This work is supported by R01-HD058629 to I.E.Z.

1. Preparation of ECM Coated Culture Dishes or Coverslips
<ol>
	<li>Prepare ECM stock solution by dissolving ECM substrate in PBS (Dulbecco's Phosphate-Buffered Saline, Invitrogen, #14040-133). Filter using a 0.2 mm syringe filter (VWR, #28145-495) and freeze in aliquots at -80 &deg;C.</li>
	<li>Dilute ECM stock solution in PBS to desired concentration. For MaxGel, dilute in DMEM (Dulbecco's Modified Eagle Medium, Invitrogen, #11995-065).</li>
	<li>If immunofluorescence will be performed, sterilize 12 mm circular glass coverslips (VWR, #89015-724) by dipping in ethanol and allow to air dry. This should be done in the sterile laminar hood. If immunofluorescence will not be performed skip to step 1.5.</li>
	<li>Place sterilized coverslips in each well of a 24 well plate (Corning, #3526).</li>
	<li>Add 0.5 ml of diluted substrate solution to each well of 24 well plate and incubate at room temperature for 3 hr.</li>
	<li>Aspirate off the ECM solution and wash three times in PBS.</li>
	<li>Aspirate off PBS and use immediately or wrapped in parafilm and stored at 4 &deg;C for up to 2 weeks.</li>
</ol>
2. Preparation of Dissection Tools
Sylgard plates with added charcoal are prepared per the manufacture's protocol. The resistant black background of these plates provides support for pinning down the embryo and the color offers good contrast which facilitates visualizing the translucent embryo.
<ol>
	<li>To prepare plates weigh directly into a tri-pour beaker on a balance 150 g Part A liquid 15 g Part B and 1 g charcoal powder (World Precision Instruments, #SYLG184). Mix together using a wooden popsicle stick or tongue depressor. Pour into glass or plastic 100 cm dishes. Use a small butane torch held about 10 to 12 inches above the dishes to burn off any bubbles. Place lids on plates and let sit for at least 24 hr to set.</li>
	<li>To prepare dissection tools insert a Minutien pin (0.1 mm diameter) into a pin holder (Fine Scientific Tools, #26002-10 and #26018-17, respectively) alternatively 26 gauge needles (BD, # 309597) bent at right angles with the aid of forceps may be used as dissecting instruments. Sharpened Student Dumont #5 Forceps (Fine Science Tools, #91150-20) using a sharpening stone (Fine Scientific Tools, #29008-22).</li>
	<li>Sterilize all dissection tools with 70% ethanol before use.</li>
</ol>
3. Collection of Embryos
<ol>
	<li>Euthanize the timed pregnant mouse at 8.5 days post coitum according to animal use committee regulations and remove uterus.</li>
	<li>Place uterus in a plastic Petri dish with sterile PBS.</li>
	<li>Wash uterus once with sterile PBS.</li>
	<li>Under a dissection microscope carefully remove embryo from deciduas using a pair of forceps.</li>
	<li>Remove extraembryonic membranes and save for genotyping if needed.</li>
	<li>Stage embryo by counting somites. The ideal stages to make explants to analyze cellular behavior during neural fold elevation would be between the 6 and 10 somite stages when the neural folds are just beginning to elevate and before the major morphogenesis of the cranial mesenchyme takes place.</li>
	<li>Wash embryo in PBS. Place in a black sylgard plate with DMEM that contains no phenol red (Invitrogen, #21063-029). Remove the extraembryonic membranes and save for genotyping if needed.</li>
</ol>
4. Preparation of Explants
<ol>
	<li>Refocus the microscope at the highest possible magnification over the head region of the embryo.</li>
	<li>Make cuts with a dissection needle in each hand. Use one needle to hold the tissue in place and the other to make cuts.</li>
	<li>Cut off the head of the embryo anterior to the rhombomeres.</li>
	<li>Cut along the midline to bisect the head into two equal halves.</li>
	<li>Cut around the outer fringes of the head and midline. These cuts will remove the premigratory crest and surface ectoderm at the outer border and the prechordal plate at the midline. The remaining tissue will be the explant and will contain the migrating cranial neural crest, paraxial mesoderm and surface ectoderm.</li>
	<li>As the tissue is cut, remove dissected debris from the dish using a sterile pasteur pipette. This prevents confusing the explant with the dissected debris.</li>
	<li>Gently wash the explant by pipeting up and down with a 20 &mu;l pipette tip and pipetman. This will remove loosely attached cells and prevent jagged edges on the explants.</li>
	<li>Carefully pick up each explant in approximately 10 &mu;l of liquid using a 20 &mu;l pipette tip and pipetman and place at the center of the coated culture well.</li>
	<li>Add 20 &mu;l of culture media supplemented with 15% FBS to cover the explant.</li>
	<li>Place plate in a humidified tissue culture incubator (5% CO2, 37 &deg;C) for 1-2 hr to allow the explant to attach to the bottom of the dish. View periodically to ensure that the explant does not dry out.</li>
	<li>When the explant has attached, carefully add to each well 250 &mu;l DMEM supplemented with 10% FBS that has been warmed to 37 &deg;C in a waterbath. Once explants have attached they will not move when plate is gently shaken under microscope.</li>
	<li>Return dish to tissue culture incubator.</li>
	<li>After 24 hr, the explants will be firmly attached and cells will begin to emigrate. At this time pharmacological agents can be added to the media.</li>
	<li>48 hr after plating, cells will have migrated from the explants. The distance migrated can be visualized and imaged. We typically stop the culture at this point since explants begin to show signs of decreased viability with longer incubations. A vital stain such as acridine orange or trypan blue can also be used to check viability of the explant.</li>
	<li>At this point images can be acquired to document the distance and number of cells that migrate from the explant. Alternatively, if explants were plated on coverslips, they can be fixed and immunohistochemical analyses performed to visualize proteins of interest.</li>
</ol>
Critical steps in the protocol
<ol>
	<li>Cut explants of approximately the same size.</li>
	<li>Clear all dissection debris off the dissecting plate as you are dissecting to avoid confusing debris with the desired explanted tissue.</li>
	<li>When plating explants in wells or on coverslips be sure to place droplet of media with explant in center of well.</li>
	<li>Allow explant sufficient time to attach before flooding the well with media.</li>
	<li>Treatment conditions should be standardized for every test substrate and pharmacological agent used.</li>
	<li>Two explants can be made from each embryo (one from the left and one from the right of the neural tube).</li>
	<li>When testing migration of cells from mutant mouse lines under different experimental conditions, put one mutant explant in the control group and the other in the treated group.</li>
</ol>
Troubleshooting
<ol>
	<li>If explants fail to attach, increase the allotted time for attachment. In our experience, approximately half of the explants do not attach and larger explants adhere more efficiently.</li>
	<li>If explants attach to the periphery of the well they will be hard to image. Be sure to place explants in center of the well.</li>
	<li>Ideally explants will be approximately the same size. However, discrepancies in explant size can be corrected for using the diameter of the explants as a correction factor when quanitating migrations of cells from explants.</li>
	<li>If contamination occurs be sure sterile working conditions are being used. All tools and work surfaces should be sterilized with 70% ethanol. Pipets and Petri dishes should be new and sterile.</li>
	<li>If explants are getting lost during the dissection, periodically remove debris to prevent losing explant.</li>
	<li>Mastering the dissection is time consuming and takes practice. Optimal magnification, sharp dissection tools and needles help.</li>
</ol>

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No conflicts of interest declared.

The method applied here provides a powerful assay to examine the behavior of cranial mesenchyme cells. In addition to the static analyses presented here, live imaging experiments in bright field or in combination with expression of GFP-labeled proteins can be employed to examine the behaviors of cells in real time as they migrate from the explant. For live imaging experiments, explants could be labeled with DiI or to differentiate migration of neural crest from the paraxial mesoderm, ROSA-YFP;Mesp1-cre or ROSA-YFP;Wnt1-cre or other transgenic lines could be used. Cranial mesenchyme-ECM interactions can also be examined utilizing this explant assay. Here we plate explants on different ECM substrates that are present in the cranial mesenchyme to show that cells migrate on these to different degrees and that the ECM influences the morphology of the cells. Furthermore, explants can be embedded in a three-dimensional matrix to access the behaviors of cells in this context. This explant assay is amendable to analysis of the effect of genetic and pharmacological manipulations on cranial mesenchyme behaviors to analyze intrinsic and extrinsic cues that can regulate and modify migration. For example, we utilized this assay in our studies to discern the role of excess Hsp90 secretion in the abnormal cellular behaviors in Hectd1 mutant cranial mesenchyme15. In these experiments, we treated explants with anti-Hsp90 antibody, Hsp90 protein, geldanamycin and DMA (dimethyl amelioride) to block secretion of Hsp90 to demonstrate that the abnormal behavior of Hectd1 mutant cells is due to excess extracellular Hsp9015. Similar approaches could be used to pharmacologically dissect additional pathways underlying normal or abnormal morphogenesis of the cranial mesenchyme. Once the assay is completed, explants and migrating cells can be analyzed by a host of methods. For example, immunohistochemical analyses can be employed to examine localization of proteins critical for cell movements. Transcriptome analyses could also be employed to determine how treatments affect gene expression.
One significant limitation of this technique is that it does not model behaviors in three-dimensions as they would occur in vivo. Modification of the protocol where explants are embedded in a three-dimensional matrix (e.g. matrigel) along with expression of fluorescent protein-marked cellular compartments could be employed necessary to address this issue. Even if these modifications were performed, further experiments to correlate behaviors seen in this ex vivo assay with those in vivo would be necessary. Other limitations include the large number of embryos required to generate sufficient numbers to generate statically significant data. Most importantly, the dissection is relatively simple, it does require practice to master.

Neural tube closure in the cranial region occurs between embryonic day 8.5 (E8.5) and 9.5 in the mouse embryo. Failure to properly close the neural tube in the head results in anencephaly, a common structural birth defect in humans and is incompatible with life. The forces that drive cephalic neural tube closure are generated from both the neural tissue itself and the surrounding epidermis and mesenchyme3. In particular expansion of the cranial mesenchyme is thought to be essential for elevation of the cranial neural folds1,2. The cranial mesenchyme is rich in ECM proteins in particular glycosylated proteins such as heparin sulphate proteoglycans, chondroitin sulfates and hyaluronate4-8.
Unlike in the chicken embryo where neural crest cells emigrate from the dorsal neural tube following neural tube closure, the neural crest in the mouse embryo migrates at the same time that the neural folds begin to rise (after the 5 somite stage). Thus during neurulation in the mouse embryo, the cranial mesenchyme is composed of cells derived from the neural crest and the paraxial mesoderm. Neural crest and paraxial mesoderm populations are induced at different times in development; localized in different positions in the embryo and develop into different structures9,10. The paraxial mesoderm originates from the primitive streak and migrates to the anterior region of the embryo to underlie the presumptive neural plate. The neural crest is induced at the junction of the neural plate and epidermal ectoderm, undergoes an epithelial to mesenchyme transition and delaminates just prior to neural fold elevation in the rodent embryo. Neural crest cells migrate along stereotypic paths in the subectodermal paraxial mesoderm to the branchial arch, frontonasal and periocular mesenchyme. The paraxial mesoderm will contribute to some of the bones of the skull vault and muscles of the face; whereas the neural crest will contribute to other bones of the skull and face in addition to cranial nerves9-11. The paraxial mesoderm and neural crest lineages can be differentially marked by the Mesp1-cre and Wnt1-cre transgenic mouse lines, respectively 9.
The essential role of the cranial mesenchyme in neural tube closure has been inferred from experiments where treatment of rodent embryos with ECM disrupting agents such as hyaluronidase, chondroitinase ABC, heparitinase or Diazo-oxo-norleucine (DON) during neurulation impaired neural tube closure7,12-14. In these experiments, histological analysis of static sections following neurulation revealed associated dysmorphogeneis of the cranial mesenchyme7,12-14. However, since the teratogenic agent had access to multiple tissues, it remains to be determined if the cranial mesenchyme is really the target tissue. In support of the conclusion that this tissue is essential for neurulation, the cranial mesenchyme appears abnormal upon histological analyses in some mouse mutants with exencephaly15-17. Still, in most cases, the effect of the mutation on the cellular behavior of the cranial mesenchyme has not been addressed.
We have devised an ex vivo explant assay to directly examine the consequence of genetic mutation or pharmacological manipulation on the behavior of cranial mesenchyme cells15. This assay is similar to that published by Tzahor et al 2003 to access the differentiation potential of the cranial mesenchyme that underlies the rhombomeres18 except we have modified the explant dissection to study the migratory properties of more anterior populations of cranial mesenchyme that underlie the anterior neural plate. Our method is also a modification of explant assays performed in the chicken embryo to analyze the migratory behavior of the neural crest with key differences. Previous preparations have explanted the neural crest or the more posterior paraxial mesoderm19,20. Furthermore, during neural fold elevation in the chicken embryo, the neural crest has not yet emigrated from the dorsal neural tube and thus explants taken of the anterior paraxial mesoderm would not contain neural crest cells. In our assay, cranial mesenchyme explants consisting of paraxial mesoderm, neural crest and surface ectoderm are prepared and plated on a substrate. Experimental manipulation including isolation of explants from genetic mutants, plating explants on different ECM or pharmacological treatments can be performed. Cells migrate from the explant and the distance, number and behavior can be analyzed and compared between treatment groups. In addition, this preparation is amendable to analyses of cellular migration by live imaging techniques. After the migration experiment, explants can be fixed and subjected to immunohistochemical analyses to further elucidate the effect of treatments. On the whole, the protocol presented here is a simple ex vivo assay to investigate the behavior of the cranial mesenchyme. As a representative experiment, we utilize this assay to examine the migration of cranial mesenchyme on different extracellular substrates.

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 appearance of cells migrating from cranial mesenchyme explants is shown in Figure 2. We tested migration on various ECM substrates that are present in the cranial mesenchyme during neurulation including Fibronectin (100 mg/ml; Sigma #F1141), Laminin (100 mg/ml; Sigma #L2020), MaxGel ECM (1:500 dilution in DMEM; Sigma #E0282) and Hyaluronic acid (1 mg/ml; Sigma, # 53747). Cells fail to migrate from the explant when no ECM is present (Data not shown) and all substrates tested supported migration of the cells. Both the shape as well as the sizes of cells that migrate from the explants depends upon the substrate. This is expected as previous studies demonstrate that the mechanical forces generated by interaction of cells with different ECM affect both the shape and migratory properties of cells21.
<img alt="Figure 1" src="/files/ftp_upload/4245/4245fig1.jpg" fo:content-width="5.5in" fo:src="/files/ftp_upload/4245/4245fig1highres.jpg" /> 
 Figure 1. Preparation of explants. (A) To prepare explants, E8.5 embryos are dissected from deciduas and extraembryonic tissues. (B) Embryo heads are dissected from the body anterior to the rhombomeres. Explants are prepared by removing tissue from the midline and the outer edges of the head to remove the prechordal plate at the midline and the premigratory neural crest and surface ectoderm at the border, respectively. (C) Explants are prepared by cutting away tissue from the midline and outer edges of the head to remove the prechordal plate at the midline and the premigratory neural crest and surface ectoderm at the borders. (D) Explants are placed in a small volume of media on ECM coated dishes/coverslips and allowed to attach before the addition of more media.
<img alt="Figure 2" src="/files/ftp_upload/4245/4245fig2.jpg" /> 
 Figure 2. Migration of cranial mesenchyme cells from explants plated on different substrates. Explants were plated on Fibronectin, Laminin, MaxGel ECM or Hyaluronic acid and photographed after 48 hr in culture. Size bar = 200 mm.

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An Explant Assay for Assessing Cellular Behavior of the Cranial Mesenchyme

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 ...

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9.7K Aufrufe.  Children's National Medical Center. Der kraniale Mesenchym erfährt dramatischen morphogene Bewegungen, die wahrscheinlich stellt eine Antriebskraft für die Elevation der Neuralfalten 1,2. Hier beschreiben wir ein einfaches Ex vivo Explantation Assay, um die zellulären Verhalten der Schädelbasis Mesenchym während Neurulation charakterisieren. Dieser Test hat zahlreiche Anwendungen, darunter gefügig zu sein pharmakologischen Manipulationen und Live-Imaging-Analysen.

Serie: An Explant Assay for Assessing Cellular Behavior of the Cranial Mesenchyme

A Microfluidic Device with Groove Patterns for Studying Cellular Behavior

We describe a microfluidic device with microgrooved patterns for studying cellular behavior. This microfluidic platform consists of a top fluidic channel and a bottom microgrooved substrate. To fabricate the microgrooved channels, a top poly(dimethylsiloxane) (PDMS) mold containing the impression of the microfluidic channels was aligned and bonded to a microgrooved substrate. Using this device, mouse fibroblast cells were immobilized and patterned within microgrooved substrates (25, 50, 75, and 100 &mu;m wide). To study apoptosis in a microfluidic device, media containing hydrogen peroxide, Annexin V, and propidium iodide was perfused into the fluidic channel for 2 hours. We found that cells exposed to the oxidative stress became apoptotic. These apoptotic cells were confirmed by Annexin V that bound to phosphatidylserine at the outer leaflet of the plasma membrane during the apoptosis process. Using this microfluidic device with microgrooved patterns, the apoptosis process was observed in real-time and analyzed by using an inverted microscope containing an incubation chamber (37&deg;C, 5% CO2). Therefore, this microfluidic device incorporated with microgrooved substrates could be useful for studying the cellular behavior and performing high-throughput drug screening.

We describe a protocol for the fabrication of microfluidic devices that can enable cell capture and culture. In this approach patterned microstructures such as grooves within microfluidic channels are used to create low shear stress regions within which cell can dock.

We describe a microfluidic device with microgrooved patterns for studying cellular behavior.  This microfluidic platform consists of a top fluidic channel ...

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.

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.

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

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. ...

Electroporation of Craniofacial Mesenchyme

Electroporation is an efficient method of delivering DNA and other charged macromolecules into tissues at precise time points and in precise locations. For example, electroporation has been used with great success to study neural and retinal development in Xenopus, chicken and mouse 1-10. However, it is important to note that in all of these studies, investigators were not targeting soft tissues. Because we are interested in craniofacial development, we adapted a method to target facial mesenchyme.
When we searched the literature, we found, to our surprise, very few reports of successful gene transfer into cartilaginous tissue. The majority of these studies were gene therapy studies, such as siRNA or protein delivery into chondrogenic cell lines, or, animal models of arthritis 11-13. In other systems, such as chicken or mouse, electroporation of facial mesenchyme has been challenging (personal communications, Dept of Craniofacial Development, KCL). We hypothesized that electroporation into procartilaginous and cartilaginous tissues in Xenopus might work better. In our studies, we show that gene transfer into the facial cartilages occurs efficiently at early stages (28), when the facial primordium is still comprised of soft tissue prior to cartilage differentiation.
Xenopus is a very accessible vertebrate system for analysis of craniofacial development. Craniofacial structures are more readily visible in Xenopus than in any other vertebrate model, primarily because Xenopus embryos are fertilized externally, allowing analyses of the earliest stages, and facilitating live imaging at single cell resolution, as well as reuse of the mothers 14. Among vertebrate models developing externally, Xenopus is more useful for craniofacial analysis than zebrafish, as Xenopus larvae are larger and easier to dissect, and the developing facial region is more accessible to imaging than the equivalent region in fish. In addition, Xenopus is evolutionarily closer to humans than zebrafish (&tilde;100 million years closer) 15. Finally, at these stages, Xenopus tadpoles are transparent, and concurrent expression of fluorescent proteins or molecules will allow easy visualization of the developing cartilages. We anticipate that this approach will allow us to rapidly and efficiently test candidate molecules in an in vivo model system.

Craniofacial cartilages develop in close contact with other tissues and are difficult to manipulate in live animals. We are using electroporation to deliver molecular tools during growth of the craniofacial skeleton while bypassing early embryonic effects. This approach will allow us to efficiently test candidate molecules in vivo.

Electroporation is an efficient method of delivering DNA and other charged macromolecules into tissues at precise time points and in precise locations. ...

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 ...

Separation of Mouse Embryonic Facial Ectoderm and Mesenchyme

Orofacial clefts are the most frequent craniofacial defects, which affect 1.5 in 1,000 newborns worldwide1,2. Orofacial clefting is caused by abnormal facial development3. In human and mouse, initial growth and patterning of the face relies on several small buds of tissue, the facial prominences4,5. The face is derived from six main prominences: paired frontal nasal processes (FNP), maxillary prominences (MxP) and mandibular prominences (MdP). These prominences consist of swellings of mesenchyme that are encased in an overlying epithelium. Studies in multiple species have shown that signaling crosstalk between facial ectoderm and mesenchyme is critical for shaping the face6. Yet, mechanistic details concerning the genes involved in these signaling relays are lacking. One way to gain a comprehensive understanding of gene expression, transcription factor binding, and chromatin marks associated with the developing facial ectoderm and mesenchyme is to isolate and characterize the separated tissue compartments.
Here we present a method for separating facial ectoderm and mesenchyme at embryonic day (E) 10.5, a critical developmental stage in mouse facial formation that precedes fusion of the prominences. Our method is adapted from the approach we have previously used for dissecting facial prominences7. In this earlier study we had employed inbred C57BL/6 mice as this strain has become a standard for genetics, genomics and facial morphology8. Here, though, due to the more limited quantities of tissue available, we have utilized the outbred CD-1 strain that is cheaper to purchase, more robust for husbandry, and tending to produce more embryos (12-18) per litter than any inbred mouse strain8. Following embryo isolation, neutral protease Dispase II was used to treat the whole embryo. Then, the facial prominences were dissected out, and the facial ectoderm was separated from the mesenchyme. This method keeps both the facial ectoderm and mesenchyme intact. The samples obtained using this methodology can be used for techniques including protein detection, chromatin immunoprecipitation (ChIP) assay, microarray studies, and RNA-seq.

A protocol for separation of embryo facial ectoderm and mesenchyme is described. We use Dispase II to treat whole embryos first, dissect whole facial prominences out, and then separate the facial ectoderm and mesenchyme.

 Orofacial clefts are the most frequent  craniofacial defects, which affect 1.5 in 1,000 newborns worldwide1,2. Orofacial clefting is caused by abnormal ...

Live Imaging Assay for Assessing the Roles of Ca2+ and Sphingomyelinase in the Repair of Pore-forming Toxin Wounds

Plasma membrane injury is a frequent event, and wounds have to be rapidly repaired to ensure cellular survival. Influx of Ca2+ is a key signaling event that triggers the repair of mechanical wounds on the plasma membrane within ~30 sec. Recent studies revealed that mammalian cells also reseal their plasma membrane after permeabilization with pore forming toxins in a Ca2+-dependent process that involves exocytosis of the lysosomal enzyme acid sphingomyelinase followed by pore endocytosis. Here, we describe the methodology used to demonstrate that the resealing of cells permeabilized by the toxin streptolysin O is also rapid and dependent on Ca2+ influx. The assay design allows synchronization of the injury event and a precise kinetic measurement of the ability of cells to restore plasma membrane integrity by imaging and quantifying the extent by which the liphophilic dye FM1-43 reaches intracellular membranes. This live assay also allows a sensitive assessment of the ability of exogenously added soluble factors such as sphingomyelinase to inhibit FM1-43 influx, reflecting the ability of cells to repair their plasma membrane. This assay allowed us to show for the first time that sphingomyelinase acts downstream of Ca2+-dependent exocytosis, since extracellular addition of the enzyme promotes resealing of cells permeabilized in the absence of Ca2+.

Live Imaging Assay for Assessing the Roles of Ca2+ and Sphingomyelinase in the Repair of Pore-forming Toxin Wounds

Live imaging of cells exposed to the lipophilic dye FM1-43 allows precise determination of the kinetics by which pore-forming toxins are removed from the plasma membrane. This is a sensitive assay that can be used to assess requirements for Ca2+, sphingomyelinase and other factors on plasma membrane repair.

Plasma membrane injury is a frequent event, and wounds have to be rapidly repaired to ensure cellular survival. Influx of Ca2+ is a key signaling event ...

Measuring the Effects of Bacteria on C. Elegans Behavior Using an Egg Retention Assay

C. elegans egg-laying behavior is affected by environmental cues such as osmolarity1 and vibration2. In the total absence of food C. elegans also cease egg-laying and retain fertilized eggs in their uterus3. However, the effect of different sources of food, especially pathogenic bacteria and particularly Enterococcus faecalis, on egg-laying
	behavior is not well characterized. The egg-in-worm (EIW) assay is a useful tool to quantify the effects of different types of bacteria, in this case E. faecalis, on egg- laying behavior.
	
	EIW assays involve counting the number of eggs retained in the uterus of C. elegans4. The EIW assay involves bleaching staged, gravid adult C. elegans to remove the cuticle and separate the retained eggs from the animal. Prior to bleaching, worms are exposed to bacteria (or any type of environmental cue) for a fixed period of time. After bleaching, one is very easily able to count the number of eggs retained inside the uterus of the worms. In this assay, a quantifiable increase in egg retention after E. faecalis exposure can be easily measured. The EIW assay is a behavioral assay that may be used to screen for potentially pathogenic bacteria or the presence of environmental toxins. In addition, the EIW assay may be a tool to screen for drugs that affect neurotransmitter signaling since egg-laying behavior is modulated by neurotransmitters such as serotonin and acetylcholine5-9.

Measuring the Effects of Bacteria on C. Elegans Behavior Using an Egg Retention Assay

An egg-in-worm (EIW) assay is a useful method to quantify egg-laying behavior. Alterations in egg laying can be a behavioral response of the model organism Caenorhabditis elegans to potentially harmful environmental substances such as those produced by pathogenic bacteria.

C. elegans egg-laying behavior is  affected by environmental cues such as osmolarity1 and  vibration2. In the total absence of food C. elegans also cease ...

Single Cell Durotaxis Assay for Assessing Mechanical Control of Cellular Movement and Related Signaling Events

Durotaxis is the process by which cells sense and respond to gradients of tension. In order to study this process in vitro, the stiffness of the substrate underlying a cell must be manipulated. While hydrogels with graded stiffness and long-term migration assays have proven useful in durotaxis studies, immediate, acute responses to local changes in substrate tension allow focused study of individual cell movements and subcellular signaling events. To repeatably test the ability of cells to sense and respond to the underlying substrate stiffness, a modified method for application of acute gradients of increased tension to individual cells cultured on deformable hydrogels is used which allows for real time manipulation of the strength and direction of stiffness gradients imparted upon cells in question. Additionally, by fine tuning the details and parameters of the assay, such as the shape and dimensions of the micropipette or the relative position, placement, and direction of the applied gradient, the assay can be optimized for the study of any mechanically sensitive cell type and system. These parameters can be altered to reliably change the applied stimulus and expand the functionality and versatility of the assay. This method allows examination of both long term durotactic movement as well as more immediate changes in cellular signaling and morphological dynamics in response to changing stiffness.

Mechanical forces are important for controlling cell migration. This protocol demonstrates the use of elastic hydrogels that can be deformed using a glass micropipette and a micromanipulator to stimulate cells with a local stiffness gradient to elicit changes in cell structure and migration.

Durotaxis is the process by which cells sense and respond to gradients of tension. In order to study this process in vitro, the stiffness of the substrate ...

Assessing the Age-Specific Phagocytic Ability of Adult Drosophila melanogaster Hemocytes using an In Vivo Phagocytosis Assay

Phagocytosis is an essential function of the innate immune response. This process is carried out by phagocytic hemocytes whose primary function is to recognize a wide range of particles and destroy microbial pathogens. As organisms age, this process begins to decline, yet little is known about the underlying mechanisms or the genetic basis of immunosenescence. Here, an injection based in vivo phagocytosis assay is used to assess age related changes in different aspects of phagocytosis, such as binding, engulfment, and degradation of internalized particles, by quantifying phagocytic events in hemocytes in adult Drosophila. Drosophila melanogaster has become an ideal model to investigate age related changes in innate immune function for many reasons. For one, many genetic components and functions of the innate immune response, including phagocytosis, are evolutionarily conserved between Drosophila and mammals. Because of that, results obtained from using this protocol are likely to be widely relevant to understanding the age related changes in immune function in a variety of organisms. Additionally, we note that this method provides quantitative estimates of hemocyte phagocytic ability, which could be useful for a variety of research topics, and need not be limited to studies of aging.

Assessing the Age-Specific Phagocytic Ability of Adult Drosophila melanogaster Hemocytes using an In Vivo Phagocytosis Assay

This protocol describes an in vivo assay of phagocytosis used to assess and quantify the ability of young and aged Drosophila melanogaster hemocytes to phagocytose bacteria.

Phagocytosis is an essential function of the innate immune response. This process is carried out by phagocytic hemocytes whose primary function is to ...