Preface to Special Topic: Selected Papers from the 5th International Conference on Optofluidics     

Shih-Kang Fan  Zhenchuan Yang 《Biomicrofluidics》2016,10(1)

The 5th International Conference on Optofluidics (Optofluidics 2015) was held in Taipei, Taiwan, July 26–29, 2015. The aim of this conference was to provide a forum to promote scientific exchange and to foster closer networks and collaborative ties between leading international researchers in optics and micro/nanofluidics across various disciplines. The scope of Optofluidics 2015 was deliberately broad and interdisciplinary, encompassing the latest advances and the most innovative developments in micro/nanoscale science and technology. Topics ranged from fundamental research to its applications in chemistry, physics, biology, materials, and medicine.Approximately 300 delegates participated in Optofluidics 2015 from across the globe, including Australia, Canada, China, France, Germany, Hong Kong, India, Japan, Korea, Singapore, Taiwan, UK, and USA. In total, 242 presentations were arranged, including 10 plenary speeches, 27 keynote speeches, 65 invited talks, 33 contributed talks, and 107 poster presentations. This collection of twelve papers on this special topic spans both the fundamentals and the frontier applications of this interdisciplinary research field.Optical measurements of particle or flow and fluidic manipulation for optical applications were presented. Lin and Su¹ reported a novel method to measure the depth position of rapidly moving objects inside a microfluidic channel based on the chromatic aberration effect; the depth positions of label-free particles of diameter as small as 2 μm and erythrocytes of concentration 2 × 10³ cells/μl and velocity 2.78 mm/s were detected within a range ±25 μm in a simple and inexpensive manner. Sun and Huang² demonstrated the use of a microscopic circular polariscope to measure the flow-induced birefringence in a microfluidic device that represents the kinematics of fluid motion optically; CTAB:NaSal, CPyCl:NaSal, and CPyCl:NaSal:NaCl solutions were used to investigate the strain rate and the results were compared with the μPIV diagnosis. He et al.³ studied the fundamentals, especially the thinning and opening of the oil film within each pixel of an electrowetting display; to achieve repeatable oil movement and the resulting pixel performance, a new method to fill each pixel with a controllable oil volume using an oil-droplet emulsion created with a microfluidic device was demonstrated.This special topic includes papers also on particle manipulation. Weng et al.⁴ evaluated the size-dependent crossing frequency of dielectrophoretically driven particles; numerical simulation using a Maxwell stress tensor and a finite element method was reported to assess the size effect. In addition to electric manipulation, magnetic driving of the particles was demonstrated. Ido et al.⁵ examined microswimmers of magnetic particle chains in an oscillating magnetic field experimentally and analyzed numerically with a lattice Boltzmann method, an immersed boundary method, and a discrete particle method based on simplified Stokesian dynamics. Huang et al.⁶ described a technique to manipulate magnetic beads and achieved a great washing efficiency with zero bead loss using an appropriate electrode design and channel height of a digital microfluidic immunoassay; a model immunoassay of human soluble tumor necrosis factor receptor I (sTNF-RI) was performed to offer an improved limit of detection (3.14 pg/ml) with a small number of magnetic beads (25 beads), decreased reagent volumes (200 nl), and decreased duration of analysis (<1 h). Chiu et al.⁷ reported particle separation using cross-flow filtration enhanced with hydrodynamic focusing; label-free separation of particles of diameters 2.7 and 10.6 μm at a sample throughput 10 μl/min was performed; separation of spiked human prostate cancer cell lines (PC3) cells in whole blood was also demonstrated.Chemical sensors and biosensors are covered in this special topic. Cheng et al.⁸ measured the chemical compounds in third-hand smoke on varied clothing fibres with an analytical balance, or nicotine and 3-ethenylpyridine (3-EP) with a surface-acoustic-wave sensor composed of coated oxidized hollow mesoporous carbon nanospheres. Pu et al.⁹ described a continuous glucose monitoring microsystem consisting of a three-electrode electrochemical sensor in which the working electrode (WE) was covered with a single layer of graphene and gold nanoparticles to improve the sensor performance; the results of glucose measurement were linear below concentration 162 mg/dl with a detection limit 1.44 mg/dl. Li et al.¹⁰ implemented a microfluidic device measuring the glucose concentration with integrated fibre-optic surface plasmon resonance sensor and electrode pairs for volume quantification.Implantable devices and microneedles for drug delivery and liquid transport are addressed in this special topic. Zhang et al.¹¹ reported a flexible polyimide device seated under rabbit eyelids to deliver drug by iontophoresis; varied currents to release manganese ions (Mn²⁺) as tracers were investigated; the thermal effect on application of a current was studied. Lee et al.¹² presented a disposable Parylene microneedle array of large aspect ratio that vibrated with a piezoelectric actuator to mimic the vibrating motion of a mosquito''s proboscis and to decrease the insertion force by 40%. Song et al.¹³ demonstrated microinjection into a model organism, Caenorhabditis elegans (C. elegans) on an automated device capable of loading, immobilization, injection, and sorting; with 200 worms studied, injection speed 6.6 worm/min, injection success rate 77.5%, and sorting success rate 100% were obtained.We express our gratitude for the financial support from Ministry of Science and Technology (Taiwan), Bureau of Foreign Trade (Taiwan), National Taiwan University and Research Center for Applied Sciences of Academia Sinica, and for administrative support from Instrument Technology Research Center in making Optofluidics 2015 a successful conference. Our acknowledgements include Leslie Yeo, Frederick Kontur, Christine Urso, and all staff from Biomicrofluidics for their kind assistance during the preparation, and, most importantly, all authors who have contributed their work for this special topic.  相似文献   

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

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

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

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

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

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

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

Xin Hua  Matthew J. Marshall  Yijia Xiong  Xiang Ma  Yufan Zhou  Abigail E. Tucker  Zihua Zhu  Songqin Liu  Xiao-Ying Yu 《Biomicrofluidics》2015,9(3)

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

Polyphosphonium‐based bipolar membranes for rectification of ionic currents     

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

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

A high-throughput cellulase screening system based on droplet microfluidics     

Raluca Ostafe  Radivoje Prodanovic  W. Lloyd Ung  David A. Weitz  Rainer Fischer 《Biomicrofluidics》2014,8(4)

A new ultra-high-throughput screening assay for the detection of cellulase activity was developed based on microfluidic sorting. Cellulase activity is detected using a series of coupled enzymes leading to the formation of a fluorescent product that can be detected on a chip. Using this method, we have achieved up to 300-fold enrichments of the active population of cells and greater than 90% purity after just one sorting round. In addition, we proved that we can sort the cellulase-expressing cells from mixtures containing less than 1% active cells.Cellulases are important enzymes with numerous applications across multiple industries, including biofuel, pulp, paper, textile and laundry, food, feed, brewing, and agriculture.¹ Most cellulases have low activity and stability, so improving these properties would have substantial impact on numerous industrial processes.Enzymatic properties can be improved by protein engineering² but the limiting step is the screening process. Classical screening uses microtiter plates (MTPs), where each well contains cells expressing a single type of mutant enzyme. However, this type of screening is the bottleneck in directed evolution, because a maximum number of 10⁵ clones can be screened over the course of weeks or even months³ and large quantities of reagents and consumables are needed. High-throughput screening methods based on either fluorescence activated cell sorting (FACS)^4–7 or microfluidic devices⁸ increase the number of clones that can be screened and reduce the amount of consumables required. Here, we demonstrate the use of a high-throughput screening system for cellulases by combining lab-on-chip sorting devices with an emulsion-based fluorescent assay previously developed for use in flow cytometry.⁵Water–in-oil emulsions are needed to maintain the connection between genotype and phenotype by compartmentalizing individual cells expressing a mutant enzyme together with the components of the fluorescence assay corresponding to the enzyme activity.⁷ For FACS, double emulsions (water-in-oil-in-water) are required because the instrument''s mobile phase is an aqueous solution. Such double emulsions can be produced by stirring or agitation,^9,10 but the resulting emulsions are polydisperse and multiple water droplets may be scattered within a single oil droplet. In addition, large droplets tend to produce more fluorescence because there are more substrate molecules available for conversion into the fluorescent product. The emulsions are produced in bulk, so each droplet will be detected at a different time point from the start of the reaction. This means that increased fluorescence may result because an enzyme has worked on the substrate for a longer amount of time, and the fluorescence of the droplet may plateau before sorting as the enzyme consumes all the available substrate. Cell loading is difficult to control because the average number of cells per droplet scales with droplet volume. Also, if several inner droplets, containing cells with different activities, are encapsulated within the same outer droplet, false positives may occur upon sorting. Consequently, it is impossible to differentiate fluorescence changes due to enzyme activity from those due to other effects using polydisperse double emulsions in FACS, but it is possible to achieve plus/minus screening,⁴ separating cells with activity from those without.Droplet microfluidics overcomes many of the drawbacks of high-throughput enzyme sorting with FACS. Both the size and composition of the droplets can be tuned precisely. Furthermore, once the enzyme is mixed with the substrate, the incubation time can be controlled and all compartments will have the same conditions in terms of concentration and total number of substrate molecules. Although cell loading is still subject to Poisson statistics, the probability for cells to be loaded into a given droplet is the same and can be adjusted by tuning the input cell density. These characteristics make the microfluidic method more sensitive, flexible, and quantitative at detecting changes in enzyme activity than the FACS-based sorting of double emulsions.Here, we report a method in which droplet microfluidics is used to sort libraries containing different percentages of cells expressing cellulase activity and demonstrate enrichment of the cells expressing active cellulases. The entire process is summarized in Figure ​Figure11.Open in a separate windowFIG. 1.General overview of cellulase screening using droplet microfluidics. In the emulsification device, suspensions of yeast surface displayed libraries are co-flowed with the substrate solution at equal flow rates to a drop-forming junction where they mix. A stream of perfluorinated oil then breaks the aqueous mixture into monodisperse water-in-oil emulsions. Within each droplet, the cellulase reaction starts after compartmentalization and the fluorescent product is formed by a coupled enzymatic cascade in droplets containing cells that express the active enzyme. After a fixed incubation time, the emulsion droplets are re-injected into a microfluidic sorting device, where they are analyzed and sorted based on their fluorescence.To detect cellulase activity, we designed an assay that uses a chain of coupled enzymatic reactions to yield fluorescence corresponding to cellulase activity without needing artificial substrates (which may lead to confounding effects, such as improved binding of the enzyme specifically to the artificial compound but not the natural substrate). In this method, cellulase hydrolyzes cellulose, its natural substrate, into monosaccharides and oligosaccharides that are further detected by the enzymatic cascade⁵ (Figure ​(Figure11).Based on previous FACS experiments, no difference in activity can be detected between the positive and the negative droplets before 2 h incubation time.⁵ Based on these observations, we expected the cells to require more than 2 h of incubation in droplets for the reaction to develop.Emulsions were formed using a co-flow flow-focusing Polydimethylsiloxane device prepared by soft lithography as previously described⁸ and using fluorocarbon oil containing 1% (v/v) Krytox-PEG-Krytox detergent synthesized as reported in an earlier study.^11,14 The solutions, one containing library cells (S. cerevisiae YPH500 cells, Agilent Technologies, Santa Clara, USA) and the other with the substrate,¹⁴ were mixed at the same flow rate, giving a one-to-one mixing ratio. The library cells were a defined mixture of cells transformed with cel5A pESC-Trp (positive cells) or empty pESC-Trp (negative cells). The two solutions therefore mixed just prior to encapsulation, minimizing the chance that fluorescent products would enter neighboring droplets. The substrate solution contained carboxymethyl cellulose (CMC), which has a high viscosity. To prevent fluctuations in the flow of substrate during the emulsification process, we optimized the flow rate and the concentration of CMC and found that a CMC concentration of 0.33% (w/v) produced monodisperse emulsions.We discovered that the HOx required for the enzymatic cascade causes droplet coalescence. HOx alone was sufficient to cause the observed change in droplet stability because droplets containing only hexose oxidase in buffer exhibited the same amount of coalescence as those containing the full set of assay components. We hypothesized that the enzyme might be surface active, disturbing the emulsion interface, but emulsions of an inactivated form of the enzyme were stable (Figure 2(a)). One possible explanation is that active HOx may interact with the detergent through the active site. Adding bovine serum albumin (BSA), which is known to have a stabilizing effect,¹² to the mixture improved droplet stability (Figure 2(a)). Emulsions of the assay mixture with BSA were stable for more than 1 day at room temperature.Open in a separate windowFIG. 2.(a) Transmission light micrographs of water-in-perfluorinated-oil emulsions produced using the microfluidic emulsification devices after 2 h incubation at room temperature. The emulsions contain 3 U/ml HOx either in its native form (left image), inactivated by heating at 99 °C for 20 min (middle image), or supplemented with 1 mg/ml BSA (right image). (b) Images of the results of the agar plate Congo Red cellulase assay before and after sorting, with the percentage of positive colonies indicated. The cells expressing cellulase activity show clear hallos.The time required for the cellulase reaction to produce detectable quantities of fluorescent product was monitored using the droplet screening instrument. These devices proved to have a higher sensitivity than the FACS system because the optics are designed for the droplet size selected for the assay. We were able to detect cellulase activity just 20 min after the compartmentalization of cells. This shorter incubation time allowed us to couple the emulsification device directly to the droplet sorting device using a short piece of tubing. The rate of emulsion flow and the dimensions of the tube set the droplet incubation time.Using the optimized conditions, we used droplet microfluidics to sort cellulase-expressing cells from a set of reference libraries. The reference libraries were created by mixing different concentrations of positive S. cerevisiae YPH500 cells expressing Cel5A cellulase and negative S. cerevisiae YPH500 cells transformed with the pESC-Trp empty vector. The mixed populations were emulsified together with the assay components in water-in-perfluorinated-oil emulsions and incubated at room temperature for 20 min. The gated population was sorted and the cells were spread on yeast nitrogen base casaminoacids (YNB CAA) Glu agar plates. An aliquot of the reference library was also plated on agar plates prior to sorting. Approximately, 100 cells before and after sorting were transferred to YNB CAA CMC Gal/Raf induction plates, and the Congo red assay¹³ was used to detect cells expressing cellulase. In this assay, colonies of positive cells developed transparent halos around them.¹⁴ The results before and after sorting are presented in Figure 2(b).We enriched cellulase-expressing cells from a pool of negative cells, regardless of the starting concentration of positive cells. We were able to isolate the cellulase-expressing cells even when starting from a low percentage of active cells (0.1%). We obtained high enrichment factors of up to 300 when starting from low concentrations of positive cells, and we were able to sort to a purity of greater than 90%. These results exceed those obtained by comparable experiments using FACS.⁵In conclusion, we developed a high-throughput screening system for cellulase activity based on droplet microfluidics. We optimized the emulsification conditions to produce highly stable and monodisperse droplets. The low dispersity of the emulsion enables the sensitive, tunable, and quantitative detection of cellulase activity. In addition, we substantially reduced the reaction time needed for the development of a fluorescent product from 2 h to 20 min. As a result, we sorted reference libraries of cellulases with various ratios of positive to negative cells, and regardless of the starting population of positive cells we were always able to enrich the active population to a higher purity than that obtained by FACS.  相似文献   

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

Sasan Asiaei  Brendan Smith  Patricia Nieva 《Biomicrofluidics》2015,9(6)

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   

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

Chih-kuan Tung  Robert Riehn    Robert H. Austin 《Biomicrofluidics》2009,3(3)

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

Sampled-data distributed H∞ control of a class of 1-D parabolic systems under spatially point measurements     

Wu-Hua Chen  Shixian Luo  Wei Xing Zheng 《Journal of The Franklin Institute》2017,354(1):197-214

This paper considers the sampled-data distributed $H_{\infty }$ control problem for 1-D semilinear transport reaction equations with external disturbances. It is assumed that a finite number of point spatial state measurements are available. A Razumikhin-type approach is developed for stability and L₂-gain analysis of the closed-loop system. In contrast to Halanay?s inequality based approach, the proposed Razumikhin-type approach not only provides a subtle decay estimate of the selected Lyapunov functional, but also guarantees the $H_{\infty }$ performance index to be negative if certain conditions are satisfied. By introducing a time-dependent Lyapunov functional combined with the use of Wirtinger?s inequality, sufficient conditions for the internal exponential stability and finite L₂-gain are derived in terms of linear matrix inequalities. The obtained conditions establish a quantitative relation among the upper bounds on the spatial sampling intervals and the time sampling intervals, and L₂-gain. Two numerical examples are provided to illustrate the usefulness of the proposed theoretical results.  相似文献