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Solvent immersion imprint lithography† Cite this: Lab Chip, 2014, 14, 2072
A. E. Vasdekis,*a M. J. Wilkins,‡b J. W. Grate,c R. T. Kelly,a A. E. Konopka,b S. S. Xantheasc and T.-M. Changd We present Solvent Immersion Imprint Lithography (SIIL), a technique for polymer functionalization and microsystem prototyping. SIIL is based on polymer immersion in commonly available solvents. This was experimentally and computationally analyzed, uniquely enabling two practical aspects. The first is imprinting and bonding deep features that span the 1 to 100 μm range, which are unattainable with existing solvent-based methods. The second is a functionalization scheme characterized by a well-controlled, 3D
Received 20th February 2014, Accepted 8th April 2014 DOI: 10.1039/c4lc00226a www.rsc.org/loc
distribution of chemical moieties. SIIL is validated by developing microfluidics with embedded 3D oxygen sensors and microbioreactors for quantitative metabolic studies of a thermophile anaerobe microbial culture. Polystyrene (PS) was employed in the aforementioned applications; however all soluble polymers – including inorganic ones – can be employed with SIIL under no instrumentation requirements and typical processing times of less than two minutes.
Introduction Polydimethilsiloxane (PDMS) has become the workhorse material in microfluidics and optofluidics due to its low cost and simple processing.1–5 However, PDMS has several recognized drawbacks, such as limited compatibility with organic solvents, and high gas permeability and resulting enhanced buffer evaporation rates, especially at elevated temperatures.6 Thermoplastic polymers,7,8 as well as emerging inorganic ones,9,10 do not have these shortcomings. As a result, certain microsystem applications have been revolutionized, such as cell culture experiments,8 complex flow generation,11 nanofluidics,12 chemical synthesis13 and photonics.14 However, imprinting in thermoplastic or inorganic polymers requires substantially more complex processing and dedicated instrumentation, such as hot presses, while bonding relies mostly on thermal means that suffers from reduced success rate and shape deformations.15 These presently make inorganic and thermoplastic polymers less attractive and accessible than PDMS. Another aspect of polymer microsystems that has received little attention is their chemical functionalization. This
a Environmental Molecular Sciences Laboratory, Pacific Northwest National Laboratory, PO Box 999, Richland, WA, 99352, USA. E-mail: [email protected] b Biological Sciences Division, Pacific Northwest National Laboratory, PO Box 999, Richland, WA, 99352, USA c Physical Sciences Division, Pacific Northwest National Laboratory, PO Box 999, Richland, WA, 99352, USA d University of Wisconsin-Parkside, Box 2000, Kenosha, Wisconsin 53141, USA † Electronic supplementary information (ESI) available. See DOI: 10.1039/ c4lc00226a ‡ Present address: School of Earth Sciences and Department of Microbiology, The Ohio State University, Columbus OH, 43210, USA.
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generally requires expensive and dedicated instruments, thus hindering integrative biology and energy microsystem applications (e.g. flow through catalysis).16 Bulk-type functionalization is also desirable in optical applications. This is possible via the microfluidic integration of thin films doped with the chemical entity of interest.17,18 In this case, however, additional processing is essential (e.g. spin coating, patterning etc.), while a uniform dopant distribution is usually obtained with no ability to control it. Controlling the dopant distribution would be of substantial importance in designing novel microsystems, for example sensors with engineered sensitivity and dynamic range. With the exception of functionalization, the imprinting and bonding complexity in thermoplastic or inorganic polymers have been partially addressed by a variety of recent benchtop processes that minimize the need for instrumentation, involving polysilsesquioxane,9 as well as cyclic olefin copolymers (COC) and poly(methyl methacrylate) (PMMA).19–21 Polystyrene (PS) is another low-cost thermoplastic polymer, where costeffective imprinting approaches involve Shrinky-Dink based methods, or soft-lithography using PS solutions.7,22 However, despite such substantial initial progress, rapid and costeffective imprinting and bonding, as well as novel and enabling functionalization strategies, still remain an important goal. Clearly, progress in this direction would further enhance the use of polymer microsystems in chemical, biological and optical applications. In order to bridge the gap between the simplicity of PDMS processing and the stability of thermoplastic and inorganic polymers, as well as address the challenge of polymer functionalization, we introduce Solvent Immersion Imprint Lithography (SIIL). SIIL is based on commonly available solvents and enables complete microsystem prototyping,
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Fig. 1 (a) Schematic illustration of the SIIL protocol. (b) A 4 cm long microchannel imprinted in PS using a 40 s long immersion in acetone; the microchannel dimensions measured using a profilometer at its two far ends were 100 μm wide and 35 μm deep, shown in the inset. (c) A statistical analysis of 11 samples like the one shown in (b) determining the depth and width variation between different imprinting runs. (d) Comparison between imprinted microchannels using existing solvent-based methods (red profile) and SIIL (blue profile); for this experiment, the same imprinting conditions were employed, namely a 20" imprinting duration, acetone as a solvent, a 5 μm deep PDMS stamp and the same exerted force enabled by a 500 g weight.
including functionalization, imprinting and bonding in a single processing step (Fig. 1a). This involves immersing the polymer in a solvent, during which the solvent penetrates into the polymer forming a surface ‘gel’ layer. This process (potentially a case II sorption23) is SIIL's basic mechanism: the softened surface layer can be readily functionalized with moieties such as chemosensors, imprinted using a mold, as well as bonded to a substrate. Large area imprinting is possible with SIIL, such as the representative PS microchannel shown in Fig. 1b (also in Fig. S1†), which is approximately 4 cm long (x-axis), 100 μm wide (y-axis) and 35 μm deep (z-axis). Such large area patterning exhibited high quality with minimal variation across the entire imprinted surface as confirmed by the profilometer measurements (inset of Fig. 1b). Additionally, the statistical analysis shown in Fig. 1c revealed minimal variation in width and depth between 11 different imprinting runs with 2.9 μm (2.6%) and 1. 2 μm (3%) respectively. SIIL expands upon previously reported solvent based nanoimprinting methods, such as the Solvent Assisted Micromolding (SAMIM), the first technique to be developed to this end.24 In SAMIM, the solvent coats the PDMS stamp, which is subsequently pressed against a thin polymer film causing it to partially dissolve and conform to the stamp's pattern.24–26 Solvent vapors have also been used to the same end,
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dissolving the polymer by first propagating through the pores of the PDMS stamp.27–29 Despite the wide success of solventassisted nano-imprinting, solvent based microfluidic prototyping using solvents has received little attention due to limitations in obtaining deep features that span the 1 μm to 100 μm range. The reason for this is that by using a solvent coated stamp, the solvent contacts the polymer only upon imprinting, which leads to solvent reflow and trapping at the edges of the stamp upon contact between the polymer and the stamp and solvent. This phenomenon is absent in thin feature imprinting, however, it becomes non-trivial for deeper features as shown in Fig. 1d (also Fig. S2†). On the contrary, SIIL overcomes these shortcomings because the polymer immersion into the solvent provides better control over the formation of the surface gel, a critical aspect in imprinting deep features (Fig. 1d and S2†). Provided that it is soluble, any kind of polymer is compatible with SIIL, including inorganic ones.9 In the following sections, we present our experimental and computational investigations of the underlying polymer–solvent interactions, specifically in relation to functionalization and deep feature imprinting. PS microfluidics, cell growth microbioreactors and microstructured oxygen sensors are demonstrated by way of example here, with typical processing times of less than two minutes.
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Materials and methods
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Materials The polymers used in the experiments, namely PS and PMMA were supplied by GoodFellow (USA) at various thicknesses ranging from 1.5 mm to 0.25 mm (Product IDs: ST313120, ST311250, 137-745-63). All organic solvents were supplied by Sigma Aldrich (USA), while purified deionized water was employed where mentioned. The oxygen sensing chromophore (PtT975) employed in the optofluidic experiment was supplied by Frontier Scientific (USA) at a concentration of 0.5 mg ml−1. The stamps were made by conventional cast-moulding lithography in PDMS. The PDMS (Sylgard 184, Dow Corning, USA) was mixed with the catalyst at 1 : 10 ratio, degassed for approximately 1 h, baked at 70 °C for 1 h, removed from the hard SU8 mask and baked for an additional 1 h. The hard SU8 mask was fabricated with conventional contact mode optical lithography using the MicroChem formulations SU8-2000. The photoresist was spin coated onto Si 4′′ wafers at different speeds to control its thickness. For confocal imaging, two different chromophores were used depending on the solvent, both supplied by Sigma Aldrich; fluorescein was mixed with acetone and Nile Red with acetone and chloroform at approximately 1 mM concentration. Imaging The fluorescent imaging of the gel layer was performed in an inverted microscope (Leica DMI6000), coupled with a spinning disk confocal system (Yokogawa, CSU10). A 20× objective was used (20×/.7 NA Plan Apo DIC Optics Inclusive) and the z-scanning was performed at a 2 μm step size and the gel film thickness was estimated by intensity thresholding. For the latter the background level was estimated by the ratio of the standard deviation over the mean. Oxygen sensing was performed in a fluorescence lifetime imaging setup (LI2CAM-P, Lambert Instruments, Netherlands), integrated with the aforementioned Leica inverted microscope. For this measurement, an LED with an emission spectrum centered at 399 nm was employed modulated at 5 kHz. Microfluidics In the microfluidic experiments, a peristaltic pump, capillary forces or manual means were employed to inject fluids. The microbial cells were manually injected in the micromodels using a syringe. The filling of the PS microchannels for fluorescent imaging (see e.g. Fig. 3) was performed by capillary action, using a fluorescein ethanol solution. Flow rate control, where stated, was provided by a precision milliGAT pump (Global FIA, USA), connected to the microfluidics with nanoport fittings through PTFE tubing. Cell cultures Clostridium thermocellum ATCC 27405 cultures were grown in complex GS-2 media,30 at 60 °C under strict anaerobic conditions. In these media, cellobiose was the main carbon source.
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Batch growth rate experiments were performed in 50 mL volumes, under shaking conditions. Cell densities were measured by removing 1 mL aliquots of culture, and measuring absorbance at 600 nm using a BioRad spectrophotometer (Bio Rad, Hercules, CA). The aerobic bacterial strains, Shewanella and Flavobacteria, were grown in a bicarbonatebuffered media containing glucose as the sole carbon source.31 These were a spontaneous variant of Shewanella oneidensis MR-1 engineered to express a green-fluorescent protein (GFP), and Flavobacterium johnsoniae UW101 expressing the mStrawberry fluorescent protein.32 Batch cultures were incubated under aerobic conditions at 30 °C. Prior to growth experiments, the biomass was grown overnight and subsequently transferred at 1% inoculum into fresh media. Computational Classical molecular dynamics (MD) simulations were used to study the dissolution processes between solvents (acetone, chloroform) and polystyrene with four monomer units. The energy of the system was described by a sum of Lennard-Jones, Coulombic, and intramolecular (bond, angle, and torsion) interactions. The acetone, chloroform, and polystyrene potentials were based on an all-atom framework, with fixed partial charges and Lennard-Jones parameters assigned to each atom. The potential parameters for acetone and polystyrene were taken from the OPLS-AA force field,33 whereas for chloroform a potential modified from a previous model was used.34 The cross Lennard-Jones interaction terms between unlike atoms are obtained using the Lorentz–Berthelot combining rules. All MD simulations were carried out with the AMBER 9 package. More information is available in the ESI† section.
Results Polymer–solvent interactions SIIL is based on polymer immersion in appropriate solvents. During this step, the solvent diffuses into the polymer,35,36 the initial stages of which were modeled using molecular dynamics (MD) simulations. The inset of Fig. 2a illustrates individual acetone molecules diffusing into PS (also in Fig. S3†). To further analyze solvent diffusion, PS films were immersed into fluorescent dye solutions. In this way, the solvent transports the dye into the polymer, hence enabling the selective staining of the solvent's diffusion path. While several methods exist in the literature to investigate polymer– solvent interaction, we employed fluorescence microscopy37 in order to combine the diffusion with the imprinting and functionalization investigations. Once dried, the films were imaged by fluorescence confocal microscopy along the film depth (z-axis) (see methods and Fig. S4†). Similar procedures have been previously reported in interfacial polymer dissolution studies;38 however, in the present experiments, confocal imaging was chosen to characterize substrates that can be specifically used in microfluidic applications. Typical results are shown in the histogram of Fig. 2a for different immersion durations, illustrating that the solvent
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Fig. 2 (a) The gel and functionalization thickness dependence on the immersion duration for the PS–acetone and PS–chloroform systems; inset plots snapshots of molecular dynamics simulations of acetone (purple)/polystyrene (cyan) interface upon contact and following a 100 ns long interaction. (b) A reconstructed confocal image across the z-axis of an imprinted PS microchannel based on an 8 s immersion in a fluorescent acetone solution; the thinner fluorescent region corresponds to the imprinted area, while the blue and red lines illustrate the bottom surface of the gel and the imprinted feature respectively. Inset plots the three-dimensional image of the same film, with the blue line denoting the location of the z-scan. (c) The depth of the imprinted features in PS following immersion in different solvents and for different durations using a 50 μm deep PDMS stamp; the same pressure was exerted in all experiments by positioning a light weight (500 g) on top of the polymer slabs. The inset plots the profile of imprinted features in PS using pure acetone and 7 μm and 12 μm deep stamps.
diffusion depth increases for longer immersions and eventually saturates, as previously reported.37 The solvent diffusion depth was also found to depend on the solvent type and the underlying polymer–solvent molecular interactions. These interactions are frequently described by the Hildebrand solubility parameter, which states that the closer the solubility parameters between two substances are, the easier it is to mix them.38 The Hildebrandt parameters, δ, for chloroform and acetone are 18.7 and 20.4 (J cm−3)1/2, respectively, while the one for PS is 18.7 (J cm−3)1/2.38 The solvent diffusion depth was substantially thicker in chloroform than acetone for similar immersion durations (Fig. 2a). This occurs because of chloroform's higher thermodynamic compatibility with PS, as evidenced by their matching Hildebrandt parameters.
Solvent immersion functionalization and imprinting Solvent diffusion results in a decreased viscosity at the polymer interface. This forms a surface gel without however modifying the polymer chains and thus manifesting the mechanisms' mesoscopic nature.39 To explore the formation of a surface gel layer, PS films were immersed in a fluorescent solution and subsequently pressed against a patterned PDMS slab, as described in Fig. 1a. During this process, the PDMS pattern is transferred to the fluorescent gel of the immersed PS, while the porosity of PDMS enables the solvent evaporation. The
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latter is important as it enables rapid solvent removal from the polymer while still in contact with the PDMS. This minimizes the solvent-induced crystallization and the associated surface inhomogeneity (frequently referred to as pooling or flashing). This phenomenon is also related to the solvent vapor pressure. Solvents with higher vapor pressure evaporate faster and result in more uniform surfaces as the comparison between acetone and gamma-butyrolactone illustrates in Fig. S5.† Fig. 2b shows a 3-dimensional reconstructed confocal image of a microchannel formed in PS by immersion in a fluorescent acetone solution. The cross-sectional analysis across the z-axis revealed that the imprinting depth (red line) does not overlap with the gel thickness (blue line); rather, a residual thin fluorescent layer exists below the imprinted area. The imprinting and gel thickness mismatch is independent of the immersion duration or solvent, and suggests that the gel viscosity is not the same throughout the solvent diffusion depth.38 Additionally, the two opposite microchannel surfaces across the x–y plane exhibit a moderately stronger fluorescence than the rest of the gel, indicating that during imprinting, the solute is transported vertically to the imprinting direction. The imprinting depth in SIIL can be controlled by multiple parameters. One is controlling the thickness of the surface gel, which, as previously discussed, in turn depends on the immersion duration as well as the solvent. This solvent dependence is illustrated in Fig. 2c for a 50 μm thick PDMS stamp and a 2 s
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immersion duration in both chloroform and acetone. Despite the same immersion duration, chloroform gave rise to deeper imprinted features due to its highest thermodynamic compatibility. In the case of ethanol, no imprinting was observed even after 40 min of immersion due to its rather dissimilar solubility parameter to PS (δ = 26 (J cm−3)1/2). Longer solvent immersion also gives rise to thicker gels and thus to deeper features. This is also shown in Fig. 2c for a 2 s and 8 s immersion in acetone and employing the same 50 μm thick PDMS stamp. Another means of controlling the imprinted features is the height of the stamp's features. This is illustrated in the inset of Fig. 2c for a PS immersion in acetone and using a 7 μm and a 12 μm deep stamp, which results into equivalently different imprinting depths. We found that the control of the imprinting depth via the stamp height was substantially more practical. It is worth noting that the same solvent transportation mechanism can be used to functionalize the polymer with entities dissolved in the solvent (solute). The polymer-solvent interactions in this case determine the functionalization depth (Fig. 2a). This is discussed in more detail in the following sections.
SIIL protocol Imprinting. A polymer slab or film is initially immersed in the solvent to soften the polymer surface (Fig. 1a). For PS, a 30–40 s long immersion in acetone was generally employed, unless stated otherwise. Subsequently, a PDMS stamp is pressed against the immersed polymer for approximately 8–10 s in order to enable the pattern transfer from the stamp to the polymer. To assist the pattern transfer, light weights (500 g) with manual pressing were applied; alternatively a manual
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press can be employed. Following this, the solvent is removed from around the polymer, while the latter is kept still in contact with the PDMS stamp. This forms the type ‘A’ approach. Alternatively, the polymer can be first removed from the solvent and subsequently imprinted under atmospheric conditions (type ‘B’). This expedites the process by 1 minute; however the imprinted features may suffer from surface inhomogeneities which do not occur in the type ‘A’ approach. Similar to previously reported solvent-assisted nanoimprinting, PDMS was chosen as the stamp material for its non-adhesion properties as well as to permit excess solvent evaporation.24 Imprinting takes place via the flow driven re-organization of the softened gel under mechanical pressure. To this end, care needs to be taken to provide a uniform imprinting pressure and avoid feature distortion due to incomplete gel reflow. Bonding. Following imprinting and solvent removal, the PDMS stamp is removed and the polymer is gently pressed against a non-treated PS surface to permanently bond. Provided that the period from solvent separation until imprinting is not too long (i.e. no longer than 20 s to 25 s for PS–acetone), then the solvent evaporation from the imprinted polymer gel through the PDMS mask is incomplete and thus the surface gel still exists giving rise to some interfacial polymer chain mobility. This enables solvent exchange between the two opposite surfaces, eventually leading to their bonding.15 The two bonded PS pieces could not be manually separated, thus indicating an exceptionally strong and irreversible bond.8,15 The complete bonding and type ‘B’ imprinting process is illustrated in Video S1,† while a representative example is shown in Fig. 3a of a microfluidic channel that is 600 μm wide and 70 μm or up to 100 μm deep. Deeper features than that were not replicated with high
Fig. 3 (a) A 70 μm deep microfluidic cross-junction in polystyrene, in-filled with a fluorescent ethanol solution; the inset shows a photograph of the same structure. (b) A 25 μm wide and approximately 6 μm deep microchannel ending in a 3 μm wide indentation. (c) The magnified indentation with the inset plotting the fluorescence intensity profile across its width, at the location noted by the white dotted arrow. (d) A three dimensional reconstructed confocal image of a microchannel in-filled with a dye solution (red); the microchannel (approximately 30 μm wide) is functionalized with chromophores emitting at a different wavelength giving rise to its endogenous fluorescence (grey).
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definition. We attribute this to both the mechanical instability of PDMS at high aspect rations, as well as the limited solvent penetration in the polymer and the absence of deep enough gels to this end. Methods to address these are currently under investigation. In Fig. 3b, a smaller microchannel is illustrated, 25 μm wide and approximately 6 μm deep ending in a 3 μm wide indentation, which is further magnified in Fig. 3c. The latter indentation was the smallest feature we replicated with SIIL, limited by the resolution of the PDMS mask. Bonding to heterogeneous surfaces was also possible, including both inorganic and organic substrates, such as poly(methyl methacrylate) (PMMA) with a similar exceptionally strong bond (Fig. 3d, and S6†). Functionalization. Solutions – instead of pure solvents – can also be employed in SIIL, so that the solvent enables the solute transport into the polymer. As a result, the immersed polymer becomes functionalized with the solute, which can be dissolved sensing or catalytic molecules. In addition, the now functionalized polymer can be readily imprinted and bonded. Such an example is illustrated in Fig. 3d involving a PS microchannel imprinted using an acetone solution of a chromophore (see methods), bonded to a thin PMMA film and filled with a fluorescent ethanol solution. The image is a 3D reconstructed fluorescence confocal one, illustrating the chromophore implantation in all imprinted surfaces, including the sidewalls and bottom surface of the microchannel. This
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type of functional molecule distribution increases the overlap between the chromophore and the channel's contents, and hence enhances their interaction. This is an attractive feature for both sensing and flow-through catalysis.
Applications Microstructured oxygen sensors. SIIL was applied to prototyping oxygen sensing optofluidics – microsystems that have recently attracted substantial attention17,18 – as oxygen is one of the most important electron acceptors in biology and chemistry. However, contrary to previous reports of optical sensor integration with microfluidics,17,18 SIIL enables controlled 3D distribution of the sensing moieties. This, as already discussed, enables higher overlap between the sensor and analyte and thus offers the possibility of enhanced sensor sensitivity. Another consequence is the similarity between the sensing layer and the imprinting depths (‘ds’ and ‘dp’ respectively in Fig. 4a). This minimizes reagent consumption, but also decreases the background noise by selectively positioning the sensing molecules in close proximity to the imprinted features (i.e. ds ~ dp). Contrary, in the ds > dp case, not all molecules would efficiently interact with O2 and their unmodified lifetime ‘τi’ would negatively contribute to the background noise (Fig. 4a).
Fig. 4 (a) A schematic illustration of the sensor distribution (red) and optical excitation and collection geometry. The excitation is illustrated by the bottom blue arrows (Iexc) and the collection by the green upper ones. For oxygen sensing, the collection signal ‘Isignal’ depends on the excited state lifetimes of all chromophores embedded within the sensitized layer in the path of the optical excitation. (b) Confocal image of a bonded polystyrene microchannel formed by immersion in a fluorescent chloroform solution with an approximate depth of 50 μm; the inset shows the cross-sectional fluorescence intensity distribution across the channel. (c) A series of FLIM snapshots following the transport of gaseous oxygen diffusion in the microchannel described in (a). (d) A contour plot of the phosphorescence lifetime in response to oxygen as a function of time (x-axis) and distance (y-axis corresponds to the white line shown in Fig. 4b).
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For the sensor formation, PS films were immersed for 1 s in a chloroform solution of the oxygen sensitive dye Pt(II) meso tetra(pentafluorophenyl) porphine, leading to the 3D integration of the sensing molecule (Fig. 4b). The imprinted microchannel was bonded to a PS film containing a single inlet, and kept overnight in a nitrogen atmosphere. Oxygen was detected by reduction of the phosphorescence lifetime of the implanted sensing molecule. For this, the microchannel was sealed under strict nitrogen condition and was then positioned in a Fluorescence Lifetime Imaging (FLIM) setup. Subsequently, access of ambient gaseous O2 (approximately 12%) to the microchannel was allowed. A series of FLIM snapshots is shown in Fig. 4c and Video S2,† where the intensity of each pixel corresponds to the phosphorescence lifetime. Both the microchannel bottom surface (area ‘1’) and its walls (areas ‘2a’ and ‘2b’) were oxygen sensitive upon gas entry (t = 0.5 min). However, the bottom surface of the channel (‘1’) exhibited faster response kinetics than the channel walls (‘2’), as better visualized in the contour graph of Fig. 4d that traces the lifetime as a function of time and distance from the microchannel. The response kinetics difference is due to the different O2 diffusional barriers, with the fastest response occurring closest to O2 that necessitates shorter diffusion times. Based on the spatiotemporal analysis of the results shown in Fig. 4d, the oxygen diffusion coefficient in PS was estimated to be 4 × 10−8 cm2 s−1. In contrast to conventional oxygen sensing microfluidics, the three-dimensional distribution of the sensing moieties enabled by SIIL also offers the possibility of differential sensing, engineering the sensor's dynamic range, as well as integrated optical sensor applications.40
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Cell growth microbioreactors. Polymer microreactors were fabricated using SIIL for microbial growth studies in confined environments. Pore network microfluidic structures (or ‘micromodels’) were chosen, similar to the ones shown in Fig. 5a. This special class of microfluidics was employed due to its relevance in quantifying transport in porous media and is representative of ecological heterogeneous subsurface environments.41 The micromodels were fabricated by immersing the PS films in acetone and were subsequently imprinted with an approximately 5 μm deep PDMS stamp. The imprinted printed surface was bonded to a 250 μm thick PS film containing access ports assembled with Nanoport fittings. Micromodels with homogenous pore networks were employed and microbial cell cultures were introduced, grown and imaged. A typical image of a mixed microbial culture is shown in Fig. 5b for Shewanella and Flavobacterium expressing different fluorescent proteins.32 It was possible to determine the cell doubling time by counting the number of cells as a function of time at fixed, 174 × 130 μm square regions across the microchannel depth (Fig. 5b). Such PS micromodels were employed to investigate the growth of Clostridium thermocellum. This strain requires strict anaerobic conditions and an optimal temperature of 60 °C for growth. Such conditions are challenging for elastomeric materials, because of oxygen permeability and buffer evaporation. C. thermocellum cells were loaded in a PS micromodel in a nitrogen atmosphere. Subsequently, the micromodels were sealed at their nanoport fittings (inset of Fig. 5c) and transported to a 60 °C incubator. Cell growth was observed for over 20 h by counting cells at four different micromodel locations that exhibit less than 10% standard deviation
Fig. 5 (a) A pore-scale micromodel design, including photographs of the same structure and fluorescent images of specific details of the micromodel. (b) A mixed microbial population of Shewanella (expressing GFP) and Flavobacterium (expressing m-Orange) growing in a PS micromodel; inset shows the number of cells as a function of time. (c) The growth curves of the Clostridium thermocellum for cells grown in PS micro-scale pore models (blue) and under ideal culture conditions (red); insets show a photograph of a sealed micromodel (upper) and a series of bright field images of a single location in the micromodel at different times.
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between them. The cell-doubling time for C. thermocellum in the micromodel was approximately 4.8 h, considerably longer than the corresponding value of 2.5 h measured in batch media (Fig. 5c). This is attributed to the availability of key nutrients in a well-mixed batch system, where waste products are rapidly removed from around the cells and growth substrates are in excess. These conditions contrast with those within a micromodel. In the latter, minimal mixing around the biomass is more likely to occur, and localized chemical gradients develop through the generation of waste products and utilization of growth substrates around regions of microbial growth.42 Additionally, differences in the micromodel growth environment may lead to shifts in the metabolism and physiology of the microbial population, further contributing to the observed slower doubling time. However, it is worth noting that growth in the PS micromodels is more representative of heterogeneous subsurface conditions, where the majority of biomass is attached to soil and mineral particles and is therefore exposed to limited pore water mixing.
Discussion Polymers have enabled a plethora of photonic and microanalysis applications, including lab-on-a-chip systems. One reason for this is that a wide variety of polymer patterning techniques exist, such as optical lithography,43 micromachining,44 hotembossing,45 injection molding,46 laser ablation,47 soft lithography,48 and nanoimprinting,49 which are generally simpler than in glass and silicon.50,51 Solvents are also known to induce structural changes in polymers without the need of special instruments or high capital investments. For this reason, techniques like SAMIM have revolutionized thin film nano-scale imprinting. Attempts to prototype microsystems using solvents have been very limited and primarily focused on PMMA and requiring long, multi-step processing.20,52 SIIL expands upon these, additionally enabling deep feature imprinting and functionalization that are presently unattainable. In addition, the functionalization step results in a well-controlled three dimensional distribution of the embedded chemical moieties, which in essence increases the overlap between the chromophore and the channel's contents. Finally, SIIL enables the assembly of heterogeneous microsystems, such as organic– inorganic ones, thus opening the possibility of simple, cold, bonding procedures of substrates with different surface chemistries and thermal, electrical, or gas conductivities. SIIL's underlying mechanism are mesoscopic polymer– solvent interactions that induce similar effects as thermal imprinting, where the polymer behaves like a viscous fluid at temperatures higher than its glass transition temperature.49 Contrary to thermal imprinting however, SIIL requires no special instrumentation or expertise other than selecting the appropriate solvent.49 Regarding the latter, the first selection criterion is the solvent thermodynamic compatibility to the polymer. This, along with the immersion duration, enable and control polymer imprinting and functionalization (see Fig. 2). Another selection criterion is based on the solvent
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vapor pressure that needs to be relatively high, leading thus to rapid solvent removal from the polymer and minimal surface non-uniformity (Fig. S5†).
Conclusions In conclusion, solvent immersion lithography was introduced and validated via the demonstration of microfluidics, cell growth microreactors and oxygen sensing microstructures and optofluidics. SIIL's underlying mechanism is based upon the mesoscopic polymer–solvent interactions, a process that was experimentally and computationally investigated. In a single processing step, SIIL enables complete microsystem assembly, including the imprinting of deep features, their bonding and functionalization. The functionalization leads to a 3D chemical moiety distribution, hence enhancing their overlap with the imprinted surface, which is attractive for chemical reactions, or sensing. Additionally, the functional moiety distribution is zero at depths higher than the imprinted features, thus reducing reagent cost as well as the background noise in optical sensing. Due to the reduced instrumentation and expertise requirements, and the possibility to employ a wide range of polymers – including inorganic ones – we anticipate this paradigm to enhance existing applications as well as spark new ones in chemistry and quantitative biology. The technique can be applied to the functionalization and micro-structuring of almost any organic materials, and is synergetic to low-cost and rapid prototyping of microbioreactors, sensors – including ones for in situ applications – polymer optoelectronics and optofluidics for energy.53,54
Contributions AEV conceived SIIL for functionalization, imprinting and bonding, designed and performed the experiments and data analysis. MJW and AEK contributed with related materials and equipment related to oxygen sensing and microbiology. JWG provided oxygen sensing expertise and background on Case II sorption of solvent molecules into polymers. RTK contributed equipment related to PDMS processing. TMC and SSX performed the computational analysis based on the MD simulations. AEV, MJW, JWG and SSX wrote the paper.
Acknowledgements AEV gratefully acknowledges funding from the Pacific Northwest National Laboratory (Linus Pauling Fellowship – LDRD project ID: PN12005/2406). Part of the research was performed using EMSL, a national scientific user facility sponsored by the Department of Energy's Office of Biological and Environmental Research and located at Pacific Northwest National Laboratory (proposal ID: 48924). The experimental work was conducted under the Laboratory Directed Research and Development Program at PNNL, a multiprogram national laboratory operated by Battelle for the U.S. Department of Energy. The computational part of this work was supported by the US Department of Energy, Office of Basic Energy Sciences, Division of Chemical Sciences, Geosciences and
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Biosciences (TMC and SSX). Computer resources were provided by the National Energy Research Scientific Computing Center, which is supported by the Office of Science of the U.S. Department of Energy under contract no. DE-AC02-05CH11231. Additionally, the kind supply of the engineered Flavobacterium strain by Shicheng Chen (Michigan State University), the Yarrowia Lipolytica by Gregory Stephanopoulos (MIT) are gratefully acknowledged.
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23 A. H. Windle, in Polymer Permeability, ed. J. Comyn, Springer, Netherlands, 1986, pp. 75–118. 24 E. Kim, Y. N. Xia, X. M. Zhao and G. M. Whitesides, Adv. Mater., 1997, 9, 651–654. 25 L. J. Guo, Adv. Mater., 2007, 19, 495–513. 26 M. H. Lee, M. D. Huntington, W. Zhou, J.-C. Yang and T. W. Odom, Nano Lett., 2011, 11, 311–315. 27 D. Y. Khang and H. H. Lee, Appl. Phys. Lett., 2000, 76, 870–872. 28 K. L. Lai, M. H. Hon and I. C. Leu, J. Micromech. Microeng., 2011, 21. 29 N. E. Voicu, S. Ludwigs, E. J. W. Crossland, P. Andrew and U. Steiner, Adv. Mater., 2007, 19, 757–761. 30 E. A. Johnson, A. Madia and A. L. Demain, Appl. Environ. Microbiol., 1981, 41, 1060–1062. 31 D. R. Lovley, S. J. Giovannoni, D. C. White, J. E. Champine, E. J. P. Phillips, Y. A. Gorby and S. Goodwin, Arch. Microbiol., 1993, 159, 336–344. 32 S. Chen, M. G. Kaufman, M. Bagdasarian, A. K. Bates and E. D. Walker, Gene, 2010, 458, 1–10. 33 W. L. Jorgensen, D. S. Maxwell and J. Tirado Rives, J. Am. Chem. Soc., 1996, 118, 11225–11236. 34 T. M. Chang, L. X. Dang and K. A. Peterson, J. Phys. Chem. B, 1997, 101, 3413–3419. 35 F. Asmussen and K. Ueberreiter, J. Polym. Sci., 1962, 57, 199–208. 36 W. Limm, D. Stanton, G. P. Dimnik, M. A. Winnik and B. A. Smith, J. Appl. Polym. Sci., 1988, 35, 2099–2116. 37 K. Ueberreiter, The solution process, Academic Press, New York, NY, 1968. 38 B. A. Miller-Chou and J. L. Koenig, Prog. Polym. Sci., 2003, 28, 1223–1270. 39 T. S. Chow, Mesoscopic Physics of Complex Materials, Springer, 2000. 40 O. S. Wolfbels, Anal. Chem., 2008, 80, 4269–4283. 41 T. Long and R. M. Ford, Environ. Sci. Technol., 2009, 43, 1546–1552. 42 R. Stocker, Science, 2012, 338, 628–633. 43 W. M. Moreau, Semiconductor Lithography: Principles and Materials, Plenum, New York, 1988. 44 D. J. Harrison, K. Fluri, K. Seiler, Z. H. Fan, C. S. Effenhauser and A. Manz, Science, 1993, 261, 895–897. 45 H. Becker and U. Heim, Sens. Actuators, A, 2000, 83, 130–135. 46 R. M. McCormick, R. J. Nelson, M. G. AlonsoAmigo, J. Benvegnu and H. H. Hooper, Anal. Chem., 1997, 69, 2626–2630. 47 R. R. Gattass and E. Mazur, Nat. Photonics, 2008, 2, 219–225. 48 Y. N. Xia and G. M. Whitesides, Annu. Rev. Mater. Sci., 1998, 28, 153–184. 49 S. Y. Chou, P. R. Krauss and P. J. Renstrom, J. Vac. Sci. Technol., B: Microelectron. Nanometer Struct.–Process., Meas., Phenom., 1996, 14, 4129–4133. 50 G. M. Whitesides, Nature, 2006, 442, 368–373. 51 K. F. Jensen, MRS Bull., 2006, 31, 101–107. 52 M. T. Koesdjojo, C. R. Koch and V. T. Remcho, Anal. Chem., 2009, 81, 1652–1659. 53 N. Szita, P. Boccazzi, Z. Y. Zhang, P. Boyle, A. J. Sinskey and K. F. Jensen, Lab Chip, 2005, 5, 819–826. 54 D. Erickson, D. Sinton and D. Psaltis, Nat. Photonics, 2011, 5, 583–590.
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Solvent immersion imprint lithography† Cite this: Lab Chip, 2014, 14, 2072
A. E. Vasdekis,*a M. J. Wilkins,‡b J. W. Grate,c R. T. Kelly,a A. E. Konopka,b S. S. Xantheasc and T.-M. Changd We present Solvent Immersion Imprint Lithography (SIIL), a technique for polymer functionalization and microsystem prototyping. SIIL is based on polymer immersion in commonly available solvents. This was experimentally and computationally analyzed, uniquely enabling two practical aspects. The first is imprinting and bonding deep features that span the 1 to 100 μm range, which are unattainable with existing solvent-based methods. The second is a functionalization scheme characterized by a well-controlled, 3D
Received 20th February 2014, Accepted 8th April 2014 DOI: 10.1039/c4lc00226a www.rsc.org/loc
distribution of chemical moieties. SIIL is validated by developing microfluidics with embedded 3D oxygen sensors and microbioreactors for quantitative metabolic studies of a thermophile anaerobe microbial culture. Polystyrene (PS) was employed in the aforementioned applications; however all soluble polymers – including inorganic ones – can be employed with SIIL under no instrumentation requirements and typical processing times of less than two minutes.
Introduction Polydimethilsiloxane (PDMS) has become the workhorse material in microfluidics and optofluidics due to its low cost and simple processing.1–5 However, PDMS has several recognized drawbacks, such as limited compatibility with organic solvents, and high gas permeability and resulting enhanced buffer evaporation rates, especially at elevated temperatures.6 Thermoplastic polymers,7,8 as well as emerging inorganic ones,9,10 do not have these shortcomings. As a result, certain microsystem applications have been revolutionized, such as cell culture experiments,8 complex flow generation,11 nanofluidics,12 chemical synthesis13 and photonics.14 However, imprinting in thermoplastic or inorganic polymers requires substantially more complex processing and dedicated instrumentation, such as hot presses, while bonding relies mostly on thermal means that suffers from reduced success rate and shape deformations.15 These presently make inorganic and thermoplastic polymers less attractive and accessible than PDMS. Another aspect of polymer microsystems that has received little attention is their chemical functionalization. This
a Environmental Molecular Sciences Laboratory, Pacific Northwest National Laboratory, PO Box 999, Richland, WA, 99352, USA. E-mail: [email protected] b Biological Sciences Division, Pacific Northwest National Laboratory, PO Box 999, Richland, WA, 99352, USA c Physical Sciences Division, Pacific Northwest National Laboratory, PO Box 999, Richland, WA, 99352, USA d University of Wisconsin-Parkside, Box 2000, Kenosha, Wisconsin 53141, USA † Electronic supplementary information (ESI) available. See DOI: 10.1039/ c4lc00226a ‡ Present address: School of Earth Sciences and Department of Microbiology, The Ohio State University, Columbus OH, 43210, USA.
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generally requires expensive and dedicated instruments, thus hindering integrative biology and energy microsystem applications (e.g. flow through catalysis).16 Bulk-type functionalization is also desirable in optical applications. This is possible via the microfluidic integration of thin films doped with the chemical entity of interest.17,18 In this case, however, additional processing is essential (e.g. spin coating, patterning etc.), while a uniform dopant distribution is usually obtained with no ability to control it. Controlling the dopant distribution would be of substantial importance in designing novel microsystems, for example sensors with engineered sensitivity and dynamic range. With the exception of functionalization, the imprinting and bonding complexity in thermoplastic or inorganic polymers have been partially addressed by a variety of recent benchtop processes that minimize the need for instrumentation, involving polysilsesquioxane,9 as well as cyclic olefin copolymers (COC) and poly(methyl methacrylate) (PMMA).19–21 Polystyrene (PS) is another low-cost thermoplastic polymer, where costeffective imprinting approaches involve Shrinky-Dink based methods, or soft-lithography using PS solutions.7,22 However, despite such substantial initial progress, rapid and costeffective imprinting and bonding, as well as novel and enabling functionalization strategies, still remain an important goal. Clearly, progress in this direction would further enhance the use of polymer microsystems in chemical, biological and optical applications. In order to bridge the gap between the simplicity of PDMS processing and the stability of thermoplastic and inorganic polymers, as well as address the challenge of polymer functionalization, we introduce Solvent Immersion Imprint Lithography (SIIL). SIIL is based on commonly available solvents and enables complete microsystem prototyping,
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Fig. 1 (a) Schematic illustration of the SIIL protocol. (b) A 4 cm long microchannel imprinted in PS using a 40 s long immersion in acetone; the microchannel dimensions measured using a profilometer at its two far ends were 100 μm wide and 35 μm deep, shown in the inset. (c) A statistical analysis of 11 samples like the one shown in (b) determining the depth and width variation between different imprinting runs. (d) Comparison between imprinted microchannels using existing solvent-based methods (red profile) and SIIL (blue profile); for this experiment, the same imprinting conditions were employed, namely a 20" imprinting duration, acetone as a solvent, a 5 μm deep PDMS stamp and the same exerted force enabled by a 500 g weight.
including functionalization, imprinting and bonding in a single processing step (Fig. 1a). This involves immersing the polymer in a solvent, during which the solvent penetrates into the polymer forming a surface ‘gel’ layer. This process (potentially a case II sorption23) is SIIL's basic mechanism: the softened surface layer can be readily functionalized with moieties such as chemosensors, imprinted using a mold, as well as bonded to a substrate. Large area imprinting is possible with SIIL, such as the representative PS microchannel shown in Fig. 1b (also in Fig. S1†), which is approximately 4 cm long (x-axis), 100 μm wide (y-axis) and 35 μm deep (z-axis). Such large area patterning exhibited high quality with minimal variation across the entire imprinted surface as confirmed by the profilometer measurements (inset of Fig. 1b). Additionally, the statistical analysis shown in Fig. 1c revealed minimal variation in width and depth between 11 different imprinting runs with 2.9 μm (2.6%) and 1. 2 μm (3%) respectively. SIIL expands upon previously reported solvent based nanoimprinting methods, such as the Solvent Assisted Micromolding (SAMIM), the first technique to be developed to this end.24 In SAMIM, the solvent coats the PDMS stamp, which is subsequently pressed against a thin polymer film causing it to partially dissolve and conform to the stamp's pattern.24–26 Solvent vapors have also been used to the same end,
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dissolving the polymer by first propagating through the pores of the PDMS stamp.27–29 Despite the wide success of solventassisted nano-imprinting, solvent based microfluidic prototyping using solvents has received little attention due to limitations in obtaining deep features that span the 1 μm to 100 μm range. The reason for this is that by using a solvent coated stamp, the solvent contacts the polymer only upon imprinting, which leads to solvent reflow and trapping at the edges of the stamp upon contact between the polymer and the stamp and solvent. This phenomenon is absent in thin feature imprinting, however, it becomes non-trivial for deeper features as shown in Fig. 1d (also Fig. S2†). On the contrary, SIIL overcomes these shortcomings because the polymer immersion into the solvent provides better control over the formation of the surface gel, a critical aspect in imprinting deep features (Fig. 1d and S2†). Provided that it is soluble, any kind of polymer is compatible with SIIL, including inorganic ones.9 In the following sections, we present our experimental and computational investigations of the underlying polymer–solvent interactions, specifically in relation to functionalization and deep feature imprinting. PS microfluidics, cell growth microbioreactors and microstructured oxygen sensors are demonstrated by way of example here, with typical processing times of less than two minutes.
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Materials and methods
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Materials The polymers used in the experiments, namely PS and PMMA were supplied by GoodFellow (USA) at various thicknesses ranging from 1.5 mm to 0.25 mm (Product IDs: ST313120, ST311250, 137-745-63). All organic solvents were supplied by Sigma Aldrich (USA), while purified deionized water was employed where mentioned. The oxygen sensing chromophore (PtT975) employed in the optofluidic experiment was supplied by Frontier Scientific (USA) at a concentration of 0.5 mg ml−1. The stamps were made by conventional cast-moulding lithography in PDMS. The PDMS (Sylgard 184, Dow Corning, USA) was mixed with the catalyst at 1 : 10 ratio, degassed for approximately 1 h, baked at 70 °C for 1 h, removed from the hard SU8 mask and baked for an additional 1 h. The hard SU8 mask was fabricated with conventional contact mode optical lithography using the MicroChem formulations SU8-2000. The photoresist was spin coated onto Si 4′′ wafers at different speeds to control its thickness. For confocal imaging, two different chromophores were used depending on the solvent, both supplied by Sigma Aldrich; fluorescein was mixed with acetone and Nile Red with acetone and chloroform at approximately 1 mM concentration. Imaging The fluorescent imaging of the gel layer was performed in an inverted microscope (Leica DMI6000), coupled with a spinning disk confocal system (Yokogawa, CSU10). A 20× objective was used (20×/.7 NA Plan Apo DIC Optics Inclusive) and the z-scanning was performed at a 2 μm step size and the gel film thickness was estimated by intensity thresholding. For the latter the background level was estimated by the ratio of the standard deviation over the mean. Oxygen sensing was performed in a fluorescence lifetime imaging setup (LI2CAM-P, Lambert Instruments, Netherlands), integrated with the aforementioned Leica inverted microscope. For this measurement, an LED with an emission spectrum centered at 399 nm was employed modulated at 5 kHz. Microfluidics In the microfluidic experiments, a peristaltic pump, capillary forces or manual means were employed to inject fluids. The microbial cells were manually injected in the micromodels using a syringe. The filling of the PS microchannels for fluorescent imaging (see e.g. Fig. 3) was performed by capillary action, using a fluorescein ethanol solution. Flow rate control, where stated, was provided by a precision milliGAT pump (Global FIA, USA), connected to the microfluidics with nanoport fittings through PTFE tubing. Cell cultures Clostridium thermocellum ATCC 27405 cultures were grown in complex GS-2 media,30 at 60 °C under strict anaerobic conditions. In these media, cellobiose was the main carbon source.
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Batch growth rate experiments were performed in 50 mL volumes, under shaking conditions. Cell densities were measured by removing 1 mL aliquots of culture, and measuring absorbance at 600 nm using a BioRad spectrophotometer (Bio Rad, Hercules, CA). The aerobic bacterial strains, Shewanella and Flavobacteria, were grown in a bicarbonatebuffered media containing glucose as the sole carbon source.31 These were a spontaneous variant of Shewanella oneidensis MR-1 engineered to express a green-fluorescent protein (GFP), and Flavobacterium johnsoniae UW101 expressing the mStrawberry fluorescent protein.32 Batch cultures were incubated under aerobic conditions at 30 °C. Prior to growth experiments, the biomass was grown overnight and subsequently transferred at 1% inoculum into fresh media. Computational Classical molecular dynamics (MD) simulations were used to study the dissolution processes between solvents (acetone, chloroform) and polystyrene with four monomer units. The energy of the system was described by a sum of Lennard-Jones, Coulombic, and intramolecular (bond, angle, and torsion) interactions. The acetone, chloroform, and polystyrene potentials were based on an all-atom framework, with fixed partial charges and Lennard-Jones parameters assigned to each atom. The potential parameters for acetone and polystyrene were taken from the OPLS-AA force field,33 whereas for chloroform a potential modified from a previous model was used.34 The cross Lennard-Jones interaction terms between unlike atoms are obtained using the Lorentz–Berthelot combining rules. All MD simulations were carried out with the AMBER 9 package. More information is available in the ESI† section.
Results Polymer–solvent interactions SIIL is based on polymer immersion in appropriate solvents. During this step, the solvent diffuses into the polymer,35,36 the initial stages of which were modeled using molecular dynamics (MD) simulations. The inset of Fig. 2a illustrates individual acetone molecules diffusing into PS (also in Fig. S3†). To further analyze solvent diffusion, PS films were immersed into fluorescent dye solutions. In this way, the solvent transports the dye into the polymer, hence enabling the selective staining of the solvent's diffusion path. While several methods exist in the literature to investigate polymer– solvent interaction, we employed fluorescence microscopy37 in order to combine the diffusion with the imprinting and functionalization investigations. Once dried, the films were imaged by fluorescence confocal microscopy along the film depth (z-axis) (see methods and Fig. S4†). Similar procedures have been previously reported in interfacial polymer dissolution studies;38 however, in the present experiments, confocal imaging was chosen to characterize substrates that can be specifically used in microfluidic applications. Typical results are shown in the histogram of Fig. 2a for different immersion durations, illustrating that the solvent
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Fig. 2 (a) The gel and functionalization thickness dependence on the immersion duration for the PS–acetone and PS–chloroform systems; inset plots snapshots of molecular dynamics simulations of acetone (purple)/polystyrene (cyan) interface upon contact and following a 100 ns long interaction. (b) A reconstructed confocal image across the z-axis of an imprinted PS microchannel based on an 8 s immersion in a fluorescent acetone solution; the thinner fluorescent region corresponds to the imprinted area, while the blue and red lines illustrate the bottom surface of the gel and the imprinted feature respectively. Inset plots the three-dimensional image of the same film, with the blue line denoting the location of the z-scan. (c) The depth of the imprinted features in PS following immersion in different solvents and for different durations using a 50 μm deep PDMS stamp; the same pressure was exerted in all experiments by positioning a light weight (500 g) on top of the polymer slabs. The inset plots the profile of imprinted features in PS using pure acetone and 7 μm and 12 μm deep stamps.
diffusion depth increases for longer immersions and eventually saturates, as previously reported.37 The solvent diffusion depth was also found to depend on the solvent type and the underlying polymer–solvent molecular interactions. These interactions are frequently described by the Hildebrand solubility parameter, which states that the closer the solubility parameters between two substances are, the easier it is to mix them.38 The Hildebrandt parameters, δ, for chloroform and acetone are 18.7 and 20.4 (J cm−3)1/2, respectively, while the one for PS is 18.7 (J cm−3)1/2.38 The solvent diffusion depth was substantially thicker in chloroform than acetone for similar immersion durations (Fig. 2a). This occurs because of chloroform's higher thermodynamic compatibility with PS, as evidenced by their matching Hildebrandt parameters.
Solvent immersion functionalization and imprinting Solvent diffusion results in a decreased viscosity at the polymer interface. This forms a surface gel without however modifying the polymer chains and thus manifesting the mechanisms' mesoscopic nature.39 To explore the formation of a surface gel layer, PS films were immersed in a fluorescent solution and subsequently pressed against a patterned PDMS slab, as described in Fig. 1a. During this process, the PDMS pattern is transferred to the fluorescent gel of the immersed PS, while the porosity of PDMS enables the solvent evaporation. The
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latter is important as it enables rapid solvent removal from the polymer while still in contact with the PDMS. This minimizes the solvent-induced crystallization and the associated surface inhomogeneity (frequently referred to as pooling or flashing). This phenomenon is also related to the solvent vapor pressure. Solvents with higher vapor pressure evaporate faster and result in more uniform surfaces as the comparison between acetone and gamma-butyrolactone illustrates in Fig. S5.† Fig. 2b shows a 3-dimensional reconstructed confocal image of a microchannel formed in PS by immersion in a fluorescent acetone solution. The cross-sectional analysis across the z-axis revealed that the imprinting depth (red line) does not overlap with the gel thickness (blue line); rather, a residual thin fluorescent layer exists below the imprinted area. The imprinting and gel thickness mismatch is independent of the immersion duration or solvent, and suggests that the gel viscosity is not the same throughout the solvent diffusion depth.38 Additionally, the two opposite microchannel surfaces across the x–y plane exhibit a moderately stronger fluorescence than the rest of the gel, indicating that during imprinting, the solute is transported vertically to the imprinting direction. The imprinting depth in SIIL can be controlled by multiple parameters. One is controlling the thickness of the surface gel, which, as previously discussed, in turn depends on the immersion duration as well as the solvent. This solvent dependence is illustrated in Fig. 2c for a 50 μm thick PDMS stamp and a 2 s
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immersion duration in both chloroform and acetone. Despite the same immersion duration, chloroform gave rise to deeper imprinted features due to its highest thermodynamic compatibility. In the case of ethanol, no imprinting was observed even after 40 min of immersion due to its rather dissimilar solubility parameter to PS (δ = 26 (J cm−3)1/2). Longer solvent immersion also gives rise to thicker gels and thus to deeper features. This is also shown in Fig. 2c for a 2 s and 8 s immersion in acetone and employing the same 50 μm thick PDMS stamp. Another means of controlling the imprinted features is the height of the stamp's features. This is illustrated in the inset of Fig. 2c for a PS immersion in acetone and using a 7 μm and a 12 μm deep stamp, which results into equivalently different imprinting depths. We found that the control of the imprinting depth via the stamp height was substantially more practical. It is worth noting that the same solvent transportation mechanism can be used to functionalize the polymer with entities dissolved in the solvent (solute). The polymer-solvent interactions in this case determine the functionalization depth (Fig. 2a). This is discussed in more detail in the following sections.
SIIL protocol Imprinting. A polymer slab or film is initially immersed in the solvent to soften the polymer surface (Fig. 1a). For PS, a 30–40 s long immersion in acetone was generally employed, unless stated otherwise. Subsequently, a PDMS stamp is pressed against the immersed polymer for approximately 8–10 s in order to enable the pattern transfer from the stamp to the polymer. To assist the pattern transfer, light weights (500 g) with manual pressing were applied; alternatively a manual
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press can be employed. Following this, the solvent is removed from around the polymer, while the latter is kept still in contact with the PDMS stamp. This forms the type ‘A’ approach. Alternatively, the polymer can be first removed from the solvent and subsequently imprinted under atmospheric conditions (type ‘B’). This expedites the process by 1 minute; however the imprinted features may suffer from surface inhomogeneities which do not occur in the type ‘A’ approach. Similar to previously reported solvent-assisted nanoimprinting, PDMS was chosen as the stamp material for its non-adhesion properties as well as to permit excess solvent evaporation.24 Imprinting takes place via the flow driven re-organization of the softened gel under mechanical pressure. To this end, care needs to be taken to provide a uniform imprinting pressure and avoid feature distortion due to incomplete gel reflow. Bonding. Following imprinting and solvent removal, the PDMS stamp is removed and the polymer is gently pressed against a non-treated PS surface to permanently bond. Provided that the period from solvent separation until imprinting is not too long (i.e. no longer than 20 s to 25 s for PS–acetone), then the solvent evaporation from the imprinted polymer gel through the PDMS mask is incomplete and thus the surface gel still exists giving rise to some interfacial polymer chain mobility. This enables solvent exchange between the two opposite surfaces, eventually leading to their bonding.15 The two bonded PS pieces could not be manually separated, thus indicating an exceptionally strong and irreversible bond.8,15 The complete bonding and type ‘B’ imprinting process is illustrated in Video S1,† while a representative example is shown in Fig. 3a of a microfluidic channel that is 600 μm wide and 70 μm or up to 100 μm deep. Deeper features than that were not replicated with high
Fig. 3 (a) A 70 μm deep microfluidic cross-junction in polystyrene, in-filled with a fluorescent ethanol solution; the inset shows a photograph of the same structure. (b) A 25 μm wide and approximately 6 μm deep microchannel ending in a 3 μm wide indentation. (c) The magnified indentation with the inset plotting the fluorescence intensity profile across its width, at the location noted by the white dotted arrow. (d) A three dimensional reconstructed confocal image of a microchannel in-filled with a dye solution (red); the microchannel (approximately 30 μm wide) is functionalized with chromophores emitting at a different wavelength giving rise to its endogenous fluorescence (grey).
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definition. We attribute this to both the mechanical instability of PDMS at high aspect rations, as well as the limited solvent penetration in the polymer and the absence of deep enough gels to this end. Methods to address these are currently under investigation. In Fig. 3b, a smaller microchannel is illustrated, 25 μm wide and approximately 6 μm deep ending in a 3 μm wide indentation, which is further magnified in Fig. 3c. The latter indentation was the smallest feature we replicated with SIIL, limited by the resolution of the PDMS mask. Bonding to heterogeneous surfaces was also possible, including both inorganic and organic substrates, such as poly(methyl methacrylate) (PMMA) with a similar exceptionally strong bond (Fig. 3d, and S6†). Functionalization. Solutions – instead of pure solvents – can also be employed in SIIL, so that the solvent enables the solute transport into the polymer. As a result, the immersed polymer becomes functionalized with the solute, which can be dissolved sensing or catalytic molecules. In addition, the now functionalized polymer can be readily imprinted and bonded. Such an example is illustrated in Fig. 3d involving a PS microchannel imprinted using an acetone solution of a chromophore (see methods), bonded to a thin PMMA film and filled with a fluorescent ethanol solution. The image is a 3D reconstructed fluorescence confocal one, illustrating the chromophore implantation in all imprinted surfaces, including the sidewalls and bottom surface of the microchannel. This
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type of functional molecule distribution increases the overlap between the chromophore and the channel's contents, and hence enhances their interaction. This is an attractive feature for both sensing and flow-through catalysis.
Applications Microstructured oxygen sensors. SIIL was applied to prototyping oxygen sensing optofluidics – microsystems that have recently attracted substantial attention17,18 – as oxygen is one of the most important electron acceptors in biology and chemistry. However, contrary to previous reports of optical sensor integration with microfluidics,17,18 SIIL enables controlled 3D distribution of the sensing moieties. This, as already discussed, enables higher overlap between the sensor and analyte and thus offers the possibility of enhanced sensor sensitivity. Another consequence is the similarity between the sensing layer and the imprinting depths (‘ds’ and ‘dp’ respectively in Fig. 4a). This minimizes reagent consumption, but also decreases the background noise by selectively positioning the sensing molecules in close proximity to the imprinted features (i.e. ds ~ dp). Contrary, in the ds > dp case, not all molecules would efficiently interact with O2 and their unmodified lifetime ‘τi’ would negatively contribute to the background noise (Fig. 4a).
Fig. 4 (a) A schematic illustration of the sensor distribution (red) and optical excitation and collection geometry. The excitation is illustrated by the bottom blue arrows (Iexc) and the collection by the green upper ones. For oxygen sensing, the collection signal ‘Isignal’ depends on the excited state lifetimes of all chromophores embedded within the sensitized layer in the path of the optical excitation. (b) Confocal image of a bonded polystyrene microchannel formed by immersion in a fluorescent chloroform solution with an approximate depth of 50 μm; the inset shows the cross-sectional fluorescence intensity distribution across the channel. (c) A series of FLIM snapshots following the transport of gaseous oxygen diffusion in the microchannel described in (a). (d) A contour plot of the phosphorescence lifetime in response to oxygen as a function of time (x-axis) and distance (y-axis corresponds to the white line shown in Fig. 4b).
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For the sensor formation, PS films were immersed for 1 s in a chloroform solution of the oxygen sensitive dye Pt(II) meso tetra(pentafluorophenyl) porphine, leading to the 3D integration of the sensing molecule (Fig. 4b). The imprinted microchannel was bonded to a PS film containing a single inlet, and kept overnight in a nitrogen atmosphere. Oxygen was detected by reduction of the phosphorescence lifetime of the implanted sensing molecule. For this, the microchannel was sealed under strict nitrogen condition and was then positioned in a Fluorescence Lifetime Imaging (FLIM) setup. Subsequently, access of ambient gaseous O2 (approximately 12%) to the microchannel was allowed. A series of FLIM snapshots is shown in Fig. 4c and Video S2,† where the intensity of each pixel corresponds to the phosphorescence lifetime. Both the microchannel bottom surface (area ‘1’) and its walls (areas ‘2a’ and ‘2b’) were oxygen sensitive upon gas entry (t = 0.5 min). However, the bottom surface of the channel (‘1’) exhibited faster response kinetics than the channel walls (‘2’), as better visualized in the contour graph of Fig. 4d that traces the lifetime as a function of time and distance from the microchannel. The response kinetics difference is due to the different O2 diffusional barriers, with the fastest response occurring closest to O2 that necessitates shorter diffusion times. Based on the spatiotemporal analysis of the results shown in Fig. 4d, the oxygen diffusion coefficient in PS was estimated to be 4 × 10−8 cm2 s−1. In contrast to conventional oxygen sensing microfluidics, the three-dimensional distribution of the sensing moieties enabled by SIIL also offers the possibility of differential sensing, engineering the sensor's dynamic range, as well as integrated optical sensor applications.40
Lab on a Chip
Cell growth microbioreactors. Polymer microreactors were fabricated using SIIL for microbial growth studies in confined environments. Pore network microfluidic structures (or ‘micromodels’) were chosen, similar to the ones shown in Fig. 5a. This special class of microfluidics was employed due to its relevance in quantifying transport in porous media and is representative of ecological heterogeneous subsurface environments.41 The micromodels were fabricated by immersing the PS films in acetone and were subsequently imprinted with an approximately 5 μm deep PDMS stamp. The imprinted printed surface was bonded to a 250 μm thick PS film containing access ports assembled with Nanoport fittings. Micromodels with homogenous pore networks were employed and microbial cell cultures were introduced, grown and imaged. A typical image of a mixed microbial culture is shown in Fig. 5b for Shewanella and Flavobacterium expressing different fluorescent proteins.32 It was possible to determine the cell doubling time by counting the number of cells as a function of time at fixed, 174 × 130 μm square regions across the microchannel depth (Fig. 5b). Such PS micromodels were employed to investigate the growth of Clostridium thermocellum. This strain requires strict anaerobic conditions and an optimal temperature of 60 °C for growth. Such conditions are challenging for elastomeric materials, because of oxygen permeability and buffer evaporation. C. thermocellum cells were loaded in a PS micromodel in a nitrogen atmosphere. Subsequently, the micromodels were sealed at their nanoport fittings (inset of Fig. 5c) and transported to a 60 °C incubator. Cell growth was observed for over 20 h by counting cells at four different micromodel locations that exhibit less than 10% standard deviation
Fig. 5 (a) A pore-scale micromodel design, including photographs of the same structure and fluorescent images of specific details of the micromodel. (b) A mixed microbial population of Shewanella (expressing GFP) and Flavobacterium (expressing m-Orange) growing in a PS micromodel; inset shows the number of cells as a function of time. (c) The growth curves of the Clostridium thermocellum for cells grown in PS micro-scale pore models (blue) and under ideal culture conditions (red); insets show a photograph of a sealed micromodel (upper) and a series of bright field images of a single location in the micromodel at different times.
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between them. The cell-doubling time for C. thermocellum in the micromodel was approximately 4.8 h, considerably longer than the corresponding value of 2.5 h measured in batch media (Fig. 5c). This is attributed to the availability of key nutrients in a well-mixed batch system, where waste products are rapidly removed from around the cells and growth substrates are in excess. These conditions contrast with those within a micromodel. In the latter, minimal mixing around the biomass is more likely to occur, and localized chemical gradients develop through the generation of waste products and utilization of growth substrates around regions of microbial growth.42 Additionally, differences in the micromodel growth environment may lead to shifts in the metabolism and physiology of the microbial population, further contributing to the observed slower doubling time. However, it is worth noting that growth in the PS micromodels is more representative of heterogeneous subsurface conditions, where the majority of biomass is attached to soil and mineral particles and is therefore exposed to limited pore water mixing.
Discussion Polymers have enabled a plethora of photonic and microanalysis applications, including lab-on-a-chip systems. One reason for this is that a wide variety of polymer patterning techniques exist, such as optical lithography,43 micromachining,44 hotembossing,45 injection molding,46 laser ablation,47 soft lithography,48 and nanoimprinting,49 which are generally simpler than in glass and silicon.50,51 Solvents are also known to induce structural changes in polymers without the need of special instruments or high capital investments. For this reason, techniques like SAMIM have revolutionized thin film nano-scale imprinting. Attempts to prototype microsystems using solvents have been very limited and primarily focused on PMMA and requiring long, multi-step processing.20,52 SIIL expands upon these, additionally enabling deep feature imprinting and functionalization that are presently unattainable. In addition, the functionalization step results in a well-controlled three dimensional distribution of the embedded chemical moieties, which in essence increases the overlap between the chromophore and the channel's contents. Finally, SIIL enables the assembly of heterogeneous microsystems, such as organic– inorganic ones, thus opening the possibility of simple, cold, bonding procedures of substrates with different surface chemistries and thermal, electrical, or gas conductivities. SIIL's underlying mechanism are mesoscopic polymer– solvent interactions that induce similar effects as thermal imprinting, where the polymer behaves like a viscous fluid at temperatures higher than its glass transition temperature.49 Contrary to thermal imprinting however, SIIL requires no special instrumentation or expertise other than selecting the appropriate solvent.49 Regarding the latter, the first selection criterion is the solvent thermodynamic compatibility to the polymer. This, along with the immersion duration, enable and control polymer imprinting and functionalization (see Fig. 2). Another selection criterion is based on the solvent
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vapor pressure that needs to be relatively high, leading thus to rapid solvent removal from the polymer and minimal surface non-uniformity (Fig. S5†).
Conclusions In conclusion, solvent immersion lithography was introduced and validated via the demonstration of microfluidics, cell growth microreactors and oxygen sensing microstructures and optofluidics. SIIL's underlying mechanism is based upon the mesoscopic polymer–solvent interactions, a process that was experimentally and computationally investigated. In a single processing step, SIIL enables complete microsystem assembly, including the imprinting of deep features, their bonding and functionalization. The functionalization leads to a 3D chemical moiety distribution, hence enhancing their overlap with the imprinted surface, which is attractive for chemical reactions, or sensing. Additionally, the functional moiety distribution is zero at depths higher than the imprinted features, thus reducing reagent cost as well as the background noise in optical sensing. Due to the reduced instrumentation and expertise requirements, and the possibility to employ a wide range of polymers – including inorganic ones – we anticipate this paradigm to enhance existing applications as well as spark new ones in chemistry and quantitative biology. The technique can be applied to the functionalization and micro-structuring of almost any organic materials, and is synergetic to low-cost and rapid prototyping of microbioreactors, sensors – including ones for in situ applications – polymer optoelectronics and optofluidics for energy.53,54
Contributions AEV conceived SIIL for functionalization, imprinting and bonding, designed and performed the experiments and data analysis. MJW and AEK contributed with related materials and equipment related to oxygen sensing and microbiology. JWG provided oxygen sensing expertise and background on Case II sorption of solvent molecules into polymers. RTK contributed equipment related to PDMS processing. TMC and SSX performed the computational analysis based on the MD simulations. AEV, MJW, JWG and SSX wrote the paper.
Acknowledgements AEV gratefully acknowledges funding from the Pacific Northwest National Laboratory (Linus Pauling Fellowship – LDRD project ID: PN12005/2406). Part of the research was performed using EMSL, a national scientific user facility sponsored by the Department of Energy's Office of Biological and Environmental Research and located at Pacific Northwest National Laboratory (proposal ID: 48924). The experimental work was conducted under the Laboratory Directed Research and Development Program at PNNL, a multiprogram national laboratory operated by Battelle for the U.S. Department of Energy. The computational part of this work was supported by the US Department of Energy, Office of Basic Energy Sciences, Division of Chemical Sciences, Geosciences and
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Biosciences (TMC and SSX). Computer resources were provided by the National Energy Research Scientific Computing Center, which is supported by the Office of Science of the U.S. Department of Energy under contract no. DE-AC02-05CH11231. Additionally, the kind supply of the engineered Flavobacterium strain by Shicheng Chen (Michigan State University), the Yarrowia Lipolytica by Gregory Stephanopoulos (MIT) are gratefully acknowledged.
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