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1.
Fuchs (2010, 2012) argues that users of social media produce value and surplus value in the Marxian sense. Arvidsson and Colleoni (2012) critique this hypothesis, claiming that Marx's theory of value is irrelevant to the regime of value production on social media platforms in particular and in informational capitalism in general. They claim that the affective relations and financial speculations that generate value on social media are not dependent on labor time. This article critically engages Fuchs, and Arvidsson and Colleoni, by revisiting Marx's theory of value. Contra Fuchs, we argue that audiences do not produce value and surplus value—neither for social nor for mass media. Contra Arvidsson and Colleoni, we argue that so-called affective relations (philia) do not produce value either. Instead we demonstrate that social media generate revenue from four primary sources—by leasing advertisement space to generate advertisement rent, by selling information, by selling services to advertisers, and by generating profits from fictitious capital and speculative windfalls. All four, we argue, can be adequately explained by Marx's theory of value. 相似文献
2.
Pille Pruulmann-Vengerfeldt 《The Information Society》2013,29(5):303-310
Using Layder's domain theory (1997) as an analytical framework, this article shows how the information society can be measured through various levels of society. Layder's notions of psychobiography, situated activity, social setting, and contextual resources help identify cultural and social indicators for understanding changes in the information society. With the help of empirical indicators for each domain, this article uses the case of Estonia to show that there is often more to the information society than what is captured by traditional measures. This article calls for a context-sensitive approach, which takes into consideration social and cultural indicators. Measurements from all four domains are necessary for understanding the complexity of information-society-related issues. 相似文献
3.
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 resistors14, 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 Pfin=Vin∕Vfin×Pin, where Vin is the initial volume of air in the syringe, Vfin is the volume after expansion, and Pin 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
4.
Polymer-based microneedles have drawn much attention in transdermal drug delivery resulting from their flexibility and biocompatibility. Traditional fabrication approaches are usually time-consuming and expensive. In this study, we developed a new double drawing lithography technology to make biocompatible SU-8 microneedles for transdermal drug delivery applications. These microneedles are strong enough to stand force from both vertical direction and planar direction during penetration. They can be used to penetrate into the skin easily and deliver drugs to the tissues under it. By controlling the delivery speed lower than 2 μl/min per single microneedle, the delivery rate can be as high as 71%.Microelectromechanical systems (MEMS) technology has enabled wide range of biomedical devices applications, such as micropatterning of substrates and cells,1 microfluidics,2 molecular biology on chips,3 cells on chips,4 tissue microengineering,5 and implantable microdevices.6 Transdermal drug delivery using MEMS based devices can delivery insoluble, unstable, or unavailable therapeutic compounds to reduce the amount of those compounds used and to localize the delivery of potent compounds.7 Microneedles for transdermal drug delivery are increasingly becoming popular due to their minimally invasive procedure,8 promising chance for self-administration,9 and low injury risks.10 Moreover, since pharmaceutical and therapeutic agents can be easily transported into the body through the skin by microneedles,11, 12 the microneedles are promising to replace traditional hypodermic needles in the future. Previously, various microneedles devices for transdermal drug delivery applications have been reported. They have been successfully fabricated by different materials, including silicon,13 stainless steel,14 titanium,15 tantalum,16 and nickel.17 Although microneedles with these kinds of materials can be easily fabricated into sharp shape and offer the required mechanical strength for penetration purpose, such microneedles are prone to be damaged18 and may not be biocompatible.19 As a result, polymer based microneedles, such as SU-8,20, 21 polymethyl meth-acrylate (PMMA),22, 23 polycarbonates (PCs),24, 25 maltose,26, 27 and polylactic acid (PLA),28, 29 have caught more and more attentions in the past few years. However, in order to obtain ultra-sharp tips for penetrating the barrier layer of stratum corneum,30 conventional fabrication technologies, for instances, PDMS (Polydimethylsiloxane) molding technology,31, 32 stainless steel molding technology,33 reactive ion etching technology,34 inclined UV (Ultraviolet) exposure technology,35 and backside exposure with integrated lens technology36 are time-consuming and expensive. In this paper, we report an innovative double drawing lithography technology for scalable, reproducible, and inexpensive microneedle devices. Drawing lithography technology37 was first developed by Lee et al. They leveraged the polymers'' different viscosities under different temperatures to pattern 3D structures. However, it required that the drawing frames need to be regular cylinders, which is not proper for our devices. To solve the problem, the new double drawing lithography is developed to create sharp SU-8 tips on the top of four SU-8 pillars for penetration purpose. Drugs can flow through the sidewall gaps between the pillars and enter into the tissues under the skin surface. The experiment results indicate that the new device can have larger than 1N planar buckling force and be easily penetrated into skin for drugs delivery purpose. By delivering glucose solution inside the hydrogel, the delivering rate of the microneedles can be as high as 71% when the single microneedle delivery speed is lower than 2 μl/min.An array of 3 × 3 SU-8 supporting structures was patterned on a 140 μm thick, 6 mm × 6 mm SU-8 membrane (Fig. (Fig.1a).1a). Each SU-8 supporting structure included four SU-8 pillars and was 350 μm high. The four pillars were patterned into a tubelike shape on the membrane (Fig. (Fig.1b).1b). The inner diameter of the tube was 150 μm, while the outer diameter was 300 μm. SU-8 needles of 700 μm height were created on the top of SU-8 supporting structures to ensure the ability of transdermal penetration. Two PDMS layers were bonded with SU-8 membrane to form a sealed chamber for storing drugs from the connection tube. Once the microneedles entered into the tissue, drugs could be delivered into the body through the sidewall gaps between the pillars (Fig. (Fig.1c1c).Open in a separate windowFigure 1Schematic illustration of the SU-8 microneedles. (a) Overview of the whole device; (b) SU-8 supporting structures made of 4 SU-8 pillars; and (c) enlarged view of a single SU-8 microneedle.The fabrication process of SU-8 microneedles is shown in Fig. Fig.2.2. SU-8 microneedles fabrication started from a layer of Polyethylene Terephthalate (PET, 3M, USA) film pasted on the Si substrate by sticking the edge area with kapton tape (Fig. (Fig.2a).2a). The PET film, a kind of transparent film with poor adhesion to SU-8, was used as a sacrificial layer to dry release the final device from Si substrate. A 140 μm thick SU-8 layer was deposited on the top of this PET film. To ensure a uniform surface of this thick SU-8 layer, the SU-8 deposition was conducted in two steps coating. After exposed under 450 mJ/cm2 UV, the membrane pattern could be defined (Fig. (Fig.2b).2b). In order to ensure an even surface for following spinning process, another 350 μm SU-8 layer was directly deposited on this layer in two steps without development. With careful alignment, an exposure of 650 mJ/cm2 UV energy was performed on this 350 μm SU-8 layer to define the SU-8 supporting structures (Fig. (Fig.2c).2c). The SU-8 structure could be easily released from the PET substrate by removing the kapton tape and slightly bending the PET film. Two PDMS layers were bonded with this SU-8 structure by a method reported by Zhang et al.38 (Fig. (Fig.2d2d).Open in a separate windowFigure 2Fabrication process for SU-8 microtubes. (a) Attaching a PET film on the Si substrate; (b) exposing the first layer of SU-8 membrane without development; (c) depositing and patterning two continuous SU-8 layers as sidewall pillars; (d) releasing the SU-8 structure from the substrate and bonding it with PDMS; (e) drawing hollowed microneedles on the top of supporting structures; (f) baking and melting the hollowed microneedles to allow the SU-8 flow in the gaps between pillars; and (g) drawing second time on the top of the melted SU-8 flat surface to get microneedles.In our previous work,39 we used one time stepwise controlled drawing lithography technology for the sharp tips integration. However, since the frame used to conduct drawing process in present study is a four-pillars structure rather than a microtube, the conventional drawing process can only make a hollowed tip but not a solid tip structure (Fig. (Fig.3).3). This kind of tip was fragile and could not penetrate skin in the practical testing process. To solve the problem, we developed an innovative double drawing lithography process. After bonding released SU-8 structure with PDMS layers (Fig. (Fig.2d),2d), we used it to conduct first time stepwise controlled drawing lithography37 and got hollowed tips (Fig. (Fig.2e).2e). Briefly, the SU-8 was spun on the Si substrate and kept at 95 °C until the water inside completely vaporized. Device of SU-8 supporting structures was fixed on a precision stage. Then, the SU-8 supporting structures were immersed into the SU-8 by adjusting the precision state. The SU-8 were coated on the pillars'' surface. Then, the SU-8 supporting structures were drawn away from the interface of the liquid maltose and air. After that, the temperature and drawing speed were increased. Since the SU-8 was less viscous at higher temperature, the connection between the SU-8 supporting structures and surface of the liquid SU-8 became individual SU-8 bridge, shrank, and then broke. The end of the shrunk SU-8 bridge forms a sharp tip on the top of each SU-8 supporting structure when the connection was separated. After the hollowed tips were formed in the first step drawing process, the whole device was baked on the hotplate to melt the hollowed SU-8 tips. Melted SU-8 reflowed into the gaps between four pillars and the tips became domes (Fig. (Fig.2f).2f). Then, a second drawing process was conducted on the top of melted SU-8 to form sharp and solid tips (Fig. (Fig.2g).2g). The final fabricated device is shown in Fig. Fig.44.Open in a separate windowFigure 3A hollowed SU-8 microneedle fabricated by single drawing lithography technology (scale bar is 100 μm).Open in a separate windowFigure 4Optical images for the finished SU-8 microneedles.During the double drawing process, as long as the heated time and temperature were controlled, the SU-8 flow-in speed of SU-8 inside the gaps could be precisely determined. The relationship between baking temperature and flow-in speed was studied. As shown in Fig. Fig.5,5, the flow-in speed is positive related to the baking temperature. The explanation for this phenomena is that the SU-8''s viscosity is different under different baking temperatures.40 Generally, baked SU-8 has 3 status when temperature increases, solid, glass, and liquid. The corresponding viscosity will decrease and the SU-8 can also have higher fluidity. When the baking temperature is larger than 120 °C, the flow-in speed will increase sharply. But, if the baking temperature is higher, the SU-8 will reflow in the gaps too fast, which makes the flow-in depth hard to be controlled. There is a high chance that the whole gaps will be blocked, and no drugs can flow through these gaps any more. Considering that the total SU-8 supporting structure is only 350 μm high, we choose 125 °C as baking temperature for proper SU-8 flow-in speed and easier SU-8 flow-in depth control.Open in a separate windowFigure 5The relationship between flow-in speed and baking temperature.To ensure the adequate stiffness of the SU-8 microneedles in vertical direction, Instron Microtester 5848 (Instron, USA) was deployed to press the microneedles with the similar method reported by Khoo et al.41 As shown in Fig. Fig.6a,6a, the vertical buckling force was as much as 8.1N, which was much larger than the reported minimal required penetration force.42 However, in the previous practical testing experiments, even though the microneedles were strong enough in vertical direction, the planar shear force induced by skin deformation might also break the interface between SU-8 pillars and top tips. In our new device with four pillars supporting structure, the SU-8 could flow inside the sidewall gaps between the pillars to form anchors. These anchors could enhance microneedles'' mechanical strength and overcome the planar shear force problems. Moreover, the anchors strength could be improved by controlling the SU-8 flow-in depth. Fig. Fig.77 shows that the flow-in depth increases when the baking time increases as the baking time increases at 125 °C. Fig. Fig.6b6b shows that the corresponding planar buckling force can be improved to be larger than 1 N by increasing flow-in depth. Some sidewall gaps at bottom are kept on purpose for drugs delivery; hence, the flow-in depth is chosen as 200 μm.Open in a separate windowFigure 6(a) Measurement of the vertical buckling force. (b) The planar buckling force varies under different flow-in depth (I, II, III, and IV corresponding to the certain images in Fig. Fig.77).Open in a separate windowFigure 7Different flow-in depth inside the gaps between SU-8 pillars. (a) 0 μm; (b) 100 μm; (c) 200 μm; and (d) 350 μm (scale bar is 100 μm).The penetration capability of the 3 × 3 SU-8 microneedles array is characterized by conducting the insertion experiment on the porcine cadaver skin. 10 microneedles devices were tested and all of them were strong enough to be inserted into the tissue without any breakage. Histology images of the skin at the site of one microneedle penetration were derived to prove that the sharp conical tip was not broken during the insertion process (Fig. (Fig.8).8). It also shows penetrated evidence because the hole shape is the same as the sharp conical tip.Open in a separate windowFigure 8Histology image of individual microneedle penetration (scale bar is 100 μm).In order to verify that the drug solution can be delivered into tissue from the sidewall gaps of the microneedles, FITC (Fluorescein isothiocyanate) (Sigma Aldrich, Singapore) solution was delivered through the SU-8 microneedles after they were penetrated into the mouse cadaver skin. The representative results were then investigated via a confocal microscope (Fig. (Fig.9).9). The permeation pattern of the solution along the microchannel created by microneedles confirmed the solution delivery results. The black area was a control area without any diffused florescent solution. In contrast, the illuminated area in Fig. Fig.99 indicates the area where the solution has diffused to it. These images were taken consecutively from the skin surface down to 180 μm with 30 μm intervals. The diffusion area had a similar dimension with the inserted microneedles. It has proved that the device can be used to deliver drugs into the body.Open in a separate windowFigure 9Images of confocal microscopy to show the florescent solution is successfully delivered into the tissue underneath the skin surface. (a) 30 μm; (b) 60 μm; (c) 90 μm; (d) 120 μm; (e) 150 μm; and (f) 180 μm (scale bar is 100 μm).Due to the uneven surface of deformed skin, there is always tiny gap happened between tips of some microneedles and local surface skin. The microneedles could not be entirely inserted into the tissue. Drugs might leak to the skin surface through the sidewall gaps under certain driven pressure. Hydrogel absorption experiment was conducted to quantify the delivery rate (i.e., the ratio of solution delivered into tissues in the total delivered volume) and to optimize the delivery speed. Using hydrogel as the tissue model for quantitative analysis of microneedle releasing process was reported by Tsioris et al.43 The details are shown here. Gelatin hydrogel was prepared by boiling 70 ml DI (Deionized) water and mixing it with 7 g of KnoxTM original unflavored gelatin powder. The solution was poured into petri dish to 1 cm high. Then, the petri dish was put into a fridge for half an hour. Gelatin solution became collagen slabs. The collagen slabs were cut into 6 mm × 6 mm sections. A piece of fully stretched parafilm (Parafilm M, USA) was tightly mounted on the surface of the collagen slabs. This parafilm was used here to block the leaked solution further diffusing into the collagen slab in the delivery process. Then, the microneedles penetrated the parafilm and went into the collagen slab. Controlled by a syringe pump, 0.1 ml–0.5 mg/ml glucose solution was delivered into the collagen slab under different speeds. Methylene Blue (Sigma Aldrich, Singapore) was mixed into the solution for better inspection purpose (Fig. 10a). Then, the collagen slabs was digested in 1 mg/ml collagenase (Sigma Aldrich, Singapore) at room temperature (Fig. 10b). It took around 1 h that all the collagen slabs could be fully digested (Fig. 10d). The solution was collected to measure the glucose concentration with glucose detection kit (Abcam, Singapore). Briefly, both diluted glucose standard solution and the collected glucose solution were added into a series of wells in a well plate. Glucose assay buffer, glucose enzyme, and glucose substrate were mixed with these samples in the wells. After incubation for 30 min, their absorbance were examined by using a microplate reader at a wavelength of 450 nm. By comparing the readings with the measured concentration standard curve (Fig. 11a), the glucose concentration in the hydrogel, the glucose absorption rate in the hydrogel, and the solution delivery rate by microneedles could be measured and calculated. As shown in Fig. 11b, when the delivering speed of a single microneedle increased from 0.1 μl/min to 2 μl/min, the glucose absorption rate also increased. Most of the glucose solution from microneedles could go into the hydrogel. The delivered rate could be as high as 71%. The rest solution leaked from sidewall gaps and blocked by parafilm. However, when the delivered speed for a single microneedle was larger than 2 μl/min, the hydrogel absorption rate was saturated. More and more solution could not go into the hydrogel but leak from the sidewall gaps. Then, the delivered rate decreased. Therefore, 2 μl/min was chosen as the optimized delivery speed for the microneedle.Open in a separate windowFigure 10Glucose solution could be delivered into the hydrogel, and the collagen stabs were dissolved by collagenase.Open in a separate windowFigure 11(a) Standard curve for glucose detection; (b) glucose absorption rate and solution delivery rate in a single needle corresponding to different delivery speed.In conclusion, a drug delivery device of integrated vertical SU-8 microneedles array is fabricated based on a new double drawing lithography technology in this study. Compared with the previous biocompatible polymer-based microneedles fabrication technology, the proposed fabrication process is scalable, reproducible, and inexpensive. The fabricated microneedles are rather strong along both vertical and planar directions. It is proved that the microneedles were penetrated into the pig skin easily. The feasibility of drug delivery using SU-8 microneedles is confirmed by FITC fluorescent delivery experiment. In the hydrogel absorption experiment, by controlling the delivery speed under 2 μl/min per microneedle, the delivery rate provided the microneedle is as high as 71%. In the next step, the microneedles will be further integrated with microfluidics on a flexible substrate, forming a skin-patch like drug delivery device, which may potentially demonstrate a self-administration function. When patients need an injection treatment at home, they can easily use such a device just like using an adhesive bandage strip. 相似文献
5.
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 = μaUa/σ, and electric Bond number, Boe = E2(εD/σ). Ca and Boe describe the ratio of viscous force to interfacial tension force and the ratio of electric-field force to interfacial tension force, respectively. Here, μa (1 mPa s), σ (8.5 mN/m), ε (7.1 × 10−10 F/m), E, Ua, 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 Boe, 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 Boe 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 Boe 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
6.
《普罗米修斯》2012,30(3):349-351
John Vincent coordinates The Network, formed in May 1999 as a legacy of a project funded by the Library and Information Commission, Public Library Policy and Social Exclusion (see Muddiman, 2000). The Network’s mission is ‘to assist the cultural sector, including libraries, museums, archives and galleries, heritage and other organisations, to work towards social justice’. 相似文献
7.
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
8.
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), adenine10,11 (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
9.
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 culturing5–7
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.
Open in a separate windowaData 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
Table I.
The E. coli cultivation results under different flow rates and sterilization temperatures. a25 °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/+++ |
10.
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 complex1,7,8 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/Plac-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
11.
This research reports an improved conjugation process for immobilization of antibodies on carboxyl ended self-assembled monolayers (SAMs). The kinetics of antibody/SAM binding in microfluidic heterogeneous immunoassays has been studied through numerical simulation and experiments. Through numerical simulations, the mass transport of reacting species, namely, antibodies and crosslinking reagent, is related to the available surface concentration of carboxyl ended SAMs in a microchannel. In the bulk flow, the mass transport equation (diffusion and convection) is coupled to the surface reaction between the antibodies and SAM. The model developed is employed to study the effect of the flow rate, conjugating reagents concentration, and height of the microchannel. Dimensionless groups, such as the Damköhler number, are used to compare the reaction and fluidic phenomena present and justify the kinetic trends observed. Based on the model predictions, the conventional conjugation protocol is modified to increase the yield of conjugation reaction. A quartz crystal microbalance device is implemented to examine the resulting surface density of antibodies. As a result, an increase in surface density from 321 ng/cm2, in the conventional protocol, to 617 ng/cm2 in the modified protocol is observed, which is quite promising for (bio-) sensing applications.Microfluidics have been implemented in various bio-medical diagnostic applications, such as immunosensors and molecular diagnostic devices.1 In the last decade, a vast number of biochemical species has been detected by microfluidic-based immunosensors. Immunosensors are sensitive transducers which translate the antibody-antigen reaction to physical signals. The detection in an immunosensor is performed through immobilization of an antibody that is specific to the analyte of interest.2 The antibody is often bound to the transducing surface of the sensor covered by self-assembled monolayers (SAMs). SAMs are organic materials that form a thin, packed and robust interface on the surface of noble metals like that of gold, suitable for biosensing applications.3 Thiolic SAMs have a “head” group that shows a high affinity to being chemisorbed onto a substrate, typically gold. The SAMs'' carboxylic functional group of the “tail” end can be linked to an amine terminal of an antibody to form a SAM/antibody conjugation.3,4 The conjugation process is usually accomplished in the presence of carbodiimides, such as 1-ethyl-3-(3-dimethylaminopropyl) carbodiimide (EDC). A yield increasing additive, N-Hydroxysuccinimide (NHS), is often used to enhance the surface loading density of the antibody.4,5A typical reaction for coupling the carboxylic acid groups of SAMs with the amine residue of antibodies in the presence of EDC/NHS is depicted in Figure Figure11.4 NHS promotes the generation of an active NHS ester (k2 reaction path). The NHS ester is capable of efficient acylation of amines, including antibodies (k3 reaction path). As a result, the amide bond formation reaction, which typically does not progress efficiently, can be enhanced using NHS as a catalyst.4Open in a separate windowFIG. 1.NHS catalyzed conjugation of antibodies to carboxylic-acid ended SAMs through EDC mediation (Adapted from G. T. Hermanson, Bioconjugate Techniques, 2nd. Edition. Copyright 2008 by Elsevier4). EDC reacts with the carboxylic acid and forms o-acylisourea, a highly reactive chemical that reacts with NHS and forms an NHS ester, which quickly reacts with an amine (i.e., antibody) to form an amide.A number of groups have studied EDC/NHS mediated conjugation reactions such as the ones depicted in Figure Figure1.1. The general stoichiometry of the reaction involves a carboxylic acid (SAM), an amine (antibody), and EDC to produce the final amide (antibody conjugated SAM) and urea. However, the recommended concentration ratio of the crosslinking reagents inside the buffer, i.e., the ratio of EDC and NHS with respect to adsorbates and each other, varies from one study to another.6 The kinetics of the reactions outlined in Figure Figure11 have also been investigated,4,6–8 but only in the absence of NHS for EDC or carboxylic acids in aqueous solutions.8 A relatively recent experimental study verified the catalytic role of the yield-increasing reagent N-hydroxybenzotriazole (HOBt), which acts similarly to NHS.7 In this study, the amide formation rate (k3 reaction path, Figure Figure1)1) was found to be dependent on the concentration of the carboxylic acid and EDC in the buffer solution, and independent of the amine and catalyst reagent concentration. The same group also showed that the amide bond formation kinetics is controlled by the reaction between the carboxylic acid and the EDC to give the O-acylisourea, which they marked as the rate-determining step (k1 reaction path, Figure Figure11).The k1 reaction path, or the conjugation reaction, is usually a lengthy process and takes between 1 and 3 h.4,9 Compared to k1, the k2 and ?k3 reactions are considerably faster. Microfluidics has the potential to enhance the kinetics of these reactions using the flow-through mode.10,11 This improvement occurs because while conventional methods rely only on diffusion as the primary reagent transport mode, microfluidics adds convection to better replenish the reagents to the reaction surfaces. However, there are many fundamental fluidic and geometrical parameters that might affect the process time and reagents consumption in a microfluidics environment, such as concentration of antibodies and reagents, flow rate, channel height, and final surface density of antibodies. A model that studies the kinetics of conjugation reaction against all these parameters would therefore be helpful for the optimization of this enhanced kinetics.There are a number of reports on numerical examination of the kinetics of binding reactions in microfluidic immunoassays.12–15 All these models developed so far couple the transport of reagents, by diffusion and convection, to the binding on the reaction surface. Myszka''s model assumes a spatially homogeneous constant concentration of reagents throughout the reaction chamber, thus fails to describe highly transport-limited conditions due to the presence of spatial heterogeneity and depletion of the bulk fluid from reagents.16,17 In transport-limited conditions, the strength of reaction is superior to the rate of transport of reagents to the reaction surface.18,19 More recently, the convection effects were included in a number of studies, describing the whole kinetic spectrum from reaction-limited conditions to transport-limited reactions.20–22 Immunoreaction kinetics has also been examined with a variety of fluid driving forces, from capillary-driven flows,20 to electrokinetic flows in micro-reaction patches,21 pressure-driven flows in a variety of geometric designs.22 Despite these comprehensive numerical investigations, the fundamental interrelations between the constitutive kinetic parameters, such as concentration, flow velocity, microchannel height, and antibody loading density, have not been studied in detail. In addition, the conjugation kinetics has not yet been exclusively examined.In this paper, a previous model for immunoreaction is modified to study the antibody/SAM conjugation reaction in a microfluidic system. Model findings are used to examine the process times recommended in the literature and possible modification scenarios are proposed. The new model connects the convective and diffusive transport of reagents in the bulk fluid to their surface reaction. The conjugation reaction is studied against fluidic and geometrical parameters such as flow rate, concentration, microchannel height and surface density of antibodies. Damköhler number is used to compare the reaction and fluidic phenomena present and justify the kinetic trends observed. Model predictions are discussed and the main finding on possible overexposure of carboxylates to crosslinking reagents, in conventional protocols, is verified by comparing the resultant antibody loading densities obtained using a quartz crystal microbalance (QCM) set up. The results demonstrate an improved receptor (antibody) loading density which is quite promising for a number of (bio-) sensing applications.23,24 Major application areas include antibody-based sensors for on-site, rapid, and sensitive analysis of pathogens such as Bacillus anthracis,23
Escherichia coli, and Listeria monocytogenes, and toxins such as fungal pathogens, viruses, mycotoxins, marine toxins, and parasites.24 相似文献
12.
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, 20Herein, 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 Sample Average scores Standard deviation 1 33 790 368 2 23 269 271 3 39 598 307 4 7784 52 5 21 222 197 6 38 878 290 7 22 437 227 8 36 295 334 9 41 024 396 Negative 200 32