This work was supported by National Science Foundation Award No. ECCS 1610995. The authors would like to thank the Institute of Electronics and Nanotechnology and the Parker H. Petit Institute for Bioengineering and Bioscience staff for their support in using shared facilities. The authors also would like to thank Chia-Heng Chu for his help in preparing the manuscript.

1. Design of Coding Electrodes
Note: Figure 1a shows the 3-D structure of the micropatterned electrodes.
<ol>
	<li>Design a set of four 7-bit Gold codes for encoding the microfluidic channels23.
	<ol>
		<li>Construct two linear feedback shift-registers (LFSRs), each representing a primitive polynomial.</li>
		<li>Use the LFSRs to generate a preferred pair of 7-bit m-sequences.</li>
		<li>Cyclically shift the preferred pair of m-sequences and add them in mod 2 to generate four distinct Gold codes.</li>
	</ol>
	</li>
	<li>Design the layout of the coding electrodes (Figure 1b).
	<ol>
		<li>Place three electrode terminals, representing the positive, negative, and reference electrodes at three corners.</li>
		<li>Route positive and negative electrode traces on opposite sides of each microfluidic channel.</li>
		<li>Extend positive and negative electrodes into the microfluidic channels as electrode fingers, following the uniquely assigned Gold code (Figure 1c).</li>
		<li>Place the reference electrode in between the positive and negative electrode fingers.</li>
		<li>Place positive and negative electrode traces far from the outermost reference electrode fingers in order to minimize electrical conduction outside the coding region.</li>
	</ol>
	</li>
</ol>
2. Microfabrication of Surface Electrodes
Note: Figure 2b shows the fabrication process of surface electrodes.
<ol>
	<li>Clean a 4-inch borosilicate glass wafer in a piranha solution (98% sulfuric acid : 30% hydrogen peroxide = 5 : 1) at 120 &#176;C for 20 min to remove all the organic contaminants. Then place the wafer on a 200 &#176;C hot plate for 20 min to remove residual water.</li>
	<li>Transfer the wafer to a spinner. Dispense 2 mL negative photoresist onto the wafer and spin the wafer at a speed of 3,000 rpm for 40 s to uniformly coat the wafer with a 1.5-&#181;m photoresist layer.</li>
	<li>Place the wafer on a 150 &#176;C hot plate and bake the spun photoresist for 1 min.</li>
	<li>Expose the photoresist to 365-nm UV light (225 mJ/cm2) through a chrome mask using a mask aligner.</li>
	<li>Place the wafer on a 100 &#176;C hot plate and bake the exposed photoresist for 1 min.</li>
	<li>Develop the photoresist by immersing the wafer in a photoresist developer (RD6) for 15 s. Gently spray deionized (DI) water and wash the wafer. Dry by blowing compressed nitrogen.</li>
	<li>Place the wafer with patterned photoresist into an e-beam metal evaporator, and deposit a 20-nm-thick chrome film, followed by an 80-nm-thick gold film onto the wafer at a base pressure of 3&#215;10-6 Torr with a deposition rate of 1 &#197;/s.</li>
	<li>Immerse the metal-coated wafer into acetone in an ultrasonic bath set at a frequency of 40 kHz with 100% amplitude for 30 min at room temperature to etch the underlying photoresist and complete the lift-off process.</li>
	<li>Dice the wafer into smaller pieces using a conventional dicing saw.</li>
</ol>
3. Fabrication of the SU-8 Mold for Microfluidic Channels
Note: Figure 2a shows the fabrication process of the mold for microfluidic channels.
<ol>
	<li>Clean and bake a 4-inch silicon wafer using the same procedure described in 2.1.</li>
	<li>Transfer the wafer to a spinner. Pour 4 mL photoresist onto the wafer. Coat the wafer with photoresist.
	<ol>
		<li>Spin the wafer at 500 rpm for 15 s.</li>
		<li>Spin the wafer at 1,000 rpm for 15 s.</li>
		<li>Spin the wafer at 3,000 rpm for 60 s to obtain a uniformly coated 15-&#181;m thick photoresist layer.</li>
	</ol>
	</li>
	<li>Place the wafer face up on a cleanroom wipe soaked in acetone and remove the residual photoresist from the backside and edges of the wafer.</li>
	<li>Transfer the wafer onto a hot plate for soft baking. First, bake the wafer at 65 &#176;C for 1 min. Then quickly move the wafer to a 95 &#176;C hot plate and bake for 2 min.</li>
	<li>Expose the photoresist to 365-nm UV light (180 mJ/cm2) through a chrome mask by using a mask aligner.</li>
	<li>Bake the wafer following exposure at 65 &#176;C for 1 min and then at 95 &#176;C for 2 min.</li>
	<li>Immerse the wafer in developer and gently shake the container for 3 min. Then, rinse the wafer with isopropanol alcohol (IPA) and dry it by blowing compressed nitrogen. If a white-colored residue appears on the wafer, immerse it into the developer again and develop for longer time and dry.</li>
	<li>Bake the wafer on a 200 &#176;C hot plate for 30 min to dry it completely.</li>
	<li>Measure the thickness of the patterned photoresist using a profilometer at different locations across the wafer to ensure uniformity.</li>
	<li>Silanize the mold wafer by utilizing the technique of vapor deposition. Add 200 &#181;L of trichlorosilane in a Petri dish and place in a vacuum desiccator along with the SU-8 mold wafer for 8 h.</li>
</ol>
4. Assembly of the Microfluidic CODES Device
<ol>
	<li>Place the 4-inch silicon wafer with the mold in a 150-mm diameter Petri dish, and fix it by taping from its edges.</li>
	<li>Mix the polydimethylsiloxane (PDMS) pre-polymer and cross-linker at a ratio of 10:1, and pour 50 g of the mixture into the Petri dish. Place the Petri dish in a vacuum desiccator to degas the mixture for 1 h, and then cure it in an oven at 65 &#176;C for at least 4 h (Figure 2a).</li>
	<li>Cut out the cured PDMS layer using a scalpel and peel it off the mold wafer using a tweezer. The size of the proof-of-principle device is approximately 20 mm &#215; 7 mm. Then punch holes with a diameter of 1.5 mm through the PDMS for the inlet and outlet of the microfluidic channel using a biopsy puncher.</li>
	<li>Clean the patterned side of the PDMS part by placing it on a clean-room adhesive tape.</li>
	<li>Clean the glass substrate with surface electrodes by rinsing it with acetone, IPA, DI water and dry using compressed nitrogen.</li>
	<li>Activate the surfaces of PDMS and glass substrate in oxygen plasma for 30 s with the micromachined side of each part facing up in an RF plasma generator set at 100 mW.</li>
	<li>Align the PDMS microfluidic channel with surface electrodes on the glass substrate using an optical microscope and then bring the two plasma-activated surfaces in physical contact.</li>
	<li>Bake the device on a 70 &#176;C hot plate for 5 min, with the glass side facing the hot plate.</li>
	<li>Connect the contact pads of the electrodes with wires by soldering.</li>
</ol>
5. Preparation of the Simulated Biological Sample
<ol>
	<li>Culture the HeyA8 human ovarian cancer cells in RPMI 1640 supplemented with 10% fetal bovine serum (FBS) and 1% penicillin-streptomycin in 5% CO2 atmosphere at 37 &#176;C until they reach 80% confluence.</li>
	<li>Aspirate the media from the culture flask using a glass pipette. Dispense and then aspirate 1x phosphate buffered saline (PBS) to wash the cells.</li>
	<li>Incubate cells in 2 mL 0.05% (w/v) trypsin solution for 2 min at 37 &#176;C to suspend adherent cells. Then, add 4 mL of the culture media to neutralize the trypsin.</li>
	<li>Centrifuge the cell suspension at 100 &#215; g for 5 min to pellet the cells in a test tube. Then, aspirate supernatant completely.</li>
	<li>Re-suspend the cells in 1-2 mL 1x PBS by gently pipetting up and down to mechanically dissociate cell clumps.</li>
	<li>Draw a small amount of cell suspension into a pipette and count the number of cells using hemocytometer.</li>
	<li>Dilute the cell suspension with PBS to prepare a sample with final cell concentration of 105-106 cells/mL.</li>
</ol>
6. Running the Microfluidic CODES Device
Note: Figure 3 shows the experimental setup.
<ol>
	<li>Place the Microfluidic CODES device on the stage of an optical microscope.</li>
	<li>Apply a 400 kHz sine wave to the reference electrode on the chip using an electronic function generator.</li>
	<li>Connect positive and negative sensing electrodes to two independent trans-impedance amplifiers to convert current signals from each to voltage signals.</li>
	<li>Subtract the positive sensing electrode voltage signal from the negative sensing electrode voltage signal using a differential voltage amplifier in order to obtain a bipolar signal.</li>
	<li>Use a high-speed camera to optically record operation of the device for validation and characterization purposes.</li>
	<li>Drive the cell suspension through the Microfluidic CODES device at a constant flow rate (50-1,000 &#181;L/h) using a syringe pump.</li>
	<li>Measure the impedance modulation signal using a lock-in amplifier.
	<ol>
		<li>Connect the reference AC signal to the reference input of the lock-in amplifier. Connect the differential bipolar signal to the lock-in amplifier as input signal.</li>
		<li>Obtain the RMS amplitude of the differential signal from the lock-in amplifier output.</li>
	</ol>
	</li>
	<li>Sample the lock-in amplifier output signal at 1 MHz rate into a computer through a data acquisition board for further analysis.</li>
</ol>
7. Processing of Sensor Signals
<ol>
	<li>Transfer recorded electrical data into MATLAB for post-processing and decoding.</li>
	<li>Filter the recorded signal in the digital domain using a Butterworth filter (MATLAB built-in function) to remove the high frequency noise (&#62;2.5 kHz).</li>
	<li>Generate a template code library from sensor signals.
	<ol>
		<li>Identify representative non-overlapping code signals corresponding to each sensor in the device and extract these signal blocks from the dataset as separate waveform vectors.</li>
		<li>Normalize each template code waveform vector by its power. Use the MATLAB built-in function (bandpower) to measure the signal power.</li>
		<li>Use MATLAB function (resample) to expand the template library by digitally creating versions of normalized code signals with varying durations to accommodate variations in the cell flow speed over the electrodes.</li>
	</ol>
	</li>
	<li>Identify the signal blocks that correspond to sensor activity (threshold: SNR &#62; 12 dB) in the filtered waveform. Waveform with SNR under the threshold would be treated as noise.</li>
	<li>Decode individual blocks of sensor activity in the recorded signal by using an iterative algorithm based on the successive interference cancellation, a technique commonly employed in multi-user CDMA communication networks24,25.
	<ol>
		<li>Calculate cross-correlation of each signal block with all of the templates in the library using sliding dot product.</li>
		<li>Identify the template that produces the largest auto-correlation peak to determine the dominant individual sensor code signal. Record the both time and amplitude of the autocorrelation peak.</li>
		<li>Construct an estimate sensor code signal by scaling the identified code template based on the measured autocorrelation peak amplitude and timing information (determined in step 7.5.2).</li>
		<li>Subtract the estimated sensor code signal from the original data.</li>
		<li>Iterate the process from 7.5.1, until the residual signal does not resemble any signal in the template library, mathematically defined as the correlation coefficient being less than 0.5.</li>
	</ol>
	</li>
	<li>Refine initial sensor signal estimations from step 7.5 using an optimization process.
	<ol>
		<li>Reconstruct the signal by adding estimated individual sensor signals from each iteration.</li>
		<li>Sweep the amplitude, duration and timing of individual sensor signals around the original estimates to produce the best fit with the recorded electrical signal based on least-squares approximation26.</li>
	</ol>
	</li>
	<li>Convert amplitudes of estimated sensor signals into cell size by calibrating electrical signals against optical images.</li>
</ol>

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The authors have nothing to disclose.

Multiple resistive pulse sensors have previously been incorporated into microfluidic chips28,29,30,31,32. In these systems, resistive pulse sensors were either not multiplexed28,29,30,31 or they required individual sensors to be driv...

Accurate detection and analysis of biological particles such as cells, bacteria or viruses suspended in liquid is of great interest for a range of applications1,2,3. Well-matched in size, microfluidic devices offer unique advantages for this purpose such as high-sensitivity, gentle sample manipulation and well-controlled microenvironment4,5,6,7. In addition, microfluidic devices can be designed to employ a combination of fluid dynamics and force fields to passively fractionate a heterogeneous population of biological particles based on various properties8,9,10,11,12. In those devices, the resultant particle distribution can be used as readout but spatial information is typically accessible only through microscopy, limiting the practical utility of the microfluidic device by tying it to a lab infrastructure. Therefore, an integrated sensor that can readily report particles&#39; spatiotemporal mapping, as they are manipulated on a microfluidic device, can potentially enable low-cost, integrated lab-on-a-chip devices that are particularly attractive for the testing of samples in mobile, resource-limited settings.
Thin film electrodes have been used as integrated sensors in microfluidic devices for various applications13,14. Resistive Pulse Sensing (RPS) is particularly attractive for integrated sensing of small particles in microfluidic channels as it offers a robust, sensitive, and high-throughput detection mechanism directly from electrical measurements15. In RPS, the impedance modulation between a pair of electrodes, immersed in an electrolyte, is used as a means to detect a particle. When the particle passes through an aperture, sized on the order of the particle, the number and amplitude of transient pulses in the electrical current are used to count and size particles, respectively. Moreover, the sensor geometry can be designed with a photolithographic resolution to shape resistive pulse waveforms in order to enhance sensitivity16,17,18,19 or to estimate vertical position of particles in microfluidic channels20.
We have recently introduced a scalable and simple multiplexed resistive pulse sensing technology called Microfluidic Coded Orthogonal Detection by Electrical Sensing (Microfluidic CODES)21. Microfluidic CODES relies on an interconnected network of resistive pulse sensors, each consisting of an array of electrodes micromachined to modulate conduction in a unique, distinguishable manner, so as to enable multiplexing. We have specifically designed each sensor to produce orthogonal electrical signals similar to the digital codes used in code division multiple access22 (CDMA) telecommunication networks, so that individual resistive pulse sensor signal can be uniquely recovered from a single output waveform, even if signals from different sensors interfere. In this way, our technology compresses 2D spatial information of particles into a 1D electrical signal, permitting monitoring of particles at different locations on a microfluidic chip, while keeping both device- and system-level complexity to a minimum.
In this paper, we present a detailed protocol for experimental and computational methods necessary to use the Microfluidic CODES technology, as well as representative results from its use in analysis of simulated biological samples. Using the results from a prototype device with four multiplexed sensors as an example to explain the technique, we provide protocols on (1) the microfabrication process to create microfluidic devices with the Microfluidic CODES technology, (2) the description of the experimental setup including the electronic, optical, and fluidic hardware, (3) the computer algorithm for decoding interfering signals from different sensors, and (4) the results from detection and analysis of cancer cells in microfluidic channels. We believe that using the detailed protocol described here, other researchers can apply our technology for their research.

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microfluidic platform with multiplexed electronic detection for spatial tracking of particles

Microfluidic processing of biological samples typically involves differential manipulations of suspended particles under various force fields in order to spatially fractionate the sample based on a biological property of interest. For the resultant spatial distribution to be used as the assay readout, microfluidic devices are often subjected to microscopic analysis requiring complex instrumentation with higher cost and reduced portability. To address this limitation, we have developed an integrated electronic sensing technology for multiplexed detection of particles at different locations on a microfluidic chip. Our technology, called Microfluidic CODES, combines Resistive Pulse Sensing with Code Division Multiple Access to compress 2D spatial information into a 1D electrical signal. In this paper, we present a practical demonstration of the Microfluidic CODES technology to detect and size cultured cancer cells distributed over multiple microfluidic channels. As validated by the high-speed microscopy, our technology can accurately analyze dense cell populations all electronically without the need for an external instrument. As such, the Microfluidic CODES can potentially enable low-cost integrated lab-on-a-chip devices that are well suited for the point-of-care testing of biological samples.

A Microfluidic CODES device consisting of four sensors distributed over four microfluidic channels is shown in Figure 1b. In this system, the cross-section of each microfluidic channel was designed to be close to the size of a cell so that (1) multiple cells cannot pass over the electrodes in parallel and (2) cells remain close to the electrodes increasing the sensitivity. Each sensor is designed to generate a unique 7-bit digital code. The device was then tested using a ...

Watch this Scientific Journal Video about Microfluidic Platform with Multiplexed Electronic Detection for Spatial Tracking of Particles at JoVE.com

Microfluidic Platform with Multiplexed Electronic Detection for Spatial Tracking of Particles

We demonstrate a microfluidic platform with an integrated surface electrode network that combines resistive pulse sensing (RPS) with code division multiple access (CDMA), to multiplex detection and sizing of particles in multiple microfluidic channels.

Microfluidic processing of biological samples typically involves differential manipulations of suspended particles under various force fields in order to ...

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JoVE Journal

Bioengineering

9.2K Views.  Georgia Institute of Technology. We demonstrate a microfluidic platform with an integrated surface electrode network that combines resistive pulse sensing (RPS) with code division multiple access (CDMA), to multiplex detection and sizing of particles in multiple microfluidic channels.

Video: Microfluidic Platform with Multiplexed Electronic Detection for Spatial Tracking of Particles

A Microfluidic-based Hydrodynamic Trap for Single Particles

The ability to confine and manipulate single particles in free solution is a key enabling technology for fundamental and applied science. Methods for particle trapping based on optical, magnetic, electrokinetic, and acoustic techniques have led to major advancements in physics and biology ranging from the molecular to cellular level. In this article, we introduce a new microfluidic-based technique for particle trapping and manipulation based solely on hydrodynamic fluid flow. Using this method, we demonstrate trapping of micro- and nano-scale particles in aqueous solutions for long time scales. The hydrodynamic trap consists of an integrated microfluidic device with a cross-slot channel geometry where two opposing laminar streams converge, thereby generating a planar extensional flow with a fluid stagnation point (zero-velocity point). In this device, particles are confined at the trap center by active control of the flow field to maintain particle position at the fluid stagnation point. In this manner, particles are effectively trapped in free solution using a feedback control algorithm implemented with a custom-built LabVIEW code. The control algorithm consists of image acquisition for a particle in the microfluidic device, followed by particle tracking, determination of particle centroid position, and active adjustment of fluid flow by regulating the pressure applied to an on-chip pneumatic valve using a pressure regulator. In this way, the on-chip dynamic metering valve functions to regulate the relative flow rates in the outlet channels, thereby enabling fine-scale control of stagnation point position and particle trapping. The microfluidic-based hydrodynamic trap exhibits several advantages as a method for particle trapping. Hydrodynamic trapping is possible for any arbitrary particle without specific requirements on the physical or chemical properties of the trapped object. In addition, hydrodynamic trapping enables confinement of a "single" target object in concentrated or crowded particle suspensions, which is difficult using alternative force field-based trapping methods. The hydrodynamic trap is user-friendly, straightforward to implement and may be added to existing microfluidic devices to facilitate trapping and long-time analysis of particles. Overall, the hydrodynamic trap is a new platform for confinement, micromanipulation, and observation of particles without surface immobilization and eliminates the need for potentially perturbative optical, magnetic, and electric fields in the free-solution trapping of small particles.

In this article, we present a microfluidic-based method for particle confinement based on hydrodynamic flow. We demonstrate stable particle trapping at a fluid stagnation point using a feedback control mechanism, thereby enabling confinement and micromanipulation of arbitrary particles in an integrated microdevice.

The ability to confine and manipulate single particles in free solution is a key enabling technology for fundamental and applied science.  Methods for ...

Fluorescence detection methods for microfluidic droplet platforms

The development of microfluidic platforms for performing chemistry and biology has in large part been driven by a range of potential benefits that accompany system miniaturisation. Advantages include the ability to efficiently process nano- to femoto- liter volumes of sample, facile integration of functional components, an intrinsic predisposition towards large-scale multiplexing, enhanced analytical throughput, improved control and reduced instrumental footprints.1
In recent years much interest has focussed on the development of droplet-based (or segmented flow) microfluidic systems and their potential as platforms in high-throughput experimentation.2-4 Here water-in-oil emulsions are made to spontaneously form in microfluidic channels as a result of capillary instabilities between the two immiscible phases. Importantly, microdroplets of precisely defined volumes and compositions can be generated at frequencies of several kHz. Furthermore, by encapsulating reagents of interest within isolated compartments separated by a continuous immiscible phase, both sample cross-talk and dispersion (diffusion- and Taylor-based) can be eliminated, which leads to minimal cross-contamination and the ability to time analytical processes with great accuracy. Additionally, since there is no contact between the contents of the droplets and the channel walls (which are wetted by the continuous phase) absorption and loss of reagents on the channel walls is prevented.
Once droplets of this kind have been generated and processed, it is necessary to extract the required analytical information. In this respect the detection method of choice should be rapid, provide high-sensitivity and low limits of detection, be applicable to a range of molecular species, be non-destructive and be able to be integrated with microfluidic devices in a facile manner. To address this need we have developed a suite of experimental tools and protocols that enable the extraction of large amounts of photophysical information from small-volume environments, and are applicable to the analysis of a wide range of physical, chemical and biological parameters. Herein two examples of these methods are presented and applied to the detection of single cells and the mapping of mixing processes inside picoliter-volume droplets. We report the entire experimental process including microfluidic chip fabrication, the optical setup and the process of droplet generation and detection.

Droplet-based microfluidic platforms are promising candidates for high throughput experimentation since they are able to generate picoliter, self-compartmentalized vessels inexpensively at kHz rates. Through integration with fast, sensitive and high resolution fluorescence spectroscopic methods, the large amounts of information generated within these systems can be efficiently extracted, harnessed and utilized.

The development of microfluidic platforms for performing chemistry and biology has in large part been driven by a range of potential benefits that ...

Bridging the Bio-Electronic Interface with Biofabrication

Advancements in lab-on-a-chip technology promise to revolutionize both research and medicine through lower costs, better sensitivity, portability, and higher throughput. The incorporation of biological components onto biological microelectromechanical systems (bioMEMS) has shown great potential for achieving these goals. Microfabricated electronic chips allow for micrometer-scale features as well as an electrical connection for sensing and actuation. Functional biological components give the system the capacity for specific detection of analytes, enzymatic functions, and whole-cell capabilities. Standard microfabrication processes and bio-analytical techniques have been successfully utilized for decades in the computer and biological industries, respectively. Their combination and interfacing in a lab-on-a-chip environment, however, brings forth new challenges. There is a call for techniques that can build an interface between the electrode and biological component that is mild and is easy to fabricate and pattern.
Biofabrication, described here, is one such approach that has shown great promise for its easy-to-assemble incorporation of biological components with versatility in the on-chip functions that are enabled. Biofabrication uses biological materials and biological mechanisms (self-assembly, enzymatic assembly) for bottom-up hierarchical assembly. While our labs have demonstrated these concepts in many formats 1,2,3, here we demonstrate the assembly process based on electrodeposition followed by multiple applications of signal-based interactions. The assembly process consists of the electrodeposition of biocompatible stimuli-responsive polymer films on electrodes and their subsequent functionalization with biological components such as DNA, enzymes, or live cells 4,5. Electrodeposition takes advantage of the pH gradient created at the surface of a biased electrode from the electrolysis of water 6,7,. Chitosan and alginate are stimuli-responsive biological polymers that can be triggered to self-assemble into hydrogel films in response to imposed electrical signals 8. The thickness of these hydrogels is determined by the extent to which the pH gradient extends from the electrode. This can be modified using varying current densities and deposition times 6,7. This protocol will describe how chitosan films are deposited and functionalized by covalently attaching biological components to the abundant primary amine groups present on the film through either enzymatic or electrochemical methods 9,10. Alginate films and their entrapment of live cells will also be addressed 11. Finally, the utility of biofabrication is demonstrated through examples of signal-based interaction, including chemical-to-electrical, cell-to-cell, and also enzyme-to-cell signal transmission. 
Both the electrodeposition and functionalization can be performed under near-physiological conditions without the need for reagents and thus spare labile biological components from harsh conditions. Additionally, both chitosan and alginate have long been used for biologically-relevant purposes 12,13. Overall, biofabrication, a rapid technique that can be simply performed on a benchtop, can be used for creating micron scale patterns of functional biological components on electrodes and can be used for a variety of lab-on-a-chip applications.

This article describes a biofabrication approach: deposition of stimuli-responsive polysaccharides in the presence of biased electrodes to create biocompatible films which can be functionalized with cells or proteins. We demonstrate a bench-top strategy for the generation of the films as well as their basic uses for creating interactive biofunctionalized surfaces for lab-on-a-chip applications.

Advancements in lab-on-a-chip technology promise to revolutionize both research and medicine through lower costs, better sensitivity, portability, and ...

Cell Squeezing as a Robust, Microfluidic Intracellular Delivery Platform

Rapid mechanical deformation of cells has emerged as a promising, vector-free method for intracellular delivery of macromolecules and nanomaterials. This technology has shown potential in addressing previously challenging applications; including, delivery to primary immune cells, cell reprogramming, carbon nanotube, and quantum dot delivery. This vector-free microfluidic platform relies on mechanical disruption of the cell membrane to facilitate cytosolic delivery of the target material. Herein, we describe the detailed method of use for these microfluidic devices including, device assembly, cell preparation, and system operation. This delivery approach requires a brief optimization of device type and operating conditions for previously unreported applications. The provided instructions are generalizable to most cell types and delivery materials as this system does not require specialized buffers or chemical modification/conjugation steps. This work also provides recommendations on how to improve device performance and trouble-shoot potential issues related to clogging, low delivery efficiencies, and cell viability.

Rapid mechanical deformation of cells has emerged as a promising, vector-free method for intracellular delivery of macromolecules and nanomaterials. This protocol provides detailed steps on how to use the system for a broad range of applications.

Rapid mechanical deformation of cells has emerged as a promising, vector-free method for intracellular delivery of macromolecules and nanomaterials. This ...

Microfluidic Platform for Measuring Neutrophil Chemotaxis from Unprocessed Whole Blood

Neutrophils play an essential role in protection against infections and their numbers in the blood are frequently measured in the clinic. Higher neutrophil counts in the blood are usually an indicator of ongoing infections, while low neutrophil counts are a warning sign for higher risks for infections. To accomplish their functions, neutrophils also have to be able to move effectively from the blood where they spend most of their life, into tissues, where infections occur. Consequently, any defects in the ability of neutrophils to migrate can increase the risks for infections, even when neutrophils are present in appropriate numbers in the blood. However, measuring neutrophil migration ability in the clinic is a challenging task, which is time consuming, requires large volume of blood, and expert knowledge. To address these limitations, we designed a robust microfluidic assays for neutrophil migration, which requires a single droplet of unprocessed blood, circumvents the need for neutrophil separation, and is easy to quantify on a simple microscope. In this assay, neutrophils migrate directly from the blood droplet, through small channels, towards the source of chemoattractant. To prevent the granular flow of red blood cells through the same channels, we implemented mechanical filters with right angle turns that selectively block the advance of red blood cells. We validated the assay by comparing neutrophil migration from blood droplets collected from finger prick and venous blood. We also compared these whole blood (WB) sources with neutrophil migration from samples of purified neutrophils and found consistent speed and directionality between the three sources. This microfluidic platform will enable the study of human neutrophil migration in the clinic and the research setting to help advance our understanding of neutrophil functions in health and disease.

This protocol details an assay designed to measure human neutrophil chemotaxis from one droplet of whole blood with robust reproducibility. This approach circumvents the need for neutrophil separation and requires only a few minutes of assay preparation time. The microfluidic chip enables the repeated measure of neutrophil chemotaxis over time in infants or small mammals, where sample volume is limited.

Neutrophils play an essential role in protection against infections and their numbers in the blood are frequently measured in the clinic. Higher ...

A Microfluidic Platform for Precision Small-volume Sample Processing and Its Use to Size Separate Biological Particles with an Acoustic Microdevice

A major advantage of microfluidic devices is the ability to manipulate small sample volumes, thus reducing reagent waste and preserving precious sample. However, to achieve robust sample manipulation it is necessary to address device integration with the macroscale environment. To realize repeatable, sensitive particle separation with microfluidic devices, this protocol presents a complete automated and integrated microfluidic platform that enables precise processing of 0.15&#8211;1.5 ml samples using microfluidic devices. Important aspects of this system include modular device layout and robust fixtures resulting in reliable and flexible world to chip connections, and fully-automated fluid handling which accomplishes closed-loop sample collection, system cleaning and priming steps to ensure repeatable operation. Different microfluidic devices can be used interchangeably with this architecture. Here we incorporate an acoustofluidic device, detail its characterization, performance optimization, and demonstrate its use for size-separation of biological samples. By using real-time feedback during separation experiments, sample collection is optimized to conserve and concentrate sample. Although requiring the integration of multiple pieces of equipment, advantages of this architecture include the ability to process unknown samples with no additional system optimization, ease of device replacement, and precise, robust sample processing.

This protocol describes a system architecture for performing automated small volume (0.15&#8211;1.5 ml) particle separations using a microfluidic device, and discusses methods to optimize acoustofluidic device performance and operation.

A major advantage of microfluidic devices is the ability to manipulate small sample volumes, thus reducing reagent waste and preserving precious sample. ...

Photoactivated Localization Microscopy with Bimolecular Fluorescence Complementation (BiFC-PALM)

Protein-protein interactions (PPIs) are key molecular events to biology. However, it remains a challenge to visualize PPIs with sufficient resolution and sensitivity in cells because the resolution of conventional light microscopy is diffraction-limited to ~250 nm. By combining bimolecular fluorescence complementation (BiFC) with photoactivated localization microscopy (PALM), PPIs can be visualized in cells with single molecule sensitivity and nanometer spatial resolution. BiFC is a commonly used technique for visualizing PPIs with fluorescence contrast, which involves splitting of a fluorescent protein into two non-fluorescent fragments. PALM is a recent superresolution microscopy technique for imaging biological samples at the nanometer and single molecule scales, which uses phototransformable fluorescent probes such as photoactivatable fluorescent proteins (PA-FPs). BiFC-PALM was demonstrated by splitting PAmCherry1, a PA-FP compatible with PALM, for its monomeric nature, good single molecule brightness, high contrast ratio, and utility for stoichiometry measurements. When split between amino acids 159 and 160, PAmCherry1 can be made into a BiFC probe that reconstitutes efficiently at 37 &#176;C with high specificity to PPIs and low non-specific reconstitution. Ras-Raf interaction is used as an example to show how BiFC-PALM helps to probe interactions at the nanometer scale and with single molecule resolution. Their diffusion can also be tracked in live cells using single molecule tracking (smt-) PALM. In this protocol, factors to consider when designing the fusion proteins for BiFC-PALM are discussed, sample preparation, image acquisition, and data analysis steps are explained, and a few exemplary results are showcased. Providing high spatial resolution, specificity, and sensitivity, BiFC-PALM is a useful tool for studying PPIs in intact biological samples.

Photoactivated Localization Microscopy with Bimolecular Fluorescence Complementation BiFC-PALM

Protein-protein interactions are visualized in cells with nanometer spatial resolution by combining bimolecular fluorescence complementation (BiFC) with photoactivated localization microscopy (PALM). Described here is the use of BiFC-PALM for imaging Ras-Raf interactions in U2OS cells for visualizing the nanoscale clustering and diffusion of individual Ras-Raf complexes.

Protein-protein interactions (PPIs) are key molecular events to biology. However, it remains a challenge to visualize PPIs with sufficient resolution and ...

A Microfluidic Platform for High-throughput Single-cell Isolation and Culture

Studying the heterogeneity of single cells is crucial for many biological questions, but is technically difficult. Thus, there is a need for a simple, yet high-throughput, method to perform single-cell culture experiments. Here, we report a microfluidic chip-based strategy for high-efficiency single-cell isolation (~77%) and demonstrate its capability of performing long-term single-cell culture (up to 7 d) and cellular heterogeneity analysis using clonogenic assay. These applications were demonstrated with KT98 mouse neural stem cells, and A549 and MDA-MB-435 human cancer cells. High single-cell isolation efficiency and long-term culture capability are achieved by using different sizes of microwells on the top and bottom of the microfluidic channel. The small microwell array is designed for precisely isolating single-cells, and the large microwell array is used for single-cell clonal culture in the microfluidic chip. This microfluidic platform constitutes an attractive approach for single-cell culture applications, due to its flexibility of adjustable cell culture spaces for different culture strategies, without decreasing isolation efficiency.

Here, we present a protocol for isolating and culturing single cells with a microfluidic platform, which utilizes a new microwell design concept to allow for high-efficiency single cell isolation and long-term clonal culture.

Studying the heterogeneity of single cells is crucial for many biological questions, but is technically difficult. Thus, there is a need for a simple, yet ...

Assembly and Tracking of Microbial Community Development within a Microwell Array Platform

The development of microbial communities depends on a combination of complex deterministic and stochastic factors that can dramatically alter the spatial distribution and activities of community members. We have developed a microwell array platform that can be used to rapidly assemble and track thousands of bacterial communities in parallel. This protocol highlights the utility of the platform and describes its use for optically monitoring the development of simple, two-member communities within an ensemble of arrays within the platform. This demonstration uses two mutants of Pseudomonas aeruginosa, part of a series of mutants developed to study Type VI secretion pathogenicity. Chromosomal inserts of either mCherry or GFP genes facilitate the constitutive expression of fluorescent proteins with distinct emission wavelengths that can be used to monitor community member abundance and location within each microwell. This protocol describes a detailed method for assembling mixtures of bacteria into the wells of the array and using time-lapse fluorescence imaging and quantitative image analysis to measure the relative growth of each member population over time. The seeding and assembly of the microwell platform, the imaging procedures necessary for the quantitative analysis of microbial communities within the array, and the methods that can be used to reveal interactions between microbial species area all discussed.

The development of microbial communities depends on a combination of factors, including environmental architecture, member abundance, traits, and interactions. This protocol describes a synthetic, microfabricated environment for the simultaneous tracking of thousands of communities contained in femtoliter wells, where key factors such as niche size and confinement can be approximated.

The development of microbial communities depends on a combination of complex deterministic and stochastic factors that can dramatically alter the spatial ...

A Microfluidic Platform for Longitudinal Imaging in Caenorhabditis elegans

In the last decade, microfluidic techniques have been applied to study small animals, including the nematode Caenorhabditis elegans, and have proved useful as a convenient live imaging platform providing capabilities for precise control of experimental conditions in real time. In this article, we demonstrate live imaging of individual worms employing WormSpa, a previously-published custom microfluidic device. In the device, multiple worms are individually confined to separate chambers, allowing multiplexed longitudinal surveillance of various biological processes. To illustrate the capability, we performed proof-of-principle experiments in which worms were infected in the device with pathogenic bacteria, and the dynamics of expression of immune response genes and egg laying were monitored continuously in individual animals. The simple design and operation of this device make it suitable for users with no previous experience with microfluidic-based experiments. We propose that this approach will be useful for many researchers interested in longitudinal observations of biological processes under well-defined conditions.

A Microfluidic Platform for Longitudinal Imaging in Caenorhabditis elegans

In this article, we demonstrate live imaging of individual worms employing a custom microfluidic device. In the device, multiple worms are individually confined to separate chambers, allowing multiplexed longitudinal surveillance of various biological processes.

In the last decade, microfluidic techniques have been applied to study small animals, including the nematode Caenorhabditis elegans, and have proved ...