Bacteria under the physical constraints of periodic micro-nanofluidic junctions reveal morphological plasticity and dynamic shifting of Min patterns     

Jie-Pan Shen  Chia-Fu Chou 《Biomicrofluidics》2014,8(4)

Morphological plasticity is an important survival strategy for bacteria adapting to stressful environments in response to new physical constraints. Here, we demonstrate Escherichia coli morphological plasticity can be induced by switching stress levels through the physical constraints of periodic micro-nanofluidic junctions. Moreover, the generation of diverse morphological aberrancies requires the intact functions of the divisome- and elongasome-directed pathways. It is also intriguing that the altered morphologies are developed in bacteria undergoing morphological reversion as stresses are removed. Cell filamentation underlies the most dominant morphological phenotypes, in which transitions between the novel pattern formations by the spatial regulators of the divisome, i.e., the Min system, are observed, suggesting their potential linkage during morphological reversion.Most bacteria have evolved sophisticated systems to manage their characteristic morphology by orchestrating the spatiotemporal synthesis of the murein sacculus (peptidoglycan exoskeleton), which is known to be the stress-bearing component of cell wall and presides over de novo generation of cell shape.¹ Morphological plasticity is attributed to a bacterial survival strategy as responding to stressful environments such as innate immune effectors, antimicrobial therapy, quorum sensing, and protistan predation.² It comes of no surprise that stress-induced diversified morphology and mechanisms, ascribed to shape control and determination, have drawn great attention in both fundamental and clinical studies.^3–6 The molecular mechanism to form filamentous bacteria has been revealed that both β-lactam antibiotics³ and oxidative radicals produced by phagocytic cells⁵ trigger the SOS response, promoting cell elongation by inactivating cell division via the blockade of tubulin-like FtsZ, known as the divisome initiator. While apart from the scenario of length control by the divisome-directed filamentation, the elongasome assembled by proteins associated with actin-like MreB complex^1,7,8 helps the insertion of peptidoglycans into lateral cell wall, suggesting the role in the determination of cell diameter during cell elongation.Recently, additional mechanisms other than the divisome/elongasome-directed pathways of shape maintenance are discovered to regenerate normal morphology de novo from wall-less lysozyme-induced (LI) spheroplasts of E. coli via a plethora types of aberrant division intermediates.⁹ Similar morphological reversion from different aberrant bacterial shapes has been observed as squashed wild-type bacteria generated through sub-micron constrictions are released into connected microchambers.¹⁰ Previous work using the microfluidic approach focuses on the septation accuracy and robustness of constricted bacteria,¹¹ but the reversion process of stress-released bacteria is not well studied and analyzed. In particular, the aberrant bacterial shape is mainly branched-type with bent and curved variants in the reverting bacteria, analogous to the aberrant intermediate found in the morphological reversion of LI spheroplasts with PBP5-defective mutant.⁹ Since bacteria suffering from starvation¹² or confronting mechanical stresses exerted by phagocytosis and protistan grazing⁶ can induce morphological alterations, one could manipulate the stress levels of physical constraints by adopting repeated structures of sub-micron constricted channels (nanoslits) and microchambers,^10,11 to select and enrich bacteria converting to specified aberrant intermediates. The stress incurred by the nanoslit on bacteria is about the mechanical intervention over de novo synthesis of the cell wall, which is the major factor causing morphological aberrancy, while the second environmental stress comes from bacterial growth in the restricted space of microchamber as bacteria proliferate to full confluency, resulting in growth pressure of high population density, nutrient deficiency, and the size reduction of bacteria.Here, we report the selection of distinctive bacterial morphologies by size shrinkage in the outlet cross-section (W × H = 1.5 × 1.5 μm) of the terminal microchamber in the periodic structures of nanoslit-microchamber (Figs. 1(a) and 1(b)). The fluidic structures were micropatterned on fused silica wafers by photolithography, fabricated through reactive ion etching (RIE) and inductively coupled plasma (ICP) etching, and encapsulated by cover glasses coated with polydimethylsiloxane (PDMS) or polysilsesquioxane (PSQ) layer as described earlier.^13,14 Two days after the outgrowth of Escherichia coli (imp4213 [MC4100 ΔlamB106 imp4213]) loaded to the microfluidic device at 25 °C, bacteria started to penetrate into the nanoslit as they proliferated to full confluency in the first microchamber (Fig. 1(c)). It takes about 10 days for bacteria traversing 500 μm long (5 repeated nanoslit-microchamber units) via proliferations and being released from the outlet of the terminal microchamber. The narrowed outlet allows only bacteria with smaller diameters to be squeezed into the spacious and nutrient-rich region, thus it acts as a spatial filter to avoid the passage of branching bacteria with cross-sectional size larger than that of the outlet. The rationale of this design is to select aberrant bacteria prone to promote de novo shape regeneration other than the branched-type, which is the dominant morphology of reverting bacteria in the prior microfluidic constriction study.¹⁰ As anticipated, the stress-released bacteria through the narrowed outlet are therefore mostly filamentous (see statistical analysis for cell morphology in the supplementary material).¹⁵ However, it is noted that the aberrant morphology of lemon-like shape with tubular poles (Figs. 1(d-1), 1(d-3), and 1(d-11)) is developed about 3 h after the stress-released bacteria escaped through the outlet. Though the generation of the lemon-like aberrancy in bacteria has been reported in PBP5/7-defective E. coli mutant subjected to a high-level inhibition of both MreB and FtsZ, while the same mutant treated with low-level MreB inhibitor, together with antagonized-FtsZ, displays filamentous shape with varying diameters,¹⁶ these morphological aberrances can be observed in our system (Figs. 1(d-2) and 1(d-12)). Besides, a high-level inhibition of MreB in E. coli with an intact divisome function is known to cause round bacteria, resembling to the cell morphology of the bacteria shown in Fig. 1(d-4). Interestingly, parallel experiments using bacteria mutants carrying impaired regulatory functions in either the divisome (Min⁻) or the elongasome (MreB⁻) do not develop morphological plasticity (supplementary Fig. S1).¹⁵ Taken together, the filamentous and lemon-like variants selected from our microfluidic platform, while elaborating the morphological plasticity and reverting progression, require both the functional divisome/elongasome. Alternatively, the selection by the spatial filter does not fully exclude cells with aberrant shapes such as the branched-type with initial budding (Fig. 1(d-7)), cells with asymmetric cross-section perpendicular to the longitudinal axis (Figs. 1(d-2), 1(d-8), 1(d-9), 1(d-9′), and 1(d-10)), and those resembling to the morphological phenotypes of the division intermediates reported in the LI-spheroplasts carrying genetic defects on some non-cytoskeletal proteins (Figs. 1(d-5) and 1(d-6)). In particular, intracellular vesicles and cell autolysis are observed in some reverting bacteria (Figs. 1(d-5) and 1(d-6)), which are reminiscent to the phenomena reported in the division intermediates of the LI-spheroplasts lacking stress response system (Rcs) or some accessory proteins (PBP1B and LpoB). Unlike the bacteria grow with odd shapes under the stress of nanofluidic confinement only¹⁰ (Fig. 1(c)), all the morphological aberrancy reported here are developed in the reverting bacteria, which grow in the spacious and nutrition-rich environment and are free from physical constraints. Further investigations over the expression levels of the divisome/elongasome networks and the stress-response system in bacterial cells subjected to micro-nanofluidic junctions could be insightful in understanding their role in bacterial shape control.⁹Open in a separate windowFIG. 1.(a) Schematics of the microfluidic device used in this study with an H-shaped geometry (left upper panel), where repeated nanoslit (L×W×H = 50×10×0.4 μm)−microchamber (L×W×H = 50×50×1.5 μm) structures are bridged between two arms of the H-shaped microchannels (left lower panel and enlarged view in right panel). (b) Top-view layout of an individual channel in (a) with close view of the outlet in the terminal microchamber (orange: nanoslits; blue: microchambers). (c) Fluorescence micrograph of E. coli imp4213 penetrating a nanoslit (scale bar: 5 μm). (d) Bright-field micrographs for various cell morphology of the selected imp4213 released from the outlet (magenta arrows: cells with vesicles; scale bar: 5 μm). (e) Sequential bright-field micrographs of morphological reversion. T1–T3 indicate the time after bacteria escaping from the outlet. T1: 3 h; T2: 6 h; T3: 24 h. Scale bar: 10 μm.During the morphological reversion, the stress-released bacteria rapidly increase their size in the first 3 h after escaping from the terminal microchamber (T1 in Fig. 1(e)). Some filamentous bacteria even grow over 50 μm long, though such a morphological phenotype implicates the cessation of functional divisome. With active growth and proliferation, the progeny of stress-released bacteria increase their population but gradually reduce their size about 6 h after being released from the constriction stress (T2 in Fig. 1(e)). Fig. ​Fig.22 displays the marginal histograms for different shape factors, where Fig. 2(a) is the plot of the minimal Feret diameter (cell diameter) versus Feret diameter (cell length), i.e., the shortest versus the longest distance between any two points with parallel tangents along the cell peripheral, respectively, indicating that cell diameters are larger for reverting bacteria at T1 (mean ± S.E.M. = 1.89 ± 0.08 μm) with respect to T2 (1.51 ± 0.06 μm). Moreover, the histogram of Feret diameter depicts two major populations of the cell length for reverting bacteria at T1, which mostly resume to typical cell length at T2 (the median of Feret diameter = 3.33 μm; see statistical analysis for Fig. ​Fig.22 in the supplementary material).¹⁵ The shape factors of circularity (4π × [area]/[perimeter]²) and aspect ratio ([major axis]/[minor axis] for the cell geometry fitted to an ellipse) confirm the existence of dual populations for bacteria at T1 as well (Fig. 2(b)). About 24 h after escaping (T3 in Fig. 1(e)), almost all the progeny of stress-released bacteria regained the rod shape.Open in a separate windowFIG. 2.Marginal histograms for shape factors measured from the reverting imp4213 at T1 and T2. (a) Minimal Feret diameter (cell diameter) versus Feret diameter (cell length). (b) Circularity versus aspect ratio. N = 366 for T1 and N = 494 for T2.The bacterial size reduction of filamentous and lemon-like shape variants, though involving negative control of the divisome positioning by the spatial regulators of MinCDE system,¹⁷ is not completely understood as to how they coordinate in aberrant geometries. Besides, the filamentation of stress-released bacteria during the period of T1 to T2 implicates the inhibition of functional divisome. With minimal perturbation of the divisome by leaky expression of GFP-MinD and MinE (imp4213/P_lac-gfpmut2::minD minE), the patterning dynamics of GFP-MinD in different bacterial morphology were time-lapse imaged during morphological reversion. Intriguingly, more than the standing-wave-like pattern of MinD denoted in filamentous E. coli,¹⁸ we discovered bidirectional drifting of two standing-wave-like patterns of MinD occur in most reverting bacteria filaments (supplementary Figs. S2(a) and S2(b)).¹⁵ The bidirectional drifting in the longitudinal direction of the cells may be emanating from the cell poles (the blue upper panel of Fig. 3(a) and supplementary Fig. S2(c)¹⁵) and the cylinder region (the blue lower panel of Fig. 3(a) and supplementary Fig. S2(d)¹⁵). Furthermore, the MinD pattern transitions from the standing to traveling waves are occasionally observed (the lower panel of Figs. 3(a) and supplementary Fig. S2(e)¹⁵). Notably, the standing-wave-like MinD patterns exhibit bidirectional drifting along the cell longitudinal direction and intermittently change directions, implying the competition between coexisting MinD patterns can be supported under filamentous geometry. Despite there have been observations of multiple wave-packet of traveling waves in filamentous cells,¹⁹ the mixture of distinct wave-like MinD patterns have never been experimentally reported. While most intriguingly, multiple drifting movements of wave-like MinD patterns potentiate the mitigation of periodic minima in time-averaged Min gradient in the reverting filamentous bacteria, suggesting the disability of proper divisome positioning for recovering the typical rod shape. Apart from the wave-like movements, amoeba-like motion of Min proteins has been shown in vitro upon synthetic minimal system, but never been verified in vivo.²⁰ Strikingly, here amoeba-like motion of MinD is the dominant mode in lemon-like bacteria and the transitions between wave-like patterns and amoeba-like motion are supported even under filamentous geometry (Figs. 3(b) and 3(c), Multimedia view).Open in a separate windowFIG. 3.Kymographs for GFP-MinD dynamics in selected imp4213 cells during morphological reversion: (a) Mixture modes of standing wave packets and traveling wave. The left panel is the stacked fluorescence micrograph displaying cell shape (scale bar = 5 μm). The kymograph is derived from the filamentous cell indicated by the green arrow (scale bar: 120 s horizontal; 5 μm vertical), where the lower panel follows the upper panel in time. The yellow windows indicate bidirectional-drifting standing wave packets, while the green indicates traveling waves (see also supplementary Fig. S2).¹⁵ (b) Sequential fluorescence micrographs of GFP-MinD in lemon-shape imp4213 show amoeba-like motion, with the first left a bright-field image (scale bar: 10 μm). (c) Mixed modes of amoeba-like motion and waves in selected filamentous imp4213 cell indicated by the green arrow in the left panel (scale bar = 5 μm). The filamentous cells depicted in (a) and (c) locate at the top region while the lemon-shape cell in (b) at the central region of the movie (time stamp in min:s). (Multimedia view) [URL: http://dx.doi.org/10.1063/1.4892860.1]In summary, we have demonstrated that the development of bacterial morphological plasticity can be stress-induced by periodic physical constraints with intact functions of the divisome and elongasome-directed pathways. Through size exclusion, the constricted outlet structure designed in our microfluidic device is useful in selecting bacteria with plethora morphological aberrancies other than the branched type. Interestingly, disparate morphological changes, rather than those being directly induced under a stressful environment, can be generated in the stress-released bacteria experiencing morphological reversion. Further, the discovery of novel transitions between the Min patterns in most reverting bacteria implicates its regulatory effect of cell filamentation. However, by exploiting the micro-nanofluidic approach, further investigations of the mechanism underlying the development of morphological plasticity in bacteria adapting to physical constraints are expected in future studies to gain more insights into the molecular basis of shape generation.  相似文献   

Magnetographic array for the capture and enumeration of single cells and cell pairs     

C. Wyatt Shields  IV  Carissa E. Livingston  Benjamin B. Yellen  Gabriel P. López  David M. Murdoch 《Biomicrofluidics》2014,8(4)

We present a simple microchip device consisting of an overlaid pattern of micromagnets and microwells capable of capturing magnetically labeled cells into well-defined compartments (with accuracies >95%). Its flexible design permits the programmable deposition of single cells for their direct enumeration and pairs of cells for the detailed analysis of cell-cell interactions. This cell arraying device requires no external power and can be operated solely with permanent magnets. Large scale image analysis of cells captured in this array can yield valuable information (e.g., regarding various immune parameters such as the CD4:CD8 ratio) in a miniaturized and portable platform.The emergent need for point-of-care devices has spurred development of simplified platforms to organize cells across well-defined templates.¹ These devices employ passive microwells, immunospecific adhesive islands, and electric, optical, and acoustic traps to manipulate cells.^2–6 In contrast, magnetic templating can control the spatial organization of cells through its ability to readily program ferromagnetic memory states.⁷ While it has been applied to control the deposition of magnetic beads,^8–13 it has not been used to direct the deposition of heterogeneous cell pairs, which may help provide critical insight into the function of single cells.^14,15 As such, we developed a simple magnetographic device capable of arraying single cells and pairs of cells with high fidelity. We show this magnetic templating tool can use immunospecific magnetic labels for both the isolation of cells from blood and their organization into spatially defined wells.We used standard photolithographic techniques to fabricate the microchips (see supplementary material¹⁶). Briefly, an array of 10 × 30 μm cobalt micromagnets were patterned by a photolithographic liftoff process and overlaid with a pattern of dumbbell-shaped microwells formed in SU-8 photoresist (Fig. 1(a)). The micromagnets were designed to produce a predominantly vertical field in the microwells by aligning the ends of the micromagnet at the center of each well of the dumbbell. These features were deposited across an area of ≈400 mm² (>50 000 well pairs per microchip) (Fig. 1(b)). Depending on the programmed magnetization state with respect to the external field, magnetic beads or cells were attracted to one pole and repelled by the other pole of each micromagnet, leading to a biased deposition (Fig. 1(c)).¹²Open in a separate windowFIG. 1.Magnetographic array for single cell analysis. (a) SEM image of the dumbbell-shaped well pairs for capturing magnetically labelled cells. (b) Photograph of the finished device. (c) An array of well pairs displaying a pitch of 60 × 120 μm before (top) and 10 min after the deposition of magnetic beads (bottom).To demonstrate the capability of the array to capture cells into a format amenable for rapid image processing, we organized CD3+ lymphocytes using only hand-held permanent magnets. We isolated CD3+ lymphocytes from blood via positive selection using anti-CD3 magnetic nanoparticles (EasySep™, STEMCELL Technologies) with purities confirmed by flow cytometry (97.8%; see supplementary material¹⁶). We then stained 1 × 10⁶ CD3+ cells with anti-CD8 Alexa-488 and anti-CD4 Alexa-647 (5 μl of each antibody in 100 μl for 20 min; BD Bioscience) to determine the CD4:CD8 ratio, a prognostic ratio for assessing the immune system.^17,18Variably spaced neodymium magnets (0.5 in. × 0.5 in. × 1 in.; K&J Magnetics, Inc.) were fixed on either side of the microchip to generate a tunable magnetic field (0–400 G; Fig. 2(a)). Using this setup, fluorescently labeled cells were deposited, and the populations of CD4+ and CD8+ cells were indiscriminately arrayed, imaged, and enumerated using ImageJ. The resulting CD4:CD8 ratio of 1.84 ± 0.18 (Fig. 2(b)) was confirmed by flow cytometry with a high correlation (5.4% difference; Fig. 2(c)), indicating the magnetographic microarray can pattern cells for the rapid and accurate assessment of critical phenotypical parameters without complex equipment (e.g., function generators or flow cytometers).Open in a separate windowFIG. 2.CD8 analysis of CD3+ lymphocytes. (a) Photograph of the magnetographic device activated by permanent magnets (covered with green tape). The CD4:CD8 ratio determined by the (b) magnetographic microarray and (c) and (d) flow cytometry was 1.84 and 1.74, respectively.More complex operations, such as the programmed deposition of cell pairs, can be achieved by leveraging the switchable, bistable magnetization of the micromagnets for the detailed studies of cell-cell interactions (Figs. 3(a)–3(d)).¹² For these studies, a 200 G horizontal field generated from an electromagnetic coil was used to magnetize the micromagnets.¹⁹ We then captured different concentrations of magnetic beads as surrogates for cells (8.4 μm polystyrene, Spherotech, Inc.) and found that higher bead concentrations did not affect the capture accuracy (>95%; see supplementary material¹⁶).Open in a separate windowFIG. 3.Programmed pairing of magnetic beads and CD3+ lymphocytes. (a) Schematic of the magnetographic cell pair isolations. (b) Polarized micromagnets isolate cells of one type to one side in a vertical magnetic field and then cells of a second type to the other side when the field is reversed. (c) Fluorescent image of magnetically trapped green stained (top) and red stained (bottom) cell pairs. (d) SEM image of magnetically labeled cells in the microwells. (e) Capture accuracy of magnetic bead pairs. (Each color (and shape) represents the field strength of the reversed field.) (f) Change in the capture accuracy (loss) of initially captured beads after reversing the magnetic field. The capture accuracy of (g) magnetically labeled cell pairs and (h) the second magnetically labeled cell (for (e)–(h): n = 5; time starts from the deposition of the second set of cells or beads).The opposite side of each micromagnet was then populated with the second (yellow fluorescent) bead by reversing the direction of the applied magnetic field. We tested several field strengths (i.e., 10, 25, 40, or 55 G) to optimize the conditions for isolating the desired bead in the opposite well without ejecting the first bead. If the field strength was too large, the previously deposited beads could be ejected from their wells due to the repulsive magnetic force overcoming gravity.¹² As shown in Figure 3(e), increasing the field strength from 10 to 25 G significantly increased the capture accuracy at 60 min from the deposition of the second bead (p < 0.01), but increases from 25 to 55 G did not affect the capture accuracy (p > 0.10). As shown in Figure 3(f), higher field strengths (i.e., 40 and 55 G) resulted in lower capture accuracies compared to lower field strengths (i.e., 10 and 25 G) (p < 0.01), which was primarily due to ejection of the initially captured beads when the micromagnets reversed their polarity.We then arranged pairs of membrane dyed (calcein AM, Invitrogen; PKH26, Sigma) magnetically labeled CD3+ lymphocytes. First, red stained cells (150 μl of 2 × 10⁴ cells/ml) were deposited on the microchip in the presence of 250 G vertical magnetic field. After 20 min, the field was reversed (i.e., to 40, 55, and 70 G) and green stained cells (150 μl of 2 × 10⁴ cells/ml) were deposited on the microchip with images taken in 10 min intervals. Fluorescence images were overlaid (Fig. 3(c)) and the capture accuracy of cell pairs was determined (ImageJ).As seen in Figure 3(g), the capture accuracy of pairs of CD3+ lymphocytes was lower than that of magnetic beads (Fig. 3(e)). However, as shown in Figure 3(h), the second set of cells (green fluorescent) exhibited an average capture accuracy of 91.8% ± 1.9%. This indicates that the lower capture accuracy of cell pairs was either due to the ejection of initially captured (red fluorescent) cells or the migration of initially captured cells through the connecting channel, resulting from their relatively high deformability compared to magnetic beads.In summary, we developed a simple device capable of organizing magnetic particles, cells, and pairs of cells into well-defined compartments. A major advantage of this system is the use of specific magnetic labels to both isolate cells and program their deposition. While the design of this device does not enable dynamic control of the spacing between captured cell pairs as does some dielectrophoresis-based devices,²⁰ it can easily capture cells with high fidelity using only permanent magnets and has clinical relevance in the assessment of immune parameters. These demonstrations potentiate a relatively simple and robust device where highly organized spatial arrangement of cells facilitates rapid and accurate analyses towards a functional and low-cost point-of-care device.  相似文献   

Modulating patterns of two-phase flow with electric fields     

Dingsheng Liu  Bejan Hakimi  Michael Volny  Joelle Rolfs  Robbyn K. Anand  Frantisek Turecek  Daniel T. Chiu 《Biomicrofluidics》2014,8(4)

This paper describes the use of electro-hydrodynamic actuation to control the transition between three major flow patterns of an aqueous-oil Newtonian flow in a microchannel: droplets, beads-on-a-string (BOAS), and multi-stream laminar flow. We observed interesting transitional flow patterns between droplets and BOAS as the electric field was modulated. The ability to control flow patterns of a two-phase fluid in a microchannel adds to the microfluidic tool box and improves our understanding of this interesting fluid behavior.Microfluidic technologies have found use in a wide range of applications, from chemical synthesis to biological analysis to materials and energy technologies.^1,2 In recent years, there has been increasing interest in two-phase flow and droplet microfluidics, owing to their potential for providing a high-throughput platform for carrying out chemical and biological analysis and manipulations.^3–8 Although droplets may be generated in many different ways, such as with electric fields or extrusion through a small nozzle,^9–12 the most common microfluidic methods are based on the use of either T-junctions or flow-focusing geometries with which uniform droplets can be formed at high frequency in a steady-state fashion.^13,14 Various operations, such as cell encapsulation, droplet fusion, splitting, mixing, and sorting, have also been developed, and these systems have been demonstrated for a wide range of applications, including cell analysis, protein crystallization, and material synthesis.^1–17In addition to forming discrete droplets, where a disperse phase is completely surrounded by a continuous phase, it is also possible in certain situations to have different phases flow side-by-side. In fact, multi-stream laminar flow, either of the same phase or different phases, has been exploited for both biochemical analysis and microfabrication.^1,2,18–20 Beads-on-a-string (BOAS) is another potential flow pattern, which has been attracting attentions in microfluidics field. BOAS flow, owing to its special flow structures, may be particularly useful in some applications, such as optical-sensor fabrication.²¹ In BOAS flow, queues of droplets are connected by a series of liquid threads, which makes them look like a fluid necklace with regular periods.^21–25 The BOAS pattern is easily found in nature, such as silk beads and cellular protoplasm, and is often encountered in industrial processes as well, such as in electrospinning and anti-misting.^21,22 In general, it is thought that BOAS structure occurs mostly in viscoelastic fluids²² and is an unstable structure, which evolves continually and breaks eventually.^21–29Flow patterns determine the inter-relations of fluids in a microdevice and are an important parameter to control. Common methods for adjusting microfluidic flow patterns include varying the fluid flow rates, fluid properties, and channel geometries. Additionally, the application of an electric field can be a useful supplement for adjusting microfluidic flow patterns, although most work in this area has been focused on droplets and in some cases also on multi-stream laminar flows.^30–33 Here, in addition to forming droplets and two-phase laminar flow with electro-hydrodynamic actuation, we also observed a new stable flow pattern in a non-viscoelastic fluid, BOAS flow. Such flow patterns may find use in controlling the interactions between droplets, such as limited mixing by diffusion between neighboring droplets.To generate droplets, we used the flow-focusing geometry (Figure 1(a)), in which aqueous phase (water) was flown down the middle channel and droplets were pinched off by the oil phase (1-octanol) from the two side channels at the junction; Figure 1(b) shows the droplets formed after the junction. To apply electric field along the main channel where the droplets were formed, we patterned a pair of electrodes upstream and downstream of the junction (Figure 1(a); for experimental details, please see Ref. ³⁴ for supplementary material). The average electric field strength may be calculated from the voltages applied and the distance (1.7 mm) between the two electrodes. When a high voltage was applied along the channel between the two electrodes, the aqueous-oil interface at the flow-focusing junction became charged and behaved like a capacitor. As a result, more negative charges were drawn back upstream towards the positive electrode, and left behind more positive charges at the aqueous-oil interface, which then became encapsulated into the aqueous droplets dispersed in the oil phase.Open in a separate windowFIG. 1.(a) Schematic of the setup. (b) Micrograph showing droplet generation in a flow-focusing junction. The scale bar represents 40 μm.The positively charged aqueous-oil interface was stretched under an applied electric field, and by adjusting the voltage and/or the two-phase flow-rate ratio, we found interestingly that various flow patterns emerged. We tested different combinations of applied voltages and flow-rate ratios and found that most of them resulted in similar flow patterns and transitions between flow patterns.Figure ​Figure22 illustrates the effects of varying the applied voltages on droplets at a fixed liquid flow rate. With increasing electric-field strength and force, we found it was easier for the aqueous phase to overcome interfacial tension and form droplets. For example, as the voltage increased from 0.0 kV to 0.8 kV (average field strength increased from 0 to 0.47 V/μm), droplet-generation frequencies became slightly higher, and the formed droplets were smaller in volume. Additionally, droplets gradually became more spherical in shape at higher voltages.Open in a separate windowFIG. 2.Images showing the effects of applied voltage on droplet shape and flow pattern. Oil-phase flow rate, 0.5 μl/min; aqueous-phase flow rate, 0.2 μl/min. The scale bar represents 40 μm.As the voltage increased further (e.g., up to 1.0 kV in Figure ​Figure3),3), the distance between neighboring droplets became smaller, and the aqueous-oil interface at the junction was stretched further toward the downstream channel. At a threshold voltage (1 kV here with corresponding average field strength of 0.59 V/μm), the tip of the aqueous-oil interface would catch up with the droplet that just formed, and the tip of the interface of this newly captured droplet would in turn catch up with the interface of the droplet that formed before it. Consequently, a series of threads would connect all the droplets flowing between the two electrodes, thus resulting in a BOAS flow pattern.Open in a separate windowFIG. 3.Series of images showing the reversibility and synchronicity of a transitional flow pattern between droplets and BOAS (bead-on-a-string). Voltage applied, 1.00 kV (corresponding field strength of 0.59 V/μm); oil-phase flow rate, 0.5 μl/min; aqueous-phase flow rate, 0.2 μl/min. The scale bar represents 40 μm.At voltages near the threshold value, the flow pattern was not stable, but oscillated between droplets flow and BOAS flow. Figure ​Figure33 is a series of images captured by a high-speed camera that show the flow in this transition region. In Figures 3(a) and 3(b), the string of BOAS became thinner over time, and then the BOAS broke into droplets (Figures 3(c) and 3(d)). The newly formed droplets, however, were not stable either. Thin liquid threads would appear and then connect neighboring droplets, and a new switching period between discrete droplets and BOAS would repeat (Figures 3(e)–3(h)). In addition to this oscillation and reversibility, the flow pattern had a synchronous behavior: all the droplets appeared connected simultaneously by liquid threads or were separated at the same time.When the voltage reached 1.3 kV, which corresponded to an average field strength of 0.76 V/μm, a stable BOAS flow was obtained (Figure 4(a)). BOAS structures are thought to be present mostly in viscoelastic fluids,²² because viscoelasticity is helpful in enhancing the growth of beads and in delaying breakup of the string; thus, the viscoelastic filament has much longer life time than its Newtonian counterpart. Here, with the help of electric field, regular BOAS structures are realized in a non-viscoelastic fluid (water) in microchannels.Open in a separate windowFIG. 4.(a) Micrograph showing BOAS flow in a channel. (b) Profile of the top-half of the BOAS flow recorded continuously at a cross-section (shown in Figure 4(a)) of a channel. Voltage applied, 1.30 kV (corresponding field strength of 0.76 V/μm); oil-phase flow rate, 0.5 μl/min; aqueous-phase flow rate, 0.2 μl/min. The scale bar represents 40 μm.Microenvironment and electric fields alter the common evolution of BOAS structure observed in macroscopic or unbound environments. The BOAS structure formed in our experiments is not a stationary pattern, but a steady-state flowing one. Electric-field force prevents liquid strings from breaking between beads, and thus plays a similar role as elastic force in viscoelastic fluids. Figure 4(b) shows the dynamic BOAS profile, obtained at a fixed plane (shown in Figure 4(a)) perpendicularly across the channel as the BOAS structure passed through it. Droplets and liquid-thread diameters were nearly constant during the sampling time. The longer term experiments (over 3 min) showed there were slight variations of the two diameters in time, but the essential BOAS structure still remained qualitatively the same as a whole.When the voltage was further increased, the string diameter became larger and the droplet diameter became smaller. Because of the low flow-rate ratio (0.4) between the aqueous phase and oil phase used in the experiment depicted in Figure ​Figure4,4, the flow did not further develop into a multi-stream laminar flow, as would be expected at a higher voltage, and instead became unstable and irregular. When the flow-rate ratio was increased to 1.0 and the voltage was adjusted to 3.0 kV (corresponding field strength of 1.76 V/μm), we observed a stable multi-stream laminar flow (Figure ​(Figure5).5). The aqueous stream flowed in the channel center surrounded by the oil phase on the sides. This experiment showed that higher electric-field strengths alone would not give rise to another stable flow pattern (i.e., multi-stream laminar flow), but a suitable flow-rate ratio of aqueous phase to oil phase is required for the formation of stable two-phase laminar flow.Open in a separate windowFIG. 5.Micrograph showing multi-stream two-phase laminar flow in the channel. Voltage applied, 3.00 kV (corresponding field strength of 1.76 V/μm); oil-phase flow rate, 0.5 μl/min; aqueous-phase flow rate, 0.5 μl/min. The scale bar represents 40 μm.The flow patterns we observed may be described by a phase diagram (Figure ​(Figure6),6), which depends on two dimensionless numbers: capillary number, Ca = μ_aU_a/σ, and electric Bond number, Bo_e = E²(εD/σ). Ca and Bo_e describe the ratio of viscous force to interfacial tension force and the ratio of electric-field force to interfacial tension force, respectively. Here, μ_a (1 mPa s), σ (8.5 mN/m), ε (7.1 × 10⁻¹⁰ F/m), E, U_a, and D are, respectively, the aqueous-phase viscosity, aqueous-oil interfacial tension, aqueous-phase permittivity, electric field strength, aqueous-phase velocity, and the hydraulic diameter of the channel at the junction. Figure ​Figure66 shows clearly that at higher Ca, flow pattern changes gradually from droplet to BOAS and to multi-stream laminar flow with increasing Bo_e, which indicates the increasing importance of the electric-field force compared with the interfacial tension force. At lower Ca, flow pattern and transition show similar trend with increasing Bo_e as in the higher Ca case, except that multi-stream laminar flow is not observed. The relatively higher viscous force at higher Ca may be necessary for transitioning to the multi-stream laminar flow regime. In addition, Figure ​Figure66 shows that the BOAS window at the lower Ca is smaller than that at the higher Ca.Open in a separate windowFIG. 6.Phase diagram showing different flow patterns in the Ca and Bo_e space. Hollow symbols: oil-phase flow rate, 0.5 μl/min; aqueous-phase flow rate, 0.5 μl/min. Solid symbols: oil-phase flow rate, 0.5 μl/min; aqueous-phase flow rate, 0.2 μl/min.In summary, we showed the ability to use electric fields to generate and control different flow patterns in two-phase flow. With the aid of an applied field, we were able to generate BOAS flow patterns in a non-viscoelastic fluid, a pattern that typically requires a viscoelastic fluid. The BOAS structure was stable and remained as long as the applied electric field was on. We also report transitional flow patterns, those between droplets and BOAS exhibited both good reversibility as well as synchronicity. And with a suitable flow-rate ratio between the two phases, BOAS flow could be transitioned into a stable two-phase laminar flow by applying a sufficiently high field strength. Finally, a phase diagram was presented to describe quantitatively the flow-pattern regimes using capillary number and electric Bond number. The phenomena we report here on the properties of two-phase flow under an applied electric field may find use in developing a different approach to exert control over droplet based or multi-phase laminar-flow based operations and assays, and also aid in understanding the physics of multi-phase flow.  相似文献   

Polyphosphonium‐based bipolar membranes for rectification of ionic currents     

Erik O. Gabrielsson  Magnus Berggren 《Biomicrofluidics》2013,7(6)

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.  相似文献   

Microfluidic sterilization     

Rui Zhang  Jie Huang  Fei Xie  Baojun Wang  Ming Chu  Yuedan Wang  Haichao Li  Wei Wang  Haixia Zhang  Wengang Wu  Zhihong Li 《Biomicrofluidics》2014,8(3)

Nowadays, microfluidics is attracting more and more attentions in the biological society and has provided powerful solutions for various applications. This paper reported a microfluidic strategy for aqueous sample sterilization. A well-designed small microchannel with a high hydrodynamic resistance was used to function as an in-chip pressure regulator. The pressure in the upstream microchannel was thereby elevated which made it possible to maintain a boiling-free high temperature environment for aqueous sample sterilization. A 120 °C temperature along with a pressure of 400 kPa was successfully achieved inside the chip to sterilize aqueous samples with E. coli and Staphylococcus aureus inside. This technique will find wide applications in portable cell culturing, microsurgery in wild fields, and other related micro total analysis systems.Microfluidics, which confines fluid flow at microscale, attracts more and more attentions in the biological society.^1–4 By scaling the flow domain down to microliter level, microfluidics shows attractive merits of low sample consumption, precise biological objective manipulation, and fast momentum/energy transportation. For example, various cell operations, such as culturing^5–7 and sorting,^8–10 have already been demonstrated with microfluidic approaches. In most biological applications, sterilization is a key sample pre-treatment step to avoid contamination. However, as far as the author knew, this important pre-treatment operation is generally achieved in an off-chip way, by using high temperature and high pressure autoclave. Actually, microfluidics has already been utilized to develop new solution for high pressure/temperature reactions. The required high pressure/temperature condition was generated either by combining off-chip back pressure regulator and hot-oil bath,^11,12 or by integrating pressure regulator, heater, and temperature sensor into a single chip.¹³ This work presented a microfluidic sterilization strategy by implementing the previously developed continuous flowing high pressure/temperature microfluidic reactor.Figure ​Figure11 shows the working principle of the present microfluidic sterilization chip. The chip consists of three zones: sample loading (a microchannel with length of 270 mm and width of 40 μm), sterilization (length of 216 mm and width of 100 μm), and pressure regulating (length of 42 mm and width of 5 μm). Three functional zones were separated by two thermal isolation trenches. The sample was injected into the chip by a syringe pump and experienced two-step filtrations (feature sizes of 20 μm and 5 μm, not shown in Figure ​Figure1)1) at the entrance to avoid the channel clog. All channels had the same depth of 40 μm. According to the Hagen–Poiseuille relationship,¹⁵ the pressure regulating channel had a large flow resistance (around 1.09 × 10¹⁷ Pa·s/m³, see supplementary S1 for details¹⁶) because of its small width, thereby generated a high working pressure in the upstream sterilization channel under a given flow rate. The boiling point of the solution will then be raised up by the elevated pressure in the sterilization zone followed by the Antoine equation.¹⁶ By integrating heater/temperature sensors in the pressurized zone, a high temperature environment with temperature higher than 100 °C can thereby be realized for aqueous sample sterilization. The sample was collected from the outlet and cultured at 37 °C for 12 h. Bacterial colony was counted to evaluate the sterilization performance.Open in a separate windowFIG. 1.Working principle of the present microfluidic sterilization. Only microfluidic channel, heater, and temperature sensor were schematically shown. The varied colour of the microchannel represents the pressure and that of the halation stands for the temperature.Fabrication of this chip has been introduced elsewhere.¹⁴ The fabricated chip and the experimental system are shown in Figure ​Figure2.2. There were two inlets of the chip. While, in the experiment, only one inlet used and connected to the syringe pump. The backup one was blocked manually. The sample load zone was arranged in between of the sterilization zone and the pressure regulating zone based on thermal management consideration. A temperature control system (heater/temperature sensor, power source, and multi-meter) was setup to provide the required high temperature. The heater and the temperature sensor were microfabricated Pt resistors. The temperature coefficient of resistance (TCR) was measured as 0.00152 K⁻¹.Open in a separate windowFIG. 2.The fabricated chip and the experimental system. (a) Two chips with a penny for comparison. The left chip was viewed from the heater/temperature sensor side, while the right one was observed from the microchannel side (through a glass substrate). (b) The experimental system.Thermal isolation performance of the present chip before packaging with inlet/outlet was shown in Figure ​Figure3,3, to show the thermal interference issue. The results indicated that when the sterilization zone was heated up to 140 °C, the pressure regulating zone was about 40 °C. At this temperature, the viscosity of water decreases to 0.653 mPa·s from 1.00 mPa·s (at 20 °C), which will make the pressure in the sterilization zone reduced from 539 kPa (calculated at 20 °C and flow rate of 4 nl/s) to 387 kPa. The boiling point will then decrease to 142.8 °C, which will guarantee a boiling-free sterilization. In the cases without the thermal isolation trenches, the temperature of the pressure regulating zone reached as high as 75 °C because of the thermal interference from the sterilization zone, as shown in Figure ​Figure3.3. The pressure in the sterilization zone was then reduced to 268 kPa (calculated at flow rate of 4 nl/s) and the boiling temperature was around 130 °C, which was lower than the set sterilization temperature. Detail calculation can be found in supplementary S2.¹⁶Open in a separate windowFIG. 3.The temperature distribution of the chips (before packaged) with and without thermal isolation trenches (powered at 1 W). The data were extracted from the central lines of infrared images, as shown as inserts.Bacterial sterilization performance of the present chip was tested and the experimental results were shown in Figure ​Figure4.4. E. coli with initial concentration of 10⁶/ml was pumped into and flew through the chip with the sterilization temperatures varied from 25 °C to 120 °C at flow rates of 2 nl/s and 4 nl/s. The outflow was collected and inoculated onto the SS agar plate evenly with inoculation loops. The population of bacteria in the outflow was counted based on the bacterial colonies after incubation at 37 °C for 12 h. Typical bacterial colonies were shown in Figure ​Figure4.4. The low flow rate case showed a better sterilization performance because of the longer staying period in the sterilization channel. The population of E. coli was around 1.25 × 10⁴/ml after a 432 s-long, 70 °C sterilization (at flow rate of 2 nl/s). While at the flow rate of 4 nl/s, the cultivation result indicated the population was around 3.8 × 10⁴/ml because the sterilization time was shorten to 216 s. A control case, where the solution flew through an un-heated chip at 2 nl/s, was conducted to investigate the effect of the shear stress on the sterilization performance (see the supplementary S3 for details¹⁶). As listed in Table ​TableI,I, the results indicated that the shear stress did not show any noticeable effect on the bacterial sterilization. When the chip was not heated, i.e., the case with the largest shear stress because of the highest viscosity of fluid, the bacterial cultivation was nearly the same as the off-chip results (no stress). The temperature has the most significant effect on the sterilization performance. No noticeable bacteria proliferation was observed in the cases with the sterilization temperature higher than 100 °C, as shown in Figure ​Figure44.

Table I.

The E. coli cultivation results under different flow rates and sterilization temperatures. ^a

	25 °C	70 °C	100 °C	120 °C	25 °C b
2 nl/s	1.89/+++	1.38/+	1.16/−	1.04/−	0/+++
4 nl/s	3.78/+++	2.76/+	2.32/−	2.08/−	0/+++

Open in a separate window^aData in the table are shear stress (Pa)/population of bacteria, where “+++” indicates a large proliferation, “+” means small but noticeable proliferation, “−” represents no proliferation.^bOff-chip control group.Open in a separate windowFIG. 4.Sterilization performance of the present chip with E. coli and S. aureus as test bacteria. All the original population was 10⁶/ml. Inserted images showed the images of the culture disk after bacteria incubation.Sterilization of another commonly encountered bacterium, Staphylococcus aureus, with initial population of 10⁶/ml was also tested in the present chip, as shown in Figure ​Figure4.4. Similarly, no noticeable S. aureus proliferation was found when the sterilization temperature was higher than 100 °C.In short, we demonstrated a microfluidic sterilization strategy by utilizing a continuous flowing high temperature/pressure chip. The population of E. coli or S. aureus was reduced from 10⁶/ml to an undetectable level when the sterilization temperature of the chip was higher than 100 °C. The chip holds promising potential in developing portable microsystem for biological/clinical applications.  相似文献   

Black silicon as a platform for bacterial detection     

Jennifer S. Hartley  M. Myintzu Hlaing  Gediminas Seniutinas  Saulius Juodkazis  Paul R. Stoddart 《Biomicrofluidics》2015,9(6)

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.  相似文献   

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

Shoupeng Liu  Yu Yan  Yunshan Wang  Satyajyoti Senapati  Hsueh-Chia Chang 《Biomicrofluidics》2013,7(6)

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.  相似文献   

An on-chip study on the influence of geometrical confinement and chemical gradient on cell polarity     

Wenfu Zheng  Yunyan Xie  Kang Sun  Dong Wang  Yi Zhang  Chen Wang  Yong Chen  Xingyu Jiang 《Biomicrofluidics》2014,8(5)

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You cannot tell a book by looking at the cover: Cryptic complexity in bacterial evolution     

Qiucen Zhang  Julia Bos  Grigory Tarnopolskiy  James C. Sturm  Hyunsung Kim  Nader Pourmand  Robert H. Austin 《Biomicrofluidics》2014,8(5)

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

Jason Riordon  Michael Nash  Wenyang Jing  Michel Godin 《Biomicrofluidics》2014,8(1)

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.  相似文献   

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

Anil Haraksingh Thilsted  Vahid Bazargan  Nina Piggott  Vivien Measday  Boris Stoeber 《Biomicrofluidics》2012,6(4)

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.  相似文献   

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

Adam R. Abate  David A. Weitz 《Biomicrofluidics》2011,5(1)

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}  相似文献   

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

Yunshan Wang  Ting-Chou Chang  Paul R. Stoddart  Hsueh-Chia Chang 《Biomicrofluidics》2014,8(2)

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.  相似文献   

An integrated microfluidic device for rapid serodiagnosis of amebiasis     

Wang Zhao  Li Zhang  Wenwen Jing  Sixiu Liu  Hiroshi Tachibana  Xunjia Cheng  Guodong Sui 《Biomicrofluidics》2013,7(1)

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