The authors extend thanks to colleagues at the University of Arkansas for Medical Sciences (Jerry Ware and Sung W. Rhee), the University of Pennsylvania (Tim Stalker and Lawrence Brass), the University of Kentucky (Sidney W. Whiteheart and Smita Joshi), and the National Institute of Bioimaging and Bioengineering of the National Institutes of Health (Richard D. Leapman and Maria A. Aronova) from whom we have learned much. The authors express appreciation to the American Heart Association and the National Heart Lung and Blood Institute of the National Institutes of Health (R01 HL119393, R56 HL119393, R01 155519 to BS and subawards from NIH grants R01 HL146373 and R35 HL150818) for financial support.

Experiments were reviewed and approved by the Institutional Animal Care and Use Committee (IACUC) at the University of Arkansas for Medical Sciences. Here, 8 - 12-week-old, wild-type male and female C57BL/6 mice were used. These mice are young adults with little accumulation of fat. The same procedures are applicable to mice mutants for various proteins important to hemostasis, such as von Willebrand factor or platelet glycoprotein VI (GPVI)5,6. All equipment, surgical instruments, reagents, and other materials used are shown in Figure 1 and listed in the Table of Materials.
1. Jugular vein/femoral artery puncture wound1,7
<ol>
	<li>Anesthetization of the mouse
	<ol>
		<li>Place the mouse in the induction chamber and turn the isoflurane vaporizer to a high flow rate setting (500 mL/min, 3% isoflurane). Isoflurane is chosen because it has little effect on blood vessel diameter. 
		CAUTION: Long-term exposure to isoflurane can cause adverse health effects. The use of a low-flow vaporizer, anesthesia gas capture canisters, and good room ventilation can greatly reduce exposure.</li>
		<li>When the mouse appears asleep, pinch one of the mouse&#39;s toes to see if the mouse reacts. If so, place the mouse back in the induction chamber for a few minutes and then try the pinch test again. If the mouse does not react, place it in a supine position on the heating plate (previously covered with aluminum foil).</li>
		<li>Turn the vaporizer down to a low flow rate (100 mL/min, 1.75% isoflurane). Place the mouse&#39;s nose in the nose cone. Extend each limb and tape down with small pieces of adhesive tape.</li>
		<li>Insert the rectal temperature probe and set the temperature to 37 &#176;C on the temperature controller. Tape down the temperature probe cord to prevent the probe from accidentally being pulled out.</li>
	</ol>
	</li>
	<li>Mouse incision
	<ol>
		<li>Using the clippers, shave the mouse&#39;s neck and chest (for jugular vein) or inner right leg (for femoral artery). Wipe away the fur clippings with an alcohol pad. If needed, go over the area again with the clippers to remove any remaining fur.</li>
		<li>Wipe the area again with a fresh alcohol pad to clear away all the clippings. It is very important that the area is as clean as possible. Confirm the surgical plane of anesthesia via a toe-pinch.</li>
		<li>Using scissors and blunt forceps, lift and cut the skin over the right jugular vein. Cut a round window in the skin large enough to expose the jugular vein and some of the surrounding tissue.</li>
	</ol>
	</li>
	<li>Cleaning of the jugular vein (same principles apply to the femoral artery)
	<ol>
		<li>Using the blunt dissection technique, gently push aside adipose and lymph nodes as well as connective tissue. 
		NOTE: Some general cleaning around the vessel takes place at this step; however, careful, thorough cleaning will take place under the microscope after the jugular vein is fixed in paraformaldehyde for 24 h.</li>
		<li>Fill a 20 mL syringe with normal saline warmed to 37 &#176;C. Load the syringe into the syringe infusion pump. Attach the 27G butterfly needle and associated tubing to the syringe.</li>
		<li>Set the syringe pump to a flow rate of 0.5 mL/min. The gentle stream of saline will wash away blood from the punctured vein/artery. Position the stream of saline a few mm to the side of the puncture site, not directly over it.</li>
	</ol>
	</li>
	<li>Puncturing the jugular vein/femoral artery
	<ol>
		<li>Start the timer. Obtain a 30G hypodermic needle (nominal 300 &#181;m diameter) for the jugular vein and a 33G hypodermic needle (200 &#181;m diameter) for the femoral artery. The use of a smaller needle gives only a slightly longer bleeding time for the higher-pressure arterial situation.</li>
		<li>Turn the bevel facing up. Position the needle at a 25&#176; angle over the jugular vein/femoral artery. Note the time.</li>
		<li>Quickly puncture the vessel, making sure to pierce the top of the vessel only. Be careful not to puncture the bottom of the vessel as well. Note the time when the bleeding stops completely.</li>
		<li>Wait for the appropriate amount of time to obtain thrombus formation (Figure 2A) before starting perfusion fixation (i.e., either 1, 5, 10, or 20 min to obtain different stages of thrombus development). Euthanize the animal by exsanguination or follow local institutional guidelines for euthanasia.</li>
	</ol>
	</li>
	<li>Performing transcardial perfusion fixation
	<ol>
		<li>Set up the peristaltic pump before the surgery begins. Place two 50 mL conical tubes in a test tube rack. Fill one tube with 4% paraformaldehyde fixative solution prepared in sodium cacodylate buffer solution (0.2 M sodium cacodylate, 0.15 M sodium chloride at pH 7.4) for jugular puncture studies and fill the other tube with sodium cacodylate buffer solution. 
		NOTE: The conditions detailed here are sufficient for the low-pressure case of the jugular and are then modified for the higher-pressure case of an artery. For the higher-pressure system studies with arteries, i.e., the femoral artery, the paraformaldehyde fixative is supplemented with 0.1% glutaraldehyde, and external drip fixation is combined with perfusion fixation.</li>
		<li>Place the uptake end of the pump tubing into the tube containing the buffer solution. Attach a 27G butterfly needle with associated tubing to the output end of the pump tubing and place the needle in a clean beaker.</li>
		<li>Set the peristaltic pump to a flow rate of 10 mL/min. Turn on the perfusion pump.</li>
		<li>When buffer solution begins to flow from the needle into the beaker, turn the pump off. The tubing and needle are now primed for the transcardial perfusion.</li>
		<li>Expose the chest cavity by making a midline cut in the skin from the top of the sternum to a few millimeters past the bottom of the sternum using dissecting scissors and blunt forceps. Make transverse incisions along the base of the rib cage, starting at the midline and cutting from medial to lateral on each side.</li>
		<li>Immediately before starting the transcardial perfusion, quickly cut open the rib cage and reflect each side to expose the heart.</li>
		<li>Turn on the peristaltic pump. Place the tip of the 27G butterfly needle into the base of the left ventricle.</li>
		<li>While holding the needle secure in the left ventricle, cut a slit in the right auricle using sharp dissecting scissors to allow blood and perfusion fluid to flow out. Note the time on the timer.</li>
		<li>Perfuse with buffer solution for 3 min to flush the blood out of the circulatory system. While still holding the needle secure in the left ventricle, turn off the peristaltic pump and move the uptake end of the peristaltic pump tubing over to the 50 mL tube containing the fixative solution.</li>
		<li>Note the time on the timer. Turn the peristaltic pump on again and perfuse with the fixative solution for 7 min. Turn the peristaltic pump off.</li>
		<li>Take the needle out of the left ventricle and place it in a waste beaker. Place the uptake end of the pump tubing into a beaker of distilled water.</li>
		<li>Turn the peristaltic pump on and let approximately 20 mL of water flow through the tubing and out the 27G needle into the waste beaker to clean the tubing. Be sure to let all the water exit the tubing and needle before turning the pump off.</li>
	</ol>
	</li>
</ol>
2. Collection of samples for electron microscopy
<ol>
	<li>After perfusion fixation supplemented with an initial 2 min external fixative in the case of femoral artery puncture wounds, carefully dissect the portion of the jugular vein/femoral artery containing the puncture site and thrombus (Figure 2A). With sharp scissors, make a diagonal cut across the distal end of the vessel segment and make a straight cut across the proximal end of the vessel segment. 
	NOTE: Making cuts in this manner helps to orient the vessel later and to identify the direction of blood flow in the vessel.</li>
	<li>Submerge the vessel segment in fresh, room temperature, 4% paraformaldehyde in sodium cacodylate buffer solution (0.2 M sodium cacodylate, 0.15 M sodium chloride at pH 7.4) and place at room temperature for 1 h.</li>
	<li>Transfer the fixed tissue to 4 &#176;C for 24 h. The next day, take a 35 mm culture dish containing a ~5 mm thick silicone mat (prepared in advance according to the manufacturer&#39;s instructions). Pour enough sodium cacodylate buffer solution into the dish to submerge the fixed vein segment.</li>
	<li>While viewing under a light microscope (~15x magnification), carefully transfer the tissue to the 35 mm dish containing buffer solution and pin each end of the vessel to the silicone mat using stainless steel minutien pins and fine forceps.</li>
	<li>Clean the jugular vein/femoral artery segment carefully using a pair of micro scissors and very fine forceps.
	<ol>
		<li>For jugular vein only: Remove one of the pins and, using the micro scissors, cut the vessel wall opposite the puncture/thrombus site lengthwise. Gently open the vessel and, using 4 to 5 pins (each cut in half with wire cutters), pin it to the silicone mat with the intraluminal side facing up.</li>
	</ol>
	</li>
	<li>Aspirate the buffer solution, quickly replace it with 1% paraformaldehyde solution in sodium cacodylate buffer solution and place the lid on the dish. Do not let the tissue dry out at any time. Always keep it submerged in fluid.</li>
	<li>Store the tissue at 4 &#176;C until it is time to process it for electron microscopy (Figure 2B). Process some of the jugular vein/femoral artery puncture wound samples for imaging by SBF-SEM4 and others for imaging by WA-TEM8,9. The WA-TEM steps are described in detail below. 
	CAUTION: Carry out all work involving formaldehyde, propylene oxide, OsO4, and 2,4,6-Tris(dimethylaminomethyl)phenol (DMP-30) in a chemical fume hood. These are hazardous chemicals. Exposure to OsO4 can cause blindness and death. 
	NOTE: All rinse and stain volumes are at least 2x the volume of the sample. The 1x phosphate-buffered saline (PBS) can be substituted for the 0.1 M sodium cacodylate buffer. During the fixation procedure, it is very important never to let the samples dry out, as this would significantly affect the ultra-structure. Use polypropylene tubes.</li>
</ol>
3. Preparation of sample for montaged wide-area transmission electron microscopy (WA-TEM)
NOTE: This step represents a decision point at which the investigator is committing to preparing for WA-TEM. This preparative procedure does not support SBF-SEM. For SBF-SEM volume EM, all staining must be done before embedding in plastic. Please see4 for an SBF-SEM preparation protocol.
<ol>
	<li>Wash the tissue sample in the 35 mm dish 2x with Hank&#39;s balanced salt solution (HBSS) or PBS at room temperature (RT). Keep the sample pinned, allowing for tracking the orientation of the sample through these preparative steps.</li>
	<li>Fix with 0.8 mL of 0.05% malachite green/ 2.5% glutaraldehyde/ 0.1 M sodium cacodylate (pH 6.7 - 7.0) for 20 min on ice.</li>
	<li>Remove fixative and wash sample 4x for 5 min each with 0.1 M sodium cacodylate. Remove 0.1 M sodium cacodylate.</li>
	<li>Postfix with 0.8 mL of 0.8% K3Fe(CN)6 or K4Fe(CN)6 / 1.0% OsO4 K3Fe(CN)6 in 0.1 M sodium cacodylate. Place a cover over the dish to keep the light out (OsO4 is light-sensitive). Stain for 20 min to 1 h at RT.</li>
	<li>Remove the OsO4 and rinse 4x for 5 min each with 0.1 M sodium cacodylate buffer. Remove sodium cacodylate buffer and stain with 1% tannic acid for 20 min on ice.</li>
	<li>Remove tannic acid and wash for 5 min with 0.1 M sodium cacodylate buffer 1x and 2x for 5 min each with molecular-grade distilled (DI) water.</li>
	<li>Stain with 0.5% uranyl acetate (UA) for 1 h RT or overnight at 4 &#176;C in the dark.</li>
	<li>Remove the uranyl acetate and rinse 5x for 5 min each with molecular-grade distilled water. Before the last wash and while the sample is still submerged in DI water, cut the silicone mat around the pinned sample with a razor blade. Lift this small piece of silicone mat out of the plate and place it into a polypropylene tube. 
	NOTE: It is important the sample remains pinned to the silicone piece while in the tube. Switching to polypropylene tubes in this step is necessary because the propylene oxide used in subsequent steps will degrade the polystyrene of the 35 mm plates.</li>
	<li>After removing the last buffer, briefly rinse 1x with 25% ethanol. Discard the 25% ethanol and incubate with 25% ethanol for 3 min on ice.</li>
	<li>Remove the 25% ethanol. Briefly rinse 1x with 50% ethanol. Discard the 50% ethanol and incubate with 50% ethanol for 3 min on ice.</li>
	<li>Remove the 50% ethanol and briefly rinse 1x with 75% ethanol. Discard the 75% ethanol and incubate with 75% ethanol for 3 min on ice.</li>
	<li>Remove the 75% ethanol and briefly rinse 1x with 95% ethanol. Discard the 95% ethanol and incubate with 95% ethanol for 3 min on ice.</li>
	<li>Remove the 95% ethanol and wash 3x, at least 10 min each time, with 100% ethanol (200 proof). Remove 100% ethanol and add propylene oxide (PO). Rinse 3x at least 10 min each, with PO.</li>
	<li>Prepare the resin mix as described below.
	<ol>
		<li>Warm the resin components at 60 &#176;C to liquify fully. Thoroughly mix 12.5 mL of Embed-218, 7.5 mL of Araldite 502, and 27.5 mL of DDSA together in a 50 mL tube at 60 &#176;C. This mixture can be kept for 6 months at 4 &#176;C. Warm it to 60 &#176;C before using. Aliquot the required amount prior to use.</li>
	</ol>
	</li>
	<li>Embed the samples as described below.
	<ol>
		<li>Place an aliquot of the resin mix in a tube and add 20 &#181;L/mL of DMP-30 activator. Mix thoroughly by rotating at 60 &#176;C; the color should darken.</li>
		<li>Dilute this aliquot of activated resin by 50% in PO and allow the resin and PO to mix completely.</li>
		<li>Remove the last PO rinse from the sample-containing tube and replace it with the 50% resin/PO mixture, nearly filling the tube completely. Rotate the sample overnight at room temperature.</li>
		<li>The following day, mix a new aliquot of resin (1 mL per sample) with 20 &#181;L/mL of DMP-30.</li>
	</ol>
	</li>
	<li>Place the sample in a polypropylene dish with a small volume of 50% resin/PO mix from the sample tube under a dissecting microscope (~6x magnification) and carefully take the pins out of the tissue. Transfer only the tissue to another polypropylene dish containing a small amount of 100% resin with a DMP-30 activator.</li>
	<li>Fill a silicone embedding mold with the 100% resin with the DMP-30 activator and carefully place the sample in the mold. Place mold in oven and bake at 60 &#176;C for at least 48 h . The prepared sample is shown in Figure 2B.</li>
	<li>Perform montaged wide-area transmission electron microscopy (WA-TEM) imaging as described in10,11. See Figure 3 for a depiction of the process going from an initial puncture to a 3-dimensionally rendered 1-min thrombus. The inclusion of a detailed protocol for montaged WA-TEM electron microscopy imaging is not included here and is beyond the scope of the present protocols on sample preparation. 
	NOTE: WA-TEM is an underappreciated capability of most modern transmission electron microscopes with a computerized stage. Shareware software supporting these functions10,11 is available from the group of Dr. David Mastronarde, University of Colorado, Boulder (http://bio3d.colorado.edu/SerialEM/, http://bio3d.colorado.edu/imod/.)</li>
</ol>

Puncture Wound Hemostasis and Montaged Electron Microscopy 

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The authors have no conflicts of interest related to this study.

We present a detailed puncture wound procedure for producing hemostatic thrombi in jugular veins and femoral arteries, their in situ perfusion fixation, and sample processing for montaged wide-area transmission electron microscopy. The overall procedures are useful for generating hemostatic thrombi for ultrastructural analysis and for comparing bleeding times in experimental mice, for example, mice treated with different types and dosages of pharmaceuticals. It is also useful for comparing bleeding times in cont...

The formation of a puncture wound thrombus that leads to bleeding cessation is one of the most essential events in life1. Yet despite that essentiality, knowledge of what occurs structurally during thrombus formation, be it in a vein, an artery, an atherosclerotic event, or an occlusive clot, has been limited by resolution and imaging depth. Conventional light microscopy is limited in depth when compared to a fully formed puncture wound thrombus, 200 to 300 &#181;m in Z1,&#160;and in resolution level when compared to the size of platelet organelles and their spacing, often less than 30 nm2. Two-photon light microscopy can yield the needed depth of imaging but does not improve resolution significantly. The most recent advances in light microscopy, for example, super-resolution techniques, are still resolution limited, in practice ~20 nm in XY and twice that in Z, and depth limited, no more than conventional light microscopy. Furthermore, super-resolution light microscopy, like much of research light microscopy, is based on fluorescence microscopy, a technique that is inherently biased to a small set of candidate proteins for which good antibodies exist or good tagged constructs3. In conclusion, conventional scanning electron microscopy can, at most, visualize the surface of the forming platelet-rich thrombus.
To overcome these technical limitations to characterizing thrombus structure, we had three goals. First, reproducibly produce a defined puncture wound in a mouse vein or artery that could then be readily stabilized in situ by chemical fixation. Second, apply a preparative procedure that emphasizes membrane preservation, a goal consistent with the aim of defining the position of individual platelets within the forming thrombus. Third, use an unbiased visualization technique that, in a single image, could be scaled between nanometer to near millimeter scale.
Montaged, wide-area electron microscopy was chosen as a major end visualization technique for a single important reason: in electron microscope imaging, one sees a vast array of features within a cell that outlines its organelles and features within the organelles. Small objects such as ribosomes can be recognized. This range of features is seen because the electron-dense heavy metal stains, uranyl, lead, and osmium, that are used for electron microscopy to yield contrast bind to a wide range of molecules. In an electron microscope image, one sees much of what is there, while with immunofluorescence and protein tagging approaches, one only sees what lights up. This means, for example, the antigen sites present on a given individual protein species. In the case of a tagged molecule, often a protein, it is the site(s) where that protein is. All other molecules are dark and not lit up. However, is this choice of electron microscopy practical? A puncture wound thrombus has a size of 300 by 500 &#181;m and, at a pixel size of 3 nm, that is an image of 100,000 by 167,000 pixels. A high-quality electron microscope camera has 4000 by 4000 pixels. That means that approximately 1000 frames must be stitched/blended to give a single image. That is a possibility that has been present in most electron microscopes manufactured in the last 15 years. The microscope stage is computerized, and the images can be stitched together with a computer. That is the rationale that led to choices underlying the formulation of the presented protocol.
Conclusively, we present below a series of steps that give a reproducible wound in the vein or artery of mice that then, following in situ fixation steps and later embedding steps, can be visualized by montaged, wide-area transmission electron microscopy at nm scale and in the stitched image visualized at near mm scale, the scale of the actual in situ fixed thrombus. Scalability of this kind is required for understanding thrombus formation as both a problem in hematology/health and as a developmental biology system in which platelets are the major cell type. These advances deliver a major virtue of electron microscopy, namely, one sees what is there, not only what lights up. For a detailed protocol on the preparation of samples for serial block face scanning electron microscopy (SBF-SEM), the reader is directed to a recent article by Joshi et al.4.

<table><tbody><tr><td>0.9% Normal Saline Solution</td><td>Medline</td><td>BHL2F7123HH</td><td></td></tr><tr><td>27G x 3/4 EXELint scalp vein set</td><td>Medline</td><td>NDA26709</td><td></td></tr><tr><td>30G x 1/2 EXELint hypodermic needles</td><td>Medline</td><td>NDA264372</td><td></td></tr><tr><td>33G x 1/2 EXELint specialty hypodermic needles</td><td>Medline</td><td>NDA26393</td><td></td></tr><tr><td>50 mL Conical Tubes</td><td>Fisher Scientific</td><td>06-443-20</td><td></td></tr><tr><td>Alcohol Prep Pads (70% Isopropyl Alcohol)</td><td>Medline</td><td>MDS090670Z</td><td></td></tr><tr><td>Aluminum Foil</td><td>Fisher Scientific</td><td>01-213-100</td><td></td></tr><tr><td>Animal Heating Plate</td><td>Physitemp Instruments</td><td>HP-1M</td><td></td></tr><tr><td>Araldite GY 502</td><td>Electron Microscopy Sciences</td><td>10900</td><td></td></tr><tr><td>Axiocam 305 Color R2 Microscopy Camera</td><td>Carl Zeiss Microscopy</td><td>426560-9031-000</td><td></td></tr><tr><td>BD Luer-Lok Syringes, 20 mL</td><td>Medline</td><td>B-D303310Z</td><td></td></tr><tr><td>Calcium Chloride</td><td>Fisher Scientific</td><td>C79-500</td><td></td></tr><tr><td>Cell Culture Dishes 35mm x 10mm</td><td>Corning Inc.</td><td>430165</td><td></td></tr><tr><td>Cotton Tipped Applicators</td><td>Medline</td><td>MDS202055H</td><td></td></tr><tr><td>DMP-30 Activator</td><td>Electron Microscopy Sciences</td><td>13600</td><td></td></tr><tr><td>Dodecenyl Succinic Anhydride/ DDSA</td><td>Electron Microscopy Sciences</td><td>13700</td><td></td></tr><tr><td>Dressing Forceps, 5&quot;, curved, serrated, narrow tipped</td><td>Integra Miltex</td><td>6-100</td><td></td></tr><tr><td>Dressing Forceps, 5&quot;, standard, serrated</td><td>Integra Miltex</td><td>6-6</td><td></td></tr><tr><td>EMBED 812 Resin</td><td>Electron Microscopy Sciences</td><td>14900</td><td></td></tr><tr><td>Ethyl Alcohol, anhydrous 200 proof</td><td>Electron Microscopy Sciences</td><td>15055</td><td></td></tr><tr><td>Fisherbrand 4-Way Tube Rack</td><td>Fisher Scientific</td><td>03-448-17</td><td></td></tr><tr><td>Fisherbrand Digital Timer</td><td>Fisher Scientific</td><td>14-649-17</td><td></td></tr><tr><td>Fisherbrand Single Syringe Infusion Pump</td><td>Fisher Scientific</td><td>7801001</td><td></td></tr><tr><td>Gauze Sponges 2&quot; x 2&quot;- 4 Ply</td><td>Medline</td><td>NON26224H</td><td></td></tr><tr><td>Glutaraldehyde (10% Solution)</td><td>Electron Microscopy Sciences</td><td>16120</td><td></td></tr><tr><td>Isoflurane Liquid Inhalant Anesthesia, 100 mL</td><td>Medline</td><td>66794-017-10</td><td></td></tr><tr><td>Jeweler-Style Micro-Fine Forceps, Style 5F</td><td>Integra Miltex</td><td>17-305</td><td>Need 2 pairs.</td></tr><tr><td>L/S Pump Tubing, Silicone, L/S 15; 25 Ft</td><td>VWR</td><td>MFLX96410-15</td><td></td></tr><tr><td>L-Aspartic Acid</td><td>Fisher Scientific</td><td>BP374-100</td><td></td></tr><tr><td>Lead Nitrate</td><td>Fisher Scientific</td><td>L-62</td><td></td></tr><tr><td>Malachite Green 4</td><td>Electron Microscopy Sciences</td><td>18100</td><td></td></tr><tr><td>Masterflex L/S Easy-Load II Pump Head</td><td>VWR</td><td>MFLX77200-62</td><td></td></tr><tr><td>Masterflex L/S Variable Speed Digital Drive</td><td>VWR</td><td>MFLX07528-10</td><td></td></tr><tr><td>MSC Xcelite 5&quot; Wire Cutters</td><td>Fisher Scientific</td><td>50-191-9855</td><td></td></tr><tr><td>Osmium Tetroxide 4% Aqueous Solution</td><td>Electron Microscopy Sciences</td><td>19150</td><td></td></tr><tr><td>Paraformaldehyde (16% Solution)</td><td>Electron Microscopy Sciences</td><td>15710</td><td></td></tr><tr><td>Physitemp Temperature Controller</td><td>Physitemp Instruments</td><td>TCAT-2LV</td><td></td></tr><tr><td>Potassium Ferrocyanide</td><td>Sigma-Aldrich</td><td>P-8131</td><td></td></tr><tr><td>Propylene Oxide, ACS Reagent</td><td>Electron Microscopy Sciences</td><td>20401</td><td></td></tr><tr><td>Pyrex Glass Beakers</td><td>Fisher Scientific</td><td>02-555-25B</td><td></td></tr><tr><td>Rectal Temperature Probe for Mice</td><td>Physitemp Instruments</td><td>RET-3</td><td></td></tr><tr><td>Scotch Magic Invisible Tape, 3/4&quot; x 1000&quot;</td><td>3M Company</td><td>305289</td><td></td></tr><tr><td>Sodium Cacodylate Buffer 0.2M, pH 7.4</td><td>Electron Microscopy Sciences</td><td>11623</td><td></td></tr><tr><td>SomnoFlo Low Flow Electronic Vaporizer</td><td>Kent Scientific</td><td>SF-01</td><td></td></tr><tr><td>SomnoFlo Starter Kit for Mice</td><td>Kent Scientific</td><td>SF-MSEKIT</td><td></td></tr><tr><td>Stainless Steel Minutien Pins</td><td>Fine Science Tools</td><td>26002-10</td><td></td></tr><tr><td>Stereomicroscope steREO Discovery.V12</td><td>Carl Zeiss Microscopy</td><td>495015-9880-010</td><td></td></tr><tr><td>Sylgard 184 Silicone Elastomer</td><td>World Precision Instruments</td><td>SYLG184</td><td>silicone mat</td></tr><tr><td>Tannic Acid</td><td>Electron Microscopy Sciences</td><td>21700</td><td></td></tr><tr><td>Thiocarbohydrazide (TCH)</td><td>Sigma-Aldrich</td><td>88535</td><td></td></tr><tr><td>Uranyl Acetate</td><td>Electron Microscopy Sciences</td><td>22400</td><td></td></tr><tr><td>Vannas Spring Micro Scissors</td><td>Fine Science Tools</td><td>15000-08</td><td></td></tr><tr><td>Von Graefe Eye Dressing Forceps, 2.75&quot;, Curved, Serrated</td><td>Integra Miltex</td><td>18-818</td><td>Need 2 pairs.</td></tr><tr><td>Wagner Scissors</td><td>Fine Science Tools</td><td>14068-12</td><td></td></tr><tr><td>Wahl MiniFigura Animal Trimmer</td><td>Braintree Scientific</td><td>CLP-9868</td><td></td></tr><tr><td>Zen Lite Software</td><td>Carl Zeiss Microscopy</td><td>410135-1001-000</td><td></td></tr></tbody></table>

puncture wound hemostasis and preparation of samples for montaged wide-area electron microscopy analysis

Hemostasis, the process of normal physiological control of vascular damage, is fundamental to human life. We all suffer minor cuts and puncture wounds from time to time. In hemostasis, self-limiting platelet aggregation leads to the formation of a structured thrombus in which bleeding cessation comes from capping the hole from the outside. Detailed characterization of this structure could lead to distinctions between hemostasis and thrombosis, a case of excessive platelet aggregation leading to occlusive clotting. An imaging-based approach to puncture wound thrombus structure is presented here that draws upon the ability of thin-section electron microscopy to visualize the interior of hemostatic thrombi. The most basic step in any imaging-based experimental protocol is good sample preparation. The protocol provides detailed procedures for preparing puncture wounds and platelet-rich thrombi in mice for subsequent electron microscopy. A detailed procedure is given for in situ fixation of the forming puncture wound thrombus and its subsequent processing for staining and embedding for electron microscopy. Electron microscopy is presented as the end imaging technique because of its ability, when combined with sequential sectioning, to visualize the details of the thrombus interior at high resolution. As an imaging method, electron microscopy gives unbiased sampling and an experimental output that scales from nanometer to millimeters in 2 or 3 dimensions. Appropriate freeware electron microscopy software is cited that will support wide-area electron microscopy in which hundreds of frames can be blended to give nanometer-scale imaging of entire puncture wound thrombi cross-sections. Hence, any subregion of the image file can be placed easily into the context of the full cross-section.

Quantitation of drug effects on puncture wound bleeding time 
Puncture wound bleeding times provide a physiological model of a drug risk that can be readily carried out in mice. Outcomes that come from a puncture wound experiment are predictive. Here, we show a dabigatran dose-response bleeding curve. Dabigatran, a thrombin inhibitor, is used as an oral direct-acting anti-coagulant, a so-called DOAC12. The jugular vein puncture wound model was used to assess the risk inherent...

Author Spotlight: Revealing Platelet Dynamics Through Advances in Structural Hematology

A puncture wound procedure for hemostatic thrombus formation is presented here. The formed thrombi are large and are hundreds of microns in diameter. Hence, volume imaging approaches are appropriate. We suggest montaged wide-area transmission electron microscopy as a high-resolution approach available to many and detail a preparative protocol.

Puncture Wound Hemostasis and Preparation of Samples for Montaged Wide-Area Electron Microscopy Analysis

Hemostasis, the process of normal physiological control of vascular damage, is fundamental to human life. We all suffer minor cuts and puncture wounds ...

puncture-wound-hemostasis-preparation-samples-for-montaged-wide-area

Research

JoVE Journal

Medicine

219 Views.  University of Arkansas for Medical Sciences. A puncture wound procedure for hemostatic thrombus formation is presented here. The formed thrombi are large and are hundreds of microns in diameter. Hence, volume imaging approaches are appropriate. We suggest montaged wide-area transmission electron microscopy as a high-resolution approach available to many and detail a preparative protocol. 

Video: Author Spotlight: Revealing Platelet Dynamics Through Advances in Structural Hematology

Sample Preparation Method of Scanning and Transmission Electron Microscope for the Appendages of Woodboring Beetle

This report described sample preparation methods that scanning and transmission electron microscope observations, demonstrated by preparing appendages of the woodboring beetle, Chlorophorus caragana Xie &#38; Wang (2012), for both types of electron microscopy. The scanning electron microscopy (SEM) sample preparation protocol was based on sample chemical fixation, dehydration in a series of ethanol baths, drying, and sputter-coating. By adding Tween 20 (Polyoxyethylene sorbitan laurate) to the fixative and the wash solution, the insect body surface of woodboring beetle was washed more cleanly in SEM. This study&#39;s transmission electron microscopy (TEM) sample preparation involved a series of steps including fixation, ethanol dehydration, embedding in resin, positioning using fluorescence microscopy, sectioning, and staining. Fixative with Tween 20 enabled penetrate the insect body wall of woodboring beetle more easily than it would had been without Tween 20, and subsequently better fixed tissues and organs in the body, thus yielded clear transmission electron microscope observations of insect sensilla ultrastructures. The next step of this preparation was determining the positions of insect sensilla in the sample embedded in the resin block by using fluorescence microscopy to increase the precision of target sensilla positioning. This improved slicing accuracy.

In order to observe ultrastructure of insect sensilla, scanning and transmission electron microscopy (SEM and TEM, respectively) sample preparation protocol were presented in the study. Tween 20 was added into the fixative to avoid sample deformation in SEM. Fluorescence microscopy was helpful for improving slicing accuracy in TEM.

This report described sample preparation methods that scanning and transmission electron microscope observations, demonstrated by preparing appendages of ...

Environment

Generation of a Three-dimensional Full Thickness Skin Equivalent and Automated Wounding

In vitro models are a cost effective and ethical alternative to study cutaneous wound healing processes. Moreover, by using human cells, these models reflect the human wound situation better than animal models. Although two-dimensional models are widely used to investigate processes such as cellular migration and proliferation, models that are more complex are required to gain a deeper knowledge about wound healing. Besides a suitable model system, the generation of precise and reproducible wounds is crucial to ensure comparable results between different test runs. In this study, the generation of a three-dimensional full thickness skin equivalent to study wound healing is shown. The dermal part of the models is comprised of human dermal fibroblast embedded in a rat-tail collagen type I hydrogel. Following the inoculation with human epidermal keratinocytes and consequent culture at the air-liquid interface, a multilayered epidermis is formed on top of the models. To study the wound healing process, we additionally developed an automated wounding device, which generates standardized wounds in a sterile atmosphere.

The goal of this protocol is to build up a three-dimensional full thickness skin equivalent, which resembles natural skin. With a specifically constructed automated wounding device, precise and reproducible wounds can be generated under maintenance of sterility.

In vitro models are a cost effective and ethical alternative to study cutaneous wound healing processes. Moreover, by using human cells, these models ...

Bioengineering

Murine Excisional Wound Healing Model and Histological Morphometric Wound Analysis

The murine excisional wound model has been used extensively to study each of the sequentially overlapping phases of wound healing: inflammation, proliferation and remodeling. Murine wounds have a histologically well-defined and easily recognizable wound bed over which these different phases of the healing process are measurable. Within the field, it is common to use an arbitrarily defined &#8220;middle&#8221; of the wound for histological analyses. However, wounds are a three-dimensional entity and often not histologically symmetrical, supporting the need for a well-defined and robust method of quantification to detect morphometric defects with a small effect size. In this protocol, we describe the procedure for creating bilateral, full-thickness excisional wounds in mice as well as a detailed instruction on how to measure morphometric parameters using an image processing program on select serial sections. The two-dimension measurements of wound length, epidermal length, epidermal area, and wound area are used in combination with the known distance between sections to extrapolate the three-dimension epidermal area covering the wound, overall wound area, epidermal volume and wound volume. Although this detailed histological analysis is more time and resource consuming than conventional analyses, its rigor increases the likelihood of detecting novel phenotypes in an inherently complex wound healing process.

This protocol describes how to generate bilateral, full-thickness excisional wounds in mice and how to subsequently monitor, harvest, and prepare the wounds for morphometric analysis. Included is an in-depth description of how to use serial histological sections to define, precisely quantify and detect morphometric defects.

The murine excisional wound model has been used extensively to study each of the sequentially overlapping phases of wound healing: inflammation, ...

Developmental Biology

An Unbiased Approach of Sampling TEM Sections in Neuroscience

Investigations of the ultrastructural features of neurons and their synapses are only possible with electron microscopy. Especially for comparative studies of the changes in densities and distributions of such features, an unbiased sampling protocol is vital for reliable results. Here, we present a workflow for the image acquisition of brain samples. The workflow allows systematic uniform random sampling within a defined brain region, and the images can be analyzed using a disector. This technique is much faster than extensive examination of serial sections but still presents a feasible approach to estimate the densities and distributions of ultrastructure features. Before embedding, stained vibratome sections were used as a reference to identify the brain region under investigation, which helped speed up the overall specimen preparation process. This approach was used for comparative studies investigating the effect of an enriched-housing environment on several ultrastructural parameters in the mouse brain. Based on the successful use of the workflow, we adapted it for the purpose of elemental analysis of brain samples. We optimized the protocol in terms of the time of user-interaction. Automating all the time-consuming steps by compiling a script for the open source software SerialEM helps the user to focus on the main work of acquiring the elemental maps. As in the original workflow, we paid attention to the unbiased sampling approach to guarantee reliable results.

We introduce a novel workflow for electron microscopy investigations of brain tissue. The method allows the user to examine neuronal features in an unbiased fashion. For elemental analysis, we also present a script that automatizes most of the workflow for randomized sampling.

Investigations of the ultrastructural features of neurons and their synapses are only possible with electron microscopy. Especially for comparative ...

Neuroscience

Proteomic Sample Preparation from Formalin Fixed and Paraffin Embedded Tissue

Preserved clinical material is a unique source for proteomic investigation of human disorders. Here we describe an optimized protocol allowing large scale quantitative analysis of formalin fixed and paraffin embedded (FFPE) tissue. The procedure comprises four distinct steps. The first one is the preparation of sections from the FFPE material and microdissection of cells of interest. In the second step the isolated cells are lysed and processed using 'filter aided sample preparation' (FASP) technique. In this step, proteins are depleted from reagents used for the sample lysis and are digested in two-steps using endoproteinase LysC and trypsin. 
After each digestion, the peptides are collected in separate fractions and their content is determined using a highly sensitive fluorescence measurement. Finally, the peptides are fractionated on 'pipette-tip' microcolumns. The LysC-peptides are separated into 4 fractions whereas the tryptic peptides are separated into 2 fractions. In this way prepared samples allow analysis of proteomes from minute amounts of material to a depth of 10,000 proteins. Thus, the described workflow is a powerful technique for studying diseases in a system-wide-fashion as well as for identification of potential biomarkers and drug targets.

Archival formalin fixed and paraffin embedded (FFPE) clinical samples are valuable material for investigation of diseases. Here we demonstrate a sample preparation workflow allowing in-depth proteomic analysis of microdissected FFPE tissue.

Preserved clinical material is a unique source for proteomic investigation of human disorders. Here we describe an optimized protocol allowing large scale ...

Biology

Cryo-electron Microscopy Specimen Preparation By Means Of a Focused Ion Beam

Here we present a protocol used to prepare cryo-TEM samples of Aspergillus niger spores, but which can easily be adapted for any number of microorganisms or solutions. We make use of a custom built cryo-transfer station and a modified cryo-SEM preparation chamber2. The spores are taken from a culture, plunge-frozen in a liquid nitrogen slush and observed in the cryo-SEM to select a region of interest. A thin lamella is then extracted using the FIB, attached to a TEM grid and subsequently thinned to electron transparency. The grid is transferred to a cryo-TEM holder and into a TEM for high resolution studies. Thanks to the introduction of a cooled nanomanipulator tip and a cryo-transfer station, this protocol is a straightforward adaptation to cryogenic temperature of the routinely used FIB preparation of TEM samples. As such it has the advantages of requiring a small amount of modifications to existing instruments, setups and procedures; it is easy to implement; it has a broad range of applications, in principle the same as for cryo-TEM sample preparation. One limitation is that it requires skillful handling of the specimens at critical steps to avoid or minimize contaminations.

Cryo Electron Microscopes, either Scanning (SEM) or Transmission (TEM), are widely used for characterization of biological samples or other materials with a high water content1. A SEM/Focused Ion Beam (FIB) is used to identify features of interest in samples and extract a thin, electron-transparent lamella for transfer to a cryo-TEM.

Here we present a protocol used to prepare cryo-TEM samples of Aspergillus niger spores, but which can easily be adapted for any number of microorganisms ...

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 ...

Ferric Chloride-Induced Arterial Thrombosis and Sample Collection for 3D Electron Microscopy Analysis

Cardiovascular diseases are a leading cause of mortality and morbidity worldwide. Aberrant thrombosis is a common feature of systemic conditions like diabetes and obesity, and chronic inflammatory diseases like atherosclerosis, cancer, and autoimmune diseases. Upon vascular injury, usually the coagulation system, platelets, and endothelium act in an orchestrated manner to prevent bleeding by forming a clot at the site of the injury. Abnormalities in this process lead to either excessive bleeding or uncontrolled thrombosis/insufficient antithrombotic activity, which translates into vessel occlusion and its sequelae. The FeCl3-induced carotid injury model is a valuable tool in probing how thrombosis initiates and progresses in vivo. This model involves endothelial damage/denudation and subsequent clot formation at the injured site. It provides a highly sensitive, quantitative assay to monitor vascular damage and clot formation in response to different degrees of vascular damage. Once optimized, this standard technique can be used to study the molecular mechanisms underlying thrombosis, as well as the ultrastructural changes in platelets in a growing thrombus. This assay is also useful to study the efficacy of antithrombotic and antiplatelet agents. This article explains how to initiate and monitor FeCl3-induced arterial thrombosis and how to collect samples for analysis by electron microscopy.

The present protocol describes how to use a FeCl3-mediated injury to induce arterial thrombosis, and how to collect and prepare arterial injury samples at various stages of thrombosis for electron microscopy analysis.

Cardiovascular diseases are a leading cause of mortality and morbidity worldwide. Aberrant thrombosis is a common feature of systemic conditions like ...

Miniaturized Sample Preparation for Transmission Electron Microscopy

Due to recent technological progress, cryo-electron microscopy (cryo-EM) is rapidly becoming a standard method for the structural analysis of protein complexes to atomic resolution. However, protein isolation techniques and sample preparation methods for EM remain a bottleneck. A relatively small number (100,000 to a few million) of individual protein particles need to be imaged for the high-resolution analysis of proteins by the single particle EM approach, making miniaturized sample handling techniques and microfluidic principles feasible.
A miniaturized, paper-blotting-free EM grid preparation method for sample pre-conditioning, EM grid priming and post processing that only consumes nanoliter-volumes of sample is presented. The method uses a dispensing system with sub-nanoliter precision to control liquid uptake and EM grid priming, a platform to control the grid temperature thereby determining the relative humidity above the EM grid, and a pick-and-plunge-mechanism for sample vitrification. For cryo-EM, an EM grid is placed on the temperature-controlled stage and the sample is aspirated into a capillary. The capillary tip is positioned in proximity to the grid surface, the grid is loaded with the sample and excess is re-aspirated into the microcapillary. Subsequently, the sample film is stabilized and slightly thinned by controlled water evaporation regulated by the offset of the platform temperature relative to the dew-point. At a given point the pick-and-plunge mechanism is triggered, rapidly transferring the primed EM grid into liquid ethane for sample vitrification. Alternatively, sample-conditioning methods are available to prepare nanoliter-sized sample volumes for negative stain (NS) EM.
The methodologies greatly reduce sample consumption and avoid approaches potentially harmful to proteins, such as the filter paper blotting used in conventional methods. Furthermore, the minuscule amount of sample required allows novel experimental strategies, such as fast sample conditioning, combination with single-cell lysis for &#34;visual proteomics,&#34; or &#34;lossless&#34; total sample preparation for quantitative analysis of complex samples.

An instrument and methods for the preparation of nanoliter-sized sample volumes for transmission electron microscopy is presented. No paper-blotting steps are required, thus avoiding the detrimental consequences this can have for proteins, significantly reducing sample loss and enabling the analysis of single cell lysate for visual proteomics.

Due to recent technological progress, cryo-electron microscopy (cryo-EM) is rapidly becoming a standard method for the structural analysis of protein ...

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 ...