 2 for 10 min. Automated extraction, labeling, and elution For experiments conducted on the integrated microdevices shown in Fig. 1b, platinum wires were inserted into the solution-filled reservoirs to provide electrical contact. Two high-voltage power supplies provided all applied potentials. A custom-designed voltage-switching box controlled by LabView was used to apply potentials to the microchips. Reservoirs 1 and 2 were filled with bicarbonate buffer, and reservoirs 3 to 6 were filled with elution solution (85 % ACN and 15 % bicarbonate buffer), dye, HSP90 (20 nmol L−1 ), and rinsing solution (50 % ACN and 50 % bicarbonate buffer), respectively. The sequence of voltages applied for the different operation steps is shown in Fig. 2. Fluorescence data collection and analysis Retention and elution were monitored via CCD detection, by measuring the background-subtracted fluorescence intensity after rinsing and elution. A Nikon Eclipse TE300 inverted microscope equipped with a CCD camera (Coolsnap HQ; Roper Scientific, Sarasota, FL, USA) was used for imaging. A 488 nm blue laser (JDSU, Shenzhen, China) with a 10× expander was directed to a 10×, 0.45 NA objective on the microscope. For fluorescence monitoring, the detection point was positioned either next to reservoir 2 (Fig. 1a, b), or directly on the monolith. The collected CCD images were analyzed in V++ Precision Digital Imaging software (Auckland, New Zealand). Photomultiplier tube (PMT) detection was also utilized, in which the detection point was positioned next to reservoir 2. Collected fluorescence went through a D600/60 band-pass filter (Chroma, Rockingham, VT, USA) and was detected with a Hamamatsu PMT (HC120-05; Bridgewater, NJ, USA); out-of-focus light was blocked by a 1000 μm diameter pinhole. The PMT voltage output was processed with a preamplifier (SR-560; Stanford Research Systems, Sunnyvale, CA, USA) and an analog-to-digital converter (PCI 6035E; National Instruments, Austin, TX, USA) and was recorded by LabView software running on a Dell computer. Results and discussion Optimization of the monoliths Thermally bonded COC devices with monoliths formed from different monomers were prepared. COC was chosen as the substrate material because of its stability in common organic solvents, for example the ACN used in this study for sample elution. Poly(methyl methacrylate) dissolves in ACN whereas polydimethylsiloxane requires additional surface modification and swells in solvents [42–44]. Additives, for example UV absorbers used to stabilize polymers such as COC, may affect the amount of UV reaching the channel during monolith polymerization; however, we were always able to apply sufficient radiation to the channels to form monoliths in the 12– 15 min reaction time. During monolith polymerization Tween 20 was added as surfactant to increase the through pore size by affecting phase separation by emulsification. The surfactant content selected was less than 30 %, because bubbles were produced in monoliths prepared with higher surfactant content when voltage was applied, which hindered flow of solution in the microchannel [45]. A total porogen content of 55 % was selected, because monolith rigidity was too low if higher porogen content was used, as reported by Pagaduan et al. [45]. In this work, monoliths were prepared from four different monomers (MMA, BMA, OMA, and LMA). Figure 3 shows SEM images of monoliths prepared with the different monomers. For monoliths prepared from MMA (Fig. 3a), evenly packed nodules with diameters of 500–2000 nm were observed. Through pores formed by the voids between these nodules were in the same size range. For monoliths prepared from the other three monomers, nodules with much smaller sizes were observed (Fig. 3b–d), which resulted in more surface area and hence more binding capacity. For BMA monoliths (Fig. 3b), through pores with sizes of several hundreds of nanometers were observed. Uniform material was found only within the central section of the monolith; most of the channel contained discrete porous clusters of different lengths. This is consistent with the observations of Ramsey and Collins [46], which were explained on the basis of localized fluid flow during in-situ photopolymerization. For 740 R. Yang et al. Author's personal copy monoliths prepared from OMA and LMA (Fig. 3c, d), different sizes of through pores formed by agglomerates of nodules with dimensions of ~100 nm were observed, which is favorable, because irregular pores enhance convective transport as liquids flow through the monolith [47]. On application of voltage for rinsing and elution, none of the monoliths moved, in agreement with results reported by Ladner et al. [48] and Nge et al. [39]. Complicated column pretreatments, for example photografting, were therefore avoided [48]. Figure 4 shows the background-subtracted fluorescence signal after both retention and elution of BSA on monoliths prepared from different monomers. We observed that the retention of BSA after rinsing with 50 % ACN increased with carbon chain length for monoliths prepared from MMA, BMA, and OMA, consistent with monomer hydrophobicity. For monoliths prepared from a mixture of MMA and LMA, retention of BSA was comparable with that obtained on monoliths prepared from OMA; this is explained by the combined hydrophobicity of MMA and LMA. For monoliths prepared from a mixture of BMA and LMA, higher retention was observed, because of the greater hydrophobicity of BMA compared with MMA. Fluorescent intensities on MMA, BMA, and OMA monoliths after elution with 85 % ACN were very low (Fig. 4), indicating that the BSA retained by the column was eluted almost completely under these conditions. In contrast, BSA fluorescence on both types of mixed LMA monolith after elution with 85 % ACN was readily detectable (Fig. 4), indicating stronger interaction between BSA and these monoliths. In addition, for LMA mixed monoliths, buffer flow through the column was limited, requiring higher voltage to achieve adequate flow. We note that optimum sample preconcentration in our system consists in high protein retention on the monolith after rinsing with 50 % ACN, followed by complete removal of protein during the 85 % ACN elution step. On the basis of these considerations, we chose monoliths prepared from OMA for subsequent work. Retention results provide further insights into optimization of these monoliths. Figure 5 shows a comparison of elution in 85 % ACN of FITC-labeled BSA from monoliths prepared with 20, 30, and 40 % (w/w) OMA (relative to the total weight of monolith pre-polymer solution). For the monolith prepared with 20 % (w/w) OMA, two overlapping peaks were observed during elution. The first large peak is attributed to unreacted Fig. 2 Schematic diagram of automated device operation: (a) loading, (b) labeling, (c) rinsing, and (d) elution. Reservoirs 1 and 2 were filled with bicarbonate buffer, and reservoirs 3 to 6 were filled with elution solution (85 % ACN and 15 % bicarbonate buffer), fluorescent label, HSP90, and rinsing solution (50 % ACN and 50 % bicarbonate buffer), respectively. (a) For sample loading, reservoirs 1 and 5 were grounded, and +650 V and +200 V were applied to reservoirs 2 and 3, respectively. (b) For sample labeling, reservoirs 1 and 4 were grounded, and +650 V and +200 V were applied to reservoirs 2 and 3, respectively. (c) For rinsing, reservoirs 1 and 6 were grounded, and +650 V and +250 V were applied to reservoirs 2 and 4, respectively. (d) For elution, reservoirs 1 and 3 were grounded, and +650 V and +250 V were applied to reservoirs 2 and 4, respectively. The solution flow directions are indicated by arrows On chip preconcentration and fluorescence labeling of model proteins 741 Author's personal copy fluorescent dye, and the second (smaller) peak to FITClabeled BSA, suggesting that both BSA and FITC were retained on the monolith after the 50 % ACN rinse. For the monolith prepared with 30 % (w/w) OMA, a single peak of BSA was observed, indicating successful retention of BSA with limited retention of fluorescent dye after the 50 % ACN rinse. For the monolith prepared with 40 % (w/w) OMA, no distinct protein or dye peak was observed, which we attribute to stronger interaction between protein and monolith with increased monomer content, such that essentially no protein was eluted even with 85 % ACN. On the basis of these experiments we chose an OMA monomer concentration of 30 % (w/w) as best suited to protein retention and elution. Retention and elution with OMA monoliths Figure 6 shows the background-subtracted fluorescence signal, indicative of retention of fluorescent dyes and labeled proteins on OMA monoliths after 50 % ACN rinsing. Retention of the fluorescent dyes (Alexa Fluor 488 TFP ester and FITC) on the OMA monolith was less than retention of Fig. 3 SEM images of monoliths prepared from (a) MMA (scale bar for inset: 2 μm), (b) BMA, (c) OMA, and (d) LMA Fig. 4 Background-subtracted fluorescent signal obtained from 200 ng mL−1 FITC-labeled BSA on monoliths prepared from different monomers, after rinsing with 50 % ACN (solid bars) and after eluting with 85 % ACN (striped bars) Fig. 5 Elution of FITC-labeled BSA with 85 % ACN from monoliths prepared with different concentrations of OMA. From top to bottom: 20, 30, and 40 % (w/w) OMA in the polymerization mixture. Chromatograms are offset vertically 742 R. Yang et al. Author's personal copy proteins (HSP90 and BSA), which is consistent with results reported by Nge et al. [39]. Previous studies have shown that preconditioning of monolithic columns affects the retention of amino acids and proteins [49]. Nge et al. [39] showed that protein retention increased when a BMA monolith was rinsed with 30 % ACN just before sample loading. This pre-rinse helped remove impurities and activate and/or hydrate the monolith surface to provide adequate contact with the liquid sample [50]. It is apparent from Fig. 6 that good retention of proteins on OMA monoliths was achieved without any preconditioning with ACN; this might be explained by the different hydrophobicity of the BMA and OMA monoliths. Figure 7 shows 85 % ACN elution profiles of labeled proteins and their corresponding fluorescent dyes that were retained on an OMA monolith after rinsing with 50 % ACN. Elution of HSP90 and Alexa Fluor 488 TFP ester (Fig. 7a) was apparent from a large peak of HSP90 at approximately 20 s and a small peak for Alexa Fluor 488 TFP ester at approximately 5 s. The labeled HSP90 was retained on the monolith after rinsing with 50 % ACN but was eluted with 85 % ACN, whereas most of the Alexa Fluor 488 TFP ester was rinsed off with 50 % ACN, consistent with their retention in Fig. 6. In a different experiment performed under the same conditions and after rinsing with 50 % ACN (Fig. 7b), a large peak of labeled BSA was eluted after approximately 20 s by 85 % ACN, and small peaks corresponding to FITC were observed at approximately 12 s on elution with 85 % ACN, again confirming successful retention and elution of protein separate from fluorescent label with an OMA monolith. Off and on-chip labeling of HSP90 with Alexa Fluor 488 TFP ester Figure 8 shows elution profiles for HSP90 labeled off and onchip with Alexa Fluor 488 TFP ester. The elution of HSP90 labeled on-chip with Alexa Fluor TFP 488 ester was similar to that for the protein labeled off-chip. Protein peaks in both Fig. 6 Background-subtracted fluorescence from retention of dyes and proteins on an OMA monolith after rinsing with 50 % ACN Fig. 7 Elution profiles in 85 % ACN of fluorescent dyes and labeled proteins from OMA monolithic columns. (a) HSP90 (top) and Alexa Fluor 488 TFP ester (bottom); (b) BSA (top) and FITC (bottom). Traces are offset vertically Fig. 8 Elution profiles in 85 % ACN for HSP90 labeled on-chip (top) and off-chip (bottom). Traces are offset vertically On chip preconcentration and fluorescence labeling of model proteins 743 Author's personal copy samples appeared at approximately the same time (~25 s). A small peak at ~8 s was obtained from the on-chip labeled sample; this was attributed to unconjugated fluorescent dye, because of the shorter incubation time for on-chip labeling (15 min versus overnight for off-chip labeling). Longer protein loading times resulted in broader eluted peaks; in addition, longer labeling times required us to use longer rinse times to adequately remove the unattached fluorophore. Minor variations in elution times of peaks occur because experiments were performed with different devices. Although laminar flow in microfluidic channels generally limits mixing of fluids, in our devices use of a monolith (with tortuous flow paths) for retention and labeling facilitates mixing [37] and thus reaction of fluorophore and protein. The results in Fig. 8 show that on-chip labeling can be integrated with automated SPE in a single microfluidic device. Automated extraction, labeling, and elution To test the feasibility of automated and integrated on-chip SPE and fluorescence labeling, a six-reservoir microchip with an OMA monolith in the microchannel (Fig. 1b) was used. Automated loading, retention, rinsing, and elution of 10 mg mL−1 Alexa Fluor 488 TFP ester by itself, and onchip HSP90 loading, retention, fluorescent labeling with Alexa Fluor 488 TFP ester, rinsing, and elution were conducted according to the procedures outlined in Fig. 2. As shown in Fig. 9a, for the Alexa Fluor 488 TFP ester solution, a single peak at ~17 s was observed in the rinsing step with 50 % ACN and a small peak was observed at ~5 s during elution with 85 % ACN, indicating that nearly all of the dye was eluted from the monolith during rinsing. For on-chip labeling of HSP90 (Fig. 9b), a peak at ~15 s was observed in the 50 % ACN rinse step, similar to that observed in Fig. 9a when Alexa Fluor 488 TFP ester was loaded. A minor peak at ~28 s may indicate a small amount of labeled protein being eluted during the rinsing step. During 85 % ACN elution of the on-chip labeled HSP90 (Fig. 9b), a single peak at ~24 s was observed, indicating that HSP90 was successfully retained, labeled, and then eluted automatically with the microfluidic system. Conclusions Reversed-phase, polymeric monoliths in cyclic olefin copolymer microfluidic devices were prepared and optimized. In addition, a model protein (HSP90) was loaded, retained, and fluorescently labeled on-chip; unreacted dye was then automatically eluted separately from the labeled protein. The combination of SPE and on-chip labeling could potentially address important sample-preparation needs, for example preconcentration and pretreatment. The ease of monolith preparation and rapid on-chip labeling could also reduce analysis time and effort compared with other techniques. In addition, this approach could be further integrated with other samplepreparation and separation techniques to achieve enhanced specificity for more complicated bioanalysis. In these experiments we were able to demonstrate proof of concept of SPE and labeling using polymeric monoliths; however, quantification of protein biomarkers will require more work. Several characteristics of the device can be further modified to achieve better quantification. For example, the ratio of monomer to porogen can be adjusted to change the porosity of the column, which affects the surface area, flow rate, and the resulting retention and elution. In addition, experimental conditions, for example the maximum voltage that can be applied without solvent evaporation as a result of Joule heating, are also affected by surface area and porosity. Moreover, column length can be altered to vary loading capacity. After this optimization, it should be possible for quantitative experiments to be conducted, and corresponding calibration methods to be established. Importantly, the monoliths reported in this work have potential to be integrated with upstream immunoaffinity extraction and downstream electrophoresis separation. We have previously demonstrated integration of immunoaffinity extraction and electrophoresis separation for cancer-relevant Fig. 9 Rinsing (with 50 % ACN) and elution (with 85 % ACN) of (a) 10 mg mL−1 Alexa Fluor 488 TFP ester and (b) on-chip-labeled HSP90 in an integrated and automated microdevice. Traces are offset vertically 744 R. Yang et al. Author's personal copy proteins in blood serum [21, 51]. Therefore, in future studies biofluids could be loaded in a device and first passed through an affinity column in which target biomarkers would be extracted via antibody–antigen interaction. Subsequently, the extracted biomarkers could be released and passed through a monolithic column similar to those optimized in this work for preconcentration and fluorescence labeling. 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Proc SPIE 8031 (Pt. 2, Micro- and Nanotechnology Sensors, Systems, and Applications III):80311 V/80311–80311 V/80317 46. Ramsey JD, Collins GE (2005) Integrated microfluidic device for solid-phase extraction coupled to micellar electrokinetic chromatography separation. Anal Chem 77(20):6664–6670 47. Svec F (2004) Organic polymer monoliths as stationary phases for capillary HPLC. J Sep Sci 27(17–18):1419–1430 48. Ladner Y, Cretier G, Faure K (2010) Electrochromatography in cyclic olefin copolymer microchips: A step towards field portable analysis. J Chromatogr A 1217(51):8001–8008 49. Augustin V, Jardy A, Gareil P, Hennion M-C (2006) In situ synthesis of monolithic stationary phases for electrochromatographic separations: Study of polymerization conditions. J Chromatogr A 1119(1– 2):80–87 50. Marchiarullo DJ (2009) Development of microfluidic technologies for on-site clinical and forensic analysis: extraction, amplification, separation, and detection. Dissertation, University of Virginia 51. Yang W, Sun X, Wang H-Y, Woolley AT (2009) Integrated microfluidic device for serum biomarker quantitation using either standard addition or a calibration curve. Anal Chem 81(19):8230–8235 Rui Yang received a B.Sc. degree in Applied Chemistry from Beijing Institute of Technology, and a M.Sc. in Analytical Chemistry from Brigham Young University. Her master’s research focused on the fabrication and optimization of polymer microfluidic devices for bioanalysis Jayson V. Pagaduan is a Biochemistry Ph.D. candidate in the Department of Chemistry and Biochemistry at Brigham Young University. He is working on microfluidics with Professor Adam Woolley to develop point-ofcare diagnostics for different diseases 746 R. Yang et al. Author's personal copy Ming Yu is an analytical chemist born in China who graduated with a Ph.D. from Colorado State University. Her research interest lies in optimization of miniaturized biosensors. Dr Yu is now teaching at Utah Valley University Adam T. Woolley is Professor and Associate Chair in the Department of Chemistry and Biochemistry at Brigham Young University in Provo, Utah, USA. His research focuses on three general topics: the development of novel and sophisticated integrated microfluidic systems for preterm birth biomarker quantification, the design of simple miniaturized biomolecular assays, and biotemplated fabrication for nanoelectronic systems



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