Name: Scanning Electron Microscopy of Macerated Tissue to Visualize the Extracellular Matrix (Video) | JoVE Journal
Uploaded: 2016-06-14T15:00:00.000+00:00
Duration: 10 min 21 s
Description: Learn scanning electron microscopy of macerated tissue to visualize cardiac extracellular matrix structure. This method reveals matrix organization in decellularized samples for fibrosis and cardiomyopathy research.

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5 minutes

This study was funded by grants from the National Institutes of Health (NIH), Heart, Lung, and Blood Institute (NIHLB): K01-HL-121045, K08-HL-094703, 5T32HL007411-35, P20 HL101425, U01 HL100398. 
Imaging and tissue processing (after NaOH maceration) were performed through the use of the Vanderbilt University Medical Center (VUMC) Cell Imaging Shared Resource (CISR) (supported by NIH grants CA68485, DK20593, DK58404, DK59637 and EY08126). We are especially grateful to the VUMC CISR core directors (Dr. Sam Wells and Dr. W. Gray (Jay) Jerome) for valuable technical advice and also for providing core space and resources for the purposes of filming the technique highlighted in this paper. 
We would like to extend our deepest appreciation to Dr. Yan Ru Su and Ms. Kelsey Tomasek in the Cardiology Core Lab for Translational and Clinical Research at Vanderbilt University for providing technical expertise and for collecting human tissue samples used in this study.

Ethics Statement: The protocols for animal handling were approved by the Vanderbilt Institutional Animal Care and Use Committee (IACUC, protocols number M/10/117 (swine) and M/10/219 (mice) and conducted according to AAALAC-International standards. Usage of banked human cardiac tissues was approved by the Vanderbilt University Medical Center IRB (protocol number 100887).
1. Sample Collection and Storage
<ol>
	<li>Make fresh 4% glutaraldehyde in 0.1 M phosphate buffer (PB) solution. 
	CAUTION: Glutaraldehyde is toxic, wear gloves and work in a fume hood.
	<ol>
		<li>Make a 0.2 M stock solution of PB, pH 7.4 using 21.8 g Na2HPO4 and 6.4 g NaH2PO4. Quantum satis (Q.s.) to 1,000 ml dH2O.</li>
		<li>Add 400 &#181;l of 50% glutaraldehyde to 2.5 ml of PB stock solution and 2.1 ml dH2O.</li>
	</ol>
	</li>
	<li>Immerse a chunk (no larger than 2 cm2) of tissue into the 4% glutaraldehyde solution. Note: Smaller pieces can also be used but should be of sufficient size (no smaller than 5 mm2) to be easily visualized by eye in order to facilitate later steps of the protocol.</li>
	<li>Incubate at room temperature for 1 hr, then store indefinitely at 4 &#176;C.</li>
</ol>
2. Decellularization of Heart Tissue
<ol>
	<li>Make fresh 10% aqueous NaOH solution using 10 g NaOH pellets/100 ml dH2O. 
	CAUTION: NaOH solutions are corrosive and can cause alkali-burns, wear gloves.</li>
	<li>Rinse tissue in dH2O.</li>
	<li>Incubate in 10% NaOH solution at room temperature for 6 - 10 days (until tissue changes from reddish-brown to off-white or white).</li>
	<li>Rinse in dH2O until tissue becomes transparent.</li>
	<li>Immerse tissue in 1% tannic acid for 4 hr at room temperature. Use 1 ml 5% stock solution per 4 ml dH2O. 
	CAUTION: Tannic acid is a strong irritant, wear gloves.</li>
	<li>Rinse in dH2O overnight.</li>
</ol>
3. Osmication and Dehydration of Heart Tissue (in Fume Hood for Safety)
<ol>
	<li>Make a 0.2 M stock solution of sodium cacodylate buffer using 21.4 g sodium cacodylate, 10.0 g calcium chloride and 450 ml dH2O. Mix, then add hydrochloric acid as needed to adjust pH to 7.4. Q.s. to 500 ml with dH2O. 
	CAUTION: Sodium cacodylate and hydrochloric acid are toxic, wear gloves and work in a fume hood.</li>
	<li>Make 2% aqueous stock solution of osmium tetroxide in fume hood for safety by dissolving 1 g osmium tetroxide crystal in 50 ml dH2O. 
	CAUTION: Osmium tetroxide is a severe inhalation hazard; vapor fixation of mucus membranes or eyeballs is possible, hence handle only in the fume hood with gloves. A splash guard is recommended.</li>
	<li>Rinse tissue in 0.1 M sodium cacodylate buffer (mix stock solution 1:1 with dH2O) for 5 min on rotator (or gentle agitation).</li>
	<li>Repeat previous step twice, for a total of three buffer rinses.</li>
	<li>Immerse tissue in 1% osmium tetroxide in 0.1 M sodium cacodylate buffer (mix stock sodium cacodylate and stock osmium tetroxide 1:1) on rotator for 1 hr.</li>
	<li>Rinse tissue in 0.1 M sodium cacodylate buffer 3 times for 5 min each on rotator.</li>
	<li>Rinse tissue using increasing concentrations of ethanol (30%, 50%, 75%, 85%, 95%, and finally 100%) for 15 min each on rotator.</li>
</ol>
4. Cross Section Surface Preparation for SEM
<ol>
	<li>Transfer tissue in 100% ethanol to a shallow Petri dish also containing 100% ethanol.</li>
	<li>Hold two very sharp razor blades such that the flats of the sides are in contact with one another and the cutting edges cross to form two sides of an equilateral triangle above the specimen. To accomplish this, use the left hand to place one blade on the far right side of the sample and cut leftward. At the same time, use the right hand to place the second blade on the far left of the sample and slice rightward. Thus, the blades will be slid against each other from opposite directions to make a single smooth cut.</li>
	<li>Slide the flat blade sides against each other to make a very clean cut of the specimen with minimal distortion or tearing force, preferably exposing as large a surface area as possible without damaging the specimen.</li>
	<li>Repeat for each specimen to expose the cleanest possible cross section surfaces for examination in the SEM.</li>
</ol>
5. Critical Point Drying (CPD) of Heart Tissue
<ol>
	<li>Use a spatula or tweezers to transfer tissue to the sample holder of the critical point dryer (CPD), ensuring that tissue remains in 100% ethanol at all times. Ensure that the holder is immersed in ethanol and that the transfer is achieved with the tissue exposed to air for no more than a few seconds.</li>
	<li>Operate CPD per user&#39;s manual to complete 3 purge-and-fill cycles to replace ethanol with liquid carbon dioxide.</li>
	<li>Operate CPD per user&#39;s manual to achieve critical point drying of the samples and return them to atmospheric pressure with controlled venting of CO2.</li>
</ol>
6. Mounting of Heart Tissue Specimens for SEM
<ol>
	<li>Prepare an SEM sample stub for each specimen, by adhering a carbon adhesive tab to the top surface of the aluminum stub.</li>
	<li>With the aid of a stereomicroscope, carefully adhere the specimen to the adhesive tab with the cross section surface of interest facing up (away from the tab, visible), and as close as possible to parallel with the plane of the sample stub surface. Do not probe or touch the surface of interest.</li>
	<li>Break a wooden applicator stick to achieve a tapered brush, ideal for paint application. Apply silver or carbon paint at the base and sides of the specimen, to improve adherence to the stub.</li>
	<li>Extend a very thin line of silver or carbon paint up to an edge of the surface of interest, to provide a charge path from the surface of interest to ground.</li>
	<li>Apply 2 or 3 small dabs of silver or carbon paint around the perimeter of the carbon tab, to provide a conductive path from the carbon tab surface to the metal stub and thus to ground.</li>
	<li>Allow conductive paint to dry for two hrs.</li>
	<li>Operate sputter coater per user&#39;s manual to apply a relatively heavy coating (target range of 30 - 40 nm) of gold-palladium alloy or gold. Pump specimen chamber to approximately 0.1 mbar; set Timer to 40 sec. Open Set valve to the 8:00 position (moderate Argon gas flow). Press Start to initiate sputter coating at 30 mA. Purple glow discharge should be visible in the specimen chamber.</li>
</ol>
7. SEM Examination of Heart Tissue Specimens
<ol>
	<li>Perform scanning electron microscopy at relatively low accelerating voltage to minimize imaging problems associated with poor charge dissipation in the sample (charging). Suggested initial imaging condition are: 5 kV accelerating voltage, 10 mm working distance.</li>
	<li>With assistance of an experienced operator, employ increased working distance to extend depth of field in imaging conditions requiring focus of fibers in multiple focal planes, or fibers extending for considerable length in the z dimension.
	<ol>
		<li>For the microscope used here (see Table of Materials), in the user interface access the Navigation tab at upper right.</li>
		<li>Access the Coordinates tab from the Stage menu. To increase working distance, enter a larger value in millimeters for the Z coordinate then click on the Go To tab to move the sample stage to the entered working distance.</li>
	</ol>
	</li>
	<li>With assistance of an experienced operator, employ specimen tilt and rotation to position the surface of interest orthogonal to the electron beam. Additional tilt of 10 to 30 degrees from this position may improve observation and documentation of the matrix structure.
	<ol>
		<li>For the microscope used here (see Table of Materials), in the user interface, click and hold the right mouse button, then slide left or right to focus the specimen near the periphery of the prepared surface plane of interest, noting the focused working distance.</li>
		<li>Navigate with the Manual User Interface joystick to move near the opposing edge of the surface and repeat focus. If working distance is not approximately equal to the first position, tilt specimen to achieve approximate agreement at both locations using the Coordinates tab of the Stage menu (see 7.2) and enter a tilt value in the T coordinate.</li>
		<li>Rotate specimen ninety degrees (enter value into R Coordinate field) and repeat the process until all positions are focused at approximately the same working distance. 
		Note: Variable pressure SEM may be employed to improve charge dissipation if available. High vacuum is the standard operating mode for scanning electron microscopy.</li>
	</ol>
	</li>
</ol>

The authors have nothing to disclose.

Cross section surface preparation is the most critical step during the protocol. To preserve fine structure, dehydrated specimens must remain in 100% ethanol at all times until introduced to the critical point drying process. Therefore the slicing of the specimens to achieve surfaces for EM examination must be done while specimens are submersed in ethanol in a shallow dish. It is also critical that the exposed surface is not touched or probed during subsequent handling. No major modifications are anticipated for applicat...

Fibrosis disrupts the normal myocardial collagen network, which is critical for normal mechanistic functions of cardiomyocytes 1,2 as well as for inter-cellular communication, intracellular signaling, and cell survival 3. The development of fibrosis is a major determinant of cardiac function, and fibrotic remodeling of the cardiac interstitium arising from a variety of etiologies leads to increased left ventricular stiffness and diastolic dysfunction 4. Myocardial fibrosis may also lead to arrhythmias, and whether the progression of fibrotic remodeling is a general or local phenomenon, it is highly associated with a poor prognosis in patients with ischemic and non-ischemic cardiomyopathy 5. Likewise, the absence of myocardial fibrosis is a strong predictor of ventricular functional recovery and long-term survival 6. 
The hallmark of fibrosis is the deposition of excess collagen, which has the tensile strength of steel 7 and can adversely affect cardiomyocyte function on multiple levels. Mechanical forces resulting from an excessively collagenous matrix can lead to cardiomyocyte atrophy 8,9, passive tissue stiffness 10, tonic contraction-induced myocardial stiffness 11-13, and reduced delivery of oxygen to the remaining population of cardiomyocytes. Gap junction coupling of cardiomyocytes and myoFbs can also compromise the heart's electrical characteristics, creating a greater risk for the development of arrhythmias 14-16. Perivascular fibrosis alters vasomotor reactivity of intramural coronary arteries and arterioles 17 and contributes to luminal narrowing that reduces the supply of oxygen and thus the survival of cardiomyocytes 17-22. Pathogenic fibrotic and electrical remodeling, emanating from an initial site of ischemic injury or energy imbalance, inevitably progresses to heart failure. 
Cardiomyocyte necrosis initiates the fibrotic response, and subsequent adverse fibrotic remodeling can occur irrespective of etiology. Finding a way to control cardiac fibrosis would be clinically beneficial for the treatment of ischemic and idiopathic cardiomyopathies, hypertensive heart disease, hypertrophic cardiomyopathy, valvular heart disease and dystrophinopathies 23-42. Regardless of how the fibrotic disease process begins, soluble, profibrotic factors can cross the interstitial space and provoke activation of interstitial and adventitial fibroblasts at sites remote to the initial fibrotic scar, creating a cascade effect that ultimately leads to heart failure. The optimum scenario would be to exploit the fibrillogenic process using a targeted therapeutic that can be applied during the compensative hypertrophic stage of cardiomyopathy before it progresses to systolic pump failure, diastolic heart failure, or other end-stage outcomes. The ultimate goal would be to reverse fibrosis so that dead cardiomyocytes can be replaced and heart function restored completely. 
The importance of the matrix is widely understood, yet methods to study the matrix are limited mainly to quantitative measurements of major structural components, particularly collagen, and relative levels of different matrix and matricellular proteins. This protocol highlights a rarely used technique that is useful for assessing qualitative differences in the cardiac matrix. This technique has been recently used to compare and contrast fundamental differences in heart matrices from different etiologies of heart disease (in human explants), to examine hearts from post-infarcted pigs treated with the glial growth factor (GGF) isoform of neuregulin-1&#946;, relative to untreated animals 43, and to probe for differences in the matrices of cardiac tissues from mdx mice (a commonly used animal model of Duchenne Muscular Dystrophy) at different ages and compared to wild-type controls. This technique was first introduced by Drs. Caulfield and Borg in 1979 44, but few studies have since employed this powerful technique 45-47, re-introduced here with only slight modification. This methodology is a valuable research tool, because it provides qualitative information about extracellular matrix ultrastructure that might otherwise be overlooked when simply measuring matrix component content and/or level of fibrosis.

<table><tbody><tr><td>Calcium Chloride</td><td>Electron Microscopy Sciences</td><td>12340</td><td>100 g</td></tr><tr><td>Carbon Adhesive</td><td>Electron Microscopy Sciences</td><td>12664</td><td>30 g</td></tr><tr><td>Carbon Adhesive Tabs</td><td>Electron Microscopy Sciences</td><td>77825</td><td>order to fit stubs</td></tr><tr><td>Double Edge Razor Blades Stainless Steel</td><td>Ted Pella, Inc</td><td>121-6</td><td>250/pkg</td></tr><tr><td>Ethanol</td><td>Electron Microscopy Sciences</td><td>15055</td><td>450 ml</td></tr><tr><td>Gluteraldehyde, 50% Solution</td><td>Electron Microscopy Sciences</td><td>16310</td><td>EM grade, distillation purified</td></tr><tr><td>Hydrochloric Acid</td><td>Electron Microscopy Sciences</td><td>16760 or 16770</td><td>100 ml</td></tr><tr><td>Monosodium Phosphate NaH&lt;sub&gt;2&lt;/sub&gt;PO&lt;sub&gt;4&lt;/sub&gt;</td><td>Sigma-Aldrich</td><td>S9251-250G</td><td>250 g</td></tr><tr><td>Osmium Tetroxide</td><td>Electron Microscopy Sciences</td><td>19100</td><td>1 g</td></tr><tr><td>Silver Conductive Adhesive</td><td>Electron Microscopy Sciences</td><td>12686-15</td><td>15 g</td></tr><tr><td>Sodium hydroxide (NaOH)</td><td>Sigma-Aldrich</td><td>S8045-1KG</td><td>1 kg</td></tr><tr><td>Sodium Phosphate Dibasic (Na&lt;sub&gt;2&lt;/sub&gt;HPO&lt;sub&gt;4&lt;/sub&gt;)</td><td>Sigma-Aldrich</td><td>S3264-500G</td><td>500 g</td></tr><tr><td>Tannic Acid, 5% Aqueous&amp;nbsp;</td><td>Electron Microscopy Sciences</td><td>21702-5</td><td>500 ml</td></tr><tr><td>Trihydrate Sodium Cacodylate</td><td>Electron Microscopy Sciences</td><td>12300</td><td>100 g</td></tr><tr><td>Gold-palladium Alloy or Gold</td><td>Refining Systems, Inc.&amp;nbsp;</td><td>varies</td><td>specific to the sputter coater make and model</td></tr><tr><td>Critical Point Dryer</td><td>Electron Microscopy Sciences</td><td>850</td><td></td></tr><tr><td>Plain Wooden Applicators</td><td>Fisher Scientific</td><td>23-400-102</td><td></td></tr><tr><td>Quanta 250 Environmental SEM</td><td>FEI</td><td>Q250 SEM</td><td></td></tr><tr><td>Sputter Coater</td><td>Cressington Scientific Instruments Ltd.</td><td>Model 108</td><td></td></tr><tr><td>Alluminum SEM Sample Stubs</td><td>Electron Microscopy Sciences</td><td>75220-12</td><td>specific to the miscroscope</td></tr></tbody></table>

scanning electron microscopy of macerated tissue to visualize the extracellular matrix

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.

The highlighted technique was applied to cardiac tissues from an unused human heart transplant donor (Figure 1), explanted tissues from transplant recipients, hearts from wild-type and dystrophic mice (Figure 3), and in post-myocardial infarction heart samples from a swine model of heart injury (Figure 2). As shown in Figure 1, the human cardiac matrix is an intricate weave of cross-linked proteins that display a honeycom...

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 ...

The overall goal of this method is to visualize the structure of the cardiac matrix in the absence of cells or other non-matrix materials. This method can answer key questions in the field of fibrosis, especially how differences in the content of the matrix can affect the organization of the matrix, and how these changes might affect different cardiomyopathies or other fibrotic illnesses. The main advantage of this technique is that it provides a qualitative view of the matrix that can be used to augment quantitative measures.Dr.Galindo and Dr.Sawyer first had the idea to use this technique after they found that regular treatment of post-infarct swine had changes in gene expression that were similar to changes in matrix content after treatment. Individuals new to this method may struggle if they don't have previous SEM sample preparation experience, due to the fine motor skills required. The tissue sample should be large enough to be easily seen, and stored in four percent glutaraldehyde solution at four degrees Celsius.Remove the tissue from cold storage and rinse it off in distilled water. Then, immerse the tissue in a solution of 10 percent sodium hydroxide, made fresh from pellets in water. Be sure to take proper precautions.Keep the tissue in the sodium hydroxide solution for six to 10 days at room temperature. Proceed when the color of the tissue becomes off-white or white. Then, rinse the tissue off with distilled water until it becomes transparent.Next, put on gloves and immerse the tissue in one percent tannic acid for four hours at room temperature. After the tannic acid bath, rinse the tissue in distilled water overnight. In preparation, in a fume hood wearing gloves, make a 0.2 molar stock solution of sodium cacodylate buffer and a two percent aqueous stock solution of osmium tetroxide.Use a splash guard, as osmium tetroxide is a severe inhalation hazard. Now, make a 0.1 molar dilution of the sodium cacodylate buffer, and incubate the tissue in it for five minutes with gentle agitation. Change the buffer twice to wash the tissue a total of three times with sodium cacodylate.Next, make a one-to-one mixture of the stock sodium cacodylate and stock osmium tetroxide. Incubate the tissue in the new mixture on a rotator for an hour. Later, return the tissue to 0.1 molar sodium cacodylate and incubate it with gentle agitation for five minutes.Change this bath solution twice over 15 minutes. Then, dehydrate the tissue using a series of ethanol baths with increasing concentrations. Let the tissue soak for 15 minutes in each bath with gentle agitation.Next, transfer tissue and solution to a shallow Petri dish filled with 100 percent ethanol. Next, with care, hold two very sharp razor blades such that the flats of the sides are in contact with one another, and the cutting edges cross to form two sides of an equilateral triangle above the specimen. Now, use your dominant hand to place the second blade on the far left of the sample, and slice rightward.Thus, slide the blades against each other from opposite directions to make a single smooth cut with minimal distortion or tearing force. The goal is to expose as large a surface area as possible without damaging the specimen. Repeat the process until enough satisfactory cross-sections are prepared for examination by SEM.Using a spatula or tweezers, transfer the tissue to the sample holder of the CPD, while keeping the tissue immersed in 100 percent ethanol. The tissue cannot be exposed to the air for more than a few seconds. Use the CPD to replace the ethanol with liquid carbon dioxide, and perform the drying cycle as indicated by the manufacturer.Then, prepare an SEM sample stub for each specimen. Adhere a carbon adhesive tab to the top surface of the aluminum stub. Under a stereoscope, carefully adhere the specimen to the adhesive tab with the cross-section surface of interest facing up, away from the tab, and maximally planar with the stub surface.Do not probe or touch the surface of interest. For the smallest, most difficult specimens, break the wooden applicator stick to use as a spatula. Also, work over a large piece of filter paper to avoid losing a dropped specimen.For improved adherence, use a broken applicator stick to make a tapered brush, and apply silver or carbon paint to the base and sides of the specimen. Extend a very thin line of paint up to an edge of the surface of interest. This will make a charge path from the surface of interest to the ground.Then, apply two or three small dabs of paint around the perimeter of the carbon tab to provide a conductive path from the tab's surface to the metal stub. This functions as the ground. Set the specimen aside and allow the conductive paint to dry for two hours.Once dried, operate a sputter coater to apply a relatively heavy coat of gold palladium alloy or gold onto the specimen. A 30 to 40 nanometer thick coating is desired. Perform the scanning electron microscopy at a relatively low accelerating voltage to minimize problems associated with poor charge dissipation in the sample.Use the variable pressure mode if it is available. First, improve the working distance to provide enough depth of field to focus on fibers in the Z dimension. To do so, open the Navigation tab at upper right.Then, access the Coordinates tab from the stage menu. Therein, enter a larger value for the Z coordinate, such as 20 millimeters. Finally, press Go To tab to execute the change.Next, position the surface of interest orthogonally to the electron beam. Click and hold the right mouse button, then slide it left or right to focus the specimen near the periphery of the prepared surface plane of interest. Take note of the focused working distance.Then, using the manual user interface joystick, move the view to the opposing edge and repeat the focusing method. If the working distance has changed, tilt the specimen to achieve an approximate agreement at both locations. Use the Coordinates tab of the Stage menu and enter a tilt value in the T coordinate.Now, rotate the specimen 90 degrees by entering this value into the R coordinate field. Then, repeat the tilting process until all the positions are focused at approximately the same working distance. The described technique was applied to cardiac tissues from an unused human heart transplant donor.The human cardiac matrix is an intricate weave of cross-linked proteins that has a honeycomb-like pattern in a cross-section. Each honeycomb structure is approximately 40 microns wide, normally circumventing a single myocyte. When connected by intercalated discs, several cardio myocytes can be envisioned as a rod running lengthwise through a tunnel-like shape.In addition to providing structural information, an SEM of decellularized tissues can allow meaningful qualitative assessment in response to injury or non-injurious forms of cardiomyopathy, such as infarcted cardiac tissues from a swine. In a separate study, the GGF2 isoform of NRG-1 beta was shown to be a potential therapeutic treatment for heart failure. After watching this video, you should have a good idea of how to process tissue, heart tissue, for the scanning electron microscope, including the technique for exposing the surface of interest and for mounting the specimen.Following this procedure, other techniques such as mass spec, RNA sequencing, and various other assays can be used in order to measure extracellular matrix proteins and other responsible for signaling and other organizational observational phenomena. Also, don't forget that osmium tetroxide is a dangerous chemical, due to the potential for vapor fixation of the mucous membranes. Always remember to work in a fume hood with nitrile gloves, using a splash guard or hood sash.

scanning-electron-microscopy-macerated-tissue-to-visualize

Research

JoVE Journal

Biology

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Learn scanning electron microscopy of macerated tissue to visualize cardiac extracellular matrix structure. This method reveals matrix organization in decellularized samples for fibrosis and cardiomyopathy research.

Scanning Electron Microscopy of Macerated Tissue to Visualize the Extracellular Matrix

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