Diffraction-limited ultrasensitive molecular nano-arrays with singular nano-cone scattering

Large-library fluorescent molecular arrays remain limited in sensitivity (1 × 10⁶ molecules) and dynamic range due to background auto-fluorescence and scattering noise within a large (20–100 μm) fluorescent spot. We report an easily fabricated silica nano-cone array platform, with a detection limit of 100 molecules and a dynamic range that spans 6 decades, due to point (10 nm to 1 μm) illumination of preferentially absorbed tagged targets by singular scattering off wedged cones. Its fluorescent spot reaches diffraction-limited submicron dimensions, which are 10⁴ times smaller in area than conventional microarrays, with comparable reduction in detection limit and amplification of dynamic range.Commercially available fluorescent micro-arrays based on target labeling, northern blot, or enzyme-linked immunosorbent assay (ELISA) are limited to a detection threshold of 1 to 10 × 10⁶ molecules per fluorescent spot,^{1, 2, 3, 4, 5, 6, 7, 8, 9, 10, 11, 12, 13, 14, 15, 16, 17, 18, 19, 20, 21, 22, 23} thus requiring cell culturing or Polymerase Chain Reaction (PCR) amplification for many applications. The low sensitivity is often due to broad illumination, which creates auto-fluorescence noise. Even if point illumination and pin-hole filtering of non-focal plane noise are implemented in a confocal setup, the large and non-uniform fluorescent spots create scattering noise over each 20–100 μm element, which degrades the detection limit.⁴ Smaller spots can, in theory, be introduced by nano-sprays and nano-imprinting. However, directing the targets to such small areas then becomes problematic. Real-time PCR is, in principle, capable of detecting a single molecule but is limited in its target number⁵ and is hence slow/expensive for large-library assays. A large-library platform with much better detection limit than the current fluorescent microarrays would transform many screening assays. Ideally, this platform would not use the confocal configuration. Instead, it would direct the target molecules to a submicron spot and illuminate them with a nearby point source that does not require scanning.A promising platform is the optical fiber bundle array,⁶ with more than 10⁴ fibers and targets, in principle. With its endoscopic configuration, these fiber bundles are most convenient for in situ and real-time biosensing modalities in microfluidic biochips and microfluidic 3-D cell cultures. Consequently, the optical sensing is typically carried out in the transmission mode, with the optical signals transmitted through the optical fibers to a detector. Microwell arrays at the distal end of imaging fiber, with molecular targets captured and transported to the microwells by microbeads, are the most popular among these optical fiber arrays. Although detection limit better than 1 × 10⁶ molecules per bead has been reported, the bar-coded beads limit the target number of this platform.^{7, 8}Our previous work^{9, 10} has shown that plasmonics at nanotips can enhance local electric field by three orders of magnitude. However, conduction loss and quenching of fluorescence^{11, 12} by the metal substrates limit the use of plasmonic enhanced fluorescence for large-library assays. Only nano-molar sensitivity has been demonstrated using plasmonics from metal coated nanocone tips.^{13, 14} In this paper, we will extend the conical fiber array platform not by tip plasmonics but by another optical phenomenon with induced dipoles: singular scattering off dielectric wedges and tips.¹⁵ Instead of the surface plasmon resonance on metallic nanostructures,¹⁶ field focusing at the cone tip by the dielectric media (the silica fiber) is used to produce a localized and singularly large scattering intensity at the tip. Singular scattering from a wedge or a cone has been known for decades.^{17, 18} It is only recently that numerical simulation¹⁹ has revealed that field focusing by this singular scattering can effect a five-order intensity enhancement that is frequency independent. This intense tip scattering produces a local light source at the tip that does not suffer from conduction loss. Unlike plasmonic metal nanostructures, the dielectric tip would also not quench the fluorescent reporters excited by the light source. In fact, it will help scatter the fluorescent signal, with Rayleigh scattering intensity scaling with respect to wavelength. We hence utilize this phenomenon for diffraction-limit fluorescent sensing/imaging for the first time here.The local light source due to tip scattering minimizes background auto-fluorescence and scattering noise, provided the target molecules preferentially diffuse towards the dielectric vertices. If the targets do not preferentially hybridize with probes at the vertices, there would be significant target loss, with a concomitant loss in sensitivity, because the vertex regions are just a small fraction of the total area. Fortunately, like electromagnetic radiation at the electrostatic limit of the Maxwell equations for sharp (sub-wavelength) vertices,²⁰ the steady-state diffusion of molecules also obey the Laplace equation and so do the DC or AC electric potentials that drive electrophoresis and dielectrophoresis of the molecules.²¹ Hence, the diffusive, electrophoretic, and dielectrophoretic fluxes of target molecules are also singularly large at the vertices and there will be preferential hybridization there until the tip is saturated. Previously, we have demonstrated preferential diffusive transport of colloids to channel corners²² and dieletrophoretic trapping of bacteria²³ and DNA molecules²⁴ around sharp nanostructures like carbon nanotubes. Hence, dielectric nanotips fabricated by low-cost techniques can potentially provide the smallest fluorescent spot, which can preferentially capture target molecules and whose fluorescent image is limited in size only by the diffraction limit, without a confocal configuration.Although the scattering singularity is stronger at the conic tip, the total increase in scattering area of this singularity of measure zero is not as high as that of a sharp wedge, thus rendering the signal relatively weak. We hence employ a well-defined multi-wedged silica cone fabricated by wet-etching, with the wedges introduced by non-uniform stress formed during the fiber assembly process, to produce maximum scattering at the tip where three to four wedges converge (see inset of Fig. ​Fig.1A).1A). Using the reflection mode to fully exploit this singular scattering to excite fluorescent reporters at the tip and transmit the resulting signal, we report a nanocone array that can detect down to 100 molecules per cone tip with a large dynamic range from femtomolar to nanomolar concentrations. Although quantification for a single target is reported in this preliminary report, multi-target assays can readily be developed.Open in a separate windowFigure 1(A) A SEM image of the silica cone array where the single cone inset image shows three wedges converging into a 10 nm junction at the tip. (B) The optical setup of measurement. (C) The diffraction-limited fluorescent spot images.Amine-modified 35-base oligo-probes were functionalized onto both unetched silica fibers (as a control) and etched conic silica tips. The sample of 35-base ssDNA targets (corresponding to a primer for a segment of the Serotype 2 dengue genome) with a 5′ tagged Cy3 fluorophore was inserted into a microfluidic chip housing the fiber bundle (Fig. ​(Fig.1B)1B) and left overnight (see the supplementary material²⁵ for exact sequence). After a standard rinsing protocol, fluorescent images were taken with an Olympus IX-71 fluorescent microscope for target concentrations ranging from 1 fM to 1 nM. A typical fluorescent image after hybridization is shown in Fig. ​Fig.1c,1c, where each micron-sized bright spot corresponds to a single tip in the cone array. The intensity profile shown in the supplementary material²⁵ indicates a fluorescent spot smaller than 1 μm, indicating that the fluorescent light source is sub-wavelength and the resolution is close to diffraction limit. The size of this bright spot at the conic tip does not vary much with respect to the concentration but its intensity does, as shown in Fig. ​Fig.2A.2A. It was found that for flat fibers, only concentrations higher than 1 nM produced significant signals above the background. However, for etched conic fibers, 10 fM is clearly distinguishable from the background, which indicates that an improvement of sensitivity up to five orders can be realized by simply etching the flat surface into cone arrays. It also suggests very little target loss due to preferential hybridization onto the cone at sub-nM concentrations. We estimated the number of molecules per cone from the total number of molecules in target solution divided by the number of pixels on each fiber (10⁴), which suggests less than 100 molecules per cone for a 10 fM bulk concentration, four orders better than any existing technology.Open in a separate windowFigure 2(A) Fluorescent intensity of etched conic fiber and unetched fiber for different concentrations of target molecules from 1 fM to 1 nM. (B) Fluorescent intensity increases linearly with exposure time. Non-target molecules with 1 μM concentration do not produce significant signal compared to lower concentrations of target molecules such as 1 nM and 10 nM (see the supplementary material²⁵ for details of image analysis).Selectivity of the platform was also examined. Fig. ​Fig.2B2B presents the fluorescent intensity of the tips for non-target (1 μM) and target (1 nM and 10 nM) at different exposure times, which shows that fluorescent intensity increases linearly with exposure time. Beyond 5 s, saturation of images prevents further increase in the signal. For non-target, the intensity is much lower than 1 nM Target and 10 nM Target, which means non-target do not bind to the probes at the wedged tip as effectively as target molecules. Non-specific binding can be further removed by using more stringent buffers and higher flow rates.²⁶ This platform can be extended to detect 70 000 targets, in theory, by functionalizing different probes onto each cones using localized photochemistry via masking, micro-mirror directed illumination, or direct laser writing. Extension to ELISA type protein assays is also straight forward. Integration of a transmission-mode optical fiber endoscope into a microfluidic biochip and into a 3-D cell culture for real-time monitoring of multiple molecular targets at near-single molecule resolution is currently underway.     

Rare cell isolation and profiling on a hybrid magnetic/size-sorting chip

We present a hybrid magnetic/size-sorting (HMSS) chip for isolation and molecular analyses of circulating tumor cells (CTCs). The chip employs both negative and positive cell selection in order to provide high throughput, unbiased CTC enrichment. Specifically, the system utilizes a self-assembled magnet to generate high magnetic forces and a weir-style structure for cell sorting. The resulting device thus can perform multiple functions, including magnetic depletion, size-selective cell capture, and on-chip molecular staining. With such capacities, the HMSS device allowed one-step CTC isolation and single cell detection from whole blood, tested with spiked cancer cells. The system further facilitated the study of individual CTCs for heterogeneity in molecular marker expression.Circulating tumor cells (CTCs) have emerged as an important biomarker in clinical practice as well as in fundamental research.^{1, 2} CTCs, shed from primary tumors, have been shown to be an early harbinger of tumor expansion and metastasis³ and have been used to predict disease progression, response to treatment, relapse, and overall survival.^{4, 5, 6} Recent work has shown that CTCs display distinct proteomic and genetic profiles; for example, CTCs in pancreatic cancer, have increased RNA expression of Wnt, implicating this pathway in metastasis.⁷ Proteomic characterization of proliferative markers such as Ki-67, and hormonal markers such as androgen receptor in prostate cancer, also have been shown to be predictive of treatment outcome.^{8, 9}Despite such clinical potential of CTCs, their routine detection and characterization still remains a significant technical challenge.¹⁰ The task requires screening of a large number of cells (e.g., > 10⁷ cells in 10 ml blood) and enrichment of heterogeneous targets against a complex biological background. Two main methods of CTC isolation are typically used: positive and negative selection. In positive selection, CTCs are directly isolated from blood via size-based filtration^{11, 12, 13, 14, 15, 16, 17, 18, 19, 20} or antibody-based capture.^{1, 8, 21} Negative depletion reduces abundant blood cells, often by immunomagnetic separation, for downstream CTC enrichment.²² Both approaches have been used for high throughput CTC isolation from whole blood (SI Table 1).²³ Each method, however, has its own inherent limitations. Positive enrichment could be biased by its selection criteria (e.g., cell size and cell surface markers). Negative selection, albeit unbiased, often requires complex sample processing (e.g., multiple washing steps for CTC isolation) that could result in cell loss.We hypothesized that both positive and negative selection could be combined in a single platform to enable (1) highly efficient and unbiased CTC purification, and (2) in-situ molecular analyses of collected cells. As a proof-of-concept, we herein describe a hybrid magnetic/size-sorting (HMSS) system that integrates magnetic and size-based isolation into a compact microfluidic chip. The HMSS first uses a magnetic filter to deplete leukocytes through immunomagnetic capture. Samples then pass through a size-sorter region that traps individual cells at predefined locations. Since abundant leukocytes are removed by the magnetic filter, the size-sorter could have a low size cut-off (∼5 μm), which allows for the unbiased capture of even small cancer cells. Furthermore, molecular probes can be introduced to perform on-chip, multiplexed analyses at single-cell resolution. We evaluated the utility of the developed system by capturing and profiling tumor cells in whole blood. The HMSS offers the advantages of both negative and positive selection and thereby differs from the recently reported iChip system²⁴ which can operate only in either a negative or a positive selection mode.     

Plasmonic hotspots of dynamically assembled nanoparticles in nanocapillaries: Towards a micro ribonucleic acid profiling platform

Plasmonic hot spots, generated by controlled 20-nm Au nanoparticle (NP) assembly, are shown to suppress fluorescent quenching effects of metal NPs, such that hair-pin FRET (Fluorescence resonance energy transfer) probes can achieve label-free ultra-sensitive quantification. The micron-sized assembly is a result of intense induced NP dipoles by focused electric fields through conic nanocapillaries. The efficient NP aggregate antenna and the voltage-tunable NP spacing for optimizing hot spot intensity endow ultra-sensitivity and large dynamic range (fM to pM). The large shear forces during assembly allow high selectivity (2-mismatch discrimination) and rapid detection (15 min) for a DNA mimic of microRNA.Irregular expressions of a panel of microRNAs (miRNA) in blood and other physiological fluids may allow early diagnosis of many diseases, including cancer and cardiovascular diseases.¹ However, quantifying all relevant miRNAs (out of 1000), with similar sequences over 22 bases² and large variations in expression level (as much as 100 fold) at small copy numbers, requires a new molecular diagnostic platform with high-sensitivity, high-selectivity, and large dynamic range. Current techniques for miRNA profiling, such as Northern blotting,³ microarray-based hybridization,⁴ and real-time quantitative polymerase chain reaction⁵ are expensive and complex. A simple and rapid miRNA array would allow broad distribution of molecular diagnostic devices for cancer and chronic diseases, eventually into homes for frequent prescreening of many diseases.At their low concentrations in untreated samples, optical sensing of miRNA is most promising. Plasmonically excited Raman scattering (SERS) and fluorescence sensors from metallic nanoparticles (NPs) or surfaces have enhanced the sensitivity of optical molecular sensors by orders of magnitude.^{6, 7, 8, 9} However, probe-less SERS sensing or fluorescent sensing of unlabeled targets are insufficiently specific for miRNA targets in heterogeneous samples. Plasmonic detection is also very compatible with FRET probes whose donor dye offers small light sources to excite fluorescently labelled targets upon hybridization.^{7, 10}A particular family of FRET reporters does offer label-free sensing: hairpin oligo probes whose end-tagged fluorophores are quenched by the Au NP to which they are functionalized.¹¹ The fluorescent signal is only detected when the hairpin is broken by the hybridizing target nucleic acid or protein (for an aptamer probe), and the more rigid paired segment separates the end fluorophore from the quenching surface to produce a fluorescent signal. It is often hoped that plasmonics on the metal surface will enhance the intensity to overcome the quenching effect, if the linearized hairpin is within the NP plasmonic penetration length. However, since fluorescent quenching decays slowly (linearly) with fluorophore-metal spacing¹⁰ whereas the plasmonic intensity decays exponentially from a flat surface, careful experimentation shows that quenching dominates and the hairpin probe actually produces a larger intensity on non-metallic surfaces,¹⁰ on which it can not function as a label-free probe. Hence, only μM limit-of-detection (LOD) has been achieved with this technique on single NPs or on flat metal surfaces,¹² with expensive laser excitation and confocal detection.Plamonic hot spots formed between metal nanostructures and sharp nanocones can further amplify the plasmonic field.^{13, 14} The hot spot intensity decays algebraically with respect to the separation or cone tip distance and hence should dominate the linear decay of the metal quenching effect at some optimum separation.¹⁵ It is hence possible that plasmonic hot spots may allow much lower LOD with inexpensive optical instruments—ideally light-emitting diode light source and miniature camera. However, the dimension of the gaps, cones, and wedges needs to be at nanoscale, and the cost is now transferred to fabrication of such hot-spot substrates like bow-ties, double crescents, bull-eyes, etc.¹⁶ Low-cost wet-etching techniques for addressable nanocones that sustain converging plasmonic hot spots¹⁷ have been reported but the fabricated nanocones are often too non-uniform to allow precise quantification. NP monolayers have been shown to exhibit plasmonic hot spots and fluorescence enhancement.^{18, 19} However, the enhancement only occurs within a range of spacing between aggregated NPs, which is difficult to control and the location or even the existence of the hotspots are not known a priori.Higher sensitivity is expected if a minimum number of NPs are used in an assembly at a known location and if the NP assembly can produce crystal-like aggregates with controllable NP spacing. Induced DC and AC NP dipoles (related to dielectrophoresis) have been used to assemble NP crystals by embedded micro-electrodes to provide the requisite high field.^{20, 21} The resulting NP crystals are ideal for plasmonic hot spots, since the spacing of the regimented NP crystal can be controlled by the applied voltage. Conic nanocapillaries^{22, 23} will be used here for such field-induced NP assembly because the submicron-tip can focus the electric field into sufficient high intensity for NP assembly without embedded-electrodes. Because the field is highest at the tip due to field focusing, the micron-sized crystal would be confined to a small volume, which will be shown to be less than typical confocal volumes, at a known location. So long as the hotspots are regimented, the quantification of target molecules is determined by the total fluorescent intensity and is hence insensitive to the exact geometry of the nanocapillary.Fluorescent microscope equipped with tungsten lamp light source and normal CCD camera from Q Imaging were used for simultaneous optical and ion current measurements, as shown in Fig. ​Fig.1a.1a. The nanocapillaries were pulled from commercial glass capillaries using laser-assisted capillary puller. SEM image of a typical pulled glass nanocapillary in Fig. ​Fig.1b1b shows an inner diameter of 111 nm and cone angle of 7.3°. The capillary was inserted into a Polydimethylsiloxane chip with two reservoirs. The 20 nm Au NPs, functionalized with fluorescently labelled dsDNA, were injected into the base reservoir. With SEM imaging (Fig. S3 in the supplementary material²⁴), the functionalized DNA is found to prevent NP aggregation even in high ionic-strength Phosphate buffered saline buffer. The NP solution is then driven into the capillary through the tip by applying a positive voltage. Fig. ​Fig.1c1c shows the ion current evolution over 2 h at +1 V packing voltage. The ion current increases rapidly in the first 10 min, then at a much slower rate. The rise of current indicates assembly of conductive Au NP assembly at the tip. This was confirmed by the strong fluorescence signal at the tip region during the packing process (inset of Fig. ​Fig.1c).1c). The one-micron region (corresponding to roughly an aggregate volume of one attoliter) near the capillary tip shows a fluorescence signal after 1 min and also appeared as a dark spot in the transmission image (supplementary material, Fig. S1²⁴). This spot darkens with longer packing time but does not grow in size, consistent with the monotonically increasing ion current with increased packing density of the NP assembly. As contrast, a strong fluorescence appeared after only 1 min of packing, but the signal became weaker after 15 min (supplementary material, Fig. S1²⁴). This reduction in fluorescence is not due to bleaching of fluorophores because we took 2 images in 15 min at 5 s exposure each and control experiments show significant bleaching only beyond an exposure time of 100 s (see supplementary material).²⁴ Instead, the non-monotonic dependence of the fluorescence intensity with respect to time is because of the optimal hotspot spacing for highest plasmonic intensity at about 5–20 nm,^{25, 26, 27} which is reached at about 10 min.Open in a separate windowFigure 1Plasmonic hotspots generated at the tip of a nano-capillary. (a) Schematic of the experimental set up. (b) SEM image of glass nanocapillary shows opening at the tip with a diameter of 111 nm. (c) Current evolution during packing of fluorescently labeled gold particles at +1 V. Inset shows strong fluorescence only after 1 min of packing.The FRET probe is designed to exploit the plasmonic hotspot.²⁴ We first electrophoretically drove the target molecules in the tip side reservoir into the nano-capillary by applying a negative voltage of −1 V. During this process, the targets are trapped within the capillary and hybridize with the hairpin probes on the Au NP in the nanocapillary. Fluorescence of the unquenched hybridized probes is too weak to be detected by our detector as shown in Fig. ​Fig.2b.2b. A reverse positive voltage of +1 V was then applied to the capillary to pack the Au NPs to the tip. Due to plasmonic hot spots of aggregated gold nanoparticles, the fluorescence signal is significantly enhanced at the tip and can be detected by our CCD camera, as shown in Fig. ​Fig.2c2c.Open in a separate windowFigure 2(a) Schematics of designed hairpin probe on gold particle. (b) Before packing gold particles, probe fluorescence signal was too weak to be detect. (c) After packing for 3 minutes, a strong fluorescence signal appears at the NP aggregate. (d) Normalized intensity (average of all pixels above a threshold (15 au) normalized with respect to the average over all pixels (with 0-250 au)) as a function of packing voltage for different samples. Black, 1 nM target ; blue, 10 pM target; purple, 10 nM 2-mismatch non-target. (e) Intensity dependence on target concentration. Measured normalized intensity before packing (black) and after packing (red), for three independent experiments with different nano-capillaries at each concentration. NT stands for non-target at 10 nM as a reference.For the same packing time, the fluorescence intensity increases initially but saturates after 10 min time of trapping (supplementary material, Fig. S2(a)²⁴). Over 10 min of trapping with a negative voltage, we found the fluorescence intensity exhibits a maximum at a packing time of 3 min (supplementary material, Fig. S2(b)²⁴). In later experiments, we used 10 min trapping time and 3 min packing time as standards.Fig. ​Fig.2d2d shows the fluorescence intensity is sensitive to the positive packing voltage at different concentration of target and non-target molecules. For target samples (1 nM and 10pM), the optimal voltage is about 1 V. We suspect that with larger voltage, the NPs are packed too tightly such that the NP spacing is smaller than the optimal distance for plasmonic hotspots. The fluorescence intensity for a nontarget with two mismatches is 7 times lower than the target even with a 10 times higher concentration (10 nM). Moreover, the optimal voltage for the non-target miRNA is reduced to 0.5 V instead 1 V for the target miRNA. Strong shear during electrophoretic packing has probably endowed this high selectivity.²⁰Using the protocol above, the LOD and dynamic range of the target was determined (Fig. ​(Fig.2e).2e). The intensity at each concentration is measured with three independent experiments with different nanocapillaries to verify insensitivity with respect to the nanocapillary. The intensity increases monotonically with respect to the concentration from 1fM to 1pM. Beyond 1pM, the fluorescence signal saturates, presumably because all hairpin probes at the tip have been hybridized. At 1 fM, the fluorescent intensity is still well above the background measured from the non-target sample. Note both auto-fluorescence of gold nanoparticles and free diffusing non-target DNA molecules contribute to the background. Given the volume of tip side reservoir (∼50 μl), there are about 30 000 target molecules in the reservoir at 1 fM. However, with a short 10 min trapping time, we estimate only a small fraction of these molecules, less than 100, have been transferred from the tip reservoir into the nanocapillary.     

Syringe-vacuum microfluidics: A portable technique to create monodisperse emulsions

We present a simple method for creating monodisperse emulsions with microfluidic devices. Unlike conventional approaches that require bulky pumps, control computers, and expertise with device physics to operate devices, our method requires only the microfluidic device and a hand-operated syringe. The fluids needed for the emulsion are loaded into the device inlets, while the syringe is used to create a vacuum at the device outlet; this sucks the fluids through the channels, generating the drops. By controlling the hydrodynamic resistances of the channels using hydrodynamic resistors and valves, we are able to control the properties of the drops. This provides a simple and highly portable method for creating monodisperse emulsions.Droplet-based microfluidic devices use micron-scale drops as “test tubes” for biological reactions.^{1, 2, 3} With the devices, the drops are loaded with cells, incubated to stimulate cell growth, picoinjected to introduce additional reagents, and sorted to extract rare specimens.^{4, 5, 6} This allows biological reactions to be performed with greatly enhanced speed and efficiency over conventional approaches: by reducing the drop volume, only picoliters of reagent are needed per reaction, while through the use of microfluidics, the reactions can be executed at rates exceeding hundreds of kilohertz. This combination of incredible speed and efficient reagent usage is attractive for a variety of applications in biology, particularly those that require high-throughput processing of reactions, including cell screening, directed evolution, and nucleic acid analysis.^{7, 8} The same advantages of speed and efficiency would also be beneficial for applications in the field, in which the amount of material available for testing is limited, and results are needed with short turnaround. However, a challenge to using these techniques in field applications is that the control systems developed to operate the devices are intended for use in the laboratory: to inject fluids, mechanical pumps are needed, while computers must adjust flow rates to maintain optimal conditions in the device.^{9, 10, 11, 12} In addition to significantly limiting the portability of the system, these qualities make them impractical for use outside the laboratory. For droplet-based microfluidic techniques to be useful for applications in the field, a general, robust, and portable system for operating them is needed.In this paper, we introduce a general, robust, and portable system for operating droplet-based microfluidic devices. In this system, which we call syringe-vacuum microfluidics (SVM), we load the reagents needed for the emulsion into the inlets of a microfluidic drop maker; using a standard plastic syringe, we generate a vacuum at the outlet of the drop maker,¹³ sucking the reagents through the channels, generating drops, and transporting them to different regions for visualization and analysis. By controlling the vacuum strength and channel resistances using hydrodynamic resistors^{14, 15, 16} and single-layer membrane valves,^{17, 18} we are able to specify the flow rates in different regions of the device and to adjust them in real time. No pumps, control computers, or electricity is needed for these operations, making the entire system portable and of potential use for field applications. To characterize the adjustability and precision of this system, we vary channel resistances and vacuum pressures while measuring the effects on drop size and production frequency. We also show how to use this to form drops of many distinct reagents simultaneously using only a single vacuum syringe.Monodisperse drop formation is the central operation in droplet-based microfluidics but can be quite challenging due to the need for precise, steady pumping of reagents; forming monodisperse drops with controlled properties is thus a stringent demonstration of the effectiveness of a control system. While there are many geometries available for microfluidic drop formation,¹⁹ in this discussion we use a simple cross-junction for its proven ability to form uniform emulsions at high rates of speed,^{20, 21} a schematic of which is shown in Fig. ​Fig.1.1. The devices are fabricated in poly(dimethylsiloxane) (PDMS) using soft lithography.²² The drop formation channels have dimensions of 25 μm in width and 25 μm in height. To enable production of aqueous drops in oil, which are the most useful for biological assays, we require hydrophobic devices, which we achieve using an Aquapel chemical treatment: we flow Aqualpel through the channels for a few seconds, flush with air, and then bake the devices for 20 min at 65 °C. After this treatment, the channels are permanently hydrophilic, as is needed for forming aqueous-in-oil emulsions. To introduce reagents into the device, we use 200 μl plastic pipette tips inserted into the channel inlets. To apply the suction, we use a 10 ml Bectin-Dickenson plastic syringe coupled to the device through a 16 G needle and PE∕5 tubing. The other end of the tubing is inserted into the outlet of the device.Open in a separate windowFigure 1Schematic of the microfluidic drop maker for use with SVM. To form water drops in oil, the device must be hydrophobic, which we achieve by treating the channels with Aquapel. The water and surfactant-containing oil are loaded into pipette tips inserted into the device inlets at the locations indicated. To pump the fluids through the drop maker, a syringe applies a vacuum to the outlet; this sucks the fluids through the drop maker, forming drops. The drops are collected into the suction syringe, where they can be stored, incubated, and reintroduced into a microfluidic device for additional processing.To begin forming drops, we fill the device with HFE-7500 fluorocarbon oil, displacing trapped air bubbles that could restrict flow and interfere with drop formation. Pipette tips containing reagents are then inserted into the device inlets, as shown in Fig. ​Fig.11 and pictured in Fig. ​Fig.2a;2a; during this step, care must be taken to not trap air bubbles under the pipette tips, as they would restrict flow. For the fluids, we use distilled water for the droplet phase and HFE-7500 with the ammonium salt of Krytox 157 FSL at 1.8 wt % for the continuous phase. The suction syringe is then connected to the device outlet; to initiate drop formation, the piston is pulled outward and locked in place with a 1 in. binder clip, as shown in Fig. ​Fig.2a.2a. This expands the air in the syringe, generating a vacuum that is transferred to the device through tubing. Since the inlet reagents are open to the atmosphere and thus maintained at a pressure of 1 atm, this creates a pressure differential through the device that pumps the fluids. As the fluids flow through the cross-channel, forces are generated that create drops, as shown in Fig. ​Fig.2b2b (enhanced online). Due to the very steady flow, the drops are highly monodisperse, as shown in Fig. ​Fig.2c.2c. After they are formed, the drops flow out of the device through the suction tube and are collected into the syringe. Depending on the emulsion formulation, drops may coalesce on the metal needle of the syringe; if so, an Upchurch fitting should be used to couple the tubing instead. The collected drops can be stored in the syringe, incubated, and reintroduced into additional microfluidic devices, as needed for the assay.Open in a separate windowFigure 2Photograph of the microfluidic drop formation device with pipette tips containing emulsion reagents and vacuum syringe for pumping (a). Distilled water is used for the droplet phase and HFE-7500 fluorocarbon oil with fluorinated surfactant for the continuous phase. The vacuum applies a pressure differential through the device that pumps the fluids through the drop maker (b) forming drops. The drops are monodisperse, due to the controlled properties of drop formation in microfluidics (c). The scale bars denote 50 μm (enhanced online).In many biological applications, drop size must be precisely controlled. This is essential, for example, when encapsulating molecules or cells in the drops, in which the number encapsulated depends on the drop size.^{3, 23, 24} With SVM, the drop size can be precisely controlled. Our strategy to accomplish this is motivated by the physics of microfluidic drop formation. In microfluidic devices, the capillary number of the flow is normally small, Ca<0.1; as a consequence, the drop formation physics follows a plugging∕squeezing mechanism, in which the drop size depends on the flow rate ratio of the dispersed-to-continuous phase.^{20, 25} By adjusting this ratio, we can thus control the drop size. To adjust this ratio, we use hydrodynamic resistor channels.^{14, 15, 16} These channels are analogous to electronic resistors in that for a fixed pressure drop (voltage) the flow rate through them (current) is inversely proportional to their resistance. By making the resistors longer or shorter, we adjust their resistance, thereby controlling the flow rate.To use resistors to control the drop size, we place three on the inlets of the cross-junction, at the locations indicated in Fig. ​Fig.3a.3a. In this configuration, the flow rate ratio depends on the resistances of the central and side resistors: shortening the side resistors increases the continuous phase flow rate with respect to the dispersed phase, thereby reducing the ratio and, consequently, the drop size, whereas lengthening it increases the drop size. By varying the ratio, we produce drops over a range of sizes, as shown in Fig. ​Fig.3b3b (enhanced online). The drop size is linear in the resistance ratio, indicating that it is linear in the flow rate ratio, as is expected for plugging∕squeezing drop formation [Fig. ​[Fig.3b3b].^{20, 25} This behavior is identical to that of pump-driven fluidics, demonstrating that SVM affords similar control.Open in a separate windowFigure 3Drop properties can be controlled using resistor channels. The resistors are placed on the inlets of the drop maker at the locations indicated in (a). The resistors enable the flow rates of the inner and continuous phases to be controlled. By varying the length ratio of the inlet resistors, we control the flow rate ratio in the drop maker. This allows the drop volume to be controlled, as shown by drop volume plotted as a function of inlet resistor length ratio in (b); varying this ratio does not significantly affect the drop formation frequency, as shown in (c). By varying the length of the outlet resistor, we control the total flow rate through the device; this allows us to form drops of constant volume, but at a different formation frequency, as shown by the plots of volume and frequency as a function of the inverse of the outlet resistor length in (d) and (e), respectively. The measured hydrodynamic resistance of a resistor channel with water as a function of length is shown as inset into (d) (enhanced online).We can also control the frequency of the drop formation using resistor channels. We place a resistor on the outlet of the device; this sets the total flow rate through the device, thereby adjusting drop frequency, as shown in Fig. ​Fig.3e3e (enhanced online). To confirm that the size and frequency control are independent, we plot size as a function of the outlet resistance and frequency as a function of the resistance ratio [Figs. ​[Figs.3c,3c, ​,3d];3d]; both are constant as a function of these parameters, again demonstrating independent control. Frequency can also be adjusted by changing the strength of the vacuum, which can be accomplished by loading a prescribed volume of air into the syringe before expansion. In this case, the vacuum pressure applied is P_fin=V_in∕V_fin×P_in, where V_in is the initial volume of air in the syringe, V_fin is the volume after expansion, and P_in is the initial pressure, which is 1 atm. By loading a prescribed volume of air into the syringe before connecting it to the device and pulling the piston, the expansion factor can be reduced, thereby lowering the vacuum strength.The flow rates through the microfluidic device depend on the applied pressure differential, which, in turn, depends on the value of the ambient pressure. Since ambient pressure may vary due to differences in altitude, the drop formation may also vary. However, since ambient pressure variations affect the inner and outer phase flows equally, this should alter the total flow rate but not the flow rate ratio. Consequently, we expect it to alter drop formation frequency but not drop size because while the frequency depends on absolute flow rate [as illustrated by Fig. ​Fig.3e],3e], drop size depends on the flow rate ratio [as illustrated in Fig. ​Fig.3b].3b]. Based on normal variations in atmospheric pressure on the surface of the Earth, we expect this to produce differences in the drop formation frequency of ∼25%, for example, when operating a device at sea level compared to at the top of a moderately sized mountain.Resistor channels allow drop properties to be controlled, equivalent to what is possible with pump-driven flow; however, they do not allow real-time control because their dimensions are fixed during the fabrication. Real-time control is often needed, for example, as it is when performing reactions in drops for the first time, in which the optimal drop size is not known. To enable real-time control, we must adjust flow rates, which can be achieved using the fluidic analog of electronic potentiometers. Single-layer membrane valves are analogous fluidic components, consisting of a control channel that abuts a flow channel.^{17, 18} By pressurizing the control channel, the thin PDMS membrane between these channels is deflected laterally, constricting the flow channel, thereby increasing its hydrodynamic resistance and reducing its flow rate.¹⁸ To use these membrane valves to vary drop size, we replace the inlet resistors with inlet valves, as shown in Fig. ​Fig.4a.4a. To set the flow rate through a path, we actuate the valve with a defined pressure. To actuate the valves, we use air-filled syringes: a 1 ml syringe is filled with air and connected to the valve control channel through tubing; an additional component, a three-way stopcock is inserted between the syringe and needle, allowing the pressure to be locked in after optimal actuation conditions are obtained. We use one syringe to control the dispersed phase valves and another to control the continuous phase valves. The valves are pressurized by compressing the air in the syringes to a defined degree using the marked graduations; this is achieved by pressing the piston to a defined graduation mark, compressing the air contained within it, thus increasing pressure. The stopcock is then switched to the off position, locking in the actuation. This simple scheme allows precise actuation of the valves, for accurate, defined flow rates in the drop maker, and controlled drop size, as shown in Figs. ​Figs.4b,4b, ​,4c4c (enhanced online). The drop size can be varied at a rate of several hertz without noticeable loss of control; moreover, changing the drop size does not affect the frequency, indicating that, again, these properties are independent, as shown by the constant drop frequency with varying pressure ratio in Fig. ​Fig.4d4d.Open in a separate windowFigure 4Single-layer membrane valves allow the drop size to be varied in real time to screen for optimal reaction conditions. The valves are positioned on the inner and side inlets, as indicated in (a). By adjusting the actuation pressures of the valves, we vary the flow rates in the drop maker, thereby changing the drop size (b), as shown by the plot of drop volume as a function of the actuation pressure ratio in (c). Varying the inlet resistance ratio does not significantly alter drop formation frequency, as shown by frequency as a function of the pressure ratio in (d). A movie of drop formation during actuation of the valves are available in the supplemental material (Ref. ²⁹). The scale bars denote 100 μm (enhanced online).Another useful attribute of SVM is that it readily lends itself to parallel drop formation²⁶ because the pressure that pumps the fluids through the channels is supplied by the atmosphere and is applied evenly over the whole outer surface of the device. This allows fluids to be introduced at equal pressures from different inlets, for forming drops with identical properties in different drop makers. To illustrate this, we use a parallel drop formation device to emulsify eight distinct reagents simultaneously; the product of this is an emulsion library, consisting of drops of identical size in which different drops encapsulate distinct reagents, useful for certain biological applications of droplet-based microfluidics.⁷ The microfluidic device consists of eight T-junction drop makers.²⁵ The drop makers share one oil inlet and outlet but each has its own inner-phase inlet, as shown in Fig. ​Fig.5.5. The oil and outlet channels are wide, ensuring negligible pressure drop through them, so that all T-junctions are operated under the same flow conditions. A distinct reagent fluid is introduced into the inner phase of each T-junction, for which we use eight concentrations of the dye Alexa Fluor 680 in water. After loading these solutions into the device through pipette tips, a syringe applies the vacuum to the outlet, sucking the reagents through the T-junctions, forming drops, as shown by the magnified images of the T-junctions during drop formation in Fig. ​Fig.5.5. Since the drop makers are identical and operated under the same flow conditions, the drops formed are of the same size, as shown in the magnified images in Fig. ​Fig.55 and in a movie available in the supplemental material.²⁹Open in a separate windowFigure 5Parallel drop formation device consisting of eight T-junction drop makers. The drop makers share a common oil inlet and outlet, both of which are wide to ensure even pressure distribution to all drop makers; support posts prevent these channels from collapsing under the suction. Each drop maker has its own inner-phase inlet, allowing emulsification of a distinct reagent. Since the drop maker dimensions and pressure differentials are constant through all drop makers, the drops formed are of the same size, as shown in the magnified images. The drops are ∼35 μm in diameter.To verify that the dye solutions are successfully encapsulated, we image a sample of the collected drops with a fluorescent microscope. The drops are confined in a monolayer between two glass plates so they can be individually imaged. They are of the same size but have distinct fluorescence intensities, as shown in Fig. ​Fig.6a.6a. To quantify these differences, we measure the intensity of each drop and plot the results as a histogram [see Fig. ​Fig.6b].6b]. There are eight peaks in the histogram, corresponding to the eight dye concentrations, demonstrating that all dyes are encapsulated successfully. The peak areas are also similar, demonstrating that drops of different types are formed in equal amounts due to the uniformity of the parallel drop formation.Open in a separate windowFigure 6Fluorescent microscope image of emulsion library created with parallel T-junction device (a). In this demonstration, eight concentrations of Alexa Fluor 680 dye are emulsified simultaneously, producing an emulsion library of eight elements. The drops are of the same size but encapsulate distinct concentrations of the dye solution, as demonstrated by the eight peaks in the intensity histograms in (b). The scale bar denotes 100 μm.SVM is a simple, accessible, and highly controlled way to form monodisperse emulsions for biological assays. It allows controlled amounts of different reagents to be encapsulated in individual drops, drop size to be precisely controlled, and the ability to form drops of different reagents at the same time, in a parallel drop formation device. These properties should make SVM useful for biological applications of monodisperse emulsions;^{1, 2, 3} the portability of SVM should also make it useful for applications in the field, particularly when no electrical power source is available. The parallel emulsification technique should also be useful for particle templating from drops, in which the particles must be of the same size but composed of distinct materials.^{26, 27, 28, 29}     

Enhancing conjugation rate of antibodies to carboxylates: Numerical modeling of conjugation kinetics in microfluidic channels and characterization of chemical over-exposure in conventional protocols by quartz crystal microbalance

This research reports an improved conjugation process for immobilization of antibodies on carboxyl ended self-assembled monolayers (SAMs). The kinetics of antibody/SAM binding in microfluidic heterogeneous immunoassays has been studied through numerical simulation and experiments. Through numerical simulations, the mass transport of reacting species, namely, antibodies and crosslinking reagent, is related to the available surface concentration of carboxyl ended SAMs in a microchannel. In the bulk flow, the mass transport equation (diffusion and convection) is coupled to the surface reaction between the antibodies and SAM. The model developed is employed to study the effect of the flow rate, conjugating reagents concentration, and height of the microchannel. Dimensionless groups, such as the Damköhler number, are used to compare the reaction and fluidic phenomena present and justify the kinetic trends observed. Based on the model predictions, the conventional conjugation protocol is modified to increase the yield of conjugation reaction. A quartz crystal microbalance device is implemented to examine the resulting surface density of antibodies. As a result, an increase in surface density from 321 ng/cm², in the conventional protocol, to 617 ng/cm² in the modified protocol is observed, which is quite promising for (bio-) sensing applications.Microfluidics have been implemented in various bio-medical diagnostic applications, such as immunosensors and molecular diagnostic devices.¹ In the last decade, a vast number of biochemical species has been detected by microfluidic-based immunosensors. Immunosensors are sensitive transducers which translate the antibody-antigen reaction to physical signals. The detection in an immunosensor is performed through immobilization of an antibody that is specific to the analyte of interest.² The antibody is often bound to the transducing surface of the sensor covered by self-assembled monolayers (SAMs). SAMs are organic materials that form a thin, packed and robust interface on the surface of noble metals like that of gold, suitable for biosensing applications.³ Thiolic SAMs have a “head” group that shows a high affinity to being chemisorbed onto a substrate, typically gold. The SAMs'' carboxylic functional group of the “tail” end can be linked to an amine terminal of an antibody to form a SAM/antibody conjugation.^3,4 The conjugation process is usually accomplished in the presence of carbodiimides, such as 1-ethyl-3-(3-dimethylaminopropyl) carbodiimide (EDC). A yield increasing additive, N-Hydroxysuccinimide (NHS), is often used to enhance the surface loading density of the antibody.^4,5A typical reaction for coupling the carboxylic acid groups of SAMs with the amine residue of antibodies in the presence of EDC/NHS is depicted in Figure ​Figure11.⁴ NHS promotes the generation of an active NHS ester (k₂ reaction path). The NHS ester is capable of efficient acylation of amines, including antibodies (k₃ reaction path). As a result, the amide bond formation reaction, which typically does not progress efficiently, can be enhanced using NHS as a catalyst.⁴Open in a separate windowFIG. 1.NHS catalyzed conjugation of antibodies to carboxylic-acid ended SAMs through EDC mediation (Adapted from G. T. Hermanson, Bioconjugate Techniques, 2nd. Edition. Copyright 2008 by Elsevier⁴). EDC reacts with the carboxylic acid and forms o-acylisourea, a highly reactive chemical that reacts with NHS and forms an NHS ester, which quickly reacts with an amine (i.e., antibody) to form an amide.A number of groups have studied EDC/NHS mediated conjugation reactions such as the ones depicted in Figure ​Figure1.1. The general stoichiometry of the reaction involves a carboxylic acid (SAM), an amine (antibody), and EDC to produce the final amide (antibody conjugated SAM) and urea. However, the recommended concentration ratio of the crosslinking reagents inside the buffer, i.e., the ratio of EDC and NHS with respect to adsorbates and each other, varies from one study to another.⁶ The kinetics of the reactions outlined in Figure ​Figure11 have also been investigated,^4,6–8 but only in the absence of NHS for EDC or carboxylic acids in aqueous solutions.⁸ A relatively recent experimental study verified the catalytic role of the yield-increasing reagent N-hydroxybenzotriazole (HOBt), which acts similarly to NHS.⁷ In this study, the amide formation rate (k₃ reaction path, Figure ​Figure1)1) was found to be dependent on the concentration of the carboxylic acid and EDC in the buffer solution, and independent of the amine and catalyst reagent concentration. The same group also showed that the amide bond formation kinetics is controlled by the reaction between the carboxylic acid and the EDC to give the O-acylisourea, which they marked as the rate-determining step (k₁ reaction path, Figure ​Figure11).The k₁ reaction path, or the conjugation reaction, is usually a lengthy process and takes between 1 and 3 h.^4,9 Compared to k₁, the k₂ and ?k₃ reactions are considerably faster. Microfluidics has the potential to enhance the kinetics of these reactions using the flow-through mode.^10,11 This improvement occurs because while conventional methods rely only on diffusion as the primary reagent transport mode, microfluidics adds convection to better replenish the reagents to the reaction surfaces. However, there are many fundamental fluidic and geometrical parameters that might affect the process time and reagents consumption in a microfluidics environment, such as concentration of antibodies and reagents, flow rate, channel height, and final surface density of antibodies. A model that studies the kinetics of conjugation reaction against all these parameters would therefore be helpful for the optimization of this enhanced kinetics.There are a number of reports on numerical examination of the kinetics of binding reactions in microfluidic immunoassays.^12–15 All these models developed so far couple the transport of reagents, by diffusion and convection, to the binding on the reaction surface. Myszka''s model assumes a spatially homogeneous constant concentration of reagents throughout the reaction chamber, thus fails to describe highly transport-limited conditions due to the presence of spatial heterogeneity and depletion of the bulk fluid from reagents.^16,17 In transport-limited conditions, the strength of reaction is superior to the rate of transport of reagents to the reaction surface.^18,19 More recently, the convection effects were included in a number of studies, describing the whole kinetic spectrum from reaction-limited conditions to transport-limited reactions.^20–22 Immunoreaction kinetics has also been examined with a variety of fluid driving forces, from capillary-driven flows,²⁰ to electrokinetic flows in micro-reaction patches,²¹ pressure-driven flows in a variety of geometric designs.²² Despite these comprehensive numerical investigations, the fundamental interrelations between the constitutive kinetic parameters, such as concentration, flow velocity, microchannel height, and antibody loading density, have not been studied in detail. In addition, the conjugation kinetics has not yet been exclusively examined.In this paper, a previous model for immunoreaction is modified to study the antibody/SAM conjugation reaction in a microfluidic system. Model findings are used to examine the process times recommended in the literature and possible modification scenarios are proposed. The new model connects the convective and diffusive transport of reagents in the bulk fluid to their surface reaction. The conjugation reaction is studied against fluidic and geometrical parameters such as flow rate, concentration, microchannel height and surface density of antibodies. Damköhler number is used to compare the reaction and fluidic phenomena present and justify the kinetic trends observed. Model predictions are discussed and the main finding on possible overexposure of carboxylates to crosslinking reagents, in conventional protocols, is verified by comparing the resultant antibody loading densities obtained using a quartz crystal microbalance (QCM) set up. The results demonstrate an improved receptor (antibody) loading density which is quite promising for a number of (bio-) sensing applications.^23,24 Major application areas include antibody-based sensors for on-site, rapid, and sensitive analysis of pathogens such as Bacillus anthracis,²³ Escherichia coli, and Listeria monocytogenes, and toxins such as fungal pathogens, viruses, mycotoxins, marine toxins, and parasites.²⁴     

Polyphosphonium‐based bipolar membranes for rectification of ionic currents

Bipolar membranes (BMs) have interesting applications within the field of bioelectronics, as they may be used to create non-linear ionic components (e.g., ion diodes and transistors), thereby extending the functionality of, otherwise linear, electrophoretic drug delivery devices. However, BM based diodes suffer from a number of limitations, such as narrow voltage operation range and/or high hysteresis. In this work, we circumvent these problems by using a novel polyphosphonium-based BM, which is shown to exhibit improved diode characteristics. We believe that this new type of BM diode will be useful for creating complex addressable ionic circuits for delivery of charged biomolecules.Combined electronic and ionic conduction makes organic electronic materials well suited for bioelectronics applications as a technological mean of translating electronic addressing signals into delivery of chemicals and ions.¹ For complex regulation of functions in cells and tissues, a chemical circuit technology is necessary in order to generate complex and dynamic signal gradients with high spatiotemporal resolution. One approach to achieve a chemical circuit technology is to use bipolar membranes (BMs), which can be used to create the ionic equivalents of diodes^{2, 3, 4, 5} and transistors.^{6, 7, 8} A BM consists of a stack of a cation- and an anion-selective membrane, and functions similar to the semiconductor PN-junction, i.e., it offers ionic current rectification^{9, 10} (Figure ​(Figure1a).1a). The high fixed charge concentration in a BM configuration make them more suited in bioelectronic applications as compared to other non-linear ionic devices, such as diodes constructed from surface charged nanopores¹¹ or nanochannels,¹² as the latter devices typically suffers from reduced performance at elevated electrolyte concentration (i.e., at physiological conditions) due to reduced Debye screening length.¹³ However, a unique property of most BMs, as compared to the electronic PN-junction and other ionic diodes, is the electric field enhanced (EFE) water dissociation effect.^{10, 14} This occurs above a threshold reverse bias voltage, typically around −1 V, as the high electric field across the ion-depleted BM interface accelerates the forward reaction rate of the dissociation of water into H⁺ and OH⁻ ions. As these ions migrate out from the BM, there will be an increase in the reverse bias current. The EFE water dissociation is a very interesting effect and is commonly used in industrial electrodialysis applications,¹⁵ where highly efficient water dissociating BMs are being researched.¹⁶ Also, BMs have also been utilized to generate H⁺ and OH⁻ ions in lab-on-a-chip applications.^{2, 17} However, the EFE water dissociation effect diminishes the diode property of BMs when operated outside the ±1 V window, which is unwanted in, for instance, chemical circuits and addressing matrices for delivery of complex gradients of chemical species. The effect can be suppressed by incorporating a neutral electrolyte inside the BM,^{10, 18} for instance, poly(ethylene glycol) (PEG).^{2, 6, 7} However, as previously reported,² the PEG volume will introduce hysteresis when switching from forward to reverse bias, due to its ability to store large amounts of charges. This was circumvented by ensuring that only H⁺ and OH⁻ are present in the diode, which recombines into water within the PEG volume. Such diodes are well suited as integrated components in chemical circuits for pH-regulation, due to the in situ formed H⁺ and OH⁻, but are less attractive if, for instance, other ions, e.g., biomolecules, are to be processed or delivered in and from the circuit. Furthermore, a PEG electrolyte introduces additional patterning layers, making device downscaling more challenging. This is undesired when designing complex, miniaturized, and large-scale ionic circuits. Thus, there is an interest in BM diodes that intrinsically do not exhibit any EFE water dissociation, are easy to miniaturize, and that turn off at relatively high speeds. It has been suggested that tertiary amines in the BM interface are important for efficient EFE water dissociation,^{19, 20, 21} as they function as a weak base and can therefore catalyze dissociation of water by accepting a proton. For example, anion-selective membranes that have undergone complete methylation, converting all tertiary amines to quaternary amines, shows no EFE water dissociation,¹⁹ although this effect was not permanent, as the quaternization was reversed upon application of a current. Similar results were found for anion-selective membranes containing alkali-metal complexing crown ethers as fixed charges.²¹ Also, EFE water dissociation was not observed or reduced in BMs with poor ion selectivity, for example, in BMs with low fixed-charge concentration⁵ or with predominantly secondary and tertiary amines in the anion-selective membrane,²² as the increased co-ion transport reduces the electric field at the BM interface. However, due to decreased ion selectivity, these membranes show reduced rectification. In this work, we present a non-amine based BM diode that avoids EFE water dissociation, enables easy miniaturization, and provides fast turn-off speeds and high rectification.Open in a separate windowFigure 1(a) Ionic current rectification in a BM. In forward bias, mobile ions migrate towards the interface of the BM. The changing ion selectivity causes ion accumulation, resulting in high ion concentration and high conductivity. At high ion concentration, the selectivity of the membranes fails (Donnan exclusion failure), and ions start to pass the BM. In reverse bias, the mobile ions migrate away from the BM, eventually giving a zone with low ion concentration and low conductivity. Reverse bias can cause EFE water dissociation, producing H⁺ and OH^- ions. (b) Chemical structures of PSS, qPVBC, and PVBPPh₃. (c) The device used to characterize the BMs and the BM1A, BM2A, and BM1P designs. The BM interfaces are 50 × 50 μm.An anion-selective phosphonium-based polycation (poly(vinylbenzyl chloride) (PVBC) quaternized by triphenylphospine, PVBPPh₃) was synthesized and compared to the ammonium-based polycation (PVBC quaternized by dimethylbenzylamine, qPVBC) previously used in BM diodes² and transistors,^{7, 8} when included in BM diode structures together with polystyrenesulfonate (PSS) as the cation-selective material (Figure ​(Figure1b).1b). Three types of BM diodes were fabricated using standard photolithography patterning (Figure ​(Figure1c),1c), either with qPVBC (BM1A and BM2A) or PVBPPh₃ (BM1P) as polycation and either with (BM2A) or without PEG (BM1A and BM1P). Poly(3,4-ethylenedioxythiophene):poly(styrenesulfonate) (PEDOT:PSS) electrodes covered with aqueous electrolytes were used to convert electronic input signals into ionic currents through the BMs, according to the redox reaction PEDOT⁺:PSS⁻ + M⁺ + e⁻ ↔ PEDOT⁰ + M⁺:PSS⁻.The rectifying behavior of the diodes was evaluated using linear sweep voltammetry (Figure ​(Figure2).2). The BM1A diode exhibited an increase in the reverse bias current for voltages lower than −1 V, a typical signature of EFE water dissociation,^{10, 14} which decreased the current rectification at ±4 V to 6.14. No such deviation in the reverse bias current was observed for BM2A and BM1P, which showed rectification ratios of 751 and 196, respectively. In fact, for BM1P, no evident EFE water dissociation was observed even at −40 V (see inset of Figure ​Figure2).2). Thus, the PVBPPh₃ polycation allows BM diodes to operate at voltages beyond the ±1 V window with maintained high ion current rectification.Open in a separate windowFigure 2Linear sweep voltammetry from −4 to +4 V (25 mV/s) for the BM diodes. The inset shows BM1P scanning from −40 V to +4 V (250 mV/s).The dynamic performance of the diodes was tested by applying a square wave pulse from reverse bias to a forward bias voltage of 4 V with 5–90 s pulse duration (Figure ​(Figure3).3). To access the amount of charge injected and extracted during the forward bias and subsequent turn off, the current through the device was integrated. For BM2A, we observed that the fall time increased with the duration of the forward bias pulse. This hysteresis is due to the efficient storage of ions in the large PEG volume, with no ions crossing the BM due to Donnan exclusion failure.² As a result, during the initial period of the return to reverse bias, a large amount of charge needs to be extracted in order to deplete the BM. After a 90 s pulse, 90.6% of the injected charge during the forward bias was extracted before turn-off. This may be contrasted with BM1P, where the fall time is hardly affected by the pulse duration, and the extracted/injected ratio is only 15.4% for a 90 s pulse. For this type of BM, the interface quickly becomes saturated with ions during forward bias, leading to Donnan exclusion failure and transport of ions across the BM.⁴ Thus, less charge needs to be extracted to deplete the BM, allowing for faster fall times and significantly reduced hysteresis.Open in a separate windowFigure 3Switching characteristics (5, 10, 20, 30, 60, or 90 s pulse) and ion accumulation (at 90 s pulse) of the BM2A and BM1P diodes. BM1A showed similar characteristics as BM1P when switched at ±1V (see supplementary material).²⁴Since the neutral electrolyte is no longer required to obtain high ion current rectification over a wide potential range, the size of the BM can be miniaturized. This translates into higher component density when integrating the BM diode into ionic/chemical circuits. A miniaturized BM1P diode was constructed, where the interface of the BM was shrunk from 50 μm to 10 μm. The 10 μm device showed similar IV and switching characteristics as before (Figure ​(Figure4),4), but with higher ion current rectification ratio (over 800) and decreased rise/fall times (corresponding to 90%/–10% of forward bias steady state) from 10 s/12.5 s to 4 s/4 s. Since the overlap area is smaller, a probable reason for the faster switching times is the reduced amount of ions needed to saturate and deplete the BM interface. In comparison to our previous reported low hysteresis BM diode,² this miniaturized polyphosphonium-based devices shows the same rise and fall times but increased rectification ratio.Open in a separate windowFigure 4(a) Linear sweep voltammetry and (b) switching performance of a BM1P diode with narrow junction.In summary, by using polyphosphonium instead of polyammonium as the polycation in BM ion diodes the EFE water dissociation can be entirely suppressed over a large operational voltage window, supporting the theory that a weak base, such as a tertiary amine, is needed for efficient EFE water dissociation.^{17, 18} As no additional neutral layer at the BM interface is needed, ion diodes that operate outside the usual EFE water dissociation window of ±1 V can be constructed using less active layers, fewer processing steps and with relaxed alignment requirement as compared to polyammonium-based devices. This enables the fabrication of ion rectification devices with an active interface as low as 10 μm. Furthermore, the exclusion of a neutral layer improves the overall dynamic performance of the BM ion diode significantly, as there is less hysteresis due to ion accumulation. Previously, the hysteresis of BM ion diodes has been mitigated by designing the diode so that only H⁺ and OH⁻ enters the BM, which then recombines into water.² Such diodes also show high ion current rectification ratio and switching speed but are more complex to manufacture, and their application in organic bioelectronic systems is limited due to the H⁺/OH⁻ production. By instead using the polyphosphonium-based BM diode, reported here, we foresee ionic, complex, and miniaturized circuits that can include charged biomolecules as the signal carrier to regulate functions and the physiology in cell systems, such as in biomolecule and drug delivery applications, and also in lab-on-a-chip applications.     

Preface to Special Topic: Papers from the 2009 Conference on Advances in Microfluidics and Nanofluidics, The Hong Kong University of Science & Technology, Hong Kong, 2009

The inaugural conference on Advances in Microfluidics and Nanofluidics was held at the Hong Kong University of Science and Technology on 5–7 January 2009 and brought together leading researchers from across a wide variety of disciplines from North America, Europe, Asia, and Oceania. This Special Topic section forms the second of the two issues dedicated to original contributions covering both fundamental physicochemical aspects of microfluidics and nanofluidics as well as their applications to the miniaturization of chemical and biological systems that were presented at the conference.In the last five years, we have observed rapid growth in the microfluidics and nanofluidics community in Asia, owing largely to the substantial strategic investments by both government and industry in the region to promote the microfabrication and nanotechnology sectors.¹ The organization of a regular meeting focusing on activities in the Asia-Pacific rim region was, therefore, timely, particularly to enhance dissemination of research of the highest quality within the region and to promote collaboration between researchers in the Asian community with their counterparts from Europe and the USA.Biomicrofluidics is, therefore, proud to be closely involved with the organization of the first of such conferences, Advances in Microfluidics and Nanofluidics 2009, which was kindly hosted by the Hong Kong University of Science and Technology (HKUST). As reported in the preface to the first of the two issues dedicated to invited reviews and original contributions associated with the conference,² the meeting, which took place over three days in the breathtaking HKUST campus overlooking Clearwater Bay in Hong Kong, was a tremendous success. Together with our colleagues, the Biomicrofluidics editors are busy putting in place arrangements for a follow-up meeting in January 2011. Given the overwhelming response and positive feedback we’ve had to date, we believe that Advances in Microfluidics and Nanofluidics will form a regular event in the calendar of the Asian microfluidics and nanofluidics community in the future.It was particularly pleasing to observe the translation of fundamental and theoretical work into advanced applied chip-based platforms for a variety of practical chemical and biological applications in the talks presented at the conference. The collection of articles in this second part, in fact, provides a gist of the flavor of the multidisciplinary research spanning the entire fundamental to applied research spectrum, which is exactly the scope which the journal intends to cover.Electrokinetics continues to be a dominant theme in this issue and within the microfluidics and nanofluidics community. The article by Ng et al.³ provides experimental evidence that might put to rest a longstanding area of debate within the electrokinetics community on the role of Faradaic charging in driving electro-osmotic flow, first proposed by Ben and Chang.⁴ In other electrokinetics papers, the role of interfaces is explored, for example, electrowetting on the superhydrophobic nanostructured surfaces of a lotus leaf⁵ and droplet manipulation in an immiscible dielectric liquid continuum under an electric field.⁶In addition, the characterization of the surface charge density of the nanopores etched in organic foils is reported by Xue et al.,⁷ which provides a deeper understanding of the mechanisms by which ions are transported in nanochannels, whereas Wei and Hsiao⁸ present a stochastic simulation to model the condensation of linear polyelectrolyte molecules under electric fields, in which they show the marked increase in the mobility of the polyelectrolyte chain during its unfolding in free-solution electrophoresis.Continuing along the theme of numerical simulations, particulate transport in converging-diverging microchannels was studied using a Lagrangian-Eulerian finite-element model,⁹ and slip arising in Couette flows over superhydrophobic surfaces was studied using a hybrid multiscale simulation that interfaces molecular dynamics simulations in the near-wall region with the continuum fluid model in the bulk.¹⁰ In other numerical studies, drop coalescence¹¹ and nanotube transport¹² were studied.Complementing these fundamental studies is the use of multiphase flows in microfluidic channels to engineer scaffolds for tissue engineering in which the bubbles trapped in liquid droplets transported in microchannels were employed to produced the pores of the scaffold.¹³ Other practical microfluidics applications, such as chip-based enhancement of DNA hybridization through a genetic-bead-based protocol¹⁴ and an automated ELISA chip for chemical-biological analysis with an enhancement in the detection range and time,¹⁵ also constitute papers in this Special Topic section.We hope you enjoy reading the papers in this Special Topic section and that it provides you with a feel for the broad multidisciplinary spectrum across fundamental and applied microfluidic and nanofluidic research that the conference, as well as the journal, intends to span. Do watch out for the conference announcement for the next Advances in Microfluidics and Nanofluidics meeting in 2011 on the Biomicrofluidics website (http://bmf.aip.org)—hope to see you there!     

Quantifying the volume of single cells continuously using a microfluidic pressure-driven trap with media exchange

We demonstrate a microfluidic device capable of tracking the volume of individual cells by integrating an on-chip volume sensor with pressure-activated cell trapping capabilities. The device creates a dynamic trap by operating in feedback; a cell is periodically redirected back and forth through a microfluidic volume sensor (Coulter principle). Sieve valves are positioned on both ends of the sensing channel, creating a physical barrier which enables media to be quickly exchanged while keeping a cell firmly in place. The volume of individual Saccharomyces cerevisiae cells was tracked over entire growth cycles, and the ability to quickly exchange media was demonstrated.Measuring cell growth is of primary interest to researchers who seek to study the effects of drugs, nutrients, disease, and environmental stress. This has traditionally been accomplished by monitoring the optical transmittance of large ensembles of cells and applying the Beer-Lambert Law.^1,2 Such population-scale measurements provide important culture statistics, but averaging obscures the behaviour of individual cells. In addition, these techniques often require cell synchronicity in order to correlate growth with specific points in the cell cycle, but synchronicity typically decays rapidly in many cell lines including Saccharomyces cerevisiae (yeast) cultures.³ Researchers have thus adopted methods that study the growth of individual cells. Quantifying cellular growth is especially challenging since proliferating cells such as yeast or Escherichia coli are irregularly shaped, and will only increase in size by a factor of two.⁴ Growth will affect the mass, volume, and density of the cell; having access to each of these characteristics is important in obtaining a complete picture of this process. Time-lapse fluorescence microscopy can provide valuable information as to the cell cycle progression of individual cells,⁵ but 2D optics requires geometric assumptions, and, thus, can provide an incomplete picture of growth.^6,7Microfluidic lab-on-chip devices with integrated sensors can provide high-resolution growth tracking of individual cells, either through mass, volume, or density monitoring.^4,7,8 Recently, a microfluidic mass sensor was used to track the buoyant mass of individual cells using a suspended microchannel resonator (SMR).^4,9 Monitoring growth can also be accomplished by tracking volume using microfluidic volume sensors⁷ operating on the Coulter principle.¹⁰ Trapping can be achieved by either (1) cycling the target back and forth through the sensor (pressure-driven⁴ and electrokinetic⁷) or (2) holding a cell in place (posts,¹¹ chevron structure,¹² and E-Field¹³). The former, dynamic approach, allows a single cell to be sampled periodically by reversing flow directions after a cell is detected. Simple in its implementation, this technique also has the ability to compensate for a drifting baseline current resulting from parasitic ionic changes within the sensing channel or other sources of noise. On the other hand, static traps allow cells to be held in place while the buffer is rapidly exchanged.¹² The ability to dynamically change cellular growth conditions during an experiment can lead to significant insight into the behaviour of cells in environments of varying salinity,¹⁴ oxidative,^15,16 or osmotic conditions,¹⁷ as well as the effect of nutrients¹⁸ and drugs.¹⁹In this work, we propose a device capable of tracking growth using high-resolution volume measurements, combining the best attributes of both types of measurement systems; continuous baseline correction and the ability to rapidly exchange cell media. This is accomplished by using a pressure-driven, feedback-based dynamic trap, whereby a cell is cycled back and forth through the sensor within a microfluidic channel. On-chip sieve valves positioned at both ends of the sensing channel are able to selectively capture a cell while the solution is being replaced. As proof of principle, the volume of several individual yeast cells was monitored over the course of their respective growth cycles, and the ability to quantify growth response to media exchange was demonstrated.Devices were fabricated using multilayered soft lithography with polydimethylsiloxane (PDMS) molding.²⁰ The completed device is pictured in Figure 1(a); full fabrication protocols are presented as supplementary material.²¹ To maximize measurement sensitivity, it is optimal to choose a channel width and height slightly larger than the dimensions of the target cell.²² However, yeast cells are asymmetrically shaped and tend to tumble as they traverse the sensor. Preliminary testing suggested this effect could be mitigated by having cells flow along trajectories far from the electrodes (through buoyancy), where electric field is more uniform. Thus, a channel height of 20 μm was chosen as a compromise. Channel height increases to 28 μm in the wider part of the central and bypass channels, a result of using a mold made out of reflowed photoresist.²³ Channel width was set at 25 μm through the sensor, and widens to 80 μm at the sieve valves to facilitate valve actuation, which requires a high width to height ratio.²⁰ The fluidic layer is integrated in a 35 μm thick PDMS spin-coated layer, above which sits a 50 μm tall valve channel in a 4 mm PDMS layer. Tubing connects I1 and I2 to a common inlet vial, V1 and V2 to vials filled with deionised water and O1 and O2 connect to empty vials (not pictured). Inlet pressures I1 and I2, and valve pressures V1 and V2 are controlled with manual regulators (SMC IR2000-N02-R and SMC IR2010-N02-R); outlet pressures are computer-controlled (SMC ITV-1011). This pressure scheme is detailed elsewhere.²⁴ Current pulses caused by transiting particles/cells (Figure 1(d)) were acquired by applying a 50 kHz, 220 mV AC voltage between a pair of electrodes and measuring the drawn current. This frequency is sufficiently elevated to avoid the electrical double layer capacitance at the electrode-electrolyte interface,²⁵ but low enough to avoid sensitivity to cell impedance or substrate.²⁶ The electrical setup used for these experiments has been described previously.^24,27 A temperature controller maintains the device at 30 °C.Open in a separate windowFIG. 1.(a) Micrograph of the microfluidic device. Two parallel bypass channels are connected by a sensing channel with sensing electrodes. Pressure is applied at inlets (I1, I2) and outlets (O1, O2) to control flow conditions. Valves (V1, V2) are positioned over each end of the sensing channel. Food coloring is used to highlight the valve (red) and fluidic layers (blue). (b) Flow mode: valves are unpressurized, and cells flow freely through the device. (c) Trapping mode: valves are pressurized to capture a cell within the central channel. Pressure-driven flow cycles the cell back and forth across the sensor. (d)Typical current pulses measured for a yeast cell.The cell capture, media exchange, and detection process occurs as follows. A cell suspension is loaded into the bypass channel and made to flow through the central sensing channel by imposing a pressure gradient (Figure 1(b)). Cells flowing through the sensor are observed optically; once a cell of interest is observed (a cell without a bud), valves are sealed (V1 = V2 = 35 psi). This stops all flow through the sensor, and enables bypass channels to be flushed and replaced with fresh media. After 2 min, valve channels are pressurized to 24 psi where they compress the channel to a sufficient height to physically restrict the passage of yeast cells, while allowing the media to flow through the central channel (Figure 1(c)). The pressure gradient between bypasses causes the media in the central channel to be flushed out, while the target cell is physically trapped. Replacing the media in the central channel takes 2 min. At this stage, a pressure-driven feedback-based dynamic trap can be initiated. In this dynamic trap mode, the pressure settings at O1 and O2 are adjusted to redirect the cell back and forth through the sensor, based on current pulses measured from cells transiting through the sensor. Through custom LabView® software, these outlet pressure settings are feedback-adjusted to maintain a speed of 250 μm/s in both directions at a detection frequency of 30 cells/min (Figure 1(d)). To minimize the effects of channel stretching/shrinking, the sum of pressures at O1 and O2 is held constant. This precaution was taken since the sensing channel structured within the flexible PDMS polymer will alter its geometry based on internal pressure.²⁸ The short central channel ensures steady nutrient replenishment from the bypasses. For example, a glucose molecule takes ∼4 min to diffuse from the bypass to the electrodes. In practice, Taylor-Aris dispersion will reduce this replenishment time considerably. Based on video analysis, 25% of the central channel''s media is replenished every pressure reversal (video presented as supplementary material²¹). Polystyrene microspheres of 3.9 ± 0.3 μm, 5.6 ± 0.2 μm, and 8.3 ± 0.7 μm (NIST size standards) were used to calibrate the sensor, and obtain the current pulse-to-volume calibration for every solution (supplementary material²¹). The validity of this calibration method is discussed elsewhere.²⁹ Care was taken to limit trajectory-based variations in signal: the device is positioned with electrodes at the top of the sensing channel, and with the negatively buoyant cells/particles flowing along the bottom. Based on previous experimental and theory work, we found that signal amplitude can vary as much as 3.5 fold for different heights.²⁷ The effect of trajectory on current pulse amplitude has also been reported elsewhere.^30,31 In this work, buoyancy is used to ensure that the cell flows along a trajectory at the same distance from the electrodes for every measurement.Saccharomyces cerevisiae (BY4743 Mat a/alpha, genotype: his3Δ1/his3Δ1 leu2Δ0/leu2Δ0 LYS2/lys2Δ0 met15Δ0/MET15 ura3Δ0/ura3Δ0 ade2::LEU2/ade2::URA3) was cultured to exponential phase at 30 °C in an incubator/shaker in yeast bacto-peptone (YPD) with 2% w/v glucose, supplemented with 0.2 M NaCl, 0.05% bovine serum albumin (BSA) and 42 mg/l adenine. Sodium chloride was added to enable the current pulse measurement, at a concentration where cells are viable;³² BSA was used to prevent cell agglomeration; adenine was supplemented since this particular yeast mutant does not produce its own supply. A cell suspension was introduced into the device, from which a cell at the early stages of its cell cycle was captured, and dynamically trapped for 100 min. Three typical cell growth results are shown in Figure 2(a). Since the culture was not synchronized, this leads to variability between “initial” cell volumes: there is a 27% difference in initial volume between the cells identified by red squares and green triangles. This is caused by (1) optical limits, whereby cells chosen for study are not all at the exact same cell cycle stage and (2) differences in the age of the mother cell: the more buds a mother cell has produced, the larger it becomes.³³ On average, captured yeast cell demonstrated a doubling time consistent with growth rates under ideal incubator/shaker conditions; nutrient depletion, electric field, and shear stresses are not affecting growth. Optical inspection of budding cells confirms that most growth is occurring at the daughter cell, as expected.³³ An elevated signal-to-noise ratio allows for high resolution volumetric measurements (4 μm³); cell asymmetry⁷ and trajectory variability^27,30,31 lead to a relative standard deviation of 6% for cells and 4% for microspheres of similar size. While mass or protein synthesis methods have indicated linear³⁴ or exponential^4,6,35,36 growth curves, volume-based methods have suggested sigmoidal patterns.^7,37 Prior to daughter cell emergence, and later in the cycle as the daughter cell emerges, volumetric growth rate declines.³⁸ In this work, it is difficult to ascertain with mathematical rigor the shape of the growth profile; however, for each cell, volume increases steadily throughout the growth cycle before declining near the end of the cycle.Open in a separate windowFIG. 2.(a) Growth curves for 3 cells trapped in succession. Simultaneous optical and electrical measurements allow cell cycle stage to be correlated with volume. Pictures of cell corresponding to the red squares are presented in 15 min increments. A cell is cycled through the sensor every 2 s. For clarity, each data point for yeast volume represents the average of data points over a period of 5 min, with standard deviation. (b) Demonstration of an interrupted growth cycle, where YPD + 0.2 M NaCl was replaced with 0.2 M NaCl at 40 min, and then again returned to YPD + 0.2 M NaCl at 80 min. The media exchange process takes 4 min.To demonstrate our ability to easily exchange media while maintaining a trap, the solution was exchanged 40 min into a yeast growth cycle; culture media was replaced with a pure saline solution 0.2 M NaCl + 0.05% BSA, and then replaced again with culture media at 80 min (Figure 2(b)). Cell growth is halted temporarily while in saline solution, before resuming normal growth thereafter. The cell cycle time is extended by this period. The cell volume drifts downward after the initial solution change at 40 min. Though this drift lies within our uncertainty bounds, cellular responses to osmotic shock on similar timescales have been documented elsewhere.³⁹ This result demonstrates an ability to quickly exchange cell media, and observe cellular response.In conclusion, we have demonstrated a microfluidic device capable of maintaining a dynamic, pressure-driven cell trap, which can monitor cellular volume over the cell cycle. Concurrent optical microscopy allows for real-time visual inspection of the cells. In addition, sieve valve integration provides for the exchange of media or the addition of drugs. Such a platform could also be key in cancer cell cytotoxicity assays,⁴⁰ where growth response to anticancer drugs could be monitored.     

An integrated microfluidic device for rapid serodiagnosis of amebiasis

A microfluidic device was successfully fabricated for the rapid serodiagnosis of amebiasis. A micro bead-based immunoassay was fabricated within integrated microfluidic chip to detect the antibody to Entamoeba histolytica in serum samples. In this assay, a recombinant fragment of C terminus of intermediate subunit of galactose and N-acetyl-_D-galactosamine-inhibitable lectin of Entamoeba histolytica (C-Igl, aa 603-1088) has been utilized instead of the crude antigen. This device was validated with serum samples from patients with amebiasis and showed great sensitivity. The serodiagnosis can be completed within 20 min with 2 μl sample consumption. The device can be applied for the rapid and cheap diagnosis of other infectious disease, especially for the developing countries with very limited medical facilities.Entamoeba histolytica is the causative agent of amebiasis and is globally considered a leading parasitic cause of human mortality.¹ It has been estimated that 50 × 10⁶ people develop invasive disease such as amebic dysentery and amebic liver abscess, resulting in 100 000 deaths per annum.^{2, 3} High sensitive diagnosis method for early stage amebiasis is quite critical to prevent and cure this disease. To date, various serological tests have been used for the immune diagnosis of amebiasis, such as the indirect fluorescent antibody test (IFA) and enzyme-linked immunosorbent assay (ELISA).We have recently identified a 150-kDa surface antigen of E. histolytica as an intermediate subunit (Igl) of galactose and N-acetyl-_D-galactosamine-inhibitable lectin.^{4, 5} In particular, it has been shown that the C-terminus of Igl (C-Igl, aa 603-1088) was an especially useful antigen for the serodiagnosis of amebiasis. ELISA using C-Igl is more specific than the traditional ELISA using crude antigen.⁶ However, the ELISA process usually takes several hours, which is still labor-intensive and requires experienced operators to perform. More economic and convenient filed diagnosis methods are still in need, especially for the developing countries with limited medical facilities.Among all the bioanalytical techniques, microfluidics has been attracting more and more attention because of its low reagent/power consumption, the rapid analysis speed as well as easy automation.^{7, 8, 9, 10, 11} Especially with the development of the fabrication technique, microfluidics chip can include valves, mixers, pumps, heating devices, and even micro sensors, so many traditional bioanalytical methods can be performed in the microfluidics. Qualitative and quantitative immune analysis on the microfluidic chip was successfully proved by plenty of research with improved sensitivity, shorten reaction time, and less sample consumption.^{8, 10, 11, 12, 13, 14, 15, 16, 17} Moreover, with the intervention of other physical, chemical, biology, and electronic technology, microfluidic technique has been successfully utilized in protein crystallization, protein and gene analysis, cell capture and culturing and analysis as well as in the rapid and quantitative detection of microbes.^{13, 14, 15, 16, 17, 18, 19, 20}Herein, we report a new integrated microfluidic device, which is capable of rapid serodiagnosis of amebiasis with little sample consumption. The microfluidic device was fabricated from polydimethysiloxane (PDMS) following standard soft lithography.^{21, 22} The device was composed of two layers (shown in Figure ​Figure1)1) including upper fluidic layer (in green and blue) and bottom control layer (in red).Open in a separate windowFigure 1Structure illustration of microfluidic chip.To create the fluidic layer and the control layer, two different molds with different patterns have fabricated by photolithographic processes. The mold to create the fluidic channels was made by positive photoresist (AZ-50 XT), while the control pneumatic mold was made by negative photoresist (SU8 2025). For the chip fabrication, the fluidic layer is made from PDMS (RTV 615 A: B in ratio 5:1), and the pattern was transferred from the respective mold. The control layer is made from PDMS (RTV 615 A:B in ratio 20:1). The two layers were assembled and bonded together accurately, and there is elastic PDMS membrane about 30 μm thick between the fluidic layer channels and control layer.^{21, 22} The elastic membrane at the intersection can deform to block the fluid inside the fluidic channels, functioning as valves under the pressures introduced though control channels. There are two types of channels in fluidic layer, the rectangular profiled (in green, 200 μm wide, 35 μm thick) channel and round profiled channels (in blue, 200 μm wide, 25 μm center height). Because of the position of the valves on the fluidic channels, two types of valves (Figure ​(Figure2a)2a) were built, working as a standard valve and a sieve valve. The standard valves (on blue fluidic channels) can totally block the fluid because of the round profile of fluidic channel; the sieve valve can only half close because of the rectangular profile. The sieve valve can be used to trap the microspheres (beads) filled inside the green fluidic channels, while letting the fluid pass through. By this sieve valve, a micro column (in green) is constructed, where the entire ELISA reaction happens. The micrograph of the fabricated micro device is shown in Figure ​Figure2b.2b. The channels were filled with food dyes in different colors to show the relative positions of the channels. The pressures though different control channels are individually controlled by solenoid valves, connected to a computer through relay board. By programming the status (on/off) of various valves at different time periods, all the microfluidic chip operation can be digitally controlled by the computer in manual, semi-automatic, or automatic manner.Open in a separate windowFigure 2(a) Structure illustration of micro column, standard valve and sieve valve; (b) photograph of the microfluidic chip.To validate this device, 12 patient serum samples were collected. Sera from 9 patients (Nos. 1–9) with an amebic liver abscess or amebic colitis were used as symptomatic cases. The diagnosis of these patients was based on their clinical symptoms, ultrasound examination (liver abscess) and endoscopic or microscopic examination (colitis). We also identified the clinical samples using PCR amplification of rRNA genes.²⁴ As negative control, sera obtained from 3 healthy individuals with no known history of amebiasis were mixed into pool sera. The serum was positive for E. histolytica with a titer of 1:64 (borderline positive), as determined by an indirect fluorescent-antibody (IFA) test.^{23, 24} In our previously study, the sensitivity and specificity of the recombinant C-Igl in the ELISA were 97% and 99%.^{6, 25} In the current study, the serodiagnosis of amebiasis was also examined by ELISA using C-Igl.²⁶ The cut-off for a positive result was defined as an ELISA value > 3 SD above the mean for healthy negative controls²⁷ (shown in Figure ​Figure3).3). The seropositivity to C-Igl was 100% in patients with amebiasis.Open in a separate windowFigure 3ELISA reactivity of sera from patients against C-Igl. ELISA plate was coated with 100 ng per well of C-Igl. Serum samples from patients and healthy controls were used at 1:400 dilutions. The dashed line indicates the cut-off value. Data are representative of results from three independent experiments.In the diagnosis process with microfluidic chip, the 4 micro immuno-columns filled with C-Igl-coated microspheres were the key components of the device. The C-Igl was prepared in E. coli as inclusion bodies. After expression, the recombinant protein was purified and analyzed by SDS-PAGE. The apparent molecular mass was 85 kDa.²⁶The immune-reaction mechanism is illustrated in Figure ​Figure4.4. The anti-His monocolonal antibody was immobilized onto the microspheres (beads, 9 μm diameter) coated with protein A. The C-Igl was then immobilized onto the beads through the binding between the His tag and C-Igl. For the diagnosis, the microspheres immobilized with C-Igl and blocked by 5% BSA were preloaded into the columns for the rapid analysis of the patient serum samples. Generally, serum samples which were diluted 100 times were first loaded into the reaction column and incubated at room temperature for 5 min. After being washed by PBS buffer, FITC-conjugated goat anti-human polyclonal antibody was added into the column for 4 min incubation. The fluorescence image can be collected by the fluorescence microscope after the micro column was washed with PBS buffer. From loading diluted serum samples into column to collecting fluorescence images, the total time to complete the immunoassay is less than 10 min. The final fluorescence results were analyzed by Image Pro Plus 6.0.Open in a separate windowFigure 4Schematic representation of the ELISA in the chip.Different reaction conditions have been investigated to find the optimized ones. For each patient, 2 μl sample is enough for the analysis. The designed microfluidic chip with 4 micro columns is capable for 4 parallel analyses at the same time. More micro columns can be integrated into the device if more parallel tests are needed.Different incubating time for the diagnosis has also been investigated and no significant difference has been found for various time periods. It is enough to incubate the chip for only 5 min. The total diagnosis time for one sample is less than 10 min. The detection result appeared as the fluorescence intensity of the reaction column. As shown in Figure ​Figure5,5, the negative sample showed relatively low fluorescence intensity, because little FITC-conjugated goat anti-human polyclonal antibody could attach to the surface of microspheres; on the contrast, the positive sample showed much brighter fluorescence. The fluorescence intensity can be transferred to digital data (Table

Flow manipulation and cell immobilization for biochemical applications using thermally responsive fluids

A flow redirection and single cell immobilization method in a microfluidic chip is presented. Microheaters generated localized heating and induced poly(N-isopropylacrylamide) phase transition, creating a hydrogel that blocked a channel or immobilized a single cell. The heaters were activated in sets to redirect flow and exchange the fluid in which an immobilized cell was immersed. A yeast cell was immobilized in hydrogel and a 4′,6-diamidino-2-phenylindole (DAPI) fluorescent stain was introduced using flow redirection. DAPI diffused through the hydrogel and fluorescently labelled the yeast DNA, demonstrating in situ single cell biochemistry by means of immobilization and fluid exchange.The ability to control microfluidic flow is central to nearly all lab-on-a-chip processes. Recent developments in microfluidics either include microchannel based flow control in which microvalves are used to control the passage of fluid,¹ or are based on discrete droplet translocation in which electric fields or thermal gradients are used to determine the droplet path.^{2, 3} Reconfigurable microfluidic systems have certain advantages, including the ability to adapt downstream fluid processes such as sorting to upstream conditions and events. This is especially relevant for work with individual biomolecules and high throughput cell sorting.⁴ Additionally, reconfigurable microfluidic systems allow for rerouting flows around defective areas for high device yield or lifetime and for increasing the device versatility as a single chip design can have a variety of applications.Microvalves often form the basis of flow control systems and use magnetic, electric, piezoelectric, and pneumatic actuation methods.⁵ Many of these designs require complicated fabrication steps and can have large complex structures that limit the scalability or feasability of complex microfluidic systems. Recent work has shown how phase transition of stimuli-responsive hydrogels can be used to actuate a simple valve design.⁶ Beebe et al. demonstrated pH actuated hydrogel valves.⁷ Phase transition of thermosensitive poly(N-isopropylacrylamide) (PNIPAAm) using a heater element was demonstrated by Richter et al.⁸ Phase transition was also achieved by using light actuation by Chen et al.⁹ Electric heating has shown a microflow response time of less than 33 ms.¹¹ Previous work¹⁰ showed the use of microheaters to induce a significant shift in the viscosity of thermosensitive hydrogel to block microchannel flow and deflect a membrane, stopping flow in another microchannel. Additionally, Yu et al.¹² demonstrated thermally actuated valves based on porous polymer monoliths with PNIPAAm. Krishnan and Erickson¹³ showed how reconfigurable optically actuated hydrogel formation can be used to dynamically create highly viscous areas and thus redirect flow with a response time of  ～ 2?s. This process can be used to embed individual biomolecules in hydrogel and suppress diffusion as also demonstrated by others.^{15, 16} Fiddes et al.¹⁴ demonstrated the use of hydrogels to transport immobilized biomolecules in a digital microfluidic system. While the design of Krishnan and Erickson is highly flexible, it requires the use of an optical system and absorption layer to generate a geometric pattern to redirect flow.This paper describes the use of an array of gold microheaters positioned in a single layer polydimethylsiloxane (PDMS) microfluidic network to dynamically control microchannel flow of PNIPAAm solution. Heat generation and thus PNIPAAm phase transition were localized as the microheaters were actuated using pulse width modulation (PWM) of an applied electric potential. Additionally, hydrogel was used to embed and immobilise individual cells, exchange the fluid parts of the microfluidic system in order to expose the cells to particular reagents to carry out an in situ biochemical process. The PDMS microchannel network and the microheater array are shown in Figure ​Figure11.Open in a separate windowFigure 1A sketch of the electrical circuit and a microscope image of the gold microheaters and the PDMS microchannels. The power to the heaters was modulated with a PWM input through a H-bridge. For clarity, the electrical circuit for only the two heaters with gelled PNIPAAm is shown (H1 and V2). There are four heaters (V1-V4) in the “vertical channels” and three heaters (H1-H3) in the “horizontal” channel.The microchannels were fabricated using a patterned mould on a silicon wafer to define PDMS microchannels, as described by DeBusschere et al.¹⁷ and based on previous work.¹⁰ A 25 × 75 mm glass microscope slide served as the remaining wall of the microchannel system as well as the substrate for the microheater array. The gold layer had a thickness of 200 nm and was deposited and patterned using E-beam evaporation and photoresist lift-off.²¹ The gold was patterned to function as connecting electrical conductors as well as the microheaters.It was crucial that the microheater array was aligned with an accuracy of  ～ 20μm with the PDMS microchannel network for good heat localization. The PDMS and glass lid were treated with plasma to activate the surface and alignment was carried out by mounting the microscope slide onto the condenser lens of an inverted microscope (TE-2000 Nikon Instruments). While imaging with a 4× objective, the x, y motorized stage aligned the microchannels to the heaters and the condenser lens was lowered for the glass substrate to contact the PDMS and seal the microchannels.Local phase transition of 10% w/w PNIPAAm solution in the microchannels was achieved by applying a 7 V potential through a H-bridge that received a PWM input at 500 Hz which was modulated using a USB controller (Arduino Mega 2650) and a matlab (Mathworks) GUI. The duty cycle of the PWM input was calibrated for each microheater to account for differences in heater resistances (25?Ω to 52?Ω) due to varying lengths of on-chip connections and slight fabrication inconsistencies, as well as for different flow conditions during device operation. Additionally, thermal cross-talk between heaters required decreasing the PWM input significantly when multiple heaters were activated simultaneously. This allowed confining the areas of cross-linked PNIPAAm to the microheaters, allowing the fluid in other areas to flow freely.By activating the heaters in sets, it was possible to redirect the flow and exchange the fluid in the central area. Figure ​Figure22 demonstrates how the flow direction in the central microchannel area was changed from a stable horizontal flow to a stable vertical flow with a 3 s response time, using only PNIPAAm phase transition. Constant pressures were applied to the inlets to the horizontal channel and to the vertical channels. Activating heaters V1-4 (Figure ​(Figure2,2, left) resulted in flow in the horizontal channel only. Likewise, activating heaters H1 and H2 allowed for flow in the vertical channel only. In this sequence, the fluid in the central microchannel area from one inlet was exchanged with fluid from the other inlet. Additionally, by activating heater H3, a particle could be immobilised during the exchange of fluid as shown in Figure ​Figure33 (top).Open in a separate windowFigure 2Switching between fluid from the horizontal and the vertical channel using hydrogel activation and flow redirection with a response time of 3 s. A pressure of 25 mbar was applied to the inlet of the horizontal channel and a pressure of 20 mbar to the vertical channel. The flow field was determined using particle image velocimetry, in which the displacement of fluorescent seed particles was determined from image pairs generated by laser pulse exposure. Processing was carried out with davis software (LaVision).Open in a separate windowFigure 3A series of microscope images near heater H3 showing: (1a)-(1c) A single yeast cell captured by local PNIPAAm phase transition and immobilized for 5 min before being released. (2a) A single yeast cell was identified for capture by embedding in hydrogel. (2b) The cell as well as the hydrogel displayed fluorescence while embedded due to the introduction of DAPI in the surrounding region. (2c) The diffusion of DAPI towards the cell as the heating power of H3 is reduced after 15 min, showing a DAPI stained yeast cell immobilized.Particle immobilisation in hydrogel and fluid exchange in the central area of the microfluidic network were used to carry out an in situ biochemical process in which a yeast cell injected through one inlet was stained in situ with a 4′,6-diamidino-2-phenylindole (DAPI) solution (Invitrogen), which attached to the DNA of the yeast cell.¹⁸ A solution of yeast cells with a concentration of 5 × 10⁷cells/ml suspended in a 10% w/w PNIPAAm solution was injected through the horizontal channel. A solution of 2μg/l DAPI in a 10% w/w PNIPAAm solution was injected through the vertical channel. A single yeast cell was identified and captured near the central heater, and by deactivating the heaters in the vertical channel, DAPI solution was introduced in the microchannels around the hydrogel. After immobilising the cell for 15 min, the heater was deactivated, releasing the cell in the DAPI solution. This process is shown in Figure ​Figure33 (bottom). The sequence of the heater activation and deactivation in order to immobilize the cell and exchange the fluid is outlined in the supplementary material.²¹Eriksen et al.¹⁵ demonstrated the diffusion of protease K in the porous hydrogel matrix,¹⁹ and it was therefore expected that DAPI fluorescent stain (molecular weight of 350 kDa, Ref. ²⁰) would also diffuse. DAPI diffusion is shown in Figure 3(2b) in which the yeast cell shows fluorescence while embedded in the hydrogel. The yeast cell was released by deactivating the central heater and activating all the others to suppress unwanted flow in the microchannel. As a result, the single cell was fully immersed in the DAPI solution. Immobilization of a single cell allows for selection of a cell that exhibits a certain trait and introduction of a new fluid while maintaining the cell position in the field of view of the microscope such that a biochemical response can be imaged continuously.In summary, a microfluidic chip capable of local heating was used to induce phase transition of PNIPAAm to hydrogel, blocking microchannel flow, and thereby allowing for reconfigurable flow. Additionally, the hydrogel was used to embed and immobilise a single yeast cell. DAPI fluorescent stain was introduced using flow redirection, and it stained the immobilized cell, showing diffusion into the hydrogel. The versatile design of this microfluidic chip permits flow redirection, and is suitable to carry out in situ biochemical reactions on individual cells, demonstrating the potential of this technology for forming large-scale reconfigurable microfluidic networks for biochemical applications.     

Complementary metal oxide semiconductor compatible fabrication and characterization of parylene-C covered nanofluidic channels with integrated nanoelectrodes

Nanochannels offer a way to align and analyze long biopolymer molecules such as DNA with high precision at potentially single basepair resolution, especially if a means to detect biomolecules in nanochannels electronically can be developed. Integration of nanochannels with electronics will require the development of nanochannel fabrication procedures that will not damage sensitive electronics previously constructed on the device. We present here a near-room-temperature fabrication technology involving parylene-C conformal deposition that is compatible with complementary metal oxide semiconductor electronic devices and present an analysis of the initial impedance measurements of conformally parylene-C coated nanochannels with integrated gold nanoelectrodes.No two cells are exactly alike, either in terms of their genome, the genomic epigenetic modification of the genome, or the expressed proteome.¹ The genomic heterogeneity of cells is particularly important from an evolutionary perspective since it represents the stages of evolution of a population of cells under stress.² Because of the important variances in the genome that occur from cell to cell, it is critical to develop genomic analysis technologies which can do single-cell and single molecule genomic analysis as an electronic “direct read” without intervening amplification steps.^{3, 4, 5, 6, 7, 8} In this paper, we present a technique which uses conformal coverage of nanochannels containing nanoelectrodes using a room-temperature deposition of parylene-C, a pin-hole-free, excellent electrical insulator with low autofluorescence.⁹ This procedure should open the door to integration of many kinds of surface electronics with nanochannels. One of the most difficult aspects in introducing electronics into nanochannel technology is the sealing of nanochannel so that the electrodes are not compromised by harsh chemicals or high temperatures. There are various methods to form nanochannels containing nanoelectrodes, including wafer bonding techniques,¹⁰ removal of sacrificial materials,¹¹ and nonuniform sputtering deposition.¹² Methods employing a sacrificial layer removal show the greater compatibility to electronic integration, but current methods to remove sacrificial materials require either high temperatures¹¹ or harsh chemicals.^{13, 14}The basic device consisted of 12 mm long, 100 nm wide, 100 nm high nanochannels interrogated by 22 pairs of 30 nm wide gold nanoelectrodes. The outline of the fabrication process is shown in Fig. ​Fig.1.1. The fabrication process was carried out on a standard 4 in. single-side polished p-type ⟨100⟩ silicon wafer with 100 nm of dry thermal oxide on the top as an insulating layer, which also helped the wetting of the nanochannels. The first step involved nanofabrication of the 25 nm thick nanoelectrodes on the SiO₂ top of the wafer using electron beam lithography (EBL). External gold connection pads were constructed using standard metal lift-off techniques and photolithography to connect to the nanoelectrodes. A Raith E-Line e-beam writing system (Raith USA, Ronkonkoma, NY) was used to expose polymethyl methacrylate (PMMA) for metal lift-off. Figure ​Figure1a1a shows a scanning electron microscopy (SEM) image of the nanoelectrodes. The 100 nm sealed nanochannels were constructed using sacrificial removal techniques. We used EBL to expose a 100 nm thick film of PMMA over the gold nanolines in the region around the nanolines, leaving behind lines of unexposed sacrificial layer of PMMA. We next evaporated 25 nm of SiO₂ over the nanolines to improve the surface wetting properties of nanochannel and then conformally coated with 4 μm thick of parylene-C [poly(chloro-p-xylylene)] using a Specialty Coating Systems model PDS 2010 parylene coating system (SCS Systems, Indianapolis, IN). Access holes for the gold electrodes and the feeding channels were etched through by oxygen plasma and 1:10 buffered oxide etchant. To avoid autofluorescence induced in parylene by an active plasma¹⁵ and ambient UV radiation,¹⁶ it is important not to expose the remaining parylene with plasma and to keep the samples in the dark. The sacrificial removal of PMMA in the nanochannels was done in four steps: (1) soaking the chip in 55 °C 1165 MicroChem resist remover (MicroChem, Newton, MA) for 36 h, (2) room-temperature soaking in 1,2-dichloroethane for 12 h, (3) soaking in room-temperature acetone for 12 h, and (4) drying the nanochannels by critical point drying (CPD-030, BAL-TEC AG, Principality of Liechtenstein), which served to prevent the collapse of the nanochannel resulting from surface tension of the acetone.Open in a separate windowFigure 1(a) SEM image of gold nanoelectrodes; scale bar is 200 nm. (b) 100×100 nm² PMMA nanoline is written over the gold nanoelectrodes by exposure of the surrounding PMMA. (c) Parylene-C conformal coating over the PMMA nanoline. PMMA is dissolved and parylene-C etched by reactive ion etching.Conductance measurements were done using ac techniques. The ac impedance Z_tot of an insulating ionic fluid such as water between electrodes is a complex subject.¹⁷ The most general model for the complex impedance of an electrode in ionic solution is typically modeled as the Randle circuit, which is shown in Fig. ​Fig.22.¹⁷ There are two major contributions to the imaginary part of the impedance: the capacitance of the double layer (C_dl), which is purely imaginary and has no dc conductance, and the impedance due to charge transfer resulting in electrochemical reactions at the electrode∕electrolyte interface, which can be modeled as a contact resistor (R_CT), which is given by the Butler–Volmer equation, which describes the I-V characteristic curve when electrochemical reactions occur at the electrode,¹⁸ in series with a complex Warburg impedance (Z_W) which represents injected charge transport near the electrode;¹⁹ more details can be found in Ref. ²⁰. Since we applied a 10 mV rms ac voltage with no dc offset in our measurements, electrochemical reactions are negligible, which means no electrochemical charge transfer occurred and as a result R_CT goes to infinity. We have drawn a gray box around the elements connected to the Warburg impedance branch of the circuit to show that they are negligible in our analysis.Open in a separate windowFigure 2The equivalent circuit of the nanoelectrodes in contact with water lying atop an insulating SiO₂ film which covers a silicon substrate. The elements in the gray boxes can be ignored in our measurements since there is no hydrolysis at low voltage, while the elements within the dotted box are coupling reactances to the underlying p-doped silicon wafer.In the case of no direct charge injection, the electrodes are coupled by the purely capacitive dielectric layer impedance C_dl to the solvent and are also coupled capacitively by the dielectric SiO₂ film capacitance C_ox to the underlying p-doped silicon semiconductor. We model the semiconductor as a purely resistive material with bulk resistivity ρ_Si. The value of C_dl∕area is on the order of ϵϵ_oκ, where ϵ is the dielectric constant of water (about 80) and κ is the Debye screening parameter of the counterions in solution: κ=ϵϵokBT∕e2Σici∞zi2,²⁰ where ci∞ is the bulk ion concentration of charge z_i. At our salt molarity of 50 mM (1∕2 Tris∕Borate∕EDTA (TBE) buffer), C_dl is approximately 30 μF∕cm² using 1∕κ∼1 nm.In Fig. ​Fig.3,3, we show the ac impedance measurements between pairs nanoelectrodes for both dry and TBE buffer wet nanochannels. The electrodes are capacitively coupled to the underlying silicon substrate through an oxide capacitor C_ox. We model the doped silicon wafer as pure resistors, so there is an R₁ that connects both C_ox, and each C_ox is connected to the ground with an R₂. Curve fitting was done by using the 3SPICE circuit emulation code (VAMP Inc., Los Angeles, CA). We therefore obtained the following parameters for the dry curve: C_ox=1.32 nF, R₁=17.5 μΩ, and R₂=32.8 kΩ. R₁ is not sensitive in the fit as long as it is smaller than the impedance of C_ox. Given ρ_Si of the wafer of 1–10 Ω cm, R₂ should be on the order of 10³ Ω, which is slightly smaller than our fitting results. The same parameters for the wafer coupling parameters were then used for fitting the impedance measurements for wet channels. For TBE buffer solution in the nanochannel, curve fitting yields C_dl=50 pF and R_sol=10⁵ Ω. However, given the dimension of our nanochannels, we should get a transverse resistance R∼10⁹ Ω. One possible explanation for this difference is that the evaporated SiO₂ film which was put over the PMMA is porous and allows buffer to penetrate the oxide film,²¹ but given that the film is only 25 nm thick this would at most increase the cross section by one order of magnitude. However, it is known that there is a high fractional presence of mobile counterions associated with the charged channel walls.²² To calculate exact conductance contribution from the surface charges is a tricky business, but since the surface-to-volume ratios in our nanochannels are much greater than the slits, a larger conductance enhancement can be expected, and more work needs to be done.Open in a separate windowFigure 3ac impedance spectra of TBE buffer solution in a transchannel measurement between adjacent pairs of nanoelectrodes separated by 135 μm. The red circles are data for a dry channel and the solid red line is the fit to the model shown in the upper right hand corner. The green squares and dashed green line are for a nanochannel wet with TBE buffer.We have presented a way to fabricate a nanochannel integrated with electrodes. This technology opens up opportunities for electronic detection of charged polymers. With our techniques to fabricate nanoelectrodes with nanochannels, it should be possible to include integrated electronics with nanofludics, allowing the electronic observation of a single DNA molecule at high spatial resolution. However, the present design has problems. Most of the ac went through the silicon wafer instead of the solution. To enhance the sensitivity, we need either to increase the ratio of current going through the liquid to the current going through the wafer or to have a circuit design that picks up the changes in C_dl and R_sol.     

Black silicon as a platform for bacterial detection

Surface-enhanced Raman scattering (SERS) shows promise for identifying single bacteria, but the short range nature of the effect makes it most sensitive to the cell membrane, which provides limited information for species-level identification. Here, we show that a substrate based on black silicon can be used to impale bacteria on nanoscale SERS-active spikes, thereby producing spectra that convey information about the internal composition of the bacterial capsule. This approach holds great potential for the development of microfluidic devices for the removal and identification of single bacteria in important clinical diagnostics and environmental monitoring applications.Plasma etching of silicon can be used to produce inexpensive, large surface area, nano-textured surfaces known as black silicon. Recently, it has been shown that black silicon nano-needles can impale bacteria¹ and that it can be used as a sensor in microfluidic devices.² When coated by a plasmonic metal, such as gold, the nano-textured surface of black silicon is ideal for use as a surface-enhanced Raman scattering (SERS) sensing platform.³ This work aims to investigate whether gold-coated black silicon nano-needles can be used to both impale bacteria and identify them by SERS. This combination of properties would promote the development of microfluidic devices for the removal and monitoring of bacteria in a wide range of medical, environmental, and industrial applications.⁴Black silicon was fabricated by a reactive ion etching technique,⁵ resulting in pyramidal-shaped spikes of height 185 ± 30 nm, full width at half height of 54 ± 10 nm, and 10 ± 2.4 nm radius of curvature at the tip. Samples were then magnetron sputter coated with 200 nm of gold, as this coating thickness was found to provide a suitable compromise between SERS enhancement and impalement efficiency. E. coli (ATCC 25922) from −80 °C stock was isolated on a nutrient agar plate (Difco nutrient broth, Becton Dickinson) for approximately 12 h. A single E. coli colony was then inoculated from the plate into 20 ml of nutrient broth media and incubated overnight at 37 °C with orbital shaking at 200 rpm. The total biomass of overnight culture was adjusted to an optical density of 0.3 at λ = 600 nm by adding fresh sterile nutrient broth (Cary 50 spectrophotometer, Agilent). The E. coli planktonic cells were washed three times by centrifugation at 12 000 rpm (Centrifuge 5804 R, Eppendorf) for 2 min. The washed cells were then re-suspended in a low strength minimum medium (Dulbecco A, phosphate buffered saline). A volume of 100 μl of solution was pipetted onto substrates and left to incubate for 1 h on the bench. Separate sets of samples were created for scanning electron microscope (SEM) imaging, live/dead staining, and SERS. Three sets were needed as each of these measurements altered the samples and left them unsuitable for further analysis.The first set of samples was washed three times with milliQ water after incubation, allowed to dry and then immediately sputter coated with gold using the Emitech K975x (operating current 35 mA, sputter time 32 s, stage rotation 138 rpm, and vacuum of 1 × 10⁻² mbar). SEM imaging was performed with a Zeiss Supra 40VP in high vacuum mode with a working distance of 5 mm and an accelerating voltage of 3 kV. Figure ​Figure11 shows an example of the different levels of impalement that occurred on the black silicon surface. All cells showed signs of damage, but in some cases, the damage was limited to the perimeter of the cell and the main body appeared whole. In other cases, the entire cell had collapsed onto the spikes.Open in a separate windowFIG. 1.A typical SEM image showing E. coli cells with different levels of impalement on gold-coated black silicon.The second set of samples was used for live/dead staining (Invitrogen BacLight Bacterial Viability Kit L7012) with 3.34 mM SYTO 9 (green fluorescence) and 20 mM propidium iodide (red fluorescence). Equal volumes of both dyes were mixed thoroughly in a tube and added to the sample in a ratio of 3 μl of mixed dye to 1 ml of bacterial suspension. After mixing, a volume of 100 μl of the solution was pipetted onto the substrates, which were then incubated at room temperature in the dark for 15 min, before the staining solution was removed by pipetting. The substrates were then washed three times with milliQ water and mounted on a microscope slide for fluorescence imaging. The substrates were not allowed to dry and were stored in phosphate buffered saline at 4 °C when not in use. An epifluorescence microscope (Olympus IX71) with a mercury lamp source and a 60× water immersion objective was used to collect live/dead images from the substrates. Two filter blocks were used to collect the images: U-MNIBA2 blue excitation narrow band delivered green emission (live) and U-MWIG2 green excitation wide band provided red emission (dead).The live/dead image in Figure ​Figure22 shows a mix of both live and dead cells on the black silicon sample. The prevalence of live cells could be due to the incomplete impalement seen under SEM for some cells. It can also be explained by the sample still being wet during live/dead staining. The cells are dried prior to imaging in the SEM and this could weaken the cell wall and allow capillary forces to draw the cells onto the spikes for impalement. This hypothesis is supported by the large number of cells on the stained sample and the presence of cell groupings and cells imaged during mid-division. If the cells were immediately impaled, then such activity would not have been visible and a greater proportion of red cells would be expected.Open in a separate windowFIG. 2.Epifluorescence image showing live (green) and dead (red) E. coli cells after incubation on gold-coated black silicon.The third set of samples was washed three times with milliQ water after incubation and allowed to dry prior to spectral analysis. SERS spectra were collected with a Renishaw inVia Raman spectrometer operating at 785 nm with a 1200 l/mm grating. Power at the sample was 150 mW focused with a 100 × /0.85 NA objective to obtain a diffraction limited laser spot. The resulting spot size (≤2 μm in diameter) is well matched to the size of the bacterial cells. Spectra were collected with three accumulations of 10 s. Data were background subtracted⁶ and normalised to unity for ease of plotting. A great deal of variability was observed in the resulting spectra, as shown in Figure ​Figure33.Open in a separate windowFIG. 3.SERS spectra of E. coli after incubation on a gold-coated black silicon substrate. The spectrum numbers represent single cells at different locations and different levels of impalement.It should be noted that E. coli SERS is known to produce a high level of variability,^7–12 depending on the experimental setup.¹³ However, the variability seen in the SERS spectra of Fig. ​Fig.33 is unusual for measurements performed under consistent experimental conditions. This increased level of variability may be related to the different levels of impalement seen in Fig. ​Fig.1,1, which results in the probing of different internal components. SERS is a surface sensitive technique, with the signal primarily arising within 2 nm of the metal surface.¹⁴ Note that unlike apertureless nanoprobes¹⁵ or conical plasmonic nanotips,¹⁶ the SERS signal in black silicon arises primarily from “hot spots” between the spikes, where the plasmon resonance field is particularly strong.¹⁷ Therefore, depending on the depth and location of impalement, different biomolecules are expected to be excited by this novel substrate.Some peaks occur in the same position for multiple spectra (e.g., peak positions 420, 893, 1001, 1285, and 1307 cm⁻¹), but there are also a lot of unique peaks. The vertical lines in Fig. ​Fig.33 indicate peaks which have appeared in the literature for SERS of E. coli.^7–12 Spectrum 3 has a high proportion of peaks matching published values. This is also the case for spectrum 5, which shares a few peak positions with spectrum 3. Preliminary peak allocations have identified carbohydrates¹¹ (420 cm⁻¹), tyrosine¹¹ (650 cm⁻¹), adenine^10,11 (706 and 735 cm⁻¹), hypoxanthine⁷ (722 and1373 cm⁻¹), phenylalanine⁹ (1001 cm⁻¹), amide III (Ref. ¹⁰) (1285 cm⁻¹), CH₂ deformation¹² (1556 cm⁻¹), and C=C¹⁰ (1587 cm⁻¹).Given the varying levels of impalement observed in the SEM, it appears that the spike shape and Au coating should be further optimized to ensure that the entire cell is consistently pierced and the internal biomolecules are more comprehensively probed. In this way, it may be possible to obtain a more reproducible SERS spectrum of the internal biomolecular constituents of single bacterial cells, thereby providing rapid identification for medical and environmental diagnostic applications. Given that SERS is insensitive to water,⁴ future work should aim to achieve impalement in an aqueous environment, so that the full capability of microfluidics can be used to separate and concentrate suspended bacteria before presenting them to the substrate for rapid analysis.⁴ This suggests a broad range of potential applications in the detection, monitoring, and control of bacterial contamination.     

Preface to Special Topic: Dielectrophoresis

This Special Topic section is on dielectrophoresis, a growing area of widespread interest and relevance to the microfluidics and nanofluidics community.There was a time when the arrival of a telegram from the local post office would foreshadow a step-function change in one’s equilibrium. An internet service provider can now deliver the same effect, as illustrated by an unexpected e-mail from Leslie Yeo inquiring if I would “be interested in guest editing a special issue of Biomicrofluidics on recent advances in dielectrophoresis (DEP).” Flattery directed towards vanity can produce interesting results—which I hope this special issue of Biomicrofluidics demonstrates. The rationale for this special issue is the belief of the journal’s Editors (Dr. Chia Chang and Dr. Leslie Yeo) that dielectrophoresis is a growing area of widespread interest and relevance to the microfluidics and nanofluidics community. Papers, both fundamental and applied, were solicited from the leaders working across this broad interdisciplinary area of research. I was delighted by the positive responses of those whose invited contributions appear in this special issue—efforts certainly not motivated by vanity but through enthusiasm for the subject. Some of those invited to contribute were unable to do so because of other demands on their time. Ongoing advances being made in DEP, especially in its various applications, will surely merit another special issue in the future and hopefully include contributions from those unable to do so now.Two of the papers in this special issue address fundamental aspects of dielectrophoresis (DEP), namely the influences on DEP from electrical double-layers and from particle-particle interactions. Consideration of electrical double layers associated with charged particle surfaces is particularly important for nanoparticles because their effective polarizabilities, associated with field-induced dynamics of the counterions and co-ions in the double layer, can dominate over the intrinsic polarizability of the particle itself. This can influence, for example, to what extent the observation of changes in the DEP crossover frequency (marking the transition between positive and negative DEP) can be relied upon in new immunoassays based on the DEP behavior of functionalized nanoparticles. By considering the electrodynamics of double layers, Basuray et al.¹ propose a theory to predict how the DEP crossover frequency will vary as a function of particle size and the ionic strength of the suspending electrolyte. In their paper, Sancho et al.² derive a theoretical model to describe how particle-particle interactions (e.g., “pearl-chaining”) influence the DEP crossover frequency value. This model also describes well the changes in electrorotation and a newly observed precession effect as particles approach each other under the influence of a rotating field.DEP at the nanoscale is also addressed in contributions from the groups of Ralph Hölzel, Junya Suehiro, and Karan Kaler. Thus, Henning et al.³ describe a new method, based on the measurement of capacitance changes between planar microelectrodes, for the automatic acquisition of the DEP properties of nanoparticles without the need for labeling protocols or visual observations. Suehiro⁴ describes how DEP can be employed as a bottom-up approach for fabricating nanomaterial-based devices such as a carbon nanotube gas sensor and a ZnO nanowire photosensor. Kaler et al.⁵ describe how the DEP manipulation of miniscule amounts of polar aqueous samples, a method known as liquid-DEP, can be used for on-chip bioassays, such as nucleic acid analysis, and through parallel sample processing offer the potential for conducting automated multiplexed assays. The use of DEP to selectively trap and separate cells has been investigated over many years, and contributions from the groups of Hywel Morgan, Ana Valero, Masau Washizu, and Gerard Markx describe the latest advances and applications. Thomas et al.⁶ describe a new automated DEP cell trap design for the isolation, concentration, separation, and recovery of human osteoblast-like cells from a heterogeneous population. Recovery of small populations of human osteoblast-like cells with a purity of 100% is demonstrated. A cell-sorting device, based on the opposition of DEP forces that discriminates between cell types according to such properties as their membrane permittivity and cytoplasm conductivity, is described by Valeroet al.⁷ The versatility of the device is demonstrated by synchronizing a yeast cell culture at a particular phase of the cell cycle. Gel et al.⁸ describe a DEP-assisted cell trapping method for fusing pairs of cells in an array of micro-orifices. This method produces not only a high yield of viable cell fusants, but also allows for subsequent study of postfusion cell development. Zhu et al.⁹ describe a DEP-based microfluidic separation system in which dead and active cells can be collected from a given cell suspension, whilst at the same time eluting dormant cells. In the second paper from Gerard Markx’s group, Zhu et al.¹⁰ demonstrate that the rate-limiting resuscitation of a colony of dormant bacteria is determined by the diffusion of a resuscitation-promoting factor into the colony interior. This study involved the artificial engineering of different sizes and shapes of bacterial aggregates using DEP forces. Finally, in my own contribution,¹¹ I have attempted to summarize the growing output of DEP publications in terms of their contributions to the theory, technology, and applications of DEP.     

Administering the Optimum Dose of l-Arginine in Regional Tumor Therapy

The purpose of this study is optimizing the l-arginine (l-Arg) doses on the basis of chemical structure in regional accessible tumor therapy to settle down a new protocol for the treatment of cancer. ³H-thymidine-based cell proliferation assay was performed in vitro on tumor cell lines of fibrosarcoma (FS), lymphosarcoma-ascitic and on normal cell line of NIH 3T3 after treatment with different concentrations of l-Arg in phosphate buffered saline (PBS). The cultures were harvested after 22 h and the incorporated radioactivity was counted to identify their histologic grades as described in earlier studies. In vivo therapy of murine tumors was conducted where FS cells injected subcutaneously at ventro-lateral position of mice. Various drug delivery schedules were injected into the centre of tumor base, once a day for 4 days. Tumor diameter and survivals were monitored where the day of sacrifice was considered for monitoring the survival period. By identifying the histologic grades of the treated cultures in vitro and in vivo by different concentrations of l-Arg, the corresponding energy of such concentrations were determined. An efficient model with a good fit (R² = 0.98) was established to describe the energy yield by l-Arg dose. The equivalence between the tumor histologic grade and energy of the l-Arg dose delivered in saline (PBS) environment is the optimum condition for regional tumor therapy achieves higher survival rate. The selective cytotoxicity to tumor cells with minimal damage to normal cells by l-Arg due to its chemical structure suggests to be considered the most promising drug for regional therapy of the accessible tumors like breast cancers of early stage with no distant metastasis.     

Preface to Special Topic: Papers from the 13th International Conference on Surface and Colloid Science (ICSCS) and the 83rd ACS Colloid and Surface Science Symposium, Columbia University, New York, 2009

This Special Topic section is a compilation of several original contributions covering both fundamental and practical aspects of electrokinetic microfluidic phenomena that were presented during the Electrokinetics and Microfluidics sessions held at the conference.Electrokinetics is currently the mechanism of choice for the manipulation of fluids as well as colloidal and biological particles at microscale and nanoscale dimensions.¹ The popularity of electrokinetics is perhaps not so surprising as electrodes are easy to fabricate and embed into microfluidic chips, thus allowing the entire fluid and particle actuation mechanism to be completely integrated into the device. In addition, driving microfluidics with electric fields is relatively straightforward and allows for precise actuation. Nevertheless, considerable challenges remain in understanding the complex mechanisms associated with the hydrodynamics of conducting and dielectric fluids and particles under the influence of electric fields. Concomitantly, there has been an exponential increase in research and development in this field along both fundamental and applied themes in the past five years.This sustained growth in the microfluidics community of electrokinetics research has led to a sequel to the first Electrokinetic Phenomena and Microfluidics session at the 82nd ACS Colloid and Surface Science Symposium in Raleigh, NC, in 2008, and which we hope will now be a regular feature at successive ACS Colloid and Surface Science meetings. This year at the combined 2009 13th International Conference on Surface and Colloid Science (ICSCS) and the 83rd ACS Colloid and Surface Science Symposium in New York, the Electrokinetics and Microfluidics symposium proved to be extremely popular, with three keynote lectures presented by Professor Howard Stone, Professor Hsueh-Chia Chang, and Professor Thomas Healy, and 44 oral presentations. In both 2008 and 2009, Biomicrofluidics has organized a special issue to cover some of the contributions reported at these meetings.²The growing interest in using electric fields to manipulate biological entities such as cells, DNA, and even single molecules is reflected in this year’s collection of papers with dielectrophoretic (DEP) phenomena comprising the bulk of the contributions. In Ref. ³, a new theory to describe Stern layer conductance along the surface of nanocolloids is proposed, forming the basis for the derivation of a more accurate prediction of the DEP crossover frequency. This theory is then employed to determine the conformation and, hence, optimum coverage of oligonucleotides on the surface of nanocolloid functionalized molecular probes during DNA hybridization under the influence of DEP, which can be exploited for biomolecular sensing. Other fundamental DEP papers include the investigation of particle motion under DEP induced optically via a photoconductor, in which Zhu et al.⁴ characterized the frequency dependence of the motion through the synchronous velocity spectra of the particles, and a numerical study of particle trapping at the throat of converging-diverging microchannels under the influence of negative DEP using a transient arbitrary Lagrangian–Eulerian finite element method.⁵ A more practical implementation is, on the other hand, reported by Yang et al.⁶ in which the negative DEP is exploited to separate colorectal cancer cells from other cells in a microfluidic device as a demonstration of a portable cancer detection tool.Continuing along the separation theme, but with regard to DNA separation using pulsed-field gel electrophoresis aided by sparse but regularly ordered microfabricated arrays of nanoposts, is a Brownian dynamics simulation model reported by Ou et al.⁷ in which DNA channeling, which predicts that the motion of DNA is undisturbed by the presence of arrays for large spacing to DNA equilibrium size ratios and when the field lines are straight, is predicted, consistent with experimental observations. In another fundamental paper, a direct numerical simulation model is presented to predict the current-voltage relationship across conducting pores along cell membranes, which is of fundamental importance in the electroporation process.⁸We hope that you will enjoy reading the contributions in this special topic and that it encourages you to participate in future Electrokinetics and Microfluidics meetings at the ACS Colloid and Surface Science Symposia, which we definitely hope will continue on a regular basis.     

New types of topological superconductors under local magnetic symmetries

We classify gapped topological superconducting (TSC) phases of one-dimensional quantum wires with local magnetic symmetries, in which the time-reversal symmetry is broken, but its combinations with certain crystalline symmetries, such as , , and , are preserved. Our results demonstrate that an equivalent BDI class TSC can be realized in the or superconducting wire, which is characterized by a chiral Z^c invariant. More interestingly, we also find two types of totally new TSC phases in the and superinducting wires, which are beyond the known AZ class, and are characterized by a helical Z^h invariant and Z^h⊕Z^c invariants, respectively. In the Z^h TSC phase, Z pairs of Majorana zero modes (MZMs) are protected at each end. In the case, the MZMs can be either chiral or helical, and even helical-chiral coexisting. The minimal models preserving or symmetry are presented to illustrate their novel TSC properties and MZMs.     

Two-dimensional and three-dimensional dynamic imaging of live biofilms in a microchannel by time-of-flight secondary ion mass spectrometry

A vacuum compatible microfluidic reactor, SALVI (System for Analysis at the Liquid Vacuum Interface), was employed for in situ chemical imaging of live biofilms using time-of-flight secondary ion mass spectrometry (ToF-SIMS). Depth profiling by sputtering materials in sequential layers resulted in live biofilm spatial chemical mapping. Two-dimensional (2D) images were reconstructed to report the first three-dimensional images of hydrated biofilm elucidating spatial and chemical heterogeneity. 2D image principal component analysis was conducted among biofilms at different locations in the microchannel. Our approach directly visualized spatial and chemical heterogeneity within the living biofilm by dynamic liquid ToF-SIMS.Mapping how metabolic pathways are interconnected and controlled at the subcellular scale within dynamic living systems continues to present a grand scientific challenge. Biofilms, consisting of aggregations of bacterial cells and extracellular polymeric substance (EPS), present an important avenue for deciphering complex microbial communities. During biofilm formation, cells assemble in a secreted polymer milieu of polysaccharides, proteins, glycolipids, and DNA.^1,2 Microfluidics provides unprecedented control over flow conditions, accessibility to real-time observation, high-throughput testing, and mimics in vivo biological environments.³ An understanding of the mechanism underlying biofilm formation and the design of advanced microfluidic experiments could address challenges such as interpreting microbial community interactions, biofouling, and resistance to antimicrobial chemicals. However, only a handful of biofilm studies used microfluidic approaches that provided hydrated chemical imaging at high spatial resolution.^4–7 Most studies utilized confocal microscopy,⁴ FTIR spectroscopy,⁵ or other approaches (e.g., high density interdigitated capacitors⁷) for biofilm monitoring. Imaging mass spectrometry has been demonstrated in biofilm studies.^8,9 A coupled microfluidic-imaging mass spectrometry approach would provide the chemical molecular spatial mapping needed to better address the scientific challenge of biofilms.Recently, we developed a portable microfluidic reactor, System for Analysis at the Liquid Vacuum Interface (SALVI),^10,11 which overcame the grand challenge of studying liquids with high volatility and liquid interfaces using surface sensitive vacuum instruments. SALVI enables direct imaging of liquid surfaces using electron or ion/molecular based vacuum techniques. Our microfluidic approach used a polydimethylsiloxane (PDMS) microchannel fully enclosed with a thin silicon nitride (SiN) membrane (100 nm thick). For visualization, 2 μm diameter holes were opened in the SiN membrane in vacuo. These detection windows were dynamically drilled using the time-of-flight secondary ion mass spectrometry (ToF-SIMS) primary ion beam (e.g., Bi⁺).¹²Unlike liquid sample holders for transmission electron microscopy and scanning transmission electron microscopy, SALVI is self-contained and portable.¹³ As a result, it can potentially be used in many finely focused analytical tool with minimal adaptation.¹⁰ The analytical performance of SALVI has been demonstrated with a variety of analytes ranging from biology to material sciences.^14,15 Unlike most microfluidic applications that are only suitable under ambient conditions (e.g., separations, cell and small amount sample manipulation, and thermal flow-sensors),^16–18 SALVI is compatible with both in situ ambient and in vacuo spectroscopy analysis and imaging.¹⁹ Biofilms have been successfully cultivated inside the microfluidic channel and imaged using correlative confocal laser scanning microscopy (CLSM) and ToF-SIMS.²⁰Our approach opens a new avenue to study biological sample in their natural state. Although ToF-SIMS has been widely used for providing molecular signatures of organic and biological molecules in complex biological systems^21,22 or lipid spatial mapping,²³ the vacuum-based ToF-SIMS generally requires solid (either dried²⁴ or cryo treated²⁵) samples. Here, we report ToF-SIMS two dimensional (2D) and three dimensional (3D) chemical images of hydrated biofilms. In situ time and space-resolved identifications of fatty acid (FA) fragments characteristic of Shewanella are illustrated by 3D images reconstructed from the ToF-SIMS depth profile time series. Principal component analysis (PCA) further elucidates biofilm chemical and spatial heterogeneity and shows the key chemical component at different depth and location of the biofilm including the biofilm-surface attachment interface.For all growth experiments, two samples were cultured simultaneously. At days 5 and 6, one sample was harvested for immediate analysis, respectively, using a ToF-SIMS V spectrometer (IONTOF GmbH, Münster, Germany). Similar results were obtained from both samples, because the biofilm-attachment surface was probed. For consistency, only day 6 data are shown here, while additional data are provided in the supplementary material.²⁸ 2D and 3D image visualizations were obtained using the IONTOF instrument software. PCA was performed using MATLAB R2012a (MathWorks, Inc., Natick, MA, USA). 2D images of .bif format were converted and integrated into a matrix. Data were pretreated by normalization to total ions, square root transformation, and then mean centering.²⁶ For m/z spectra PCA, unit mass peaks from m/z 199 to m/z 255 were used (see Figure S-2²⁸). Unit mass peaks from m/z 1–300 were also used and results are comparable (see Figure S-3²⁸). Five characteristic FA peaks (m/z 199, 213, 227, 241, and 255, corresponding to C12, C13, C14, C15, and C16 FAs) were used in image PCA.²⁷ Images representing each PC were reconstructed from the score matrix using the red, green, and blue (RGB) color scale.Using depth profiling, we drilled through the SiN membrane and collected depth-resolved images of the live biofilm (Figure 1(a)). Our analysis of the negative ToF-SIMS spectra after SiN punch-through showed Shewanella FA fragments in the m/z 195–255 range.²⁰ From the depth profile time series, we selected five regions (highlighted as I, II, III IV, and V) within the FA m/z range to visualize 2D spatially resolved images collected for 46 s (1000 scans) before (I), during (II), or after (III, IV, V) SiN membrane punch-through.²⁰ When false color 2D images of FA fragments characteristic of Shewanella biofilms were selected from the dynamic depth profiling data, differences were observed (Figure 1(b)) among the five regions. Furthermore, the biofilm images after SiN membrane punch-through (III, IV, V) displayed variations across the 2 μm diameter surfaces, with C12 (m/z 199) being distributed across regions III, IV, and V and C15 (m/z 241) FAs mostly in region V (see Figure S-4 for additional FA images²⁸). This suggested that depth-resolved chemical heterogeneities were present in the biofilm. To illustrate, we reconstructed the 2D images from depth profiling data within the biofilm region (from the beginning of III through the end of V) and show spatially resolved 3D chemical images within the entire sample (Figure 1(c) and movies S1-S3²⁸). The reconstructed 3D images revealed the heterogeneous spatial distribution overlay for C12 (red) and C15 (green) FAs during 302 s biofilm depth profiling from day 5 (Figure S-5²⁸) and day 6 (Figure 1(c)).Open in a separate windowFIG. 1.(a) ToF-SIMS depth profiling of the day 6 biofilm attached to the SiN membrane in the microfluidic channel. Five regions representing sample before SiN punch-through (I) during punch-through (II) or within the biofilm region (III, IV, and V) are illustrated. (b) 2D false color images of day 6 biofilm FAs at the five time regions highlighted in (a). (c) Reconstructed 3D day 6 biofilm images showing FA fragment distributions within the entire biofilm region (III–V, 302 s). The time axis represents depth profiling from near the SiN surface into the biofilm. (d) Spectra PCA score plot of day 6 biofilm showing the differences and similarities among selected five regions (m/z 199–255). A 95% confidence limit for each region was defined by an ellipse with the same color to the corresponding region clusters. (e) Loadings of PC1 and PC2 corresponding to (d) and the plot of PC variance contributions.Spectral PCA was used to analyze the m/z spectra. The deepest region (V) into the biofilm was the most different from the other two biofilm regions (III and IV), further confirming the heterogeneities observed in the 2D images (e.g., C12 and C15 FA fragments) contributing most to this spatial difference. In addition, C12 FA fragments played a key role in the biofilms imaged near the SiN membrane attachment surface (III and IV). When inspected individually, C12 FAs were observed throughout the entire biofilm region, suggesting that C12 FA fragments may play a role in biofilm attachment to a surface and they may be main components of EPS throughout the biofilm. In contrast, C15 FAs were more abundant deeper within the biofilm, indicating that they may be more relevant to bacteria cells themselves.Uniform sputtering rate was assumed during depth profiling. To better determine the depth and shape of the SIMS ionization crater, AFM measurements were collected using an agarose sample in the SALVI reactor as a proxy for the biofilms (Figure S-6²⁸). The AFM results showed that the 100 nm SiN was drilled through and confirmed that the biofilm interface was probed by ToF-SIMS. Ideally, real-time correlative AFM and ToF-SIMS measurements will be needed due to the self-healing property of biofilms. However, such capability is currently under development.To further analyze chemical differences within biofilms, we performed ToF-SIMS depth profiling at three locations along the microchannel; namely, the inlet, center, and outlet as illustrated in Figure S-1(b).²⁸ At each location, we defined the five regions described in Figure 1(a), and 2D image PCA analysis was conducted on the biofilm region (from the beginning of III through the end of V) to visualize the chemical distributions on day 6. Figure 2(a) shows the loading plots for the m/z peaks that contribute to each PC image (Figure 2(b)). The first three PCs explained 93.79% of the variance within the data. For PC1, the strongest positive loading fragments were C12 and C15 FAs, which are the bright red areas in three PC1 images. The C12 FAs were the main contributor to the green regions in the PC2 image. The strongest loading for PC3 in blue was C14 FAs. Compared to PC1 and PC2, PC3 played a limited contribution to the overall spatial distribution discrimination. The merged images give a demonstration of chemical spatial distribution of key components of biofilms in the liquid microenvironment.Open in a separate windowFIG. 2.(a) Image PCA loading plots illustrating the contribution of each FA peak in the day 6 biofilm at three locations within the microfluidic channel. The variance contributions of each PC are shown at the bottom. (b) Reconstructed false-color 2D PCA images in RGB corresponding to each PC scores at these locations along the microfluidic channel. The RGB composite images of the three key PCs are depicted in the bottom. Only data within the 2 μm diameter circle were considered in analysis.Our results show that SALVI and liquid ToF-SIMS studies of live biofilms offer dynamic, depth-resolved chemical mapping and produce 2D and 3D visualizations of spatial heterogeneity within a biofilm. Chemical imaging of biofilms near the attachment interface can enhance our understanding of biofilm formation in environmental, medical, and industrial settings. Our approach provides a universal portable platform and enables in situ probing of complex living biological systems potentially across multiple time and space scales. Because of the portability and vacuum compatibility, SALVI offers a valuable linkage with proteomic mass spectrometry via microfluidics and a nondestructive package for integrative in situ analysis of live biological systems in system biology.     

Changes in blood and erythrocyte membrane parameters in sodium pentosan polysulphate treated urolithiatic rats

The effect of sodium pentosan polysulphate (SPP), was studied in relation to certain blood and erythrocyte membrane parameters in calcium oxalate stone forming rats. Calcium oxalate stones were induced by feeding the rats with 3% w/w sodium glycollate. Fibrinogen, haemoglobin and serum protein levels did not show any variation with the treatment procedures. Serum mucoprotein and protein bound carbohydrates-hexosamine and sialic acid-were increased significantly in the rats receiving calculogenic (CPD) and attained nearly normal levels with SPP treatment. In contrast, hexuronic acid level was decreased in the CPD group and SPP administration increased the level of hexuronic acid in the treated groups. Erythrocyte membrane Ca²⁺-ATPase activity was increased in stone forming rats and SPP administration brought a reduction in the above enzyme activity. Changes in Membrane Mg²⁺- and Na⁺, K⁺-ATPases were minimal. Membrane cholesterol and phospholipids were also raised significantly in stone formers, SPP treatment reduced the membrane cholesterol levels in both controls and stone formers. Phospholipids were also decreased moderately. The above observations suggest that SPP is safe for administration in urolithiatic condition without adverse effects.