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 application of this technique to other tissue types for similar matrix observations, although the SEM portion of the protocol might require basic electron microscopy troubleshooting regardless of sample origin. Images collected from the fibrous samples are prone to artifacts introduced by poor charge dissipation (&#34;charging&#34;). Charging problems can usually be minimized by reducing accelerating voltage, increasing scan speed (also called dwell time) and reducing spot size of the electron beam. Integration of several scans collected quickly enough to avoid charge artifacts will produce an image of comparable signal to noise quality without the charge artifacts present in a slower, higher quality single scan image.
This technique is inherently qualitative and thus complements when considered alongside quantitative measurements (e.g., Masson&#39;s trichrome or picoserius red staining, measurement of hydroxyproline content, mass spectrometry and RNASeq) to ascertain how visualized structural differences might be related to various developmental or disease states. However, despite this limitation, the method is significant beyond cardiac fibrosis specifically, because the extracellular matrix is an essential component of almost every organ in the body. In the heart, the cardiac matrix provides critical mechanical support for continuous pumping characterized by complex stretching, twisting, and deformation, which confers optimal entry and efflux of oxygenated and deoxygenated blood 49 for the &#62;2.5 billion beats that occur over the average human lifespan. Given the extremely low regenerative capacity of cardiac tissue, a dynamic matrix that can be remodeled according to contextual needs makes logical sense. With only a slight stretch of the imagination, one might infer that there exist therapeutic targets for manipulating matrix remodeling to enhance the healing process, while limiting adverse fibrosis. Application of the demonstrated technique at a minimum showcases the sophistication and beauty of the cardiac matrix and in doing so further underscores its functional importance.
While the value of quantitative measures is a core principle for evaluation of practically all experimental studies, the technique highlighted here can be used to reveal qualitative ultrastructural variations that not only complement standard matrix measurements but might suggest alternate investigative paths to understand the fundamental biochemical alterations that underscore the qualitative changes. Anticipated future applications of this technique are its use in cardiac disease models and human tissues as a complementary tool for assessing matrix changes, as well as broadened usage to study other organs for which matrix changes are a component of the disease process.

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

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 honeycomb-like pattern when viewed in cross section. Each &#39;honeycomb&#39; structure is approximately 40 &#181;m wide, normally circumventing a single myocyte, when considering the planar view in Figure 1. When connected by intercalated discs, several cardiomyocytes can be envisioned as a rod running lengthwise through the &#39;tunnel&#39; when the metaphor is extended to three-dimensionality. Figure 1 also highlights the importance of the cutting procedure, with greater precision yielding more revelatory topographical data (Figure 1, top left) than sections that are &#34;smashed&#34; during the cutting process (Figure 1, top right).
Removal of resident cardiac cells, blood vessels, and circulatory cells prior to SEM processing uncovers additional ultra-structural details that would be less appreciable by SEM of whole heart tissue blocks. Each individual collagenous &#34;strut&#34;, for example (Figure 1, bottom panels) is apparently aligned at regular intervals and perpendicular to myofibril sarcomeres. This arrangement is suitable for helping maintain heart structure by counter-acting deformations exerted during contraction and relaxation cycles, similar to the &#39;warp and weft&#39; of textiles that helps to maintain fabric body and form against stretching.
<img alt="Figure 1" src="/files/ftp_upload/54005/54005fig1.jpg" /> 
Figure 1: Representative Scanning Electron Micrographs of the Three-dimensional Arrangement of the Extracellular Matrix in Decellularized Left Ventricular Tissue Obtained from an Unused Human Donor Heart. The top two panels show the matrix in cross section at low magnification (bars = 500 &#181;m), providing an aerial view of the architecture of normal human heart tissue. At higher magnification, one can better observe the typical honeycomb-like structure of supportive fibers that provide mechanical support for regularly spaced myofibers (middle left and right panel bars = 100 &#181;m and 50 &#181;m, respectively). Upon closer inspection, each &#39;honeycomb&#39; is composed of fibers that are organized in parallel with one another while perpendicular to resident cardiomyocytes (bottom left and right panel bars = 10 &#181;m and 5 &#181;m, respectively). <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/54005/54005fig1large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
In addition to providing structural information, SEM of decellularized tissues can allow meaningful, qualitative assessment of extracellular matrix changes in response to injury or non-injurious forms of cardiomyopathy. For example, this technique was recently used to examine the extracellular matrices of post-infarcted swine cardiac tissues 43. During the course of this large-animal experiment, designed specifically to assess the efficacy of the GGF2 isoform of NRG-1&#946; as a potential therapeutic for heart failure, post-infarcted swine which received intravenous NRG-1&#946; administration exhibited striking alterations in the cardiac matrix, compared to untreated cardiac tissues of post-infarcted animals. These results were subsequently published 43, and the technique has remained a valuable tool for building upon this initial, serendipitous discovery. Figure 2 includes example micrographs, produced during the course of that study, which highlight drastic matrix differences between untreated and NRG-1&#946;-treated matrices.
<img alt="Figure 2" src="/files/ftp_upload/54005/54005fig2.jpg" /> 
Figure 2: Representative Scanning Electron Micrographs of Left Ventricular Extracellular Matrix in Untreated and NRG-1&#946;-treated Pigs, after NaOH Maceration. The matrix in cross section highlights the regular spatial arrangement of fibers in untreated post-myocardial infarction (post-MI) pigs (top left), compared to post-MI NRG-1&#946;-treated animals (top right). When viewed in longitude, the matrix in untreated swine exhibits a thick, matte-like appearance (bottom left), whereas the matrix of NRG-1&#946;-treated pigs displays regular spatial arrangement of fibers (bottom right). White bar = 40 &#181;m (all four panels). More detailed results and figures are included in the associated manuscript 43. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/54005/54005fig2large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
Exploration of matrix changes that occur in dystrophic hearts has also yielded qualitative insights into the progression and development of an animal model of Duchenne muscular dystrophy cardiomyopathy (DMD). In mdx mice, a commonly used mouse model of DMD, there are significant and age-dependent differences among wild-type and mdx hearts when viewed by SEM after fixation and NaOH treatment. As shown in Figure 3, matrix components were relatively normal in 6 week-old mdx hearts lacking functional dystrophin relative to normal mice. More interesting, extracellular matrix organization was clearly disrupted or perhaps degraded in older dystrophin-deficient mice compared to younger mdx mice, illustrating the progressive nature of DMD in the heart. Such profound differences were not expected, because mdx mice are poorly representative of human DMD cardiomyopathy due to the fact that they exhibit a much milder cardiomyopathy phenotype and slower mortality rate than humans with DMD 48. This suggests that even small changes in heart function can be captured using the visualization technique presented in this manuscript. This methodology should also be readily adaptable to the extracellular matrices of other organs, for which there are likewise no currently available fibrosis-targeting therapies.
<img alt="Figure 3" src="/files/ftp_upload/54005/54005fig3.jpg" /> 
Figure 3: Representative Scanning Electron Micrographs of Left Ventricular Extracellular Matrix in Wild-type versus mdx Mice. The left ventricular cardiac matrix in wild-type mice (top panel) is similar to that observed in other species. The matrix in mdx mice at 6 weeks of age looks relatively normal, although slightly &#34;fluffy&#34; in appearance (middle panel). Conversely, the cardiac matrix of older mdx mice appears severely degenerated (bottom panel), indicating that the dystrophic process can be qualitatively captured using SEM in fixed, NaOH-macerated tissues. White bar = 10 &#181;m (all three panels). <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/54005/54005fig3large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

Watch this Scientific Journal Video about Scanning Electron Microscopy of Macerated Tissue to Visualize the Extracellular Matrix at JoVE.com

Scanning Electron Microscopy of Macerated Tissue to Visualize the Extracellular Matrix

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 ...

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

Nanyang Technological University

Research

JoVE Journal

Biology

9.9K Views.  Vanderbilt University Medical Center. Shown here is a method for visualizing extracellular matrix ultrastructure in decellularized cardiac tissues.

Video: Scanning Electron Microscopy of Macerated Tissue to Visualize the Extracellular Matrix

Glycoproteomics of the Extracellular Matrix: A Method for Intact Glycopeptide Analysis Using Mass Spectrometry

Fibrosis is a hallmark of many cardiovascular diseases and is associated with the exacerbated secretion and deposition of the extracellular matrix (ECM). Using proteomics, we have previously identified more than 150 ECM and ECM-associated proteins in cardiovascular tissues. Notably, many ECM proteins are glycosylated. This post-translational modification affects protein folding, solubility, binding, and degradation. We have developed a sequential extraction and enrichment method for ECM proteins that is compatible with the subsequent liquid chromatography tandem mass spectrometry (LC-MS/MS) analysis of intact glycopeptides. The strategy is based on sequential incubations with NaCl, SDS for tissue decellularization, and guanidine hydrochloride for the solubilization of ECM proteins. Recent advances in LC-MS/MS include fragmentation methods, such as combinations of higher-energy collision dissociation (HCD) and electron transfer dissociation (ETD), which allow for the direct compositional analysis of glycopeptides of ECM proteins. In the present paper, we describe a method to prepare the ECM from tissue samples. The method not only allows for protein profiling but also the assessment and characterization of glycosylation by MS analysis.

This paper describes a methodology to prepare cardiovascular tissue samples for MS analysis that allows for (1) the analysis of ECM protein composition, (2) the identification of glycosylation sites, and (3) the compositional characterization of glycan forms. This methodology can be applied, with minor modifications, to the study of the ECM in other tissues.

Fibrosis is a hallmark of many cardiovascular diseases and is associated with the exacerbated secretion and deposition of the extracellular matrix (ECM). ...

Biochemistry

Observing and Quantifying Fibroblast-mediated Fibrin Gel Compaction

Cells embedded in collagen and fibrin gels attach and exert traction forces on the fibers of the gel. These forces can lead to local and global reorganization and realignment of the gel microstructure. This process proceeds in a complex manner that is dependent in part on the interplay between the location of the cells, the geometry of the gel, and the mechanical constraints on the gel. To better understand how these variables produce global fiber alignment patterns, we use time-lapse differential interference contrast (DIC) microscopy coupled with an environmentally controlled bioreactor to observe the compaction process between geometrically spaced explants (clusters of fibroblasts). The images are then analyzed with a custom image processing algorithm to obtain maps of the strain. The information obtained from this technique can be used to probe the mechanobiology of various cell-matrix interactions, which has important implications for understanding processes in wound healing, disease development, and tissue engineering applications.

Time-lapse microscopy and image processing techniques were used to observe and analyze fibroblast-mediated gel compaction and fibrin fiber realignment in an environmentally controlled bioreactor over a 48 hr&#160;period.

Cells embedded in collagen and fibrin gels attach and exert traction forces on the fibers of the gel. These forces can lead to local and global ...

Bioengineering

Long-term Intravital Immunofluorescence Imaging of Tissue Matrix Components with Epifluorescence and Two-photon Microscopy

Besides being a physical scaffold to maintain tissue morphology, the extracellular matrix (ECM) is actively involved in regulating cell and tissue function during development and organ homeostasis. It does so by acting via biochemical, biomechanical, and biophysical signaling pathways, such as through the release of bioactive ECM protein fragments, regulating tissue tension, and providing pathways for cell migration. The extracellular matrix of the tumor microenvironment undergoes substantial remodeling, characterized by the degradation, deposition and organization of fibrillar and non-fibrillar matrix proteins. Stromal stiffening of the tumor microenvironment can promote tumor growth and invasion, and cause remodeling of blood and lymphatic vessels. Live imaging of matrix proteins, however, to this point is limited to fibrillar collagens that can be detected by second harmonic generation using multi-photon microscopy, leaving the majority of matrix components largely invisible. Here we describe procedures for tumor inoculation in the thin dorsal ear skin, immunolabeling of extracellular matrix proteins and intravital imaging of the exposed tissue in live mice using epifluorescence and two-photon microscopy. Our intravital imaging method allows for the direct detection of both fibrillar and non-fibrillar matrix proteins in the context of a growing dermal tumor. We show examples of vessel remodeling caused by local matrix contraction. We also found that fibrillar matrix of the tumor detected with the second harmonic generation is spatially distinct from newly deposited matrix components such as tenascin C. We also showed long-term (12 hours) imaging of T-cell interaction with tumor cells and tumor cells migration along the collagen IV of basement membrane. Taken together, this method uniquely allows for the simultaneous detection of tumor cells, their physical microenvironment and the endogenous tissue immune response over time, which may provide important insights into the mechanisms underlying tumor progression and ultimate success or resistance to therapy.

The extracellular matrix undergoes substantial remodeling during wound healing, inflammation and tumorigenesis. We present a novel intravital immunofluorescence microscopy approach to visualize the dynamics of fibrillar as well as mesh-like matrix components with high spatial and temporal resolution using epifluorescence or two-photon microscopy.

Besides being a physical scaffold to maintain tissue morphology, the extracellular matrix (ECM) is actively involved in regulating cell and tissue ...

Improving 2D and 3D Skin In Vitro Models Using Macromolecular Crowding

The glycoprotein family of collagens represents the main structural proteins in the human body, and are key components of biomaterials used in modern tissue engineering. A technical bottleneck is the deposition of collagen in vitro, as it is notoriously slow, resulting in sub-optimal formation of connective tissue and subsequent tissue cohesion, particularly in skin models. Here, we describe a method which involves the addition of differentially-sized sucrose co-polymers to skin cultures to generate macromolecular crowding (MMC), which results in a dramatic enhancement of collagen deposition. Particularly, dermal fibroblasts deposited a significant amount of collagen I/IV/VII and fibronectin under MMC in comparison to controls.
The protocol also describes a method to decellularize crowded cell layers, exposing significant amounts of extracellular matrix (ECM) which were retained on the culture surface as evidenced by immunocytochemistry. Total matrix mass and distribution pattern was studied using interference reflection microscopy. Interestingly, fibroblasts, keratinocytes and co-cultures produced cell-derived matrices (CDM) of varying composition and morphology. CDM could be used as &#34;bio-scaffolds&#34; for secondary cell seeding, where the current use of coatings or scaffolds, typically from xenogenic animal sources, can be avoided, thus moving towards more clinically relevant applications.
In addition, this protocol describes the application of MMC during the submerged phase of a 3D-organotypic skin co-culture model which was sufficient to enhance ECM deposition in the dermo-epidermal junction (DEJ), in particular, collagen VII, the major component of anchoring fibrils. Electron microscopy confirmed the presence of anchoring fibrils in cultures developed with MMC, as compared to controls. This is significant as anchoring fibrils tether the dermis to the epidermis, hence, having a pre-formed mature DEJ may benefit skin graft recipients in terms of graft stability and overall wound healing. Furthermore, culture time was condensed from 5 weeks to 3 weeks to obtain a mature construct, when using MMC, reducing costs.

Improving 2D and 3D Skin In Vitro Models Using Macromolecular Crowding

We present a protocol to obtain cell-derived matrices rich in extracellular matrix proteins, using macromolecular crowders (MMC). In addition, we present a protocol which incorporates MMC in 3D organotypic skin co-culture generation, which reduces culture time while maintaining maturity of construct.

The glycoprotein family of collagens represents the main structural proteins in the human body, and are key components of biomaterials used in modern ...

Visualizing Macrophage Extracellular Traps Using Confocal Microscopy

A primary method used to define the presence of neutrophil extracellular traps (NETs) is confocal microscopy. We have modified established confocal microscopy methods to visualize macrophage extracellular traps (METs). These extracellular traps are defined by the presence of extracellular chromatin with co-expression of other components such as granule proteases, citrullinated histones, and peptidyl arginase deiminase (PAD). The expression of METs is generally measured after exposure to a stimulus and compared to un-stimulated samples. Samples are also included for background and isotype control. Cells are analyzed using well-defined image analysis software. Confocal microscopy may be used to define the presence of METs both in vitro and in vivo in lung tissue.

Macrophage extracellular traps are a newly described entity. This article will concentrate on confocal microscopy methods and how they are visualized in vitro and in vivo from lung samples.

A primary method used to define the presence of neutrophil extracellular traps (NETs) is confocal microscopy. We have modified established confocal ...

Immunology and Infection

Quantitative Measurement of Invadopodia-mediated Extracellular Matrix Proteolysis in Single and Multicellular Contexts 

Cellular invasion into local tissues is a process important in development and homeostasis. Malregulated invasion and subsequent cell movement is characteristic of multiple pathological processes, including inflammation, cardiovascular disease and tumor cell metastasis1. Focalized proteolytic degradation of extracellular matrix (ECM) components in the epithelial or endothelial basement membrane is a critical step in initiating cellular invasion. In tumor cells, extensive in vitro analysis has determined that ECM degradation is accomplished by ventral actin-rich membrane protrusive structures termed invadopodia2,3. Invadopodia form in close apposition to the ECM, where they moderate ECM breakdown through the action of matrix metalloproteinases (MMPs). The ability of tumor cells to form invadopodia directly correlates with the ability to invade into local stroma and associated vascular components3. 
Visualization of invadopodia-mediated ECM degradation of cells by fluorescent microscopy using dye-labeled matrix proteins coated onto glass coverslips has emerged as the most prevalent technique for evaluating the degree of matrix proteolysis and cellular invasive potential4,5. Here we describe a version of the standard method for generating fluorescently-labeled glass coverslips utilizing a commercially available Oregon Green-488 gelatin conjugate. This method is easily scaled to rapidly produce large numbers of coated coverslips. We show some of the common microscopic artifacts that are often encountered during this procedure and how these can be avoided. Finally, we describe standardized methods using readily available computer software to allow quantification of labeled gelatin matrix degradation mediated by individual cells and by entire cellular populations. The described procedures provide the ability to accurately and reproducibly monitor invadopodia activity, and can also serve as a platform for evaluating the efficacy of modulating protein expression or testing of anti-invasive compounds on extracellular matrix degradation in single and multicellular settings.

Quantitative Measurement of Invadopodia-mediated Extracellular Matrix Proteolysis in Single and Multicellular Contexts

We describe the prototypical method for producing microscope coverslips coated with fluorescent gelatin for visualizing invadopodia-mediated matrix degradation. Computational techniques using available software are presented for quantifying the resultant levels of matrix proteolysis by single cells within a mixed population and for multicellular groups encompassing entire microscopic fields.

Cellular invasion into local tissues is a process important in development and homeostasis.  Malregulated invasion and subsequent cell movement is ...

Enrichment of Extracellular Matrix Proteins from Tissues and Digestion into Peptides for Mass Spectrometry Analysis

The extracellular matrix (ECM) is a complex meshwork of cross-linked proteins that provides biophysical and biochemical cues that are major regulators of cell proliferation, survival, migration, etc. The ECM plays important roles in development and in diverse pathologies including cardio-vascular and musculo-skeletal diseases, fibrosis, and cancer. Thus, characterizing the composition of ECMs of normal and diseased tissues could lead to the identification of novel prognostic and diagnostic biomarkers and potential novel therapeutic targets. However, the very nature of ECM proteins (large in size, cross-linked and covalently bound, heavily glycosylated) has rendered biochemical analyses of ECMs challenging. To overcome this challenge, we developed a method to enrich ECMs from fresh or frozen tissues and tumors that takes advantage of the insolubility of ECM proteins. We describe here in detail the decellularization procedure that consists of sequential incubations in buffers of different pH and salt and detergent concentrations and that results in 1) the extraction of intracellular (cytosolic, nuclear, membrane and cytoskeletal) proteins and 2) the enrichment of ECM proteins. We then describe how to deglycosylate and digest ECM-enriched protein preparations into peptides for subsequent analysis by mass spectrometry.

This protocol describes a procedure for enriching ECM proteins from tissues or tumors and deglycosylating and digesting the ECM-enriched preparations into peptides to analyze their protein composition by mass spectrometry.

The extracellular matrix (ECM) is a complex meshwork of cross-linked proteins that provides biophysical and biochemical cues that are major regulators of ...

A Rapid, Scalable Method for the Isolation, Functional Study, and Analysis of Cell-derived Extracellular Matrix

The extracellular matrix (ECM) is recognized as a diverse, dynamic, and complex environment that is involved in multiple cell-physiological and pathological processes. However, the isolation of ECM, from tissues or cell culture, is complicated by the insoluble and cross-linked nature of the assembled ECM and by the potential contamination of ECM extracts with cell surface and intracellular proteins. Here, we describe a method for use with cultured cells that is rapid and reliably removes cells to isolate a cell-derived ECM for downstream experimentation.
Through use of this method, the isolated ECM and its components can be visualized by in situ immunofluorescence microscopy. The dynamics of specific ECM proteins can be tracked by tracing the deposition of a tagged protein using fluorescence microscopy, both before and after the removal of cells. Alternatively, the isolated ECM can be extracted for biochemical analysis, such as sodium dodecyl sulphate-polyacrylamide gel electrophoresis (SDS-PAGE) and immunoblotting. At larger scales, a full proteomics analysis of the isolated ECM by mass spectrometry can be conducted. By conducting ECM isolation under sterile conditions, sterile ECM layers can be obtained for functional or phenotypic studies with any cell of interest. The method can be applied to any adherent cell type, is relatively easy to perform, and can be linked to a wide repertoire of experimental designs.

The extracellular matrix plays a major role in defining the microenvironment of cells and in modulating cell behavior and phenotype. We describe a rapid method for the isolation of cell-derived extracellular matrix, which can be adapted to different scales for microscopic, biochemical, proteomic, or functional studies.

The extracellular matrix (ECM) is recognized as a diverse, dynamic, and complex environment that is involved in multiple cell-physiological and ...

Puncture Wound Hemostasis and Montaged Electron Microscopy 

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.

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.

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

Medicine

Scanning Electron Microscopy (SEM)

Source: Laboratory of Dr. Andrew J. Steckl &#8212; University of Cincinnati
A scanning electron microscope, or SEM, is a powerful microscope that uses electrons to form an image. It allows for imaging of conductive samples at magnifications that cannot be achieved using traditional microscopes. Modern light microscopes can achieve a magnification of ~1,000X, while typical SEM can reach magnifications of more than 30,000X.&#160;Because the SEM doesn&#8217;t use light to create images, the resulting pictures it forms are in black and white.&#160;
Conductive samples are loaded onto the SEM&#8217;s sample stage. Once the sample chamber reaches vacuum, the user will proceed to align the electron gun in the system to the proper location. The electron gun shoots out a beam of high-energy electrons, which travel through a combination of lenses and apertures and eventually hit the sample. As the electron gun continues to shoot electrons at a precise position on the sample, secondary electrons will bounce off of the sample. These secondary electrons are identified by the detector. The signal found from the secondary electrons is amplified and sent to the monitor, creating a 3D image. This video will demonstrate SEM sample preparation, operation, and imaging capabilities.

Scanning Electron Microscopy SEM: Operation, Image Analysis

Source: Laboratory of Dr. Andrew J. Steckl &#8212; University of Cincinnati
A scanning electron microscope, or SEM, is a powerful microscope that uses ...

Scanning Electron Microscopy of Macerated Tissue to Visualize the Extracellular Matrix

Summary

Explore More Videos

Chapters in this video

Glycoproteomics of the Extracellular Matrix: A Method for Intact Glycopeptide Analysis Using Mass Spectrometry

Observing and Quantifying Fibroblast-mediated Fibrin Gel Compaction

Long-term Intravital Immunofluorescence Imaging of Tissue Matrix Components with Epifluorescence and Two-photon Microscopy

Improving 2D and 3D Skin In Vitro Models Using Macromolecular Crowding

Visualizing Macrophage Extracellular Traps Using Confocal Microscopy

Quantitative Measurement of Invadopodia-mediated Extracellular Matrix Proteolysis in Single and Multicellular Contexts

Enrichment of Extracellular Matrix Proteins from Tissues and Digestion into Peptides for Mass Spectrometry Analysis

A Rapid, Scalable Method for the Isolation, Functional Study, and Analysis of Cell-derived Extracellular Matrix

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

Scanning Electron Microscopy (SEM)