The authors have no acknowledgements.

NOTE: All procedures involving the use of animals are carried out according to the local legislation. Our work is approved by the Sydney Local Health District Animal Welfare Committee. The allowed procedures are described in the project license documentation and follow guidelines that ensure the welfare of the animal at all times.&#160;Ensure adherence to the legislation on animal experimentation of the country where the work is performed.
1. Protocol for Preparing EM Fixative
<ol>
	<li>For 100 ml of fixative, prepare&#160;paraformaldehyde by adding 2 g of paraformaldehyde powder to 25 ml distilled water in a conical flask, heat to 65 &#186;C, stirring constantly with a magnetic stirrer. 
	NOTE: Glutaraldehyde, paraformaldehyde and sodium cacodylate buffer are all hazardous substances and must always be handled in fumehood.</li>
	<li>Heat at 65 &#186;C for 2 min then allow to cool. If needed, add sodium hydroxide solution dropwise while stirring to clear solution completely. 
	NOTE: sodium hydroxide is hazardous, handle with care.</li>
	<li>Add 10 ml of EM grade stock&#160;glutaraldehyde, 50 ml of 0.2 M sodium cacodylate buffer and 2 g of sucrose and stir with magnetic stirrer. Add 2 ml of 1.0 M CaCl2.</li>
	<li>Adjust to pH 7.4 prior to making solution up to 100 ml with distilled water. The final fixative contains 2.5 % glutaraldehyde, 2 % Paraformaldehyde, 2 mmol CaCl2, 2 % Sucrose in 0.1 M&#160; cacodylate buffer. Ensure that the total Osmolality is 440 mOsmol/L. Use within 24 hr&#160;of preparation.</li>
</ol>
2. Perfusion Fixation of Liver
<ol>
	<li>Warm perfusion buffer and fixative to 35 - 37 &#186;C. The correct temperature of the exsanguination buffer and fixative helps to ensure tissue integrity.</li>
	<li>Anaesthetize the animal with a single intraperitoneal injection of 50 mg/kg Ketamine and 5 mg/kg Xylazine.</li>
	<li>Once the animal is under deep anaesthesia, assessed by hind limb withdrawal and tail pinch, a Y incision is made with blunt ended scissors in the animal&#8217;s abdomen to expose the liver and portal vein. 
	NOTE: Ensure that equipment is clean to avoid contamination of specimens with microorganisms and general debris which will be readily seen on SEM.</li>
	<li>Tie two very loose sutures around the portal vein one proximal to the liver, the other more distally to the liver.</li>
	<li>Cannulate the portal vein with an appropriately sized IV cannula (18 G for rats, 22 G for mice).Tighten sutures to secure cannula.</li>
	<li>Commence perfusion with phosphate buffered saline warmed to 37 &#186;C, and at a pressure of 10 cm of H20. Between 5 and 20 ml of PBS is usually required to exsanguinate the liver. 
	NOTE: Do not to allow air bubbles to enter the liver through the tubes connected to the cannula as they induce artefacts secondary to embolization and pressure; Damage to the hepatocytes, sinusoids and LSEC fenestrations occurs if the perfusion pressure is too high.</li>
	<li>Sever the abdominal and thoracic vena cava to allow the buffer to exit freely from the liver. This prevents high back-pressure damage to LSECs.</li>
	<li>Once the liver is free of blood, replace PBS with EM fixative and perfuse for approximately 5 minutes, until the liver is hardened and very pale.</li>
	<li>Discontinue perfusion and cut the fixed liver into 1 - 2 mm3 blocks using a sharp scalpel.</li>
	<li>Post-fix tissue in EM fixative for 24 - 72 hr at 4 &#186;C. 
	NOTE: In general, underfixation causes more artefacts than overfixation.</li>
	<li>Change fixative following post-fix period to 0.1 M sodium cacodylate buffer for storage at 4 &#186;C. 
	NOTE: Specimens stored in this way will be preserved for a significant period of time (up to 12 months), however timely secondary fixation and examination is recommended for best results.</li>
</ol>
3. Needle Perfusion
NOTE: Needle fixation is a variant of perfusion fixation that involves injection of fixative directly into biopsied liver tissue. The aim is for fixative to be flushed through the blood vessels in order to exsanguinate them and expose all of the tissue block to fixative. Care must be taken to keep the injecting pressure low to avoid pressure injury18,19.
<ol>
	<li>Remove a small piece of liver from the animal or subject and rinse in saline to remove blood and debris.</li>
	<li>Draw the normal saline into syringe fitted with a fine gauge needle (typically an insulin syringe).</li>
	<li>Inject the saline very slowly into the liver until blood is flushed out of the tissue. Multiple injections may be required.</li>
	<li>Draw the EM fixative into another syringe fitted with a fine gauge needle (typically an insulin syringe).</li>
	<li>Inject the fixative very slowly into the liver until it becomes hard and tan in colour. Multiple injections may be required.</li>
	<li>Cut the fixed liver into 1 - 2 mm3 blocks using a sharp scalpel. Incubate in EM fixative for 24 - 72 hr 4 &#176;C. Wash 3 times in 0.1 M sodium cacodylate buffer before commencing secondary fixation.</li>
</ol>
4. Preparation for Scanning Electron Microscopy
<ol>
	<li>Wash sample 3 times in 0.1 M sodium cacodylate buffer. To fix lipids, post fix the sample in 2% osmium tetroxide in 0.1 M sodium cacodylate buffer for 2 hr. 
	NOTE: Complete washing is very important as glutaraldehyde and osmium tetroxide cross react and generate artefacts on SEM.</li>
	<li>Rinse the samples in increasing concentrations of ethanol to commence dehydration. First rinse in 50 % ethanol for 5 minutes; then 3 times for 5 min with 70% ethanol; rinse 3 times for 5 min with 90% ethanol; rinse 2 times for 5 min in 100 % ethanol; and finally rinse 2 times for 10 min in 100% ethanol (molecular sieve).</li>
	<li>Remove all ethanol from the samples and replace with hexamethyldisilazane (HMDS) in fumehood and leave for 10 min. Remove HMDS from the tissue samples and allow to evaporate. 
	NOTE: HMDS is highly hygroscopic when cold therefore allow to reach RT before using for final dehydration step.</li>
	<li>Optionally, mount the samples immediately or place them into a desiccator until ready to mount for SEM.</li>
</ol>
5. Mounting
NOTE: Correct specimen mounting will maximize the number of clearly delineated sinusoids available for analysis under SEM.
<ol>
	<li>Label the base of the metal SEM stubs and apply double-sided conductive carbon tape to the top of the stubs.</li>
	<li>Visualize samples using a dissecting microscope to identify the surface with the best sinusoids for subsequent SEM. Place that surface upwards onto the carbon tape surface of the stub.</li>
	<li>Stick specimens firmly to the stub, specimens are now ready for sputter coating.</li>
</ol>
6. Coating
NOTE: Coating the specimen with a fine film of conductive metal (gold or platinum) in a sputter coater grounds the specimen and protects it from damage from the electron beam. If the coating is too thick structures of interest may be obscured.
<ol>
	<li>Place stubs into the vacuum chamber of automatic fine sputter coater, set mode to rotation and automatic sputter coating for 45 sec. Use platinum coating for finer detailing, however gold is also suitable.</li>
	<li>Apply a thin layer of carbon paint to the external surfaces of the mounted liver specimens, taking care not to coat the sinusoidal surfaces.</li>
	<li>Once the specimens are coated with platinum and carbon paint has dried they are ready for observation on the SEM. Samples can be stored in a desiccator.</li>
</ol>
7. Using the Scanning Electron Microscope
<ol>
	<li>Place specimens into scanning electron microscope and return microscope to vacuum.</li>
	<li>Using standard microscope operating procedures for visualisation of specimens by commencing at a low magnification, gradually increasing magnification.</li>
	<li>Ensure the microscope is adjusted appropriately for working distance, spot size and astigmatism.</li>
	<li>Check for fixation integrity. If the LSEC is mostly free from gaps and contains fenestrations measuring 30 - 250 nm diameter this would generally indicate that specimen preparation has been successful.</li>
	<li>If the sinusoids are full of gaps or devoid of fenestrations the specimen preparation has not been successful or there is significant pathology. In this case, slice the sample with a very fine razor blade, and the cut surfaces recoated and analysed for successfully fixed sinusoids.</li>
	<li>At approximately 15,000-20,000 &#215; magnification select flat LSEC surfaces that are free of debris with good fixation and where the fenestrations are clearly visible (typically containing at least 10 fenestrations).</li>
	<li>Save images of at least 10 sinusoids per liver, from different areas of the blocks, using at least two blocks per liver. Sampling 10 micrographs at 15,000 - 20,000 &#215; magnification per animal is generally sufficient for the quantification of fenestrations to be representative of the entire liver.</li>
</ol>
8. Analysis
NOTE: ImageJ software which can be downloaded free from the NIH is utilized to quantify fenestration diameter and frequency (<a href="http://imagej.nih.gov/ij">www.imagej.nih.gov/ij</a>).
<ol>
	<li>Open Image J and set the scale using the scale bar embedded in the picture (Screen shot 1).</li>
	<li>Using the polygon tool in ImageJ, trace around the entire flat area of the LSEC including fenestrated and defenestrated areas. Don&#8217;t just trace the sieve plates as this will falsely inflate porosity. Exclude any large pieces of debris that are on the cell surface that might be obscuring fenestrations.</li>
	<li>To measure fenestration diameter, trace a line through the longest diameter of each fenestration by selecting the line tool, and press &#8220;m&#8221; to measure the line, and &#8220;d&#8221; (draw) to permanently draw the line on the picture. This line is defined as the fenestration diameter. The length of the line will now be available in the results box of ImageJ (Screen shot 2).</li>
	<li>Measure all gaps which are larger holes in the LSEC cytoplasm over 250 nm, as well as fenestrations. Gaps are often artefacts reflecting poor perfusion or fixation, however gaps also can occur as a result of disease and toxicity. The data for gaps will be excluded from calculation of fenestration parameters later in the analysis.</li>
	<li>The area of gaps that are not circular should be calculated by tracing around the gaps and calculating their area on ImageJ.</li>
	<li>Paste the data from the results box in ImageJ into an EXCEL spreadsheet for further analysis.</li>
</ol>
9. Calculations
<ol>
	<li>Average fenestration diameter = average of all fenestration diameters (not including gaps, where gaps are &#8805; 250 nm).</li>
	<li>Fenestration area = &#960;r2,&#160;where r, the radius, is calculated from the individual fenestrations diameter (r=d/2), not including gaps.</li>
	<li>Porosity (%) = (&#931; (&#960;r2)/total area analyzed -&#160;&#931; (area of gaps&#160; =)) (&#181;m2)) &#215; 100.</li>
	<li>Fenestration frequency = total number of fenestrations/ (total area analyzed-&#931; (area of gaps) (&#181;m2).</li>
</ol>
10. Presentation of Fenestration Data
NOTE: Whenever possible, publications including quantification of fenestration data should include the following information
<ol>
	<li>Fenestration diameter, with a statement confirming what boundary diameters where used to define fenestrations (typically between 50 - 250 nm), Fenestration frequency (number/&#181;m2) and porosity (%).</li>
	<li>A statement confirming whether gaps have been included in the analysis, frequency distribution graph of fenestration diameter and the number of livers, blocks, images and fenestrations should be included in the analysis.</li>
</ol>

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The authors have nothing to disclose.

The ability to accurately and reproducibly measure the status of the liver sinusoidal endothelium is an important step in understanding the biology of these highly specialized cells. Newer techniques such as structured illumination microscopy32, atomic force microscopy33&#160;and d-STORM (direct Stochastic Optical Reconstrucion Microscopy) 34 will impart important information regarding the morphology of these cells in vitro but SEM remains the primary methodology to visualize and...

The liver sinusoidal endothelial cells (LSECs) are highly differentiated endothelial cells that line the wall of the hepatic sinusoid. LSECs are perforated with fenestrations that are non-diaphragmed, transcellular pores 50-250 nm in diameter. Up to 20% of the surface of LSECs is covered by fenestrations, which are usually in groups of tens to hundreds called sieve plates1-3 (Figure 1). Fenestrations allow transfer of plasma and nanosubstrates between blood and hepatocytes, creating a highly efficient ultrafiltration system. Fenestrations are dynamic structures - both their size and/or number can be altered in response to various physiological states, drugs, and disease. For example, fenestrations are larger in the fasted than in the fed state4; 2-di-iodoamphetamine increases fenestration number;5,6 and a reduction in size and number of fenestrations per cell occurs in ageing and many disease states7-13. Accurate measurement of the size and number of fenestrations is important for understanding how LSEC morphology is influenced by various disease, toxic, and physiological states; their impact on liver function; and for developing fenestration-modulating therapeutic interventions1.
The study of fenestrations is difficult. The diameter of fenestrations lies below the resolution of conventional light microscopy, so previously only observation using electron microscopy both in intact liver tissue or cultured LSECs has been possible. The scanning electron microscope (SEM) has most frequently been used to study fenestration size, frequency and porosity (the percentage of LSEC membrane that is perforated by fenestrations) because SEM allows for the observation of large areas of the endothelial surface and measurement of thousands, if not tens of thousands of fenestrations. Despite its utility, the results which are reported from SEM-based studies for LSEC parameters such as fenestration size, number, frequency and porosity vary widely in the literature (Table 1).
Fenestrations and sieve plates are fragile structures that contract, break, dilate or coalesce during specimen preparation, thus careful processing is needed to preserve their integrity. Elevated perfusion pressure14; incorrect osmolarity of the fixative and buffers15; inadequate fixation or fixation time; and speed of post-fixation dehydration and drying are all areas of processing for SEM that may produce artefacts that interfere with preservation of ultrastructure (Figure 2). Loss of fenestrations (&#8216;defenestration&#8217;) and fenestration shrinkage can occur as a result of poor fixation, resulting in reduced fenestration diameter and cell porosity. Methods to improve the preservation of specimens for SEM analysis have been described previously 15-17 and will be discussed here with additional tips on how to improve specimen preservation. The main goals of the specimen preservation are to remove blood from the sinusoids so that the surface of the LSEC can be visualized and to avoid LSEC damage from either high pressure or delayed fixation. Whole liver perfusion of fixative via the portal vein is the preferred method for liver fixation. As described in detail elsewhere16,18 perfusion must be undertaken at low pressure (eg 10 cm of H20) to avoid pressure-related perfusion artefacts and damage to the LSEC, typically manifested as large gaps within in the cell membrane. However, reasonable fixation can often be obtained using needle perfusion of liver biopsies from humans and animals, as described in detail elsewhere19. This technique involves directly injecting fixative into the tissue until blood is flushed out of the sample and the tissue is firm and fixed.&#160;Fixation of samples for electron microscopy needs to be performed as quickly as possible following cessation of blood flow to prevent ultrastructural changes occurring as a result of the livers extremely rapid autolytic processes.
We also present a method of image analysis that minimizes the inclusion of artefacts, and standardizes the measurement of fenestrations. Variation in the selection of sinusoids for micrographs, image analysis of artefacts, and measurement of cell area for porosity and fenestration frequency have led to major discrepancies in published results. A standardized approach for evaluation and measurement of fenestrations and the minimum requirements for data presentation have not been clearly addressed in the literature previously4,10,20-31.

<table border="0" cellpadding="0" cellspacing="0" width="658">
	<tbody>
		<tr height="40">
			<td height="40" style="height:40px;width:155px;">Name of material</td>
			<td style="width:100px;">Company</td>
			<td style="width:93px;">Catalogue Number</td>
			<td style="width:311px;">Comments</td>
		</tr>
		<tr height="42">
			<td height="42" style="height:42px;width:155px;">EM grade Glutaraldehyde</td>
			<td style="width:100px;">ProSciTech</td>
			<td style="width:93px;">C001</td>
			<td style="width:311px;">Store stock at -20 C until needed, avoid refreeze</td>
		</tr>
		<tr height="42">
			<td height="42" style="height:42px;width:155px;">Paraformaldehyde powder</td>
			<td style="width:100px;">Sigma Aldrich</td>
			<td>158127</td>
			<td style="width:311px;">Always prepare Paraformaldehyde fresh</td>
		</tr>
		<tr height="73">
			<td height="73" style="height:73px;width:155px;">Sodium Cacodylate powder</td>
			<td style="width:100px;">Sigma Aldrich</td>
			<td style="width:93px;">C0250</td>
			<td style="width:311px;">Prepare 0.2 M stock, pH 7.4 by dissolving powder in dH2O, used mostly at 0.1 M by preparing 1:2 dilution</td>
		</tr>
		<tr height="46">
			<td height="46" style="height:46px;width:155px;">Calcium Chloride</td>
			<td style="width:100px;">Sigma Aldrich</td>
			<td style="width:93px;">C1016</td>
			<td style="width:311px;">Prepare 1 M CaCl2by dissolving powder in dH2O</td>
		</tr>
		<tr height="63">
			<td height="63" style="height:63px;width:155px;">Osmium tretroxide</td>
			<td style="width:100px;">ProSciTech</td>
			<td style="width:93px;">C011</td>
			<td style="width:311px;">Wash ampoules in weak acid prior to use to avoid contamination. Prepare 2 % stock in glass bottle</td>
		</tr>
		<tr height="42">
			<td height="42" style="height:42px;width:155px;">Ethanol- Absolute</td>
			<td style="width:100px;">Sigma Aldrich</td>
			<td>459836&#160;</td>
			<td style="width:311px;">100 % Ethanol must be high grade and stored with Molecular Sieve</td>
		</tr>
		<tr height="40">
			<td height="40" style="height:40px;width:155px;">Other grades of Ethanol</td>
			<td style="width:100px;">Labtech</td>
			<td style="width:93px;">EL5</td>
			<td style="width:311px;">Prepare graded Ethanols with dH2O</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:155px;">Hexamethyldisilazane</td>
			<td style="width:100px;">Sigma Aldrich</td>
			<td>52619&#160;</td>
			<td style="width:311px;">Allow to reach room temperature before use</td>
		</tr>
		<tr height="105">
			<td height="105" style="height:105px;width:155px;">Cannulas</td>
			<td style="width:100px;">Terumo</td>
			<td style="width:93px;">TSROX1832C, TSROX2225C, TSROX2419C</td>
			<td style="width:311px;">18 G is suitable for most rats, 22 G is suitable for most mice, but it is good to have a few 24 G on hand in case of very small mice</td>
		</tr>
		<tr height="42">
			<td height="42" style="height:42px;width:155px;">Conductive Carbon tape</td>
			<td style="width:100px;">ProSciTech</td>
			<td>IA0201</td>
			<td style="width:311px;"></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:155px;">Carbon Paint</td>
			<td style="width:100px;">ProSciTech</td>
			<td>I003</td>
			<td style="width:311px;"></td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;width:155px;">Ketamine</td>
			<td style="width:100px;"></td>
			<td style="width:93px;"></td>
			<td style="width:311px;">Must be optained under licence</td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;width:155px;">Xylazine</td>
			<td style="width:100px;"></td>
			<td style="width:93px;"></td>
			<td style="width:311px;">Must be obtained under licence</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:155px;">Molecular Sieve</td>
			<td style="width:100px;">Sigma Aldrich</td>
			<td style="width:93px;">208647</td>
			<td style="width:311px;">Removes water from the 100 % Ethanol</td>
		</tr>
	</tbody>
</table>

a standardized method for the analysis of liver sinusoidal endothelial cells and their fenestrations by scanning electron microscopy

Liver sinusoidal endothelial cells are the gateway to the liver, their transcellular fenestrations allow the unimpeded transfer of small and dissolved substances from the blood into the liver parenchyma for metabolism and processing. Fenestrations are dynamic structures - both their size and/or number can be altered in response to various physiological states, drugs, and disease, making them an important target for modulation. An understanding of how LSEC morphology is influenced by various disease, toxic, and physiological states and how these changes impact on liver function requires accurate measurement of the size and number of fenestrations. In this paper, we describe scanning electron microscopy fixation and processing techniques used in our laboratory to ensure reproducible specimen preparation and accurate interpretation. The methods include perfusion fixation, secondary fixation and dehydration, preparation for the scanning electron microscope and analysis. Finally, we provide a step by step method for standardized image analysis which will benefit all researchers in the field.

Initial visualization at low magnification by scanning electron microscopy reveals a flat surface of the liver specimen with an exposed area large enough to observe many large liver vessels and sinusoids (Figure 1A). Ensuring correct liver block placement on the mounting stub is essential for obtaining clear images of the sinusoids and the Glisson&#8217;s capsule of the liver should be avoided for this reason (Figure 1B). Increasing the magnification allows closer inspection of the dense...

Watch this Scientific Journal Video about A Standardized Method for the Analysis of Liver Sinusoidal Endothelial Cells and Their Fenestrations by Scanning Electron Microscopy at JoVE.com

A Standardized Method for the Analysis of Liver Sinusoidal Endothelial Cells and Their Fenestrations by Scanning Electron Microscopy

The fenestrated liver sinusoidal endothelial cell is a biologically important filter system that is highly influenced by various diseases, toxins, and physiological states. These changes significantly impact on liver function. We describe methods for the standardisation of the measurement of the size and number of fenestrations in these cells.

Liver sinusoidal endothelial cells are the gateway to the liver, their transcellular fenestrations allow the unimpeded transfer of small and dissolved ...

a-standardized-method-for-analysis-liver-sinusoidal-endothelial-cells

Research

JoVE Journal

Biology

14.3K visualizações.  University of Sydney and Concord Hospital. The fenestrated liver sinusoidal endothelial cell is a biologically important filter system that is highly influenced by various diseases, toxins, and physiological states. These changes significantly impact on liver function. We describe methods for the standardisation of the measurement of the size and number of fenestrations in these cells.

Vídeo: A Standardized Method for the Analysis of Liver Sinusoidal Endothelial Cells and Their Fenestrations by Scanning Electron Microscopy

In Vivo 4-Dimensional Tracking of Hematopoietic Stem and Progenitor Cells in Adult Mouse Calvarial Bone Marrow

Through a delicate balance between quiescence and proliferation, self renewal and production of differentiated progeny, hematopoietic stem cells (HSCs) maintain the turnover of all mature blood cell lineages. The coordination of the complex signals leading to specific HSC fates relies upon the interaction between HSCs and the intricate bone marrow microenvironment, which is still poorly understood[1-2].
We describe how by combining a newly developed specimen holder for stable animal positioning with multi-step confocal and two-photon in vivo imaging techniques, it is possible to obtain high-resolution 3D stacks containing HSPCs and their surrounding niches and to monitor them over time through multi-point time-lapse imaging. High definition imaging allows detecting ex vivo labeled hematopoietic stem and progenitor cells (HSPCs) residing within the bone marrow. Moreover, multi-point time-lapse 3D imaging, obtained with faster acquisition settings, provides accurate information about HSPC movement and the reciprocal interactions between HSPCs and stroma cells.
Tracking of HSPCs in relation to GFP positive osteoblastic cells is shown as an exemplary application of this method. This technique can be utilized to track any appropriately labeled hematopoietic or stromal cell of interest within the mouse calvarium bone marrow space.

In Vivo 4-Dimensional Tracking of Hematopoietic Stem and Progenitor Cells in Adult Mouse Calvarial Bone Marrow

The nature of the interactions between hematopoietic stem and progenitor cells (HSPCs) and bone marrow niches is poorly understood. Custom hardware modifications and a multi-step acquisition protocol allow the use of two-photon and confocal microscopy to image ex vivo labeled HSPCs homed within bone marrow areas, tracking interactions and movement.

Through a delicate balance between quiescence and proliferation, self renewal and production of differentiated progeny, hematopoietic stem cells (HSCs) ...

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 ...

Scanning Electron Microscopy of Macerated Tissue to Visualize the Extracellular Matrix

Fibrosis is a component of all forms of heart disease regardless of etiology, and while much progress has been made in the field of cardiac matrix biology, there are still major gaps related to how the matrix is formed, how physiological and pathological remodeling differ, and most importantly how matrix dynamics might be manipulated to promote healing and inhibit fibrosis. There is currently no treatment option for controlling, preventing, or reversing cardiac fibrosis. Part of the reason is likely the sheer complexity of cardiac scar formation, such as occurs after myocardial infarction to immediately replace dead or dying cardiomyocytes. The extracellular matrix itself participates in remodeling by activating resident cells and also by helping to guide infiltrating cells to the defunct lesion. The matrix is also a storage locker of sorts for matricellular proteins that are crucial to normal matrix turnover, as well as fibrotic signaling. The matrix has additionally been demonstrated to play an electromechanical role in cardiac tissue. Most techniques for assessing fibrosis are not qualitative in nature, but rather provide quantitative results that are useful for comparing two groups but that do not provide information related to the underlying matrix structure. Highlighted here is a technique for visualizing cardiac matrix ultrastructure. Scanning electron microscopy of decellularized heart tissue reveals striking differences in structure that might otherwise be missed using traditional quantitative research methods.

Shown here is a method for visualizing extracellular matrix ultrastructure in decellularized cardiac tissues.

Fibrosis is a component of all forms of heart disease regardless of etiology, and while much progress has been made in the field of cardiac matrix ...

Studying the Supramolecular Organization of Photosynthetic Membranes within Freeze-fractured Leaf Tissues by Cryo-scanning Electron Microscopy

Cryo-scanning electron microscopy (SEM) of freeze-fractured samples allows investigation of biological structures at near native conditions. Here, we describe a technique for studying the supramolecular organization of photosynthetic (thylakoid) membranes within leaf samples. This is achieved by high-pressure freezing of leaf tissues, freeze-fracturing, double-layer coating and finally cryo-SEM imaging. Use of the double-layer coating method allows acquiring high magnification (&#62;100,000X) images with minimal beam damage to the frozen-hydrated samples as well as minimal charging effects. Using the described procedures we investigated the alterations in supramolecular distribution of photosystem and light-harvesting antenna protein complexes that take place during dehydration of the resurrection plant Craterostigma pumilum, in situ.

Here we describe a procedure for studying freeze-fractured plant tissues. High-pressure frozen leaf samples are freeze-fractured and double-layer coated, yielding well preserved frozen-hydrated samples that are imaged using the cryo-scanning electron microscope at high magnifications with minimal beam damage.

Cryo-scanning electron microscopy (SEM) of freeze-fractured samples allows investigation of biological structures at near native conditions. Here, we ...

Scanning Electron Microscopy (SEM) Protocols for Problematic Plant, Oomycete, and Fungal Samples

Common problems in the processing of biological samples for observations with the scanning electron microscope (SEM) include cell collapse, treatment of samples from wet microenvironments and cell destruction. Using young floral tissues, oomycete cysts, and fungi spores (Agaricales) as examples, specific protocols to process delicate samples are described here that overcome some of the main challenges in sample treatment for image capture under the SEM.
Floral meristems fixed with FAA (Formalin-Acetic-Alcohol) and processed with the Critical Point Dryer (CPD) did not display collapsed cellular walls or distorted organs. These results are crucial for the reconstruction of floral development. A similar CPD-based treatment of samples from wet microenvironments, such as the glutaraldehyde-fixed oomycete cysts, is optimal to test the differential growth of diagnostic characteristics (e.g., the cyst spines) on different types of substrates. Destruction of nurse cells attached to fungi spores was avoided after rehydration, dehydration, and the CPD treatment, an important step for further functional studies of these cells.
The protocols detailed here represent low-cost and rapid alternatives for the acquisition of good-quality images to reconstruct growth processes and to study diagnostic characteristics.

Scanning Electron Microscopy SEM Protocols for Problematic Plant, Oomycete, and Fungal Samples

Problems in the processing of biological samples for scanning electron microscopy observation include cell collapse, treatment of samples from wet microenvironments and cell destruction. Low-cost and relatively rapid protocols suited for preparing challenging samples such as floral meristems, oomycete cysts, and fungi (Agaricales) are compiled and detailed here.

Common problems in the processing of biological samples for observations with the scanning electron microscope (SEM) include cell collapse, treatment of ...

Standardized Method for High-throughput Sterilization of Arabidopsis Seeds

Arabidopsis thaliana (Arabidopsis) seedlings often need to be grown on sterile media. This requires prior seed sterilization to prevent the growth of microbial contaminants present on the seed surface. Currently, Arabidopsis seeds are sterilized using two distinct sterilization techniques in conditions that differ slightly between labs and have not been standardized, often resulting in only partially effective sterilization or in excessive seed mortality. Most of these methods are also not easily scalable to a large number of seed lines of diverse genotypes. As technologies for high-throughput analysis of Arabidopsis continue to proliferate, standardized techniques for sterilizing large numbers of seeds of different genotypes are becoming essential for conducting these types of experiments. The response of a number of Arabidopsis lines to two different sterilization techniques was evaluated based on seed germination rate and the level of seed contamination with microbes and other pathogens. The treatments included different concentrations of sterilizing agents and times of exposure, combined to determine optimal conditions for Arabidopsis seed sterilization. Optimized protocols have been developed for two different sterilization methods: bleach (liquid-phase) and chlorine (Cl2) gas (vapor-phase), both resulting in high seed germination rates and minimal microbial contamination. The utility of these protocols was illustrated through the testing of both wild type and mutant seeds with a range of germination potentials. Our results show that seeds can be effectively sterilized using either method without excessive seed mortality, although detrimental effects of sterilization were observed for seeds with lower than optimal germination potential. In addition, an equation was developed to enable researchers to apply the standardized chlorine gas sterilization conditions to airtight containers of different sizes. The protocols described here allow easy, efficient, and inexpensive seed sterilization for a large number of Arabidopsis lines.

The objective of this study was to determine the effects of bleach and chlorine gas sterilization on seed germination of a range of Arabidopsis genotypes grown on sterile media. Optimized sterilization protocols have been developed to prevent the growth of microbial contaminants while providing satisfactory seed survival.

Arabidopsis thaliana (Arabidopsis) seedlings often need to be grown on sterile media. This requires prior seed sterilization to prevent the growth of ...

Preparation of Prokaryotic and Eukaryotic Organisms Using Chemical Drying for Morphological Analysis in Scanning Electron Microscopy (SEM)

Scanning electron microscopy (SEM) is a widely available technique that has been applied to study biological specimens ranging from individual proteins to cells, tissues, organelles, and even whole organisms. This protocol focuses on two chemical drying methods, hexamethyldisilazane (HMDS) and t-butyl alcohol (TBA), and their application to imaging of both prokaryotic and eukaryotic organisms using SEM. In this article, we describe how to fix, wash, dehydrate, dry, mount, sputter coat, and image three types of organisms: cyanobacteria (Toxifilum mysidocida, Golenkina sp., and an unknown sp.), two euglenoids from the genus Monomorphina (M. aenigmatica and M. pseudopyrum), and the fruit fly (Drosophila melanogaster). The purpose of this protocol is to describe a fast, inexpensive, and simple method to obtain detailed information about the structure, size, and surface characteristics of specimens that can be broadly applied to a large range of organisms for morphological assessment. Successful completion of this protocol will allow others to use SEM to visualize samples by applying these techniques to their system.

Preparation of Prokaryotic and Eukaryotic Organisms Using Chemical Drying for Morphological Analysis in Scanning Electron Microscopy SEM

SEM analysis is an effective method to aid in species identification or phenotypic discrimination. This protocol describes the methods for examining specific morphological details of three representative types of organisms and would be broadly applicable to examining features of many organismal and tissue types.

Scanning electron microscopy (SEM) is a widely available technique that has been applied to study biological specimens ranging from individual proteins to ...

Biological Sample Preparation by High-pressure Freezing, Microwave-assisted Contrast Enhancement, and Minimal Resin Embedding for Volume Imaging

The described sample preparation technique is designed to combine the best quality of ultrastructural preservation with the most suitable contrast for the imaging modality in a focused ion beam scanning electron microscope (FIB-SEM), which is used to obtain stacks of sequential images for 3D reconstruction and modelling. High-pressure freezing (HPF) allows close to native structural preservation, but the subsequent freeze substitution often does not provide sufficient contrast, especially for a bigger specimen, which is needed for high-quality imaging in the SEM required for 3D reconstruction. Therefore, in this protocol, after the freeze substitution, additional contrasting steps are carried out at room temperature. Although these steps are performed in a microwave, it is also possible to follow traditional bench processing, which requires longer incubation times.The subsequent embedding in minimal amounts of resin allows for faster and more precise targeting and preparation inside the FIB-SEM. This protocol is especially useful for samples that require preparation by high-pressure freezing for a reliable ultrastructural preservation but do not gain enough contrast during the freeze substitution for volume imaging using FIB-SEM. In combination with the minimal resin embedding, this protocol provides an efficient workflow for the acquisition of high-quality volume data.

Here we present a protocol for combining two sample processing techniques, high-pressure freezing and microwave-assisted sample processing, followed by minimal resin embedding for acquiring data with a focused ion beam scanning electron microscope (FIB-SEM). This is demonstrated using a mouse tibial nerve sample and Caenorhabditis elegans.

The described sample preparation technique is designed to combine the best quality of ultrastructural preservation with the most suitable contrast for the ...

The CryoAPEX Method for Electron Microscopy Analysis of Membrane Protein Localization Within Ultrastructurally-Preserved Cells

Key cellular events like signal transduction and membrane trafficking rely on proper protein location within cellular compartments. Understanding precise subcellular localization of proteins is thus important for answering many biological questions. The quest for a robust label to identify protein localization combined with adequate cellular preservation and staining has been historically challenging. Recent advances in electron microscopy (EM) imaging have led to the development of many methods and strategies to increase cellular preservation and label target proteins. A relatively new peroxidase-based genetic tag, APEX2, is a promising leader in cloneable EM-active tags. Sample preparation for transmission electron microscopy (TEM) has also advanced in recent years with the advent of cryofixation by high pressure freezing (HPF) and low-temperature dehydration and staining via freeze substitution (FS). HPF and FS provide excellent preservation of cellular ultrastructure for TEM imaging, second only to direct cryo-imaging of vitreous samples. Here we present a protocol for the cryoAPEX method, which combines the use of the APEX2 tag with HPF and FS. In this protocol, a protein of interest is tagged with APEX2, followed by chemical fixation and the peroxidase reaction. In place of traditional staining and alcohol dehydration at room temperature, the sample is cryofixed and undergoes dehydration and staining at low temperature via FS. Using cryoAPEX, not only can a protein of interest be identified within subcellular compartments, but also additional information can be resolved with respect to its topology within a structurally preserved membrane. We show that this method can provide high enough resolution to decipher protein distribution patterns within an organelle lumen, and to distinguish the compartmentalization of a protein within one organelle in close proximity to other unlabeled organelles. Further, cryoAPEX is procedurally straightforward and amenable to cells grown in tissue culture. It is no more technically challenging than typical cryofixation and freeze substitution methods. CryoAPEX is widely applicable for TEM analysis of any membrane protein that can be genetically tagged.

This protocol describes the cryoAPEX method, in which an APEX2-tagged membrane protein can be localized by transmission electron microscopy within optimally-preserved cell ultrastructure.

Key cellular events like signal transduction and membrane trafficking rely on proper protein location within cellular compartments. Understanding precise ...

Array Tomography Workflow for the Targeted Acquisition of Volume Information using Scanning Electron Microscopy

Electron microscopy is applied in biology and medicine for imaging of cellular and structural details at nanometer resolution. Historically, Transmission Electron Microscopy (TEM) provided insight into cell ultrastructure, but in the recent decade, the development of modern Scanning Electron Microscopes (SEM) has changed the way of looking inside the cells. Even though the resolution of TEM is superior when protein-level structural details are needed, SEM-resolution is sufficient for the majority of the organelle-level cell biology-related questions. The advancement in technology enabled automatic volume acquisition solutions such as Serial block-face imaging (SBF-SEM) and Focused ion beam SEM (FIB-SEM). Nevertheless, to this day, these methods remain inefficient when the identification and navigation to areas of interest are crucial. Without the means for precise localization of target areas before imaging, operators need to acquire much more data than they need (in SBF-SEM), or, even worse, prepare many grids and image them all (in TEM). We propose the strategy of "lateral screening" using Array Tomography in SEM, which facilitates the localization of areas of interest, followed by automated imaging of the relevant fraction of the total sample volume. Array tomography samples are conserved during imaging, and they can be arranged into section libraries ready for repeated imaging. Several examples are shown in which lateral screening enables us to analyze structural details that are incredibly challenging to access with any other method.

We describe the preparation of ribbons of serial sections and their collection on large transfer support for use as Array Tomography samples, along with automated imaging procedures in a scanning electron microscope. The protocol allows screening, retrieval, and targeted imaging of local, rare events, and the acquisition of large data volumes.

Electron microscopy is applied in biology and medicine for imaging of cellular and structural details at nanometer resolution. Historically, Transmission ...