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

Summary

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.

Abstract

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.

Introduction

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 µm in Z¹, 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 µm 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.

Protocol

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 wound^1,7

1. Anesthetization of the mouse
   1. 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.
   2. When the mouse appears asleep, pinch one of the mouse'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).
   3. Turn the vaporizer down to a low flow rate (100 mL/min, 1.75% isoflurane). Place the mouse's nose in the nose cone. Extend each limb and tape down with small pieces of adhesive tape.
   4. Insert the rectal temperature probe and set the temperature to 37 °C on the temperature controller. Tape down the temperature probe cord to prevent the probe from accidentally being pulled out.
2. Mouse incision
   1. Using the clippers, shave the mouse'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.
   2. 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.
   3. 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.
3. Cleaning of the jugular vein (same principles apply to the femoral artery)
   1. 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.
   2. Fill a 20 mL syringe with normal saline warmed to 37 °C. Load the syringe into the syringe infusion pump. Attach the 27G butterfly needle and associated tubing to the syringe.
   3. 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.
4. Puncturing the jugular vein/femoral artery
   1. Start the timer. Obtain a 30G hypodermic needle (nominal 300 µm diameter) for the jugular vein and a 33G hypodermic needle (200 µm diameter) for the femoral artery. The use of a smaller needle gives only a slightly longer bleeding time for the higher-pressure arterial situation.
   2. Turn the bevel facing up. Position the needle at a 25° angle over the jugular vein/femoral artery. Note the time.
   3. 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.
   4. 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.
5. Performing transcardial perfusion fixation
   1. 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.
   2. 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.
   3. Set the peristaltic pump to a flow rate of 10 mL/min. Turn on the perfusion pump.
   4. 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.
   5. 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.
   6. Immediately before starting the transcardial perfusion, quickly cut open the rib cage and reflect each side to expose the heart.
   7. Turn on the peristaltic pump. Place the tip of the 27G butterfly needle into the base of the left ventricle.
   8. 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.
   9. 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.
   10. 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.
   11. 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.
   12. 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.

2. Collection of samples for electron microscopy

1. 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.
2. 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.
3. Transfer the fixed tissue to 4 °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's instructions). Pour enough sodium cacodylate buffer solution into the dish to submerge the fixed vein segment.
4. 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.
5. Clean the jugular vein/femoral artery segment carefully using a pair of micro scissors and very fine forceps.
   1. 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.
6. 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.
7. Store the tissue at 4 °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-TEM^8,9. 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.

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.

1. Wash the tissue sample in the 35 mm dish 2x with Hank'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.
2. 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.
3. Remove fixative and wash sample 4x for 5 min each with 0.1 M sodium cacodylate. Remove 0.1 M sodium cacodylate.
4. 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.
5. 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.
6. 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.
7. Stain with 0.5% uranyl acetate (UA) for 1 h RT or overnight at 4 °C in the dark.
8. 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.
9. 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.
10. Remove the 25% ethanol. Briefly rinse 1x with 50% ethanol. Discard the 50% ethanol and incubate with 50% ethanol for 3 min on ice.
11. 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.
12. 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.
13. 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.
14. Prepare the resin mix as described below.
    1. Warm the resin components at 60 °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 °C. This mixture can be kept for 6 months at 4 °C. Warm it to 60 °C before using. Aliquot the required amount prior to use.
15. Embed the samples as described below.
    1. Place an aliquot of the resin mix in a tube and add 20 µL/mL of DMP-30 activator. Mix thoroughly by rotating at 60 °C; the color should darken.
    2. Dilute this aliquot of activated resin by 50% in PO and allow the resin and PO to mix completely.
    3. 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.
    4. The following day, mix a new aliquot of resin (1 mL per sample) with 20 µL/mL of DMP-30.
16. 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.
17. 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 °C for at least 48 h . The prepared sample is shown in Figure 2B.
18. Perform montaged wide-area transmission electron microscopy (WA-TEM) imaging as described in^10,11. 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 functions^10,11 is available from the group of Dr. David Mastronarde, University of Colorado, Boulder (http://bio3d.colorado.edu/SerialEM/, http://bio3d.colorado.edu/imod/.)

Results

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 i...

Discussion

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...

Materials

List of materials used in this article

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