巴斯特螺旋预言的证实——纪念巴斯特逝世一百零三年     

尹玉英  刘春蕴 《中国科学院研究生院学报》1998,(2)

为纪念巴斯特逝世１０３年,回顾了立体化学的发展简史和巴斯特在１８６０年对立体化学基础理论所做的预言．螺旋猜想的被证实,证明巴斯特关于立体化学基础理论预言中的三个猜想和两个肯定,都是正确的．  相似文献   

Microfluidic sterilization     

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

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

Table I.

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

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

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

Antoine Béchamp: pere de la biologie. Oui ou non?     

Manchester KL 《Endeavour》2001,25(2):68-73

There is an alternative medicine lobby that, in conjunction with antivivisectionists, believes Louis Pasteur to have been a fraud [R. Bottomley's You Don't Have to Feel Unwell! (Newleaf, 1994) is a recent example]. They frame their accusations around a rivalry between Pasteur and a contemporary, Antoine Béchamp, from whom they suggest Pasteur stole his ideas and then distorted them for his own purposes. This article explores some aspects of the controversies between Béchamp and Pasteur.  相似文献   

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

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

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

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   

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

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

  相似文献   

Development of vertical SU-8 microneedles for transdermal drug delivery by double drawing lithography technology     

Zhuolin Xiang  Hao Wang  Aakanksha Pant  Giorgia Pastorin  Chengkuo Lee 《Biomicrofluidics》2013,7(6)

Polymer-based microneedles have drawn much attention in transdermal drug delivery resulting from their flexibility and biocompatibility. Traditional fabrication approaches are usually time-consuming and expensive. In this study, we developed a new double drawing lithography technology to make biocompatible SU-8 microneedles for transdermal drug delivery applications. These microneedles are strong enough to stand force from both vertical direction and planar direction during penetration. They can be used to penetrate into the skin easily and deliver drugs to the tissues under it. By controlling the delivery speed lower than 2 μl/min per single microneedle, the delivery rate can be as high as 71%.Microelectromechanical systems (MEMS) technology has enabled wide range of biomedical devices applications, such as micropatterning of substrates and cells,¹ microfluidics,² molecular biology on chips,³ cells on chips,⁴ tissue microengineering,⁵ and implantable microdevices.⁶ Transdermal drug delivery using MEMS based devices can delivery insoluble, unstable, or unavailable therapeutic compounds to reduce the amount of those compounds used and to localize the delivery of potent compounds.⁷ Microneedles for transdermal drug delivery are increasingly becoming popular due to their minimally invasive procedure,⁸ promising chance for self-administration,⁹ and low injury risks.¹⁰ Moreover, since pharmaceutical and therapeutic agents can be easily transported into the body through the skin by microneedles,^{11, 12} the microneedles are promising to replace traditional hypodermic needles in the future. Previously, various microneedles devices for transdermal drug delivery applications have been reported. They have been successfully fabricated by different materials, including silicon,¹³ stainless steel,¹⁴ titanium,¹⁵ tantalum,¹⁶ and nickel.¹⁷ Although microneedles with these kinds of materials can be easily fabricated into sharp shape and offer the required mechanical strength for penetration purpose, such microneedles are prone to be damaged¹⁸ and may not be biocompatible.¹⁹ As a result, polymer based microneedles, such as SU-8,^{20, 21} polymethyl meth-acrylate (PMMA),^{22, 23} polycarbonates (PCs),^{24, 25} maltose,^{26, 27} and polylactic acid (PLA),^{28, 29} have caught more and more attentions in the past few years. However, in order to obtain ultra-sharp tips for penetrating the barrier layer of stratum corneum,³⁰ conventional fabrication technologies, for instances, PDMS (Polydimethylsiloxane) molding technology,^{31, 32} stainless steel molding technology,³³ reactive ion etching technology,³⁴ inclined UV (Ultraviolet) exposure technology,³⁵ and backside exposure with integrated lens technology³⁶ are time-consuming and expensive. In this paper, we report an innovative double drawing lithography technology for scalable, reproducible, and inexpensive microneedle devices. Drawing lithography technology³⁷ was first developed by Lee et al. They leveraged the polymers'' different viscosities under different temperatures to pattern 3D structures. However, it required that the drawing frames need to be regular cylinders, which is not proper for our devices. To solve the problem, the new double drawing lithography is developed to create sharp SU-8 tips on the top of four SU-8 pillars for penetration purpose. Drugs can flow through the sidewall gaps between the pillars and enter into the tissues under the skin surface. The experiment results indicate that the new device can have larger than 1N planar buckling force and be easily penetrated into skin for drugs delivery purpose. By delivering glucose solution inside the hydrogel, the delivering rate of the microneedles can be as high as 71% when the single microneedle delivery speed is lower than 2 μl/min.An array of 3 × 3 SU-8 supporting structures was patterned on a 140 μm thick, 6 mm × 6 mm SU-8 membrane (Fig. ​(Fig.1a).1a). Each SU-8 supporting structure included four SU-8 pillars and was 350 μm high. The four pillars were patterned into a tubelike shape on the membrane (Fig. ​(Fig.1b).1b). The inner diameter of the tube was 150 μm, while the outer diameter was 300 μm. SU-8 needles of 700 μm height were created on the top of SU-8 supporting structures to ensure the ability of transdermal penetration. Two PDMS layers were bonded with SU-8 membrane to form a sealed chamber for storing drugs from the connection tube. Once the microneedles entered into the tissue, drugs could be delivered into the body through the sidewall gaps between the pillars (Fig. ​(Fig.1c1c).Open in a separate windowFigure 1Schematic illustration of the SU-8 microneedles. (a) Overview of the whole device; (b) SU-8 supporting structures made of 4 SU-8 pillars; and (c) enlarged view of a single SU-8 microneedle.The fabrication process of SU-8 microneedles is shown in Fig. ​Fig.2.2. SU-8 microneedles fabrication started from a layer of Polyethylene Terephthalate (PET, 3M, USA) film pasted on the Si substrate by sticking the edge area with kapton tape (Fig. ​(Fig.2a).2a). The PET film, a kind of transparent film with poor adhesion to SU-8, was used as a sacrificial layer to dry release the final device from Si substrate. A 140 μm thick SU-8 layer was deposited on the top of this PET film. To ensure a uniform surface of this thick SU-8 layer, the SU-8 deposition was conducted in two steps coating. After exposed under 450 mJ/cm² UV, the membrane pattern could be defined (Fig. ​(Fig.2b).2b). In order to ensure an even surface for following spinning process, another 350 μm SU-8 layer was directly deposited on this layer in two steps without development. With careful alignment, an exposure of 650 mJ/cm² UV energy was performed on this 350 μm SU-8 layer to define the SU-8 supporting structures (Fig. ​(Fig.2c).2c). The SU-8 structure could be easily released from the PET substrate by removing the kapton tape and slightly bending the PET film. Two PDMS layers were bonded with this SU-8 structure by a method reported by Zhang et al.³⁸ (Fig. ​(Fig.2d2d).Open in a separate windowFigure 2Fabrication process for SU-8 microtubes. (a) Attaching a PET film on the Si substrate; (b) exposing the first layer of SU-8 membrane without development; (c) depositing and patterning two continuous SU-8 layers as sidewall pillars; (d) releasing the SU-8 structure from the substrate and bonding it with PDMS; (e) drawing hollowed microneedles on the top of supporting structures; (f) baking and melting the hollowed microneedles to allow the SU-8 flow in the gaps between pillars; and (g) drawing second time on the top of the melted SU-8 flat surface to get microneedles.In our previous work,³⁹ we used one time stepwise controlled drawing lithography technology for the sharp tips integration. However, since the frame used to conduct drawing process in present study is a four-pillars structure rather than a microtube, the conventional drawing process can only make a hollowed tip but not a solid tip structure (Fig. ​(Fig.3).3). This kind of tip was fragile and could not penetrate skin in the practical testing process. To solve the problem, we developed an innovative double drawing lithography process. After bonding released SU-8 structure with PDMS layers (Fig. ​(Fig.2d),2d), we used it to conduct first time stepwise controlled drawing lithography³⁷ and got hollowed tips (Fig. ​(Fig.2e).2e). Briefly, the SU-8 was spun on the Si substrate and kept at 95 °C until the water inside completely vaporized. Device of SU-8 supporting structures was fixed on a precision stage. Then, the SU-8 supporting structures were immersed into the SU-8 by adjusting the precision state. The SU-8 were coated on the pillars'' surface. Then, the SU-8 supporting structures were drawn away from the interface of the liquid maltose and air. After that, the temperature and drawing speed were increased. Since the SU-8 was less viscous at higher temperature, the connection between the SU-8 supporting structures and surface of the liquid SU-8 became individual SU-8 bridge, shrank, and then broke. The end of the shrunk SU-8 bridge forms a sharp tip on the top of each SU-8 supporting structure when the connection was separated. After the hollowed tips were formed in the first step drawing process, the whole device was baked on the hotplate to melt the hollowed SU-8 tips. Melted SU-8 reflowed into the gaps between four pillars and the tips became domes (Fig. ​(Fig.2f).2f). Then, a second drawing process was conducted on the top of melted SU-8 to form sharp and solid tips (Fig. ​(Fig.2g).2g). The final fabricated device is shown in Fig. ​Fig.44.Open in a separate windowFigure 3A hollowed SU-8 microneedle fabricated by single drawing lithography technology (scale bar is 100 μm).Open in a separate windowFigure 4Optical images for the finished SU-8 microneedles.During the double drawing process, as long as the heated time and temperature were controlled, the SU-8 flow-in speed of SU-8 inside the gaps could be precisely determined. The relationship between baking temperature and flow-in speed was studied. As shown in Fig. ​Fig.5,5, the flow-in speed is positive related to the baking temperature. The explanation for this phenomena is that the SU-8''s viscosity is different under different baking temperatures.⁴⁰ Generally, baked SU-8 has 3 status when temperature increases, solid, glass, and liquid. The corresponding viscosity will decrease and the SU-8 can also have higher fluidity. When the baking temperature is larger than 120 °C, the flow-in speed will increase sharply. But, if the baking temperature is higher, the SU-8 will reflow in the gaps too fast, which makes the flow-in depth hard to be controlled. There is a high chance that the whole gaps will be blocked, and no drugs can flow through these gaps any more. Considering that the total SU-8 supporting structure is only 350 μm high, we choose 125 °C as baking temperature for proper SU-8 flow-in speed and easier SU-8 flow-in depth control.Open in a separate windowFigure 5The relationship between flow-in speed and baking temperature.To ensure the adequate stiffness of the SU-8 microneedles in vertical direction, Instron Microtester 5848 (Instron, USA) was deployed to press the microneedles with the similar method reported by Khoo et al.⁴¹ As shown in Fig. ​Fig.6a,6a, the vertical buckling force was as much as 8.1N, which was much larger than the reported minimal required penetration force.⁴² However, in the previous practical testing experiments, even though the microneedles were strong enough in vertical direction, the planar shear force induced by skin deformation might also break the interface between SU-8 pillars and top tips. In our new device with four pillars supporting structure, the SU-8 could flow inside the sidewall gaps between the pillars to form anchors. These anchors could enhance microneedles'' mechanical strength and overcome the planar shear force problems. Moreover, the anchors strength could be improved by controlling the SU-8 flow-in depth. Fig. ​Fig.77 shows that the flow-in depth increases when the baking time increases as the baking time increases at 125 °C. Fig. ​Fig.6b6b shows that the corresponding planar buckling force can be improved to be larger than 1 N by increasing flow-in depth. Some sidewall gaps at bottom are kept on purpose for drugs delivery; hence, the flow-in depth is chosen as 200 μm.Open in a separate windowFigure 6(a) Measurement of the vertical buckling force. (b) The planar buckling force varies under different flow-in depth (I, II, III, and IV corresponding to the certain images in Fig. ​Fig.77).Open in a separate windowFigure 7Different flow-in depth inside the gaps between SU-8 pillars. (a) 0 μm; (b) 100 μm; (c) 200 μm; and (d) 350 μm (scale bar is 100 μm).The penetration capability of the 3 × 3 SU-8 microneedles array is characterized by conducting the insertion experiment on the porcine cadaver skin. 10 microneedles devices were tested and all of them were strong enough to be inserted into the tissue without any breakage. Histology images of the skin at the site of one microneedle penetration were derived to prove that the sharp conical tip was not broken during the insertion process (Fig. ​(Fig.8).8). It also shows penetrated evidence because the hole shape is the same as the sharp conical tip.Open in a separate windowFigure 8Histology image of individual microneedle penetration (scale bar is 100 μm).In order to verify that the drug solution can be delivered into tissue from the sidewall gaps of the microneedles, FITC (Fluorescein isothiocyanate) (Sigma Aldrich, Singapore) solution was delivered through the SU-8 microneedles after they were penetrated into the mouse cadaver skin. The representative results were then investigated via a confocal microscope (Fig. ​(Fig.9).9). The permeation pattern of the solution along the microchannel created by microneedles confirmed the solution delivery results. The black area was a control area without any diffused florescent solution. In contrast, the illuminated area in Fig. ​Fig.99 indicates the area where the solution has diffused to it. These images were taken consecutively from the skin surface down to 180 μm with 30 μm intervals. The diffusion area had a similar dimension with the inserted microneedles. It has proved that the device can be used to deliver drugs into the body.Open in a separate windowFigure 9Images of confocal microscopy to show the florescent solution is successfully delivered into the tissue underneath the skin surface. (a) 30 μm; (b) 60 μm; (c) 90 μm; (d) 120 μm; (e) 150 μm; and (f) 180 μm (scale bar is 100 μm).Due to the uneven surface of deformed skin, there is always tiny gap happened between tips of some microneedles and local surface skin. The microneedles could not be entirely inserted into the tissue. Drugs might leak to the skin surface through the sidewall gaps under certain driven pressure. Hydrogel absorption experiment was conducted to quantify the delivery rate (i.e., the ratio of solution delivered into tissues in the total delivered volume) and to optimize the delivery speed. Using hydrogel as the tissue model for quantitative analysis of microneedle releasing process was reported by Tsioris et al.⁴³ The details are shown here. Gelatin hydrogel was prepared by boiling 70 ml DI (Deionized) water and mixing it with 7 g of Knox^TM original unflavored gelatin powder. The solution was poured into petri dish to 1 cm high. Then, the petri dish was put into a fridge for half an hour. Gelatin solution became collagen slabs. The collagen slabs were cut into 6 mm × 6 mm sections. A piece of fully stretched parafilm (Parafilm M, USA) was tightly mounted on the surface of the collagen slabs. This parafilm was used here to block the leaked solution further diffusing into the collagen slab in the delivery process. Then, the microneedles penetrated the parafilm and went into the collagen slab. Controlled by a syringe pump, 0.1 ml–0.5 mg/ml glucose solution was delivered into the collagen slab under different speeds. Methylene Blue (Sigma Aldrich, Singapore) was mixed into the solution for better inspection purpose (Fig. 10a). Then, the collagen slabs was digested in 1 mg/ml collagenase (Sigma Aldrich, Singapore) at room temperature (Fig. 10b). It took around 1 h that all the collagen slabs could be fully digested (Fig. 10d). The solution was collected to measure the glucose concentration with glucose detection kit (Abcam, Singapore). Briefly, both diluted glucose standard solution and the collected glucose solution were added into a series of wells in a well plate. Glucose assay buffer, glucose enzyme, and glucose substrate were mixed with these samples in the wells. After incubation for 30 min, their absorbance were examined by using a microplate reader at a wavelength of 450 nm. By comparing the readings with the measured concentration standard curve (Fig. 11a), the glucose concentration in the hydrogel, the glucose absorption rate in the hydrogel, and the solution delivery rate by microneedles could be measured and calculated. As shown in Fig. 11b, when the delivering speed of a single microneedle increased from 0.1 μl/min to 2 μl/min, the glucose absorption rate also increased. Most of the glucose solution from microneedles could go into the hydrogel. The delivered rate could be as high as 71%. The rest solution leaked from sidewall gaps and blocked by parafilm. However, when the delivered speed for a single microneedle was larger than 2 μl/min, the hydrogel absorption rate was saturated. More and more solution could not go into the hydrogel but leak from the sidewall gaps. Then, the delivered rate decreased. Therefore, 2 μl/min was chosen as the optimized delivery speed for the microneedle.Open in a separate windowFigure 10Glucose solution could be delivered into the hydrogel, and the collagen stabs were dissolved by collagenase.Open in a separate windowFigure 11(a) Standard curve for glucose detection; (b) glucose absorption rate and solution delivery rate in a single needle corresponding to different delivery speed.In conclusion, a drug delivery device of integrated vertical SU-8 microneedles array is fabricated based on a new double drawing lithography technology in this study. Compared with the previous biocompatible polymer-based microneedles fabrication technology, the proposed fabrication process is scalable, reproducible, and inexpensive. The fabricated microneedles are rather strong along both vertical and planar directions. It is proved that the microneedles were penetrated into the pig skin easily. The feasibility of drug delivery using SU-8 microneedles is confirmed by FITC fluorescent delivery experiment. In the hydrogel absorption experiment, by controlling the delivery speed under 2 μl/min per microneedle, the delivery rate provided the microneedle is as high as 71%. In the next step, the microneedles will be further integrated with microfluidics on a flexible substrate, forming a skin-patch like drug delivery device, which may potentially demonstrate a self-administration function. When patients need an injection treatment at home, they can easily use such a device just like using an adhesive bandage strip.  相似文献   

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

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

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

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

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

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

Water transport to the core–mantle boundary     

Michael J Walter 《国家科学评论(英文版)》2021,8(4)

Water is transported to Earth''s interior in lithospheric slabs at subduction zones. Shallow dehydration fuels hydrous island arc magmatism but some water is transported deeper in cool slab mantle. Further dehydration at ∼700 km may limit deeper transport but hydrated phases in slab crust have considerable capacity for transporting water to the core-mantle boundary. Quantifying how much remains the challenge.

Water can have remarkable effects when exposed to rocks at high pressures and temperatures. It can form new minerals with unique properties and often profoundly affects the physical, transport and rheological properties of nominally anhydrous mantle minerals. It has the ability to drastically reduce the melting point of mantle rocks to produce inviscid and reactive melts, often with extreme chemical flavors, and these melts can alter surrounding mantle with potential long-term geochemical consequences. At the base of the mantle, water can react with core iron to produce a super-oxidized and hydrated phase, FeO₂H_x, with the potential to profoundly alter the mantle and even the surface and atmosphere redox state, but only if enough water can reach such depths [1].Current estimates for bulk mantle water content based on the average H₂O/Ce ratio of oceanic basalts from melt inclusions and the most un-degassed basalts, coupled with mass balance constraints for Ce, indicate a fraction under one ocean mass [2], a robust estimate as long as the basalts sampled at the surface tap all mantle reservoirs. The mantle likely contains some primordial water but given that the post-accretion Earth was very hot, water has low solubility and readily degasses from magma at low pressures, and its solubility in crystallizing liquidus minerals is also very low, the mantle just after accretion may have been relatively dry. Thus, it is plausible that most or even all of the water in the current mantle is ‘recycled’, added primarily by subduction of hydrated lithospheric plates. If transport of water to the core–mantle boundary is an important geological process with planet-scale implications, then surface water incorporated into subducting slabs and transported to the core–mantle boundary may be a requirement.Water is added to the basaltic oceanic crust and peridotitic mantle in lithospheric plates (hereafter, slab crust and slab mantle, respectively) at mid-ocean ridges, at transform faults, and in bending faults formed at the outer rise prior to subduction [3]. Estimates vary but about 1 × 10¹² kg of water is currently subducted each year into the mantle [4], and at this rate roughly 2–3 ocean masses could have been added to the mantle since subduction began. However, much of this water is returned to the surface through hydrous magmatism at convergent margins, which itself is a response to slab dehydration in an initial, and large, release of water. Meta-basalt and meta-sediments comprising the slab crust lose their water very efficiently beneath the volcanic front because most slab crust geotherms cross mineral dehydration or melting reactions at depths of less than 150 km, and even if some water remains stored in minerals like lawsonite in cooler slabs, nearly complete dehydration is expected by ∼300 km [5].Peridotitic slab mantle may have much greater potential to deliver water deeper into the interior. As shown in Fig. 1a, an initial pulse of dehydration of slab mantle occurs at depths less than ∼200 km in warmer slabs, controlled primarily by breakdown of chlorite and antigorite when slab-therms cross a deep ‘trough’, sometimes referred to as a ‘choke point’, along the dehydration curve (Fig. 1a) [6]. But the slab mantle in cooler subduction zones can skirt beneath the dehydration reactions, and antigorite can transform directly to the hydrated alphabet silicate phases (Phases A, E, superhydrous B, D), delivering perhaps as much as 5 wt% water in locally hydrated regions (e.g. deep faults and fractures in the lithosphere) to transition zone depths [6]. Estimates based on mineral phase relations in the slab crust and the slab mantle coupled with subduction zone thermal models suggest that as much as 30% of subducted water may have been transported past the sub-volcanic dehydration front and into the deeper mantle [4], although this depends on the depth and extent of deep hydration of the slab mantle, which is poorly constrained. Coincidentally, this also amounts to about one ocean mass if water subduction rates have been roughly constant since subduction began, a figure tantalizingly close to the estimated mantle water content based on geochemical arguments [2]. But what is the likely fate of water in the slab mantle in the transition zone and beyond?Open in a separate windowFigure 1.(a) Schematic phase relations in meta-peridotite modified after [6,10,12]. Slab geotherms are after those in [4]. Cold slabs may transport as much as 5 wt% water past ‘choke point 1’ in locally hydrated regions of the slab mantle, whereas slab mantle is dehydrated in warmer slabs. Colder slab mantle that can transport water into the transition zone will undergo dehydration at ‘choke point 2’. How much water can be transported deeper into the mantle and potentially to the core depends on the dynamics of fluid/melt segregation in this region. (b) Schematic showing dehydration in the slab mantle at choke point 2. Migration of fluids within slab mantle will result in water dissolving into bridgmanite and other nominally anhydrous phases with a bulk storage capacity of ∼0.1 wt%, potentially accommodating much or all of the released water. Migration of fluids out of the slab into ambient mantle would also hydrate bridgmanite and other phases and result in net fluid loss from the slab. Conversely, migration of hydrous fluids into the crust could result in extensive hydration of meta-basalt with water accommodated first in nominally anhydrous phases like bridgmanite, Ca-perovskite and NAL phase, but especially in dense SiO₂ phases (stishovite and CaCl₂-type) that can host at least 3 wt% water (∼0.6 wt% in bulk crust).Lithospheric slabs are expected to slow down and deform in the transition zone due to the interplay among the many factors affecting buoyancy and plate rheology, potentially trapping slabs before they descend into the lower mantle [7]. If colder, water-bearing slabs heat up by as little as a few hundred degrees in the transition zone, hydrous phases in the slab mantle will break down to wadsleyite or ringwoodite-bearing assemblages, and a hydrous fluid (Fig. 1a). Wadselyite and ringwoodite can themselves accommodate significant amounts of water and so hydrated portions of the slab mantle would retain ∼1 wt% water. A hydrous ringwoodite inclusion in a sublithospheric diamond with ∼1.5 wt% H₂O may provide direct evidence for this process [8].But no matter if slabs heat up or not in the transition zone, as they penetrate into the lower mantle phase D, superhydrous phase B or ringwoodite in the slab mantle will dehydrate at ∼700–800 km due to another deep trough, or second ‘choke point’, transforming into an assemblage of nominally anhydrous minerals dominated by bridgmanite (∼75 wt%) with, relatively, a much lower bulk water storage capacity (< ∼0.1 wt%) [9] (Fig. 1a). Water released from the slab mantle should lead to melting at the top of the lower mantle [10], and indeed, low shear-wave velocity anomalies at ∼700–800 km below North America may be capturing such dehydration melting in real time [11].The fate of the hydrous fluids/melts released from the slab in the deep transition zone and shallow lower mantle determines how much water slabs can carry deeper into the lower mantle. Presumably water is released from regions of the slab mantle where it was originally deposited, like the fractures and faults that formed in the slab near the surface [3]. If hydrous melts can migrate into surrounding water-undersaturated peridotite within the slab, then water should dissolve into bridgmanite and coexisting nominally anhydrous phases (Ca-perovskite and ferropericlase) until they are saturated (Fig. 1b). And because bridgmanite (water capacity ∼0.1 wt%) dominates the phase assemblage, the slab mantle can potentially accommodate much or all of the released water depending on details of how the hydrous fluids migrate, react and disperse. If released water is simply re-dissolved into the slab mantle in this way then it could be transported deeper into the mantle mainly in bridgmanite, possibly to the core–mantle boundary. Water solubility in bridgmanite throughout the mantle pressure-temperature range is not known, so whether water would partially exsolve as the slab moves deeper stabilizing a melt or another hydrous phase, or remains stable in bridgmanite as a dispersed, minor component, remains to be discovered.Another possibility is that the hydrous fluids/melts produced at the second choke point in the slab mantle at ∼700 km migrate out of the slab mantle, perhaps along the pre-existing fractures and faults where bridgmanite-rich mantle should already be saturated, and into either oceanic crust or ambient mantle (Fig. 1b). If the hydrous melts move into ambient mantle, water would be consumed by water-undersaturated bridgmanite, leading to net loss of water from the slab to the upper part of the lower mantle, perhaps severely diminishing the slab’s capacity to transport water to the deeper mantle and core. But what if the water released from slab mantle migrates into the subducting, previously dehydrated, slab crust?Although slab crust is expected to be largely dehydrated in the upper mantle, changes in its mineralogy at higher pressures gives it the potential to host and carry significant quantities of water to the core–mantle boundary. Studies have identified a number of hydrous phases with CaCl₂-type structures, including δ-AlOOH, ϵ-FeOOH and MgSiO₂(OH)₂ (phase H), that can potentially stabilize in the slab crust in the transition zone or lower mantle. Indeed, these phases likely form extensive solid solutions such that an iron-bearing, alumina-rich, δ-H solid solution should stabilize at ∼50 GPa in the slab crust [12], but only after the nominally anhydrous phases in the crust, (aluminous bridgmanite, stishovite, Ca-perovskite and NAL phase) saturate in water. Once formed, the δ-H solid solution in the slab crust may remain stable all the way to the core mantle boundary if the slab temperature remains well below the mantle geotherm otherwise a hydrous melt may form instead [12] (Fig. 1a). But phase δ-H solid solution and the other potential hydrated oxide phases, intriguing as they are as potential hosts for water, may not be the likely primary host for water in slab crust. Recent studies suggest a new potential host for water—stishovite and post-stishovite dense SiO₂ phases [13,14].SiO₂ minerals make up about a fifth of the slab crust by weight in the transition zone and lower mantle [15] and recent experiments indicate that the dense SiO₂ phases, stishovite (rutile structure—very similar to CaCl₂ structure) and CaCl₂-type SiO₂, structures that are akin to phase H and other hydrated oxides, can host at least 3 wt% water, which is much more than previously considered. More importantly, these dense SiO₂ phases apparently remain stable and hydrated even at temperatures as high as the lower mantle geotherm, unlike other hydrous phases [13,14]. And as a major mineral in the slab crust, SiO₂ phases would have to saturate with water first before other hydrous phases, like δ-H solid solution, would stabilize. If the hydrous melts released from the slab mantle in the transition zone or lower mantle migrate into slab crust the water would dissolve into the undersaturated dense SiO₂ phase (Fig. 1b). Thus, hydrated dense SiO₂ phases are possibly the best candidate hosts for water transport in slab crust all the way to the core mantle boundary due to their high water storage capacity, high modal abundance and high-pressure-temperature stability.Once a slab makes it to the core–mantle boundary region, water held in the slab crust or the slab mantle may be released due to the high geothermal gradient. Heating of slabs at the core–mantle boundary, where temperatures may exceed 3000°C, may ultimately dehydrate SiO₂ phases in the slab crust or bridgmanite (or δ-H) in the slab mantle, with released water initiating melting in the mantle and/or reaction with the core to form hydrated iron metal and super oxides, phases that may potentially explain ultra-low seismic velocities in this region [1,10]. How much water can be released in this region from subducted lithosphere remains a question that is hard to quantify and depends on dynamic processes of dehydration and rehydration in the shallower mantle, specifically at the two ‘choke points’ in the slab mantle, processes that are as yet poorly understood. What is clear is that subducting slabs have the capacity to carry surface water all the way to the core in a number of phases, and possibly in a phase that has previously seemed quite unlikely, dense SiO₂.  相似文献   

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

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

  相似文献   

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

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

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

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   

Black silicon as a platform for bacterial detection     

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

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

FROM PARIS TO NARRANDERA: ADAPTATION IN THE DIFFUSION OF ANTHRAX VACCINATION IN THE AUSTRALIAN PASTORAL INDUSTRY     

《普罗米修斯》2012,30(1):32-48

In 1881 Louis Pasteur demonstrated to an incredulous world the efficacy of his vaccine for the prevention of the fatal disease anthrax in sheep and cattle. A decade later Pasteur's nephew and assistant, Adrien Loir, was manufacturing the vaccine in Sydney. But although Loir's operations were begun at the behest of local pastoralists, the diffusion of the Pasteur vaccine was not very successful in Australia. Within a few years it was virtually displaced from the market by the modified vaccine produced by two Australian men, John Alexander Gunn and John McGarvie Smith. The process of transfer and diffusion involved in that displacement is the concern of this paper. In particular, it highlights the importance of local adaptation for the successful diffusion of anthrax vaccination in Australia.  相似文献