We give special thanks to Yves Couder for many helpful discussions. We thank Atef Asnacios for the calibration of the cantilevers and discussion. We thank Lisa Willis, Elliot Meyerowitz, and Oliver Hamant&#160;for critical reading. This work was funded in part by Human Frontier Science Program grant RGP0062/2005-C; the Agence Nationale de la Recherche projects &#8216;&#8216;Growpec,&#8217;&#8217; and &#8216;&#8216;Mechastem&#8217;&#8217;.

1. Prepare Glass Slides for Mounting Sample
<ol>
	<li>Prepare imbedding agarose media: 0.7% low melting agarose in 10% mannitol (in water).</li>
	<li>Using a strong metal instrument (e.g. drill tip, lime), etch out a 0.5 x 0.5 cm area in the center&#160;of a microscopy glass slide. Or instead, glue a small piece of glass lamella&#160;(about 20 x 200 &#181;m)&#160;to the glass slide&#160;using Araldite glue. NOTE: This roughens the surface in order to facilitate the adhesion of the agarose to ensure that the agarose media sticks or fixes to the slide. This slide can be reused.</li>
	<li>Position a droplet of about 20 &#181;l&#160;of imbedding agarose media onto the etched region of the glass slide. This gives a thin film of agarose.</li>
	<li>Store the glass slide in a humid box and wait for the agarose media to solidify.</li>
</ol>
2. Dissecting and Mounting Meristem Samples
<ol>
	<li>Microdissect meristems for confocal imaging: remove the floral buds one by one from the stem by pulling them down with ultra fine tweezers. It is important to remove all the floral buds up to the buds that do not exhibit sepals (i.e. the P1 primordia14).</li>
	<li>Using a razor blade or the tip of the tweezers, cut the meristem away from the stem. The cut should be parallel to the region of the meristem surface that is to be measured with the AFM. The obtained apex should be between 300 &#181;m and 600 &#181;m high to fit under the AFM tip.</li>
	<li>Push the apex into the film of agarose media prepared at step 1.1.4. Choose a position on the glass slide/agarose and push gently such that the meristem is both directly in contact with the glass slide and protrudes out from the agarose. It is crucial to position the meristem in the agarose within the few seconds that follow its cut from the stem. NOTE: This is in order to prevent the meristem from drying. At this stage, the meristem will be standing in a crack because the agarose fractured upon the application of pressure.</li>
	<li>Prepare several meristems this way for batch imaging, providing that they are of the same height and are positioned less than 500 &#181;m from one another on the slide. Keep the prepared meristems in the humid box while dissecting further samples. 
	NOTE: It limits the variation coming from the calibration. Ideally 2 samples of each condition could be prepared and measured randomly. So far there is no observed effect of batch imaging on the measurement (stress induced response). This could be thanks to the dilution of stress signalling molecules in the agar and/or the mounting solution. Alternatively the stress response is the same regardless to the amount of sample prepared.</li>
	<li>Before moving on to the next step, make sure that the boundary between the meristem and the floral buds is dry. If not, gently remove the excess water using filter paper. This should prevent in the next stage that the melted agarose floods the sample due to capillarity forces.</li>
	<li>With a 20 &#181;l&#160;pipette, add enough melted mounting agarose to fill in the crack created at step 1.2.3. and to surround each meristem. The agarose should reach the level of the scar of the removed flower bud (primordium). To achieve this, it may be necessary to add agarose at each end of the crack.</li>
	<li>When the agarose is added, it will quickly flood into the crack; using the pipette, suck off part of the melted mounting agarose to ensure that the meristem is the highest point on the slide. This fixes the meristem so that it cannot move during imaging.</li>
</ol>
3. Mounting Root or Hypocotyl Samples
<ol>
	<li>For roots and hypocotyls, no dissection is necessary. In the thin film of agarose on the prepared glass slide, make a groove either directly above an etch in the glass or along a glass lamella. Position the root or hypocotyl in this groove ensuring that it is in contact with the glass along its whole length.</li>
	<li>Before moving on to the next step, make sure that the sample is dry at its surface. If not, gently remove the excess of water using filter paper.</li>
	<li>Add melted mounting agarose around the sample as in step 1.2.6.</li>
</ol>
4. AFM Preparation and Sensitivity Calibration (for a JPK Nanowizard AFM)
<ol>
	<li>Mount a cantilever on the AFM. The cantilever should be mounted with round shaped indenter. The radius will determine the x,&#160;y, z&#160;resolution and the indentation depth at which accurate measurement could be archived.
	<ol>
		<li>To take meso&#8212;nanoscale measurements at the epidermis&#8217;s surface, use a 50 nm radius round indenter; to take mesoscale measurements, use a 0.5 &#181;m radius round indenter; to take microscale measurements (across 2-3 cells), use a 2.5 &#181;m radius round indenter.</li>
	</ol>
	</li>
	<li>Position the laser at the tip of the cantilever. Align the laser to the middle of the captor using the mirror.</li>
	<li>Position a clean glass under the AFM. 
	NOTE: the following is set to transform the signal of the laser position on the AFM receptor into a deformation of the cantilever and, by evaluating the spring constant of the cantilever, to translate it into a force.</li>
	<li>Approach with a target force &#8220;set point&#8221; of 1 V.</li>
	<li>Select &#8220;force spectroscopy&#8221;. Do an indentation by setting the maximal &#8220;relative set point&#8221; to 2 V, the &#8220;extend speed&#8221; to 40 &#181;m/sec, and the &#8220;z length&#8221; to 5 &#181;m.</li>
	<li>From the &#8220;calibration manager&#8221; menu, select the linear part of the forward curve to fit the sensitivity using the &#8220;select fit range&#8221; button. The &#8220;relative set point&#8221; should be transformed from &#8220;volt&#8221; to &#8220;meter&#8221;.</li>
	<li>From the &#8220;calibration manager&#8221; menu, select &#8220;spring constant&#8221; to evaluate the elasticity of the cantilever using thermal fluctuations. Press &#8220;run&#8221; to obtain the spectral density plot of the cantilever. Find the resonance peak with the lowest frequency. Fit it using the &#8220;select fit range&#8221; button. 
	NOTE 1: For more details on the thermal fluctuations tuning, read Cook&#160;et al.15 
	NOTE 2: It is preferable to use a direct method to evaluate the elasticity of the cantilever using a reference cantilever (or material with known stiffness). The one used here was kindly donated by Atef Asnacios16.</li>
	<li>Add 200 &#181;l&#160;of 10% mannitol solution under the cantilever tip. Realign the laser to the middle of the captor using the mirror.</li>
	<li>Recalibrate the sensitivity: from the &#8220;calibration manager&#8221; menu, click on the &#8220;unknown&#8221; button; repeat steps 2.3 through to 2.5.</li>
</ol>
5. Data Acquisition: Apparent Young&#8217;s Modulus Cartography
<ol>
	<li>Position the mounted samples under the AFM and add a 500 &#181;l&#160;droplet of 10% mannitol solution onto the sample. The mannitol solution plasmolyzes the cells within minutes.</li>
	<li>Approach with a target force (&#8220;set point&#8221;) of 20 nN (&#8220;set point&#8221; should not be more than 3 V)</li>
	<li>Indent with a &#8220;relative set point&#8221; of 200 nN, an &#8220;extend speed&#8221; of 40 &#181;m/sec, and &#8220;z&#160;length&#8221; of 5 &#181;m.</li>
	<li>Adjust the &#8220;relative set point&#8221; in order to obtain an indentation of 100 nm for the 200 nm mounted cantilever or 0.5 &#181;m for the 1 &#181;m/5 &#181;m mounted cantilever.</li>
	<li>In &#8220;force mapping&#8221; mode, select a region to scan: for meristems, 70&#160;&#181;m&#160;x 70 &#181;m with a resolution (&#8220;pixels&#8221;) of 64 x 64; for hypocotyls and roots, 100 &#181;m&#160;x 100 &#181;m with a resolution of 64 x 64.</li>
	<li>Press scan to launch the experiments. Save the output.</li>
</ol>
6. Data Analysis: Apparent Young&#8217;s Modulus Calculations
<ol>
	<li>Open the data file in the data analysis software.</li>
	<li>Select &#8220;batch processing&#8221; to apply the same analysis to all the curves of a single map.</li>
	<li>Select &#8220;correct the height for the bending of the cantilever&#8221; so to extract the indentation depth.</li>
	<li>Select &#8220;Young&#8217;s modulus fit function&#8221; that will assume the force is proportional to the area of indentation. Set the &#8220;tip shape&#8221; to parabolic to assume that the indentation shape is a parabolic.</li>
	<li>Indicate the radius of the tip.</li>
	<li>Select preferentially the &#8220;retract&#8221; curve for the analysis because it is less sensitive to topography than the &#8220;extend&#8221; curve. However, if there is significant sticking of the probe during the &#8220;retract,&#8221; use the &#8220;extend&#8221; curve, which is insensitive to sticking.</li>
	<li>Set the Poisson ratio to 0.5. Press on analyze and save the output.</li>
</ol>

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The authors have nothing to disclose.

In plants, changing mechanical properties play a major role in directing growth and morphogenesis. To date there has been great progress in unraveling the genetic and chemical networks that control plant growth, but our knowledge of how these networks contribute to and are affected by changes in mechanical properties is rudimentary. This method should enable us to fill this gap, and so it should be of strong interest to scientists studying any aspect of plant growth or morphogenesis. We now summarize the challenges and l...

Growth in plants is achieved by the coordinated expansion of the rigid cell walls that surround each and every cell of the organism. Accumulating evidence indicates that it is through the modification of cell wall chemistry that plants locally control this expansion. The expansion is thought to be driven primarily by strain on the cell walls, caused by the cell&#8217;s high turgor pressure; this strain response to turgor pressure is governed by the mechanical properties of the cell walls1. Little is known of these mechanical properties and how they change during development. Furthermore little is known of how these mechanical properties are controlled and whether feedbacks contribute to alter cell wall chemistry in a manner that is apparently coordinated across a tissue. If we are to understand the connection between chemical and mechanical changes in plant cell walls during development, and ultimately how these microscopic interactions govern a plant&#8217;s macroscopic growth, a method that can monitor mechanical properties of cell walls in developing organs at the cellular or tissue scale is required.
The atomic force microscopy (AFM) method described here, which is based on micrometer or nanometer tissue compressions or indentations, was developed precisely to measure the mechanical properties of cell walls in developing organs simultaneously at subcellular resolutions and across entire regions of tissue. Other methods have either a resolution that is too low or too high: the extensometer is only able to measure the average mechanical properties of a whole tissue at the millimeter&#160;scale2-4, a scale that is for instance too large to measure early events in organogenesis; the microindenter can take measurements at subcellular resolution at the nanometer&#160;scale, but it is restricted to measuring isolated cells and not groups of cells or organs5-7. With the AFM, the required tissue, cellular, and subcellular resolutions can be achieved8-10. Recently several protocols have been&#160;developed specifically to measure mechanics of plants tissue that could also be used11,12.
We will present here how to evaluate the elasticity of the tissue through measurement of the apparent Young&#8217;s modulus13.
The Young modulus is commonly used to describe the stiffness of a material. During small deformation the force required to deform a material is proportional to the area of indentation. The Young modulus is this coefficient. In the case of a continuous homogenous material the same coefficient will be measured regardless of the indentation type (size and shape) but will change with the speed of the measurement. In the case of the complex structure of plants tissue, we have observed so far that the force is proportional to the deformation allowing the determination of a coefficient of proportionality that we name &#8220;apparent young modulus&#8221;. In contrast from continuous medias in the plants, this apparent young modulus is sensitive to the size of the indentation. It does not correspond to the young moduli of a pure cell wall. It best describes the elasticity of the scaffolding of the cell-wall of the tissue.

<table border="0" cellpadding="0" cellspacing="0" width="850">
	<tbody>
		<tr height="21">
			<td height="21" style="height:21px;width:216px;">AFM</td>
			<td style="width:348px;">JPK</td>
			<td style="width:119px;">NanoWizard</td>
			<td style="width:167px;">All the 3-generation are abele to do the work withe the same preferment</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:216px;">AFM stage</td>
			<td style="width:348px;">JPK</td>
			<td style="width:119px;">CellHesion</td>
			<td>Required for sample withe low topography (les then 11&#181;m between the lowest and the highest point in the aria of force scanning).</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:216px;">AFM optics</td>
			<td style="width:348px;">JPK</td>
			<td style="width:119px;">Top View Optics&#160;</td>
			<td>Very important in order to position the sample. Cold be replaces by long range a binocular or microscope</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:216px;">Stereo Microscopes&#160;</td>
			<td style="width:348px;">Leica</td>
			<td style="width:119px;">M125</td>
			<td>Any type of stereo microscopes could do.&#160;</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:216px;"></td>
			<td style="width:348px;"></td>
			<td style="width:119px;"></td>
			<td></td>
		</tr>
		<tr height="42">
			<td height="42" style="height:42px;width:216px;">150nm mounted cantilever&#160;</td>
			<td style="width:348px;">nanosensors Rue Jaquet-Droz 1Case Postale 216 CH-2002 Neuchatel Switzerland</td>
			<td style="width:119px;">R150-NCL-10</td>
			<td>To measure only the cell wall at the surface of the epidermis use</td>
		</tr>
		<tr height="42">
			<td height="42" style="height:42px;">1&#181;m mounted cantilever&#160;</td>
			<td style="width:348px;">nanosensors Rue Jaquet-Droz 1Case Postale 216 CH-2002 Neuchatel Switzerland</td>
			<td>SD-Sphere-NCH-S-10&#160;</td>
			<td>to measure the mechanics of the cell wall orthogonal to the surface of the epidermis</td>
		</tr>
		<tr height="42">
			<td height="42" style="height:42px;width:216px;">Tipless cantiliver</td>
			<td style="width:348px;">nanosensors Rue Jaquet-Droz 1Case Postale 216 CH-2002 Neuchatel Switzerland</td>
			<td>TL-NCH-20&#160;</td>
			<td>to measure the local mechanics of the tissue (2-3 cell wide) use a 5&#181;m mounted cantilever. We attached a 5&#181;m borasilicate bead to a tipless cantiliver</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:216px;">5&#181;m silicon microspheres</td>
			<td style="width:348px;">Corpuscular</td>
			<td style="width:119px;">C-SIO-5</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:216px;">Aradilte&#160;</td>
			<td style="width:348px;">Bartik S.A. 77170 Coubet France</td>
			<td style="width:119px;"></td>
			<td>Aradilte for fixing the bead to the tip les cantiliver</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:216px;"></td>
			<td style="width:348px;"></td>
			<td style="width:119px;"></td>
			<td></td>
		</tr>
		<tr height="21">
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afm-based mapping of the elastic properties of cell walls: at tissue, cellular, and subcellular resolutions

We describe a recently developed method to measure mechanical properties of the surfaces of plant tissues using atomic force microscopy (AFM) micro/nano-indentations, for a JPK&#160;AFM. Specifically, in this protocol we measure the apparent Young&#8217;s modulus of cell walls at subcellular resolutions across regions of up to 100 &#181;m&#160;x 100 &#181;m in floral meristems, hypocotyls, and roots. This requires careful preparation of the sample, the correct selection of micro-indenters and indentation depths. To account for cell wall properties only, measurements are performed in highly concentrated solutions of mannitol in order to plasmolyze the cells and thus remove the contribution of cell turgor pressure.
In contrast to other extant techniques, by using different indenters and indentation depths, this method allows simultaneous multiscale measurements, i.e. at subcellular resolutions and across hundreds of cells comprising a tissue. This means that it is now possible&#160;to spatially-temporally characterize the changes that take place in the mechanical properties of cell walls during development, enabling these changes to be correlated with growth and differentiation. This represents a key step to understand how coordinated microscopic cellular changes bring about macroscopic morphogenetic events.
However, several limitations remain: the method can only be used on fairly small samples (around 100 &#181;m in diameter) and only on external tissues; the method is sensitive to tissue topography; it measures only certain aspects of the tissue&#8217;s complex mechanical properties. The technique is being developed rapidly and it is likely that most of these limitations will be resolved in the near future.

In Figure 1 we present typical Young moduli maps of floral meristems (Figures 1A and 1B), young and old hypocotyls (Figures 1C-F), and root meristem (Figure 1G and 1H). In all experiments the indenter is hemispherical, but its radius differs so that different spatial resolutions can be achieved. Figures 1C and 1D show typical results for meso-nanoscale indenters (50 nm radius) with meso-...

Watch this Scientific Journal Video about AFM-based Mapping of the Elastic Properties of Cell Walls: at Tissue, Cellular, and Subcellular Resolutions at JoVE.com

AFM-based Mapping of the Elastic Properties of Cell Walls: at Tissue, Cellular, and Subcellular Resolutions

We describe a method to map mechanical properties of plant tissues using an atomic force microscope (AFM). We focus on how to record mechanical changes that take place in cell walls during plant development at wide-field mesoscale, enabling these changes to be correlated with growth and morphogenesis.

We describe a recently developed method to measure mechanical properties of the surfaces of plant tissues using atomic force microscopy (AFM) ...

afm-based-mapping-elastic-properties-cell-walls-at-tissue-cellular

Research

JoVE Journal

Biology

12.8K Views.  Université Paris Diderot. We describe a method to map mechanical properties of plant tissues using an atomic force microscope (AFM). We focus on how to record mechanical changes that take place in cell walls during plant development at wide-field mesoscale, enabling these changes to be correlated with growth and morphogenesis.

Video: AFM-based Mapping of the Elastic Properties of Cell Walls: at Tissue, Cellular, and Subcellular Resolutions

Actin Co-Sedimentation Assay; for the Analysis of Protein Binding to F-Actin

The actin cytoskeleton within the cell is a network of actin filaments that allows the movement of cells and cellular processes, and that generates tension and helps maintains cellular shape.  Although the actin cytoskeleton is a rigid structure, it is a dynamic structure that is constantly remodeling.  A number of proteins can bind to the actin cytoskeleton.  The binding of a particular protein to F-actin is often desired to support cell biological observations or to further understand dynamic processes due to remodeling of the actin cytoskeleton. The actin co-sedimentation assay is an in vitro assay routinely used to analyze the binding of specific proteins or protein domains with F-actin.  The basic principles of the assay involve an incubation of the protein of interest (full length or domain of) with F-actin, ultracentrifugation step to pellet F-actin and analysis of the protein co-sedimenting with F-actin.  Actin co-sedimentation assays can be designed accordingly to measure actin binding affinities and in competition assays.

Proteins bind to filamentous actin (F-actin) through distinct actin binding modules. In this video we demonstrate the procedure of actin co-sedimentation, which is an in vitro assay routinely used to analyze proteins or specific domains that bind F-actin.

The actin cytoskeleton within the cell is a network of actin filaments that allows the movement of cells and cellular processes, and that generates ...

Single Molecule Methods for Monitoring Changes in Bilayer Elastic Properties

Membrane protein function is regulated by the cell membrane lipid composition.  This regulation is due to a combination of specific lipid-protein interactions and more general lipid bilayer-protein interactions. These interactions are particularly important in pharmacological research, as many current pharmaceuticals on the market can alter the lipid bilayer material properties, which can lead to altered membrane protein function.   The formation of gramicidin channels are dependent on conformational changes in gramicidin subunits which are in turn dependent on the properties of the lipid.  Hence the gramicidin channel  current is a reporter of altered properties of the bilayer due to certain compounds.  

Membrane protein function is regulated by the cell membrane lipid composition. This video-article details how to form a patch using bilayer patch electrodes, as well as how to use gramicidin channels as reporters of altered membrane properties.

Membrane protein function is regulated by the cell membrane lipid composition.  This regulation is due to a combination of specific lipid-protein ...

Comprehensive Compositional Analysis of Plant Cell Walls (Lignocellulosic biomass) Part II: Carbohydrates

The need for renewable, carbon neutral, and sustainable raw materials for industry and society has become one of the most pressing issues for the 21st century. This has rekindled interest in the use of plant products as industrial raw materials for the production of liquid fuels for transportation2 and other products such as biocomposite materials6. Plant biomass remains one of the greatest untapped reserves on the planet4. It is mostly comprised of cell walls that are composed of energy rich polymers including cellulose, various hemicelluloses, and the polyphenol lignin5 and thus sometimes termed lignocellulosics. However, plant cell walls have evolved to be recalcitrant to degradation as walls contribute extensively to the strength and structural integrity of the entire plant. Despite its necessary rigidity, the cell wall is a highly dynamic entity that is metabolically active and plays crucial roles in numerous cell activities such as plant growth and differentiation5. Due to the various functions of walls, there is an immense structural diversity within the walls of different plant species and cell types within a single plant4. Hence, depending of what crop species, crop variety, or plant tissue is used for a biorefinery, the processing steps for depolymerisation by chemical/enzymatic processes and subsequent fermentation of the various sugars to liquid biofuels need to be adjusted and optimized. This fact underpins the need for a thorough characterization of plant biomass feedstocks. Here we describe a comprehensive analytical methodology that enables the determination of the composition of lignocellulosics and is amenable to a medium to high-throughput analysis (Figure 1). The method starts of with preparing destarched cell wall material. The resulting lignocellulosics are then split up to determine its monosaccharide composition of the hemicelluloses and other matrix polysaccharides1, and its content of crystalline cellulose7. The protocol for analyzing the lignin components in lignocellulosic biomass is discussed in Part I3.

Comprehensive Compositional Analysis of Plant Cell Walls Lignocellulosic biomass Part II: Carbohydrates

Plant biomass is a major carbon-neutral renewable resource that could be used for the production of biofuels. Plant biomass consists mainly of cell walls, a structurally complex composite material termed lignocellulosics. Here we describe a protocol for a comprehensive analysis of the content and composition of wall derived carbohydrates.

The need for renewable, carbon neutral, and sustainable raw materials for industry and society has become one of the most pressing issues for the 21st ...

One-channel Cell-attached Patch-clamp Recording

Ion channel proteins are universal devices for fast communication across biological membranes. The temporal signature of the ionic flux they generate depends on properties intrinsic to each channel protein as well as the mechanism by which it is generated and controlled and represents an important area of current research. Information about the operational dynamics of ion channel proteins can be obtained by observing long stretches of current produced by a single molecule. Described here is a protocol for obtaining one-channel cell-attached patch-clamp current recordings for a ligand gated ion channel, the NMDA receptor, expressed heterologously in HEK293 cells or natively in cortical neurons. Also provided are instructions on how to adapt the method to other ion channels of interest by presenting the example of the mechano-sensitive channel PIEZO1. This method can provide data regarding the channel&#8217;s conductance properties and the temporal sequence of open-closed conformations that make up the channel&#8217;s activation mechanism, thus helping to understand their functions in health and disease.

Described here is a procedure for obtaining long stretches of current recording from one ion channel with the cell-attached patch-clamp technique. This method allows for observing, in real time, the pattern of open-close channel conformations that underlie the biological signal. These data inform about channel properties in undisturbed biological membranes.

Ion channel proteins are universal devices for fast communication across biological membranes. The temporal signature of the ionic flux they generate ...

Isolation of Murine Valve Endothelial Cells

Normal valve structures consist of stratified layers of specialized extracellular matrix (ECM) interspersed with valve interstitial cells (VICs) and surrounded by a monolayer of valve endothelial cells (VECs). VECs play essential roles in establishing the valve structures during embryonic development, and are important for maintaining life-long valve integrity and function. In contrast to a continuous endothelium over the surface of healthy valve leaflets, VEC disruption is commonly observed in malfunctioning valves and is associated with pathological processes that promote valve disease and dysfunction. Despite the clinical relevance, focused studies determining the contribution of VECs to development and disease processes are limited. The isolation of VECs from animal models would allow for cell-specific experimentation. VECs have been isolated from large animal adult models but due to their small population size, fragileness, and lack of specific markers, no reports of VEC isolations in embryos or adult small animal models have been reported. Here we describe a novel method that allows for the direct isolation of VECs from mice at embryonic and adult stages. Utilizing the Tie2-GFP reporter model that labels all endothelial cells with Green Fluorescent Protein (GFP), we have been successful in isolating GFP-positive (and negative) cells from the semilunar and atrioventricular valve regions using fluorescence activated cell sorting (FACS). Isolated GFP-positive VECs are enriched for endothelial markers, including CD31 and von Willebrand Factor (vWF), and retain endothelial cell expression when cultured; while, GFP-negative cells exhibit molecular profiles and cell shapes consistent with VIC phenotypes. The ability to isolate embryonic and adult murine VECs allows for previously unattainable molecular and functional studies to be carried out on a specific valve cell population, which will greatly improve our understanding of valve development and disease mechanisms.

The ability to isolate heart valve endothelial cells (VECs) is critical for understanding mechanisms of valve development, maintenance, and disease. Here we describe the isolation of VECs from embryonic and adult Tie2-GFP mice using FACS that will allow for studies determining the contribution of VECs in developmental and disease processes.

Normal valve structures consist of stratified layers of specialized extracellular matrix (ECM) interspersed with valve interstitial cells (VICs) and ...

Voltage and Calcium Dual Channel Optical Mapping of Cultured HL-1 Atrial Myocyte Monolayer

Optical mapping has proven to be a valuable technique to detect cardiac electrical activity on both intact ex vivo hearts and in cultured myocyte monolayers. HL-1 cells have been widely used as a 2-Dimensional cellular model for studying diverse aspects of cardiac physiology. However, it has been a great challenge to optically map calcium (Ca) transients and action potentials simultaneously from the same field of view in a cultured HL-1 atrial cell monolayer. This is because special handling and care is required to prepare healthy cells that can be electrically captured and optically mapped. Therefore, we have developed an optimal working protocol for dual channel optical mapping. In this manuscript, we have described in detail how to perform the dual channel optical mapping experiment. This protocol is a useful tool to enhance the understanding of action potential propagation and Ca kinetics in arrhythmia development.

This article describes the technique used to perform dual channel optical mapping in cultured HL-1 atrial cell monolayers. This unique protocol allows the simultaneous visualization of both calcium (Ca) and voltage (Vm) activity in the same area for the detailed detection and analysis of electrophysiological properties of culture monolayers.

Optical mapping has proven to be a valuable technique to detect cardiac electrical activity on both intact ex vivo hearts and in cultured myocyte ...

Dissection of the Auditory Bulla in Postnatal Mice: Isolation of the Middle Ear Bones and Histological Analysis

In most mammals, auditory ossicles in the middle ear, including the malleus, incus and stapes, are the smallest bones. In mice, a bony structure called the auditory bulla houses the ossicles, whereas the auditory capsule encloses the inner ear, namely the cochlea and semicircular canals. Murine ossicles are essential for hearing and thus of great interest to researchers in the field of otolaryngology, but their metabolism, development, and evolution are highly relevant to other fields. Altered bone metabolism can affect hearing function in adult mice, and various gene-deficient mice show changes in morphogenesis of auditory ossicles in utero. Although murine auditory ossicles are tiny, their manipulation is feasible if one understands their anatomical orientation and 3D structure. Here, we describe how to dissect the auditory bulla and capsule of postnatal mice and then isolate individual ossicles by removing part of the bulla. We also discuss how to embed the bulla and capsule in different orientations to generate paraffin or frozen sections suitable for preparation of longitudinal, horizontal, or frontal sections of the malleus. Finally, we enumerate anatomical differences between mouse and human auditory ossicles. These methods would be useful in analyzing pathological, developmental and evolutionary aspects of auditory ossicles and the middle ear in mice.

We present a protocol to isolate the auditory bulla, capsule, and ossicles from postnatal mice for whole mount and histological analysis.

In most mammals, auditory ossicles in the middle ear, including the malleus, incus and stapes, are the smallest bones. In mice, a bony structure called ...

Analysis of Beta-cell Function Using Single-cell Resolution Calcium Imaging in Zebrafish Islets

Pancreatic beta-cells respond to increasing blood glucose concentrations by secreting the hormone insulin. The dysfunction of beta-cells leads to hyperglycemia and severe, life-threatening consequences. Understanding how the beta-cells operate under physiological conditions and what genetic and environmental factors might cause their dysfunction could lead to better treatment options for diabetic patients. The ability to measure calcium levels in beta-cells serves as an important indicator of beta-cell function, as the influx of calcium ions triggers insulin release. Here we describe a protocol for monitoring the glucose-stimulated calcium influx in zebrafish beta-cells by using GCaMP6s, a genetically encoded sensor of calcium. The method allows monitoring the intracellular calcium dynamics with single-cell resolution in ex vivo mounted islets. The glucose-responsiveness of beta-cells within the same islet can be captured simultaneously under different glucose concentrations, which suggests the presence of functional heterogeneity among zebrafish beta-cells. Furthermore, the technique provides high temporal and spatial resolution, which reveals the oscillatory nature of the calcium influx upon glucose stimulation. Our approach opens the doors to use the zebrafish as a model to investigate the contribution of genetic and environmental factors to beta-cell function and dysfunction.

Beta-cell functionality is important for blood-glucose homeostasis, which is evaluated at single-cell resolution using a genetically encoded reporter for calcium influx.

Pancreatic beta-cells respond to increasing blood glucose concentrations by secreting the hormone insulin. The dysfunction of beta-cells leads to ...

Super-Resolution Live Cell Imaging of Subcellular Structures

There has long been a crucial tradeoff between spatial and temporal resolution in imaging. Imaging beyond the diffraction limit of light has traditionally been restricted to be used only on fixed samples or live cells outside of tissue labeled with strong fluorescent signal. Current super-resolution live cell imaging techniques require the use of special fluorescence probes, high illumination, multiple image acquisitions with post-acquisition processing, or often a combination of these processes. These prerequisites significantly limit the biological samples and contexts that this technique can be applied to.
Here we describe a method to perform super-resolution (~140 nm XY-resolution) time-lapse fluorescence live cell imaging in situ. This technique is also compatible with low fluorescent intensity, for example, EGFP or mCherry endogenously tagged at lowly expressed genes. As a proof-of-principle, we have used this method to visualize multiple subcellular structures in the Drosophila testis. During tissue preparation, both the cellular structure and tissue morphology are maintained within the dissected testis. Here, we use this technique to image microtubule dynamics, the interactions between microtubules and the nuclear membrane, as well as the attachment of microtubules to centromeres.
This technique requires special procedures in sample preparation, sample mounting and immobilizing of specimens. Additionally, the specimens must be maintained for several hours after dissection without compromising cellular function and activity. While we have optimized the conditions for live super-resolution imaging specifically in Drosophila male germline stem cells (GSCs) and progenitor germ cells in dissected testis tissue, this technique is broadly applicable to a variety of different cell types. The ability to observe cells under their physiological conditions without sacrificing either spatial or temporal resolution will serve as an invaluable tool to researchers seeking to address crucial questions in cell biology.

Presented here is a protocol for super-resolution live-cell imaging in intact tissue. We have standardized the conditions for imaging a highly sensitive adult stem cell population in its native tissue environment. This technique involves balancing temporal and spatial resolution to allow for the direct observation of biological phenomena in live tissue.

There has long been a crucial tradeoff between spatial and temporal resolution in imaging. Imaging beyond the diffraction limit of light has traditionally ...

Quantification of Cellular Densities and Antigenic Properties using Magnetic Levitation

The described method was developed based on the principles of magnetic levitation, which separates cells and particles based on their density and magnetic properties. Density is a cell type identifying property, directly related to its metabolic rate, differentiation, and activation status. Magnetic levitation allows a one-step approach to successfully separate, image and characterize circulating blood cells, and to detect anemia, sickle cell disease, and circulating tumor cells based on density and magnetic properties. This approach is also amenable to detecting soluble antigens present in a solution by using sets of low- and high-density beads coated with capture and detection antibodies, respectively. If the antigen is present in solution, it will bridge the two sets&#160;of beads, generating a new bead-bead complex, which will levitate in between the rows of antibody-coated beads. Increased concentration of the target antigen in solution will generate a larger number of bead-bead complexes when compared to lower concentrations of antigen, thus allowing for quantitative measurements of the target antigen. Magnetic levitation is advantageous to other methods due to its decreased sample preparation time and lack of dependance on classical readout methods. The image generated is easily captured and analyzed using a standard microscope or mobile device, such as a smartphone or a tablet.

This paper describes a magnetic levitation-based method that can specifically detect the presence of antigens, either soluble or membrane-bound, by quantifying changes in the levitation height of capture beads with fixed densities.

The described method was developed based on the principles of magnetic levitation, which separates cells and particles based on their density and magnetic ...