This project was funded by the Illinois Department of Public Health and the National Science Foundation (DBI-0852416).

1. Device function
<ol>
 <li>The hypoxic insert device contains 6-pillars that nest into a standard 6-well plate. Gas flows into the pillar, across a microfluidic network at the base of the pillar, and flows back out of the device. At the base of the pillar forming the bottom wall of the microchannel is a 100 &mu;m thick gas-permeable PDMS membrane that permits oxygen diffusion between the gas microchannel and the culture media. Thus, the device operates by establishing a concentration gradient that drives the media oxygen concentration towards a desired value.</li>
 <li>The device offers a number of advantages over the hypoxic chamber and other cell oxygenation systems: 1) minimizes the oxygen diffusion path permitting faster temporal control, 2) enhances the spatial control over oxygenation, as dependent on the pillar microchannel design, 3) possesses a smaller lab footprint and higher throughput to increase experimental efficiency, and 4) adapts to common cell culture tools (e.g. multiwell plate) without the requirement for specialized knowledge and equipment.</li>
</ol>
2. Device fabrication
<ol>
 <li>First the pillar array is replica molded with PDMS in a previously machined Delran mold.</li>
 <li>Next the gas microchannel at the bottom of each pillar is fabricated using standard SU-8 photolithography and PDMS replica molding.</li>
 <li>The gas-permeable membrane is fabricated by defined spinning of PDMS on a silicon wafer to achieve the desired thickness. In this example we use a 100 &mu;m thick membrane which was fabricated by spinning 500 rpm for 10 seconds and then 900 rpm for 30 seconds.</li>
 <li>All components are bonded together after oxygen plasma treatment with a handheld plasma device (Model BD-20, Electro-Technic Products).</li>
</ol>
3. Device setup 
<ol>
 <li>Gently insert device into the plate, making sure to avoid bubbles. Angling the device during insertion helps to expel bubbles out one side. For actual cell-based experiments, this step must be done inside a sterile laminar flow hood. The chances for contamination are greatly reduced once the device is inserted, so the assembly can be carried over to the incubator in a non-sterile environment.</li>
 <li>Connect tubing from the source gas tank to the inlet and outlet ports of the device. For cell-based experiments, the culture incubator should have a hole to permit tubing entry and the tubing should be connected after carrying the device and placing it into the incubator. Be careful to avoid putting excess pressure on the device, which could deform the PDMS enough to crush the underlying cells.</li>
 <li>Make sure the precision flow regulator is closed before opening the tank top regulator to avoid overflowing the device. Start the flow of gas. Gently open precision flow regulator to desired flow rate (50-100 mL/min). Watch the value closely over the next hour and make adjustments as needed, since the pressure drop will change while the system equilibrates, altering the flow rate. To avoid formation of bubbles in the media, reduce the flow rate to between 10-20 mL/min after an initial 15 minute equilibration duration with the higher flow rate.</li>
 <li>After desired experimental duration, stop gas flow, remove the plate, and process cells accordingly (e.g. lyse, stain, count, etc.).
</li>
</ol>
4. Device Validation
<ol>
 <li>Calibration
 <ol>
 <li>Select the number and location of the positions on the fluorescent oxygen 
 (FOXY) sensor slide (depending on the validation requirements) that will be used for oxygen measurement. The slide contains a fluorescent ruthenium dye coating that is quenched by oxygen.</li>
 <li>Expose slide directly to 0, 10, and 21% oxygen from gas tank and capture images after 5 minutes for proper equilibration.</li>
 <li>Export the mean image intensity for each position and plot oxygenation concentration as dependent on fluorescent intensity.</li>
 <li>Generate calibration curve by fitting linear curves to the 0-10% line and 10-21% line.</li>
</ol>
 </li>
 <li>Heterogeneity
 <ol>
 <li>Establish multiple points spanning the width of the channel at a defined interval (e.g. every 1 mm). Note that there may be overlap of images depending on the spacing.</li>
 <li>Measure the oxygen concentration at the well surface through the device</li>
 </ol>
 </li>
 <li>Equilibration
 <ol>
 <li>Choose three points on the FOXY slide in which to measure the oxygen concentration.</li>
 <li>Immediately after capturing a first image at ambient oxygen levels, open the precision flow regulator to start gas flow into the device. Capture images at an appropriate interval to assess the duration and extent of oxygen concentration equilibration (e.g. every 10 seconds for 30 min).</li>
 </ol>
 </li>
</ol>
5. Applications
<ol>
 <li>Wound Healing
 <ol>
 <li>A day before the experiment, soak the sterilized PDMS insert in serum-free medium to reduce the inhibit gas bubble formation in the well. </li>
 <li>Culture cells to 100% confluency in a 6-well plate. </li>
 <li>Create straight scratches in the monolayers with a p200 pipette tip to simulate wounds.</li>
 <li>Aspirate the cell media, rinse with 5 mL media, and aspirate again. It is important to not disturb the monolayer of cells. </li>
 <li>Refill the wells with 4 mL serum-free medium to reduce cell proliferation.</li>
 <li>Place insert into the wells and connect each well to a corresponding oxygen concentration. </li>
 <li>Place 6-well plate with insert on heated stage at 37&deg;C.</li>
 <li>Capture time lapse images of the cells at the desired interval and total duration. MATLAB's T_Scratch, a wound measuring algorithm, can be used to analyze the unhealed surface area.</li>
	 </ol>
</li>
</ol>
6. Representative Results
<ol>
 <li>Device validation 
The hypoxic insert device exhibits great improvements over the hypoxic chamber in terms of oxygen equilibration time and extent, requiring less than 2 minutes to stabilize to 0.5% oxygen. The device membrane-well bottom gap size was the critical factor in determining equilibration efficiency, with larger gap sizes requiring more time to reach steady-state oxygen concentration values. The device also permits a great deal of control over the spatial oxygenation in a single well, allowing the formation of multiple conditions, and even generating a cyclic oxygen profile across the surface of the well.</li>
 <li>Wound healing 
Cell monolayers were exposed to 10% or 21% oxygen and the surface area of the wound was analyzed over time. Scratches exposed to 21% oxygen closed the slowest and 10% the fastest. Figure 1 shows images of a scratch over the course of 17 hours. The graphs in Figure 5 show the percent of open wound area for both oxygen concentrations for the duration of the experiment.</li>
</ol>
<a href="https://www.jove.com/files/ftp_upload/1695/1695fig1.jpg"><img src="/files/ftp_upload/1695/1695fig1tmb.jpg" alt="Figure 1" border="0" /></a> 
Figure 1. Schematic and diagrams illustrating device features. The oxygen insert device is fabricated by conventional photolithography (microfluidic network), replica molding (microfluidic network and insert scaffold), and defined spinning of PDMS (gas-permeable membrane). A) The oxygen device nested into a 6-well plate. B) Examples of 24 and 96-well pillar arrays. C) A cross-sectional schematic of a pillar. Oxygen flows into the device through the inlet and travels across a microfluidic network at the bottom of the pillar. Oxygen can freely diffuse across the gas-permeable PDMS membrane at the bottom of the pillar and dissolve into the culture media. D) A microscope image showing the various features of a single-channel pillar from above, with bonded glass posts for the equilibration studies.
<a href="https://www.jove.com/files/ftp_upload/1695/1695fig2.jpg"><img src="/files/ftp_upload/1695/1695fig2tmb.jpg" alt="Figure 2" border="0" /></a> 
Figure 2. Validation of the device with oxygen sensors. Oxygen tension within each well was characterized using a planar ruthenium oxygen sensor. All oxygen mixtures contained balanced nitrogen and 5% CO2 for media buffering. A) Plot illustrating the effect of post height, and thus oxygen diffusion distance between the membrane and cells, on the equilibration time and effectiveness. Heights were established by cut-glass posts bound to the bottom of the device. All three post sizes yield equilibration times much improved over the hypoxic chamber. Note that time is on a log scale. B) Plot depicting the rapid oxygen equilibration response time of the 0.2 mm gap device. C) Multi-position linescans were also taken across the well under the microchannel to ensure homogeneity of the oxygen concentration introduced by the device. Graph depicts the oxygen concentration measured after infusing 0%, 10%, and 21% oxygen for 10 min. D) Device effectively maintains 10% oxygen over 5 days.
<a href="https://www.jove.com/files/ftp_upload/1695/1695fig3.jpg"><img src="/files/ftp_upload/1695/1695fig3tmb.jpg" alt="Figure 3" border="0" /></a> 
Figure 3. Experimentation with more complex oxygen microchannel designs. A) Dual-condition microchannel setup yields a stable 0% and 21% oxygen profile over 14 days. B) An interdigitated and winding pattern of 500 &mu;m width microchannels extending across the pillar results in a cyclic oxygen profile. Note that the data only depicts one representative trial as microchannel alignment was difficult.
<a href="https://www.jove.com/files/ftp_upload/1695/1695fig5.jpg"><img src="/files/ftp_upload/1695/1695fig5tmb.jpg" alt="Figure 4" border="0" /></a> 
Figure 4. Timelapse images of wound closure 0, 7, and 17 hours after initial scratch. Cells were delivered 21% oxygen throughout duration of experiment.
<a href="https://www.jove.com/files/ftp_upload/1695/1695fig6.jpg"><img src="/files/ftp_upload/1695/1695fig6tmb.jpg" alt="Figure 5" border="0" /></a> 
Figure 5. Effect of oxygen concentration on wound healing rate in a scratch assay.

The device is fabricated by standard SU-8 photolithography, replica molding, and defined spinning and made entirely of polydimethylsiloxane. Gas is introduced into the device to establish a concentration gradient between the pillar microchannel and the culture media, driving the system towards a desired equilibrium oxygen concentration. The device has been shown to effectively modulate the temporal and spatial oxygenation inside a well, as well as modulate cellular behavior appropriately. The spatial patterning of oxygenation is defined by the microchannel at the base of the pillar, so a variety of designs could be implemented in crafting the photomask. Additionally, infusion of the desired gas into the gas-phase of the well is expected to improve equilibration time and extent of hypoxia. A microfluidic mixing network could be adapted to the device to provide a means to produce novel gas mixtures from only a few stock gas tanks. Finally, a mechanism for media exchange would eliminate the need for removal of the device from the multiwell plate, of which the cells may respond.
The device has applications in any in vitro or ex vivo experiment requiring control over oxygen concentration. As oxygen is an important physiological variable affecting a vast majority of signaling pathways, the areas of research that would benefit is limited by the creativity of the researcher. Some fields that would benefit from the enhanced temporal control of oxygen concentration include cancer metastasis, sleep apnea, and cardiac ischemia reperfusion injury, among many others. For example, intermittent hypoxia has been correlated with more invasive cancers, upregulating a number of metastastis-associated genes relative to continuous hypoxia and normoxia. Spatial control is also important, as oxygen gradients are critical in development, liver zonation, drug toxicity, and the stem cell niche. The device presented in this article will benefit a number of areas of research by providing a system with a smaller lab footprint, relatively simple operational requirements, and far greater control over oxygen exposure to cells.

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<td>PDMS-Sylgard 184</td>
<td>&nbsp;</td>
<td>Dow Corning</td>
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<td>Planar FOXY sensor</td>
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<td>Custom mixes</td>
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fabrication and operation of an oxygen insert for adherent cellular cultures 

Oxygen is a key modulator of many cellular pathways, but current devices permitting in vitro oxygen modulation fail to meet the needs of biomedical research. The hypoxic chamber offers a simple system to control oxygenation in standard culture vessels, but lacks precise temporal and spatial control over the oxygen concentration at the cell surface, preventing its application in studying a variety of physiological phenomena. Other systems have improved upon the hypoxic chamber, but require specialized knowledge and equipment for their operation, making them intimidating for the average researcher. A microfabricated insert for multiwell plates has been developed to more effectively control the temporal and spatial oxygen concentration to better model physiological phenomena found in vivo. The platform consists of a polydimethylsiloxane insert that nests into a standard multiwell plate and serves as a passive microfluidic gas network with a gas-permeable membrane aimed to modulate oxygen delivery to adherent cells. The device is simple to use and is connected to gas cylinders that provide the pressure to introduce the desired oxygen concentration into the platform. Fabrication involves a combination of standard SU-8 photolithography, replica molding, and defined PDMS spinning on a silicon wafer. The components of the device are bonded after surface treatment using a hand-held plasma system. Validation is accomplished with a planar fluorescent oxygen sensor. Equilibration time is on the order of minutes and a wide variety of oxygen profiles can be attained based on the device design, such as the cyclic profile achieved in this study, and even oxygen gradients to mimic those found in vivo. The device can be sterilized for cell culture using common methods without loss of function. The device's applicability to studying the in vitro wound healing response will be demonstrated.

Watch this Scientific Journal Video about Fabrication and Operation of an Oxygen Insert for Adherent Cellular Cultures at JoVE.com

Fabrication and Operation of an Oxygen Insert for Adherent Cellular Cultures

Fabrication and validation of an add-on platform that offers enhanced control over the spatial and temporal oxygenation in a 6-well plate. The device is adaptable to a number of culture systems and can be used to investigate the effects of oxygen on wound healing.

Fabrication and Operation of an Oxygen Insert for Adherent Cellular Cultures 

Oxygen is a key modulator of many cellular pathways, but current devices permitting in vitro oxygen modulation fail to meet the needs of biomedical ...

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9.7K Views.  University of Illinois. Fabrication and validation of an add-on platform that offers enhanced control over the spatial and temporal oxygenation in a 6-well plate. The device is adaptable to a number of culture systems and can be used to investigate the effects of oxygen on wound healing. 

Video: Fabrication and Operation of an Oxygen Insert for Adherent Cellular Cultures

Large Insert Environmental Genomic Library Production

The vast majority of microbes in nature currently remain inaccessible to traditional cultivation methods. Over the past decade, culture-independent environmental genomic (i.e. metagenomic) approaches have emerged, enabling researchers to bridge this cultivation gap by capturing the genetic content of indigenous microbial communities directly from the environment. To this end, genomic DNA libraries are constructed using standard albeit artful laboratory cloning techniques. Here we describe the construction of a large insert environmental genomic fosmid library with DNA derived from the vertical depth continuum of a seasonally hypoxic fjord. This protocol is directly linked to a series of connected protocols including coastal marine water sampling [1], large volume filtration of microbial biomass [2] and a DNA extraction and purification protocol [3]. At the outset, high quality genomic DNA is end-repaired with the creation of 5 -phosphorylated blunt ends. End-repaired DNA is subjected to pulsed-field gel electrophoresis (PFGE) for size selection and gel extraction is performed to recover DNA fragments between 30 and 60 thousand base pairs (Kb) in length. Size selected DNA is purified away from the PFGE gel matrix and ligated to the phosphatase-treated blunt-end fosmid CopyControl vector pCC1 (EPICENTRE <a href="http://www.epibio.com/item.asp?ID=385">http://www.epibio.com/item.asp?ID=385</a>). Linear concatemers of pCC1 and insert DNA are subsequently headfull packaged into phage particles by lambda terminase, with subsequent infection of phage-resistant E. coli cells. Successfully transduced clones are recovered on LB agar plates under antibiotic selection and archived in 384-well plate format using an automated colony picking robot (Qpix2, GENETIX). The current protocol draws from various sources including the CopyControl Fosmid Library Production Kit from EPICENTRE and the published works of multiple research groups [4-7]. Each step is presented with best practice in mind. Whenever possible we highlight subtleties in execution to improve overall quality and efficiency of library production. The whole process of fosmid library production and automated colony picking takes at least 7-10 days as there are many incubation steps included. However, there are several stopping points possible which are mentioned within the protocol.

Construction of a fosmid library with environmental genomic DNA isolated from the vertical depth continuum of a seasonally hypoxic fjord is described. The resulting clone library is picked into 384-well plates and archived for downstream sequencing and functional screening by the application of an automated colony picking system.

The vast majority of microbes in nature currently remain inaccessible to traditional cultivation methods. Over the past decade, culture-independent ...

The Microfluidic Probe: Operation and Use for Localized Surface Processing

Microfluidic devices allow assays to be performed using minute amounts of sample and have recently been used to control the microenvironment of cells. Microfluidics is commonly associated with closed microchannels which limit their use to samples that can be introduced, and cultured in the case of cells, within a confined volume. On the other hand, micropipetting system have been used to locally perfuse cells and surfaces, notably using push-pull setups where one pipette acts as source and the other one as sink, but the confinement of the flow is difficult in three dimensions. Furthermore, pipettes are fragile and difficult to position and hence are used in static configuration only. 
The microfluidic probe (MFP) circumvents the constraints imposed by the construction of closed microfluidic channels and instead of enclosing the sample into the microfluidic system, the microfluidic flow can be directly delivered onto the sample, and scanned across the sample, using the MFP. . The injection and aspiration openings are located within a few tens of micrometers of one another so that a microjet injected into the gap is confined by the hydrodynamic forces of the surrounding liquid and entirely aspirated back into the other opening. The microjet can be flushed across the substrate surface and provides a precise tool for localized deposition/delivery of reagents which can be used over large areas by scanning the probe across the surface. 
In this video we present the microfluidic probe1 (MFP). We explain in detail how to assemble the MFP, mount it atop an inverted microscope, and align it relative to the substrate surface, and finally show how to use it to process a substrate surface immersed in a buffer.

In this video we present the microfluidic probe1 (MFP). We explain in detail how to assemble the MFP, mount it atop an inverted microscope, and align it relative to the substrate surface, and finally show how to use it to process a substrate surface immersed in a buffer solution.

Microfluidic devices allow assays to be performed using minute amounts of sample and have recently been used to control the microenvironment of cells. ...

In situ Subcellular Fractionation of Adherent and Non-adherent Mammalian Cells

Protein function is intimately coupled to protein localization. Although some proteins are restricted to a specific location or subcellular compartment, many proteins are present as a freely diffusing population in free exchange with a sub-population that is tightly associated with a particular subcellular domain or structure. In situ subcellular fractionation allows the visualization of protein compartmentalization and can also reveal protein sub-populations that localize to specific structures. For example, removal of soluble cytoplasmic proteins and loosely held nuclear proteins can reveal the stable association of some transcription factors with chromatin. Subsequent digestion of DNA can in some cases reveal association with the network of proteins and RNAs that is collectively termed the nuclear scaffold or nuclear matrix. 
Here we describe the steps required during the in situ fractionation of adherent and non-adherent mammalian cells on microscope coverslips. Protein visualization can be achieved using specific antibodies or fluorescent fusion proteins and fluorescence microscopy. Antibodies and/or fluorescent dyes that act as markers for specific compartments or structures allow protein localization to be mapped in detail. In situ fractionation can also be combined with western blotting to compare the amounts of protein present in each fraction. This simple biochemical approach can reveal associations that would otherwise remain undetected.

In situ Subcellular Fractionation of Adherent and Non-adherent Mammalian Cells

In situ subcellular fractionation of mammalian cells on microscope coverslips allows the visualisation of protein localisation.

Protein function is intimately coupled to protein localization. Although some proteins are restricted to a specific location or subcellular compartment, ...

Clonogenic Assay: Adherent Cells

The clonogenic (or colony forming) assay has been established for more than 50 years; the original paper describing the technique was published in 19561. Apart from documenting the method, the initial landmark study generated the first radiation-dose response curve for X-ray irradiated mammalian (HeLa) cells in culture1. Basically, the clonogenic assay enables an assessment of the differences in reproductive viability (capacity of cells to produce progeny; i.e. a single cell to form a colony of 50 or more cells) between control untreated cells and cells that have undergone various treatments such as exposure to ionising radiation, various chemical compounds (e.g. cytotoxic agents) or in other cases genetic manipulation. The assay has become the most widely accepted technique in radiation biology and has been widely used for evaluating the radiation sensitivity of different cell lines. Further, the clonogenic assay is commonly used for monitoring the efficacy of radiation modifying compounds and for determining the effects of cytotoxic agents and other anti-cancer therapeutics on colony forming ability, in different cell lines. A typical clonogenic survival experiment using adherent cells lines involves three distinct components, 1) treatment of the cell monolayer in tissue culture flasks, 2) preparation of single cell suspensions and plating an appropriate number of cells in petri dishes and 3) fixing and staining colonies following a relevant incubation period, which could range from 1-3 weeks, depending on the cell line. Here we demonstrate the general procedure for performing the clonogenic assay with adherent cell lines with the use of an immortalized human keratinocyte cell line (FEP-1811)2. Also, our aims are to describe common features of clonogenic assays including calculation of the plating efficiency and survival fractions after exposure of cells to radiation, and to exemplify modification of radiation-response with the use of a natural antioxidant formulation.

The applicability of the clonogenic assay for evaluating reproductive viability has been established for more than 50 years. Here we demonstrate the general procedure for performing the clonogenic assay with adherent cells.

The clonogenic (or colony forming) assay has been established for more than 50 years; the original paper describing the technique was published in 19561. ...

An Analytical Tool that Quantifies Cellular Morphology Changes from Three-dimensional Fluorescence Images

The most common software analysis tools available for measuring fluorescence images are for two-dimensional (2D) data that rely on manual settings for inclusion and exclusion of data points, and computer-aided pattern recognition to support the interpretation and findings of the analysis. It has become increasingly important to be able to measure fluorescence images constructed from three-dimensional (3D) datasets in order to be able to capture the complexity of cellular dynamics and understand the basis of cellular plasticity within biological systems. Sophisticated microscopy instruments have permitted the visualization of 3D fluorescence images through the acquisition of multispectral fluorescence images and powerful analytical software that reconstructs the images from confocal stacks that then provide a 3D representation of the collected 2D images. Advanced design-based stereology methods have progressed from the approximation and assumptions of the original model-based stereology1 even in complex tissue sections2. Despite these scientific advances in microscopy, a need remains for an automated analytic method that fully exploits the intrinsic 3D data to allow for the analysis and quantification of the complex changes in cell morphology, protein localization and receptor trafficking. 
Current techniques available to quantify fluorescence images include Meta-Morph (Molecular Devices, Sunnyvale, CA) and Image J (NIH) which provide manual analysis. Imaris (Andor Technology, Belfast, Northern Ireland) software provides the feature MeasurementPro, which allows the manual creation of measurement points that can be placed in a volume image or drawn on a series of 2D slices to create a 3D object. This method is useful for single-click point measurements to measure a line distance between two objects or to create a polygon that encloses a region of interest, but it is difficult to apply to complex cellular network structures. Filament Tracer (Andor) allows automatic detection of the 3D neuronal filament-like however, this module has been developed to measure defined structures such as neurons, which are comprised of dendrites, axons and spines (tree-like structure). This module has been ingeniously utilized to make morphological measurements to non-neuronal cells3, however, the output data provide information of an extended cellular network by using a software that depends on a defined cell shape rather than being an amorphous-shaped cellular model. To overcome the issue of analyzing amorphous-shaped cells and making the software more suitable to a biological application, Imaris developed Imaris Cell. This was a scientific project with the Eidgen&ouml;ssische Technische Hochschule, which has been developed to calculate the relationship between cells and organelles. While the software enables the detection of biological constraints, by forcing one nucleus per cell and using cell membranes to segment cells, it cannot be utilized to analyze fluorescence data that are not continuous because ideally it builds cell surface without void spaces. To our knowledge, at present no user-modifiable automated approach that provides morphometric information from 3D fluorescence images has been developed that achieves cellular spatial information of an undefined shape (Figure 1). 
We have developed an analytical platform using the Imaris core software module and Imaris XT interfaced to MATLAB (Mat Works, Inc.). These tools allow the 3D measurement of cells without a pre-defined shape and with inconsistent fluorescence network components. Furthermore, this method will allow researchers who have extended expertise in biological systems, but not familiarity to computer applications, to perform quantification of morphological changes in cell dynamics.

We developed a software platform that utilizes Imaris Neuroscience, ImarisXT and MATLAB to measure the changes in morphology of an undefined shape taken from three-dimensional confocal fluorescence of single cells. This novel approach can be used to quantify changes in cell shape following receptor activation and therefore represents a possible additional tool for drug discovery.

The most common software analysis tools available for measuring fluorescence images are for two-dimensional (2D) data that rely on manual settings for ...

An Explant Assay for Assessing Cellular Behavior of the Cranial Mesenchyme

The central nervous system is derived from the neural plate that undergoes a series of complex morphogenetic movements resulting in formation of the neural tube in a process known as neurulation. During neurulation, morphogenesis of the mesenchyme that underlies the neural plate is believed to drive neural fold elevation. The cranial mesenchyme is comprised of the paraxial mesoderm and neural crest cells. The cells of the cranial mesenchyme form a pourous meshwork composed of stellate shaped cells and intermingling extracellular matrix (ECM) strands that support the neural folds. During neurulation, the cranial mesenchyme undergoes stereotypical rearrangements resulting in its expansion and these movements are believed to provide a driving force for neural fold elevation. However, the pathways and cellular behaviors that drive cranial mesenchyme morphogenesis remain poorly studied. Interactions between the ECM and the cells of the cranial mesenchyme underly these cell behaviors. Here we describe a simple ex vivo explant assay devised to characterize the behaviors of these cells. This assay is amendable to pharmacological manipulations to dissect the signaling pathways involved and live imaging analyses to further characterize the behavior of these cells. We present a representative experiment demonstrating the utility of this assay in characterizing the migratory properties of the cranial mesenchyme on a variety of ECM components.

The cranial mesenchyme undergoes dramatic morphogenic movements that likely provides a driving force for elevation of the neural folds1,2. Here we describe a simple ex vivo explant assay to characterize the cellular behaviors of the cranial mesenchyme during neurulation. This assay has numerous applications including being amenable to pharmacological manipulations and live imaging analyses.

The central nervous system is derived from the neural plate that undergoes a series of complex morphogenetic movements resulting in formation of the ...

Detecting, Visualizing and Quantitating the Generation of Reactive Oxygen Species in an Amoeba Model System

Reactive oxygen species (ROS) comprise a range of reactive and short-lived, oxygen-containing molecules, which are dynamically interconverted or eliminated either catalytically or spontaneously. Due to the short life spans of most ROS and the diversity of their sources and subcellular localizations, a complete picture can be obtained only by careful measurements using a combination of protocols. Here, we present a set of three different protocols using OxyBurst Green (OBG)-coated beads, or dihydroethidium (DHE) and Amplex UltraRed (AUR), to monitor qualitatively and quantitatively various ROS in professional phagocytes such as Dictyostelium. We optimised the beads coating procedures and used OBG-coated beads and live microscopy to dynamically visualize intraphagosomal ROS generation at the single cell level. We identified lipopolysaccharide (LPS) from E. coli as a potent stimulator for ROS generation in Dictyostelium. In addition, we developed real time, medium-throughput assays using DHE and AUR to quantitatively measure intracellular superoxide and extracellular H2O2 production, respectively.

We adapted a set of protocols for the measurement of reactive oxygen species (ROS) that can be applied in various amoeba and mammalian cellular models for qualitative and quantitative studies.

Reactive oxygen species (ROS) comprise a range of reactive and short-lived, oxygen-containing molecules, which are dynamically interconverted or ...

Development of an Insert Co-culture System of Two Cellular Types in the Absence of Cell-Cell Contact

The role of secreted soluble factors in the modification of cellular responses is a recurrent theme in the study of all tissues and systems. In an attempt to make straightforward the very complex relationships between the several cellular subtypes that compose multicellular organisms, in vitro techniques have been developed to help researchers acquire a detailed understanding of single cell populations. One of these techniques uses inserts with a permeable membrane allowing secreted soluble factors to diffuse. Thus, a population of cells grown in inserts can be co-cultured in a well or dish containing a different cell type for evaluating cellular changes following paracrine signaling in the absence of cell-cell contact. Such insert co-culture systems offer various advantages over other co-culture techniques, namely bidirectional signaling, conserved cell polarity and population-specific detection of cellular changes. In addition to being utilized in the field of inflammation, cancer, angiogenesis and differentiation, these co-culture systems are of prime importance in the study of the intricate relationships that exist between the different cellular subtypes present in the central nervous system, particularly in the context of neuroinflammation. This article offers general methodological guidelines in order to set up an experiment in order to evaluating cellular changes mediated by secreted soluble factors using an insert co-culture system. Moreover, a specific protocol to measure the neuroinflammatory effects of cytokines secreted by lipopolysaccharide-activated N9 microglia on neuronal PC12 cells will be detailed, offering a concrete understanding of insert co-culture methodology.

In multicellular organisms, secreted soluble factors elicit responses from different cell types as a result of paracrine signaling. Insert co-culture systems offer a simple way to assess the changes mediated by secreted soluble factors in the absence of cell-cell contact.

The role of secreted soluble factors in the modification of cellular responses is a recurrent theme in the study of all tissues and systems. In an attempt ...

An In Vitro Model for Studying Cellular Transformation by Kaposi Sarcoma Herpesvirus

Kaposi sarcoma (KS) is an unusual tumor composed of proliferating spindle cells that is initiated by infection of endothelial cells (EC) with KSHV, and develops most often in the setting of immunosuppression. Despite decades of research, optimal treatment of KS remains poorly defined and clinical outcomes are especially unfavorable in resource-limited settings. KS lesions are driven by pathological angiogenesis, chronic inflammation, and oncogenesis, and various in vitro cell culture models have been developed to study these processes. KS arises from KSHV-infected cells of endothelial origin, so EC-lineage cells provide the most appropriate in vitro surrogates of the spindle cell precursor. However, because EC have a limited in vitro lifespan, and as the oncogenic mechanisms employed by KSHV are less efficient than those of other tumorigenic viruses, it has been difficult to assess the processes of transformation in primary or telomerase-immortalized EC. Therefore, a novel EC-based culture model was developed that readily supports transformation following infection with KSHV. Ectopic expression of the E6 and E7 genes of human papillomavirus type 16 allows for extended culture of age- and passage-matched mock- and KSHV-infected EC and supports the development of a truly transformed (i.e., tumorigenic) phenotype in infected cell cultures. This tractable and highly reproducible model of KS has facilitated the discovery of several essential signaling pathways with high potential for translation into clinical settings.

An In Vitro Model for Studying Cellular Transformation by Kaposi Sarcoma Herpesvirus

Kaposi sarcoma (KS) is a tumor induced by infection with the oncogenic virus human herpesvirus-8/KS herpesvirus (HHV-8/KSHV). The endothelial cell culture model described here is uniquely suited for studying the mechanisms by which KSHV transforms host cells.

Kaposi sarcoma (KS) is an unusual tumor composed of proliferating spindle cells that is initiated by infection of endothelial cells (EC) with KSHV, and ...

Lens-free Video Microscopy for the Dynamic and Quantitative Analysis of Adherent Cell Culture

Here, we demonstrate that lens-free video microscopy enables us to simultaneously capture the kinetics of thousands of cells directly inside the incubator and that it is possible to monitor and quantify single cells along several cell cycles. We describe the full protocol used to monitor and quantify a HeLa cell culture for 2.7 days. First, cell culture acquisition is performed with a lens-free video microscope, and then the data is analyzed following a four-step process: multi-wavelength holographic reconstruction, cell-tracking, cell segmentation and cell division detection algorithms. As a result, we show that it is possible to gather a dataset featuring more than 10,000 cell cycle tracks and more than 2 x 106 cell morphological measurements.

Lens-free video microscopy enables us to monitor cell cultures directly inside the incubator. Here we describe the full protocol used to acquire and analyze a 2.7 day long acquisition of cultured HeLa cells, leading to a dataset of 2.2 x 106 measurements of individual cell morphology and 10584 cell cycle tracks.

Here, we demonstrate that lens-free video microscopy enables us to simultaneously capture the kinetics of thousands of cells directly inside the incubator ...