The authors thank Ronald F. Renzi and Michael S. Bartsch for their contributions to the design and development of DMF devices and the DMF hub. This research was fully supported by the Laboratory Directed Research and Development program at Sandia National Laboratories. Sandia is a multi-program laboratory managed and operated by Sandia Corporation, a wholly owned subsidiary of Lockheed Martin Corporation, for the U.S. Department of Energy's National Nuclear Security Administration under contract DE-AC04-94AL85000.

This general protocol is designed to support application of the platform to a wide variety of studies; aspects specific to the representative study described in this report are separated out in brackets. Figure 2 illustrates a representative study carried out using the protocol. Note that in the protocol, all "instruct" commands are automated through use of pre-determined scripts. Note too that Step 2 can be carried out in parallel with Step 1 (e.g. during Steps 1.7 and 1.8), and that Step 2.1 is often bypassed (because patterned/coated DMF device plates can be washed and re-used; see Step 5.2).
1. Assemble and Populate the Cell Perfusion Chambers
<ol>
	<li>Coat the interior surfaces of sterilized (autoclaved) microcapillaries (fused-silica; 679 &mu;m o.d., 534 &mu;m i.d; 15 cm long) with fibronectin, by instructing the syringe pump to draw a sterile fibronectin solution (30 &mu;l of 50 &mu;g/ml) into each microcapillary, incubating (37 &deg;C for 2 hr), instructing the syringe pump to withdraw the solution, and letting the microcapillaries dry (room temperature (RT) for 6 hr).</li>
	<li>Connect each fibronectin-coated microcapillary (perfusion chamber) to polycarbonate tubing (500 &mu;m o.d., 100 &mu;m i.d.) and the tubing to the multi-port syringe pump, using CapTite fittings (6 perfusion chambers, 8-port syringe pump).</li>
	<li>Using tape, secure the body of each perfusion chamber to a heat block set to desired temperature (37 &deg;C).</li>
	<li>Immerse the open ends of the perfusion chambers in a reservoir (microcentifuge tube or microtiter plate well) containing cells (P388D.1 murine macrophages) suspended in fresh growth medium (RPMI-1640, 10% FBS, 500 units/ml penicillin, 500 &mu;g/ml streptomycin) at desired concentration (106 cells/ml).</li>
	<li>Instruct the syringe pump to draw cells (10 &mu;l, 104 cells) into each perfusion chamber followed by enough air to move the liquid plug (~4.5 cm) to the section secured to the heat block.</li>
	<li>Allow the cells to adhere to the fibronectin-coated interior surfaces of the perfusion chambers (37 &deg;C for 1 - 2 hr). In the meantime, transfer the open ends of the perfusion chambers from the cells reservoir to a new reservoir containing fresh growth medium.</li>
	<li>After the adherence period, instruct the syringe pump to send the liquid plugs (conditioned media and unattached cells) to waste, withdraw fresh medium (10 &mu;l), and follow with enough air to position the new liquid plugs (fresh medium) over the adhered cells.</li>
	<li>Continue to carry out medium exchanges (i.e. repeat Step 1.7) at regular intervals (every 2 hr) until the cell populations are sufficiently equilibrated and expanded (16 - 24 hr of micro-scale culture generated populations of ~1,000 cells/chamber).</li>
</ol>
2. Fabricate the DMF Device and Assemble the Hub Architecture
<ol>
	<li>As described previously14,29, pattern the bottom plate of the DMF device with forty-six indium tin oxide (ITO) electrodes (drivers of droplet actuation) by photolithography and etching, and coat with 5 &mu;m of Parylene-C and 50 nm of Teflon-AF. Form the top plate of the DMF device by coating an unpatterned ITO glass substrate with Teflon-AF, as above.</li>
	<li>Using metal compression frames, fix the DMF plates into a polymer cast14,30 with recesses that maintain a 400 &mu;m spacing between the plates; this spacing, combined with the actuation electrode size (2.5 mm2), defines the droplet volume (2.5 &mu;l per electrode). Note that DMF assembly can be accomplished by simpler means24.</li>
	<li>Insert Teflon-coated transfer microcapillaries (360 &mu;m o.d., 100 &mu;m i.d.; 3.5 - 4.0 cm long) into the space between the DMF plates, positioning each to extend to the edge of its cognate actuation electrode.</li>
	<li>To the opposite end of each transfer microcapillary affix a CapTite fitting.</li>
	<li>Engage the electrical connectors to supply voltage to the ITO electrodes. Electrode activation sequences are automated through use of a computer-controlled electronic interface running pre-determined scripts.</li>
	<li>Optional: Move the assembled DMF hub onto the translational stage of a microscope (SZ-6145TR (Olympus, Japan)) equipped with a charge-coupled device (CCD) camera (DCR-HC96 (Sony, Japan)) to facilitate tracking of droplets.31</li>
</ol>
3. Connect the Perfusion Chambers to the DMF Hub and Stimulate the Cell Populations
<ol>
	<li>Remove the open ends of the perfusion chambers from the medium reservoir (see Step 1.6) and insert them into the CapTite fittings affixed to the ends of the transfer microcapillaries of the DMF hub (see Step 2.4).</li>
	<li>Instruct the syringe pump to send the liquid plugs (conditioned media) in the perfusion chambers to waste.</li>
	<li>Instruct the DMF hub to draw stimulus (E. coli BioParticles at 100 &mu;g/ml in fresh medium with 0.1% Pluronic F127 w/v) from an on-board reservoir and deliver a droplet of appropriate size (10 &mu;l) to each transfer microcapillary. The on-board reservoirs are comprised of cylindrical-shaped compartments (1.5 x 1.5 cm each) and connected to the DMF Hub via tubing.</li>
	<li>Instruct the syringe pump to draw the stimulus plugs into the transfer microcapillaries, and follow with enough air to position the plugs over the cell populations in the perfusion chambers.</li>
	<li>Incubate the cells with stimulus (37 &deg;C for 1 - 4 hr). Optional: Exchange for fresh stimulus (i.e. repeat Steps 3.2-3.4) at regular intervals.</li>
</ol>
4. Terminate Stimulation and Recover Cell Lysates for Analysis
<ol>
	<li>Optional: If a post-stimulation period is required, exchange the stimulus-containing liquid plugs for fresh medium (i.e. repeat Steps 3.2 - 3.4, substituting fresh medium for stimulus), and continue to incubate and exchange as needed.</li>
	<li>Exchange the liquid plugs in the perfusion chambers for wash buffer (Prelude Direct Lysis Module Buffer B) (i.e. repeat Steps 3.2 - 3.4, substituting wash buffer for stimulus), and incubate as needed (5 min).</li>
	<li>Exchange the liquid plugs (wash buffer) for lysis solution (Prelude Direct Lysis Module Buffer A with 0.1% Pluronic F127 w/v) (i.e. repeat Steps 3.2 - 3.4, substituting lysis solution for stimulus), and incubate as needed (5 min).</li>
	<li>Instruct the syringe pump to send the liquid plugs (cell lysates) to the DMF hub.</li>
	<li>Disassemble the DMF hub, remove the top plate of the DMF device (using care not to tilt the plate sideways, as this can result in droplet mixing), and collect the lysates using a Pipetman or syringe for off-platform analysis (gene expression profiling via qPCR).</li>
</ol>
5. Clean Platform Hardware for Re-use
<ol>
	<li>Immerse the transfer microcapillaries and CapTite fittings in 10% bleach (v/v in water) for 10 min at RT, rinse well with deionized water, and dry using nitrogen gas.</li>
	<li>Immerse the DMF device plates in 10% bleach for 10 min at RT, rinse well with isopropanol followed by deionized water, and dry using nitrogen gas followed by baking at 160 &deg;C for 10 min.</li>
	<li>The syringe pump's polycarbonate tubing, and the DMF device's polymer cast and compression frame, can be re-used without cleaning. The perfusion chamber microcapillaries are discarded after each experiment.</li>
</ol>

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The authors declare that they have no competing financial interests.

We have developed a simple and versatile automated platform for micro-scale cell culture and stimulation experiments. The platform enables us to work with small culture volumes and cell populations (1 - 20 &mu;l and 100 - 2,000 cells, per chamber); culture sizes could be further reduced through use of microcapillaries of smaller diameter. Working at these scales reduces the cost of routine studies and makes feasible studies that require use of precious reagents and/or cells. Platform-executed experiments also sh...

The study of cells maintained in culture (in vitro analysis) has provided invaluable insight into the fundamental principles and molecular mechanisms governing complex biological systems and human health. The conventional methods for culture, stimulation, and collection of cells for analysis, which utilize Petri dishes and microtiter plates, were designed for study of large populations of cells (&ge;105) in milliliter-scale culture volumes (&ge;0.1 ml). However, there are many instances in which only limited quantities of cells are available (e.g. primary cells), or small populations of cells are desirable (e.g. to reduce cell-to-cell variability across the population), or required reagents are difficult to obtain or prohibitively expensive (e.g. purified cell-secreted factors). Such issues can be successfully addressed by scaling down culture size, which has the added benefit of reducing consumption of all reagents required for in vitro analysis1,2. Unfortunately, conventional equipment and methods do not support precise and reproducible manipulation of micro-scale cultures and the microfluidics-based automated systems currently available3-11 are too complex and specialized for routine use by most laboratories.
In this report, we describe assembly and use of a simple and versatile technology platform for automated culture, stimulation, and recovery of small populations of cells (100 - 2,000 cells) in micro-scale volumes (1 - 20 &mu;l). The platform architecture (Figure 1) is modular in design: a set of fibronectin-coated microcapillaries ("cell perfusion chambers" module) serves as the site for establishment, maintenance, and stimulation of micro-scale cultures; and a digital microfluidics (DMF)12,13 device outfitted with "transfer" microcapillaries ("central hub" module)14,15 routes cells and reagents to and from the perfusion chambers. DMF enables the user to individually address multiple droplets simultaneously and to change or re-order the manipulations (i.e. reconfigure sample processing trains) without altering the device hardware. Its tremendous flexibility is evident in its recent emergence as a key technology in a wide range of applications, including cell culture16,17, enzyme assays18,19, immunoassays20,21, DNA analysis22,23, protein processing,24,25 and clinical specimen processing.26,27 Our central hub takes advantage of the flexibility inherent to DMF devices, and further enhances it through the addition of microcapillary interfaces, which provides opportunity to carry out a subset of manipulations (e.g. cell culture) in specialized peripheral modules, rather than on the DMF device itself. Compartmentalization of processing trains in this way also simplifies design of the platform architecture (no need to build a DMF device that can carry out all processing steps) and facilitates its evolution as new functions are required (simply integrate new peripheral modules as necessary). Transport of cells and reagents within the central hub is driven by electrowetting forces generated by sequential activation of electrodes within the DMF device13,28; transport to, from, and within the perfusion chambers is powered by pressure changes generated by a high-precision syringe pump. All of these fluid movements are controlled via a simple electronic interface and automated through use of pre-determined scripts.
As a representative example, we demonstrate the use of the platform for the study of transcriptional responses elicited in immune cells upon challenge with bacteria (Figure 2). Carrying out these experiments on the platform enabled us to work with small numbers of cells (~1,000 per experimental condition), minimize experiment-to-experiment variability, conserve reagents, and re-direct hands-on labor. Given the advantages that it confers, as well as its accessibility and versatility, this platform should find use in a wide variety of laboratories and applications and prove especially useful in facilitating analysis of cells and stimuli that are available in only limited quantities.

<table border="1">
 <tbody>
 <tr>
 <td>Name of Reagent/Material</td>
 <td>Company</td>
 <td>Catalog Number</td>
 <td>Comments</td>
 </tr>
 <tr>
 <td>Prelude Direct Lysis Module</td>
 <td>NuGEN</td>
 <td>1400-24</td>
 <td></td>
 </tr>
 <tr>
 <td>Trypan Blue (0.4% w/v)</td>
 <td>GIBCO</td>
 <td>15250-061</td>
 <td></td>
 </tr>
 <tr>
 <td>Cell Stripper</td>
 <td>Cellgro</td>
 <td>25-056-C1</td>
 <td></td>
 </tr>
 <tr>
 <td>Ovation PicoSL WTA</td>
 <td>NuGEN</td>
 <td>3310-048</td>
 <td></td>
 </tr>
 <tr>
 <td>Agencourt RNAClean XP</td>
 <td>Beckman Coulter Genomics</td>
 <td>A63987</td>
 <td></td>
 </tr>
 <tr>
 <td>pHrodo BioParticles</td>
 <td>Invitrogen</td>
 <td>P35361</td>
 <td></td>
 </tr>
 <tr>
 <td>CCL4 TaqMan qRT-PCR assay</td>
 <td>Applied Biosystems</td>
 <td>Mm00443111_m1</td>
 <td></td>
 </tr>
 <tr>
 <td>CCL5 TaqMan qRT-PCR assay</td>
 <td>Applied Biosystems</td>
 <td>Mm01302428_m1</td>
 <td></td>
 </tr>
 <tr>
 <td>PTGS2 TaqMan qRT-PCR assay</td>
 <td>Applied Biosystems</td>
 <td>Mm00478374_m1</td>
 <td></td>
 </tr>
 <tr>
 <td>TNF TaqMan qRT-PCR assay</td>
 <td>Applied Biosystems</td>
 <td>Mm00443258_m1</td>
 <td></td>
 </tr>
 <tr>
 <td>GAPDH TaqMan qRT-PCR assay</td>
 <td>Applied Biosystems</td>
 <td>Mm99999915_g1</td>
 <td></td>
 </tr>
 <tr>
 <td>Pluronic F127</td>
 <td>Sigma Chemical</td>
 <td>2594628</td>
 <td></td>
 </tr>
 <tr>
 <td>Fluorinert FC-40</td>
 <td>Sigma Chemical</td>
 <td>51142-49-5</td>
 <td></td>
 </tr>
 <tr>
 <td>Parylene C dimer</td>
 <td>Specialty Coating Systems</td>
 <td>28804-46-8</td>
 <td></td>
 </tr>
 <tr>
 <td>Teflon-AF</td>
 <td>DuPont</td>
 <td>AF1600</td>
 <td></td>
 </tr>
 <tr>
 <td>Polyimide tape</td>
 <td>ULINE</td>
 <td>S-11928</td>
 <td></td>
 </tr>
 <tr>
 <td>Indium tin oxide (ITO) coated glass substrates </td>
 <td>Delta Technologies </td>
 <td>CB-40IN-1107</td>
 <td></td>
 </tr>
 <tr>
 <td>DMF hub Teflon-coated fused-silica microcapillaries</td>
 <td>Polymicro Technologies</td>
 <td>TSU100375</td>
 <td></td>
 </tr>
 <tr>
 <td>Perfusion chamber microcapillaries</td>
 <td>Polymicro Technologies</td>
 <td>TSP530700</td>
 <td></td>
 </tr>
 <tr>
 <td>Tubing and microcapillary fittings</td>
 <td>Sandia National Laboratories</td>
 <td>&nbsp;</td>
 <td></td>
 </tr>
 <tr>
 <td>Polycarbonate tubing</td>
 <td>Paradigm Optics</td>
 <td>CTPC100-500-5</td>
 <td></td>
 </tr>
 <tr>
 <td>8-port precision syringe pump equipped with 30 mm (500 &mu;l capacity) syringes</td>
 <td>Hamilton Company</td>
 <td>54848-01</td>
 <td></td>
 </tr>
 <tr>
 <td>Parylene-C vapor deposition instrument</td>
 <td>Specialty Coating Systems</td>
 <td>PDS 2010 Labcoter 2</td>
 <td></td>
 </tr>
 <tr>
 <td>High-voltage function generator</td>
 <td>Trek</td>
 <td>615A-1 615-3</td>
 <td></td>
 </tr>
 <tr>
 <td>MVX10 microscope</td>
 <td>Olympus</td>
 <td></td>
 <td>Optional (facilitates tracking of droplets on DMF hub)</td>
 </tr>
 <tr>
 <td>QIClick digital CCD camera</td>
 <td>QImaging</td>
 <td>QIClick-F-CLR-12</td>
 <td>Optional (facilitates tracking of droplets on DMF hub)</td>
 </tr>
</tbody>
</table>

a versatile automated platform for micro-scale cell stimulation experiments

Study of cells in culture (in vitro analysis) has provided important insight into complex biological systems. Conventional methods and equipment for in vitro analysis are well suited to study of large numbers of cells (&ge;105) in milliliter-scale volumes (&ge;0.1 ml). However, there are many instances in which it is necessary or desirable to scale down culture size to reduce consumption of the cells of interest and/or reagents required for their culture, stimulation, or processing. Unfortunately, conventional approaches do not support precise and reproducible manipulation of micro-scale cultures, and the microfluidics-based automated systems currently available are too complex and specialized for routine use by most laboratories. To address this problem, we have developed a simple and versatile technology platform for automated culture, stimulation, and recovery of small populations of cells (100 - 2,000 cells) in micro-scale volumes (1 - 20 &mu;l). The platform consists of a set of fibronectin-coated microcapillaries ("cell perfusion chambers"), within which micro-scale cultures are established, maintained, and stimulated; a digital microfluidics (DMF) device outfitted with "transfer" microcapillaries ("central hub"), which routes cells and reagents to and from the perfusion chambers; a high-precision syringe pump, which powers transport of materials between the perfusion chambers and the central hub; and an electronic interface that provides control over transport of materials, which is coordinated and automated via pre-determined scripts. As an example, we used the platform to facilitate study of transcriptional responses elicited in immune cells upon challenge with bacteria. Use of the platform enabled us to reduce consumption of cells and reagents, minimize experiment-to-experiment variability, and re-direct hands-on labor. Given the advantages that it confers, as well as its accessibility and versatility, our platform should find use in a wide variety of laboratories and applications, and prove especially useful in facilitating analysis of cells and stimuli that are available in only limited quantities.

As a demonstration of the automated platform, we used it to carry out a study in which small populations of immune cells were grown in micro-scale cultures, challenged with bacteria, and lysed for off-platform analysis of pro-inflammatory responses (Figure 2).
Each of six cell perfusion chambers was seeded with 104 immune cells (P388D.1 murine macrophages) resuspended in 10 &mu;l of growth medium. After an adherence period (37 &deg;C for 2 hr) and a medium exchange,...

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A Versatile Automated Platform for Micro-scale Cell Stimulation Experiments

We have developed an automated cell culture and interrogation platform for micro-scale cell stimulation experiments. The platform offers simple, versatile, and precise control in cultivating and stimulating small populations of cells, and recovering lysates for molecular analyses. The platform is well suited to studies that use precious cells and/or reagents.

Study of cells in culture (in vitro analysis) has provided important insight into complex biological systems. Conventional methods and equipment for in ...

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10.5K Views.  Sandia National Laboratories. Мы разработали автоматизированную культуре клеток и допроса платформы для микро-масштабе экспериментов стимуляции клеток. Платформа предлагает простые, универсальный и точный контроль в выращивании и стимулирования небольших популяциях клеток и восстано?...

Video: A Versatile Automated Platform for Micro-scale Cell Stimulation Experiments

Organotypic Collagen I Assay: A Malleable Platform to Assess Cell Behaviour in a 3-Dimensional Context

Cell migration is fundamental to many aspects of biology, including development, wound healing, the cellular responses of the immune system, and metastasis of tumor cells. Migration has been studied on glass coverslips in order to make cellular dynamics amenable to investigation by light microscopy. However, it has become clear that many aspects of cell migration depend on features of the local environment including its elasticity, protein composition, and pore size, which are not faithfully represented by rigid two dimensional substrates such as glass and plastic 1. Furthermore, interaction with other cell types, including stromal fibroblasts 2 and immune cells 3, has been shown to play a critical role in promoting the invasion of cancer cells. Investigation at the molecular level has increasingly shown that molecular dynamics, including response to drug treatment, of identical cells are significantly different when compared in vitro and in vivo 4. 
Ideally, it would be best to study cell migration in its naturally occurring context in living organisms, however this is not always possible. Intermediate tissue culture systems, such as cell derived matrix, matrigel, organotypic culture (described here) tissue explants, organoids, and xenografts, are therefore important experimental intermediates. These systems approximate certain aspects of an in vivo environment but are more amenable to experimental manipulation such as use of stably transfected cell lines, drug treatment regimes, long term and high-resolution imaging. Such intermediate systems are especially useful as proving grounds to validate probes and establish parameters required to image the dynamic response of cells and fluorescent reporters prior to undertaking imaging in vivo 5. As such, they can serve an important role in reducing the need for experiments on living animals.

A method is described for the preparation of a 3-dimensional matrix consisting of collagen type I and primary human fibroblasts. This organotypic gel serves as a useful substrate to assess invasive cell migration because it mimics basic features of tissue stroma and is amenable to many forms of microscopy.

Cell migration is fundamental to many aspects of biology, including development, wound healing, the cellular responses of the immune system, and ...

GENPLAT: an Automated Platform for Biomass Enzyme Discovery and Cocktail Optimization

The high cost of enzymes for biomass deconstruction is a major impediment to the economic conversion of lignocellulosic feedstocks to liquid transportation fuels such as ethanol. We have developed an integrated high throughput platform, called GENPLAT, for the discovery and development of novel enzymes and enzyme cocktails for the release of sugars from diverse pretreatment/biomass combinations. GENPLAT comprises four elements: individual pure enzymes, statistical design of experiments, robotic pipeting of biomass slurries and enzymes, and automated colorimeteric determination of released Glc and Xyl. Individual enzymes are produced by expression in Pichia pastoris or Trichoderma reesei, or by chromatographic purification from commercial cocktails or from extracts of novel microorganisms. Simplex lattice (fractional factorial) mixture models are designed using commercial Design of Experiment statistical software. Enzyme mixtures of high complexity are constructed using robotic pipeting into a 96-well format. The measurement of released Glc and Xyl is automated using enzyme-linked colorimetric assays. Optimized enzyme mixtures containing as many as 16 components have been tested on a variety of feedstock and pretreatment combinations.
GENPLAT is adaptable to mixtures of pure enzymes, mixtures of commercial products (e.g., Accellerase 1000 and Novozyme 188), extracts of novel microbes, or combinations thereof. To make and test mixtures of &tilde;10 pure enzymes requires less than 100 &mu;g of each protein and fewer than 100 total reactions, when operated at a final total loading of 15 mg protein/g glucan. We use enzymes from several sources. Enzymes can be purified from natural sources such as fungal cultures (e.g., Aspergillus niger, Cochliobolus carbonum, and Galerina marginata), or they can be made by expression of the encoding genes (obtained from the increasing number of microbial genome sequences) in hosts such as E. coli, Pichia pastoris, or a filamentous fungus such as T. reesei. Proteins can also be purified from commercial enzyme cocktails (e.g., Multifect Xylanase, Novozyme 188). An increasing number of pure enzymes, including glycosyl hydrolases, cell wall-active esterases, proteases, and lyases, are available from commercial sources, e.g., Megazyme, Inc. (www.megazyme.com), NZYTech (www.nzytech.com), and PROZOMIX (www.prozomix.com).
Design-Expert software (Stat-Ease, Inc.) is used to create simplex-lattice designs and to analyze responses (in this case, Glc and Xyl release). Mixtures contain 4-20 components, which can vary in proportion between 0 and 100%. Assay points typically include the extreme vertices with a sufficient number of intervening points to generate a valid model. In the terminology of experimental design, most of our studies are "mixture" experiments, meaning that the sum of all components adds to a total fixed protein loading (expressed as mg/g glucan). The number of mixtures in the simplex-lattice depends on both the number of components in the mixture and the degree of polynomial (quadratic or cubic). For example, a 6-component experiment will entail 63 separate reactions with an augmented special cubic model, which can detect three-way interactions, whereas only 23 individual reactions are necessary with an augmented quadratic model. For mixtures containing more than eight components, a quadratic experimental design is more practical, and in our experience such models are usually statistically valid.
All enzyme loadings are expressed as a percentage of the final total loading (which for our experiments is typically 15 mg protein/g glucan). For "core" enzymes, the lower percentage limit is set to 5%. This limit was derived from our experience in which yields of Glc and/or Xyl were very low if any core enzyme was present at 0%. Poor models result from too many samples showing very low Glc or Xyl yields. Setting a lower limit in turn determines an upper limit. That is, for a six-component experiment, if the lower limit for each single component is set to 5%, then the upper limit of each single component will be 75%. The lower limits of all other enzymes considered as "accessory" are set to 0%. "Core" and "accessory" are somewhat arbitrary designations and will differ depending on the substrate, but in our studies the core enzymes for release of Glc from corn stover comprise the following enzymes from T. reesei: CBH1 (also known as Cel7A), CBH2 (Cel6A), EG1(Cel7B), BG (&beta;-glucosidase), EX3 (endo-&beta;1,4-xylanase, GH10), and BX (&beta;-xylosidase).
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GENPLAT (GLBRC Enzyme Platform) is an automated platform for discovery and optimization of enzyme cocktails for biomass degradation. It can be adapted to multiple feedstocks and mixtures of enzymes containing multiple components.

The high cost of enzymes for biomass deconstruction is a major impediment to the economic conversion of lignocellulosic feedstocks to liquid ...

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 ...

A Novel Stretching Platform for Applications in Cell and Tissue Mechanobiology

Tools that allow the application of mechanical forces to cells and tissues or that can quantify the mechanical properties of biological tissues have contributed dramatically to the understanding of basic mechanobiology. These techniques have been extensively used to demonstrate how the onset and progression of various diseases are heavily influenced by mechanical cues. This article presents a multi-functional biaxial stretching (BAXS) platform that can either mechanically stimulate single cells or quantify the mechanical stiffness of tissues. The BAXS platform consists of four voice coil motors that can be controlled independently. Single cells can be cultured on a flexible substrate that can be attached to the motors allowing one to expose the cells to complex, dynamic, and spatially varying strain fields. Conversely, by incorporating a force load cell, one can also quantify the mechanical properties of primary tissues as they are exposed to deformation cycles. In both cases, a proper set of clamps must be designed and mounted to the BAXS platform motors in order to firmly hold the flexible substrate or the tissue of interest. The BAXS platform can be mounted on an inverted microscope to perform simultaneous transmitted light and/or fluorescence imaging to examine the structural or biochemical response of the sample during stretching experiments. This article provides experimental details of the design and usage of the BAXS platform and presents results for single cell and whole tissue studies. The BAXS platform was used to measure the deformation of nuclei in single mouse myoblast cells in response to substrate strain and to measure the stiffness of isolated mouse aortas. The BAXS platform is a versatile tool that can be combined with various optical microscopies in order to provide novel mechanobiological insights at the sub-cellular, cellular and whole tissue levels.

We present in this article a novel stretching platform that can be used to investigate single cell responses to complex anisotropic biaxial mechanical deformation and quantify the mechanical properties of biological tissues.

Tools that allow the application of mechanical forces to cells and tissues or that can quantify the mechanical properties of biological tissues have ...

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 ...

Layered Alginate Constructs: A Platform for Co-culture of Heterogeneous Cell Populations

Many load bearing tissues possess structurally and functionally distinct regions, typically accompanied by different cell phenotypes with differential mechanosensing characteristics. Engineering and analysis of these tissue types remain a challenge. Layered hydrogel constructs provide an opportunity for investigating the interactions among multiple cell populations within single constructs. Alginate hydrogels are both biocompatible and allow for easy isolation of cells after experimentation. Here, we describe a method for the development of small sized dual layered alginate hydrogel discs. This process maintains high cell viability of human mesenchymal stem cells during the formation process and these layered discs can withstand unconfined cyclic compression, commonly used for stimulation of hMSCs undergoing chondrogenesis. These layered constructs can potentially be scaled up to include additional levels, and also be used to segregate cell populations initially after layering. This dual layer alginate hydrogel culture platform can be used for many different applications including engineering and analysis of cells of load bearing tissues and co-cultures of other cell types.

Engineering and analysis of load bearing tissues with heterogeneous cell populations are still a challenge. Here, we describe a method for creating bi-layered alginate hydrogel discs as a platform for co-culture of diverse cell populations within one construct.

Many load bearing tissues possess structurally and functionally distinct regions, typically accompanied by different cell phenotypes with differential ...

A High-throughput Cell Microarray Platform for Correlative Analysis of Cell Differentiation and Traction Forces

Microfabricated cellular microarrays, which consist of contact-printed combinations of biomolecules on an elastic hydrogel surface, provide a tightly controlled, high-throughput engineered system for measuring the impact of arrayed biochemical signals on cell differentiation. Recent efforts using cell microarrays have demonstrated their utility for combinatorial studies in which many microenvironmental factors are presented in parallel. However, these efforts have focused primarily on investigating the effects of biochemical cues on cell responses. Here, we present a cell microarray platform with tunable material properties for evaluating both cell differentiation by immunofluorescence and biomechanical cell&#8211;substrate interactions by traction force microscopy. To do so, we have developed two different formats utilizing polyacrylamide hydrogels of varying Young&#39;s modulus fabricated on either microscope slides or glass-bottom Petri dishes. We provide best practices and troubleshooting for the fabrication of microarrays on these hydrogel substrates, the subsequent cell culture on microarrays, and the acquisition of data. This platform is well-suited for use in investigations of biological processes for which both biochemical (e.g., extracellular matrix composition) and biophysical (e.g., substrate stiffness) cues may play significant, intersecting roles.

Cell differentiation is regulated by a host of microenvironmental factors, including both matrix composition and substrate material properties. We describe here a technique utilizing cell microarrays in conjunction with traction force microscopy to evaluate both cell differentiation and biomechanical cell&#8211;substrate interactions as a function of microenvironmental context.

Microfabricated cellular microarrays, which consist of contact-printed combinations of biomolecules on an elastic hydrogel surface, provide a tightly ...

A Versatile Method of Patterning Proteins and Cells

Substrate and cell patterning techniques are widely used in cell biology to study cell-to-cell and cell-to-substrate interactions. Conventional patterning techniques work well only with simple shapes, small areas and selected bio-materials. This article describes a method to distribute cell suspensions as well as substrate solutions into complex, long, closed (dead-end) polydimethylsiloxane (PDMS) microchannels using negative pressure. This method enables researchers to pattern multiple substrates including fibronectin, collagen, antibodies (Sal-1), poly-D-lysine (PDL), and laminin. Patterning of substrates allows one to indirectly pattern a variety of cells. We have tested C2C12 myoblasts, the PC12 neuronal cell line, embryonic rat cortical neurons, and amphibian retinal neurons. In addition, we demonstrate that this technique can directly pattern fibroblasts in microfluidic channels via brief application of a low vacuum on cell suspensions. The low vacuum does not significantly decrease cell viability as shown by cell viability assays. Modifications are discussed for application of the method to different cell and substrate types. This technique allows researchers to pattern cells and proteins in specific patterns without the need for exotic materials or equipment and can be done in any laboratory with a vacuum.

This report describes a simple, easy to perform technique, using low pressure vacuum, to fill microfluidic channels with cells and substrates for biological research.

Substrate and cell patterning techniques are widely used in cell biology to study cell-to-cell and cell-to-substrate interactions. Conventional patterning ...

DNA Nanotubes as a Versatile Tool to Study Semiflexible Polymers

Mechanical properties of complex, polymer-based soft matter, such as cells or biopolymer networks, can be understood in neither the classical frame of flexible polymers nor of rigid rods. Underlying filaments remain outstretched due to their non-vanishing backbone stiffness, which is quantified via the persistence length (lp), but they are also subject to strong thermal fluctuations. Their finite bending stiffness leads to unique, non-trivial collective mechanics of bulk networks, enabling the formation of stable scaffolds at low volume fractions while providing large mesh sizes. This underlying principle is prevalent in nature (e.g., in cells or tissues), minimizing the high molecular content and thereby facilitating diffusive or active transport. Due to their biological implications and potential technological applications in biocompatible hydrogels, semiflexible polymers have been subject to considerable study. However, comprehensible investigations remained challenging since they relied on natural polymers, such as actin filaments, which are not freely tunable. Despite these limitations and due to the lack of synthetic, mechanically tunable, and semiflexible polymers, actin filaments were established as the common model system. A major limitation is that the central quantity lp cannot be freely tuned to study its impact on macroscopic bulk structures. This limitation was resolved by employing structurally programmable DNA nanotubes, enabling controlled alteration of the filament stiffness. They are formed through tile-based designs, where a discrete set of partially complementary strands hybridize in a ring structure with a discrete circumference. These rings feature sticky ends, enabling the effective polymerization into filaments several microns in length, and display similar polymerization kinetics as natural biopolymers. Due to their programmable mechanics, these tubes are versatile, novel tools to study the impact of lp on the single-molecule as well as the bulk scale. In contrast to actin filaments, they remain stable over weeks, without notable degeneration, and their handling is comparably straightforward.

Semiflexible polymers display unique mechanical properties that are extensively applied by living systems. However, systematic studies on biopolymers are limited since properties such as polymer rigidity are inaccessible. This manuscript describes how this limitation is circumvented by programmable DNA nanotubes, enabling experimental studies on the impact of filament rigidity.

Mechanical properties of complex, polymer-based soft matter, such as cells or biopolymer networks, can be understood in neither the classical frame of ...

A Multilayer Microfluidic Platform for the Conduction of Prolonged Cell-Free Gene Expression

The limitations of cell-based synthetic biology are becoming increasingly apparent as researchers aim to develop larger and more complex synthetic genetic regulatory circuits. The analysis of synthetic genetic regulatory networks in vivo is time consuming and suffers from a lack of environmental control, with exogenous synthetic components interacting with host processes resulting in undesired behavior. To overcome these issues, cell-free characterization of novel circuitry is becoming more prevalent. In vitro transcription and translation (IVTT) mixtures allow the regulation of the experimental environment and can be optimized for each unique system. The protocols presented here detail the fabrication of a multilayer microfluidic device that can be utilized to sustain IVTT reactions for prolonged durations. In contrast to batch reactions, where resources are depleted over time and (by-) products accumulate, the use of microfluidic devices allows the replenishment of resources as well as the removal of reaction products. In this manner, the cellular environment is emulated by maintaining an out-of-equilibrium environment in which the dynamic behavior of gene circuits can be investigated over extended periods of time. To fully exploit the multilayer microfluidic device, hardware and software have been integrated to automate the IVTT reactions. By combining IVTT reactions with the microfluidic platform presented here, it becomes possible to comprehensively analyze complex network behaviors, furthering our understanding of the mechanisms that regulate cellular processes.

The fabrication process of a PDMS-based, multilayer, microfluidic device that allows in vitro transcription and translation (IVTT) reactions to be performed over prolonged periods is described. Furthermore, a comprehensive overview of the hardware and software required to automate and maintain these reactions for prolonged durations is provided.

The limitations of cell-based synthetic biology are becoming increasingly apparent as researchers aim to develop larger and more complex synthetic genetic ...