This work was supported by the National Institutes of Health grants: R01HL145753, R01HL145753-01S1, R01HL145753-03S1, and R01HL152723; LSUHSC-S CCDS Finish Line Award, COVID-19 Research Award, and LARC Research Award to MSB; LSUHSC-S Malcolm Feist Cardiovascular and AHA Postdoctoral Fellowship to CSA (20POST35210789); and LSUHSC-S Malcolm Feist Pre-doctoral Fellowship to RA.

The animal handling procedures in this protocol complied with the Guide for the Care and Use of Laboratory Animals (Eighth Edition, National Institutes of Health, Bethesda, MD) and were approved by the Animal Care and Use Committee of Louisiana State University Health Sciences Center-Shreveport. Six-month-old male mice with FVB/N background, were used for the present study. The mice were obtained from commercial sources (see Table of Materials). The mice used were housed in a well-regulated environment in cages allowing a 12 h light-dark-cycle, provided with water and a regular chow diet ad libitum, and were cared for according to the NIH Guide for the Care and Use of the Laboratory Animals. The overall process is illustrated in Figure 1.
1. Animal preparation
<ol>
	<li>Anesthetize the mice using 3% isoflurane. Open the chest by making a horizontal incision in the region above the middle abdomen and pulling the skin. Carefully, make another incision to open the chest cavity without puncturing any other organs21,22,23.
	<ol>
		<li>Perfuse24 the heart through the apex first, and then the right ventricle using cold 3% glutaraldehyde in cardioplegic solution (50 mM KCl, 5% dextrose in PBS, see Supplementary File 1) for 2 min to ensure complete myocardial relaxation.</li>
	</ol>
	</li>
	<li>Perfuse the heart using ice-cold 3% glutaraldehyde in 0.1 M of sodium cacodylate buffer (pH 7.4, step 2.1.1) for another 2 min. Using gravity pressure, use a 25 G 5/8&#34; needle to introduce the fixatives into the heart. Immediately after the heart starts to fill up with the fixative, lift up the apex of the heart and cut the vessels underneath at 1-2 mm from the heart to relieve pressure and allow the liquids to drain.</li>
	<li>Dissect the heart, remove the atria24, and drop the ventricles into ice-cold 3% glutaraldehyde/0.1 M sodium cacodylate in a Petri dish. Make a butterfly cut after 30-60 min of fixation and place it in a Petri dish containing 3% glutaraldehyde/cacodylate (see Table of Materials). Chop the heart using a surgical blade in small cubes of 1 mm3 while it is in glutaraldehyde/cacodylate solution.</li>
	<li>Fix/immerse the dissected heart tissue in glutaraldehyde/cacodylate solution for 24 h at 4 &#176;C.</li>
</ol>
2. Heart tissue processing
<ol>
	<li>After 24 h of fixation, wash the tissues in 0.1 M of sodium cacodylate buffer for 20 min.
	<ol>
		<li>Prepare 0.1 M cacodylate buffer following the steps below.
		<ol>
			<li>To prepare 0.1 M sodium cacodylate buffer, prepare a stock of 1 M sodium cacodylate buffer by dissolving sodium cacodylate (21.4 g) and of 1% calcium chloride solution (3 mL) in 90 mL of distilled water. Volume up the solution to 100 mL by adding the distilled water. Stir the solution well and leave it overnight to dissolve the solute.</li>
			<li>Next, take 10 mL of 1 M sodium cacodylate stock solution and add it to 80 mL of distilled water. Adjust the pH to 7.4 using HCl and volume it up to 100 mL by adding distilled water.</li>
		</ol>
		</li>
	</ol>
	</li>
	<li>Repeat the wash in 0.1 M sodium cacodylate buffer for another 20 min. Remove the tissues from sodium cacodylate buffer and immerse them in 2% osmium tetroxide solution for 4 h at room temperature. This process is called osmication.
	<ol>
		<li>Prepare 2% Osmium tetroxide solution following the steps below.
		<ol>
			<li>To prepare 2% Osmium tetroxide solution, take 4% Osmium tetroxide solution, 4 mL (see Table of Materials), 1 M sodium cacodylate stock solution (0.8 mL), and distilled water (3.2 mL) to make a total of 8 mL solution. 
			NOTE: This whole process and the steps from hereon must be carried out in the fume hood.</li>
		</ol>
		</li>
	</ol>
	</li>
	<li>After osmication, immerse the tissues in 2% sodium acetate solution for 10 min at room temperature.
	<ol>
		<li>Prepare 2% sodium acetate solution by dissolving 4 g of sodium acetate in 20 mL of distilled water.</li>
	</ol>
	</li>
	<li>Next, immerse the tissues in 2% uranyl acetate solution for 1 h at room temperature.
	<ol>
		<li>Prepare 2% uranyl acetate solution by dissolving 4 g of uranyl acetate in 20 mL of distilled water.</li>
	</ol>
	</li>
	<li>After uranyl acetate staining, dehydrate the tissues sequentially through the graded alcohols and acetone in the order mentioned in Supplementary File 1.</li>
</ol>
3. Heart tissue embedding
<ol>
	<li>Embed the dehydrated tissues in low viscosity epoxy resin following the steps below.
	<ol>
		<li>To make low-viscosity epoxy resin, mix 10.24 mL of vinyl cyclohexene dioxide (ERL 4221) epoxy monomer, 6.74 mL of diglycidyl ether of polypropylene glycol (DER 732), and 30.05 mL of nonenyl succinic anhydride (NSA) (see Table of Materials) in a 50 mL tube. Stir the suspension well by hand for 2 min.</li>
		<li>Add 18 drops of 2-dimethylaminoethanol (DMAE, see Table of Materials) epoxy accelerator and stir the suspension to mix the components thoroughly.</li>
		<li>Impregnate the tissues in epoxy resin in the following sequence of mixture.
		<ol>
			<li>Replace the 100% acetone with 1:1 resin: acetone suspension and immerse the tissues in it for 1 h at room temperature.</li>
			<li>Replace 1:1 resin: acetone suspension with 6:1 resin: acetone suspension and immerse the tissues in it for 3 h.</li>
			<li>Finally, replace the 6:1 resin: acetone suspension with 100% resin suspension and leave tissues immersed in it overnight at room temperature.</li>
		</ol>
		</li>
	</ol>
	</li>
	<li>Put the tissues into fresh resin in 8 mm micro molds (see Table of Materials) and cure the embedded tissues at 70 &#176;C overnight. 
	&#8203;NOTE: Ensure that the resin is hard but not brittle after curing.</li>
</ol>
4. Tissue sectioning using an ultramicrotome
<ol>
	<li>Trim the resin blocks with the tissue to no larger than 1 mm before mounting on the ultramicrotome (see Table of Materials). Place the mold precisely as possible in the segmented arm of the ultramicrotome and manually advance the sample mold toward the diamond knife.</li>
	<li>Cut the sections at a thickness of 500 nm (one-half micron) with a Histo knife (see Table of Materials), and using the EM loop tool, pick them up and place them on a glass slide.</li>
	<li>Place the glass slide on a hot plate for toluidine blue stain25 to find the area of interest.</li>
	<li>After finding the area of interest, use an Ultra 45&#176; knife (see Table of Materials) to produce pale gold ultrathin sections (100 nm). Place these ultrathin sections on the dull side of a 200 mesh copper grid.</li>
</ol>
5. Ultrathin heart section staining
<ol>
	<li>Start the staining by unmasking the antigen using an etching solution, i.e., 5% sodium metaperiodate solution.
	<ol>
		<li>Prepare 500 &#181;L of 5% sodium metaperiodate (see Table of Materials) solution in distilled water. 
		NOTE: Prepare fresh solution before use.</li>
		<li>Put 20 &#181;L of sodium metaperiodate solution on a clean paraffin film. Place the completely dried grids with tissue sections on the droplet of the etching solution. 
		NOTE: The dull side of the grid with the tissue section must face toward the etching solution.</li>
		<li>Leave the section grid on the solution for 30 min at room temperature.</li>
	</ol>
	</li>
	<li>Wash the etched tissue section by placing it on a droplet of distilled water for 60 s.</li>
	<li>Block the residual aldehydes by placing the section grids on a droplet of 0.05 M glycine solution for 10 min at room temperature. Blot the grid&#39;s edges on filter paper to remove the residual glycine solution.
	<ol>
		<li>Prepare 0.05 M glycine solution (see Table of Materials) by dissolving 3.75 mg of glycine in 1 mL of 1x PBS (pH 7.4).</li>
	</ol>
	</li>
	<li>Place the section grids in 10-20 &#181;L of blocking solution for 25 min at room temperature. 
	NOTE: Blocking solution composition: 2 &#181;L of 1% normal goat serum (NGS) + 20 &#181;L of 1% BSA (final concentration: 10%) + 178 &#181;L of 1x PBS (pH 7.4) to make a final volume of 200 &#181;L.</li>
	<li>Blot the grid edges on filter paper and place the grid section on antibody diluent for conditioning at room temperature for 10 min.</li>
	<li>Incubate the grid sections with primary antibody (Sigmar1 in this case, see Table of Materials; diluted 1:10 in antibody diluent) for 1 h 30 min in a humidified chamber.</li>
	<li>Blot dry the grid and wash the grid sections with antibody diluent twice for 5 min each.</li>
	<li>Incubate the grid sections with biotinylated secondary antibody (in this case, biotinylated goat anti-rabbit polyclonal secondary antibody, see Table of Materials; diluted 1:10 in antibody diluent) for 1 h in a humidified chamber.</li>
	<li>Blot dry the grid and wash the grid sections with antibody diluent twice for 5 min each.</li>
	<li>Incubate the grid sections in commercially available streptavidin-conjugated QD (QD655nm, see Table of Materials; diluted 1:10 in antibody diluent) for 1 h in a humidified chamber at room temperature. Prevent exposure to light by covering the chamber with aluminum foil.</li>
	<li>Blot dry the grid edges using filter paper and wash the grid sections by placing them on water droplets for 2 min.</li>
	<li>Blot the edges of the grid to dry.</li>
</ol>
6. Transmission Electron Microscopy (TEM) imaging
<ol>
	<li>Place the ultrathin sections on copper grids in a specimen quartet holder and clamp them in a position to allow the selection of grids in the electron beam.</li>
	<li>Insert the specimen holder in the microscope column and engage the pump switch to evacuate the goniometer, followed by full insertion of the specimen holder in the microscope column (see Table of Materials).</li>
	<li>Set the voltage to 80 kV before generating the beam for image observation.</li>
	<li>Focus well on the desired area, capture the image using a high-speed digital camera, and save the file in .tif format. 
	NOTE: The microscope setting to capture image in this study was accelerating voltage or high tension voltage of 80 kV and a magnification of 20,000x.</li>
</ol>

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The authors have nothing to disclose.

In the present study, QD-mediated immunolabeling was used to distinctively show the subcellular localization of Sigmar1. Using QD, Sigmar1&#39;s localization on the mitochondrial membrane, especially the inner mitochondrial membrane, was depicted in cardiac tissue. Additionally, Sigmar1 was also found to be located on sarcoplasmic/endoplasmic reticulum (S/ER) and lysosomes at the ultrastructural level, as shown in Figure 2A-D.
A critical step in this protocol is the etching or antigen unmasking step using highly concentrated sodium metaperiodate solution to unmask the antigen after glutaraldehyde fixation and osmication. This protocol used a single treatment with a high concentration (5%) of sodium metaperiodate for 30 min at room temperature. Extra care is needed in this step as a longer duration or higher concentration for sodium metaperiodate incubation will result in aggregation of structures, loss of membrane definition for the organelles, and cause perforations in the section, making it tough to visualize the protein or the structure. Alternatively, a lower concentration of (3%) metaperiodate solution in two steps for 30 min can also be used instead of 5% metaperiodate. Studies have shown this option to exhibit similar results as with 5% metaperiodate solution for one-step 30 min incubation. However, a 3% metaperiodate solution for 30 min of incubation for two times provides better control over the process26,27,28. Initially, this protocol used incubation of the sections with 10% metaperiodate solution for 30 min. However, due to too many perforations created in the tissue section by this concentration, the final concentration and incubation duration of the metaperiodate solution was tapered down and optimized to 5% for 30 min.
Another step required optimization of the fixation time with glutaraldehyde. Suboptimal fixation of tissues results in inadequate QD labeling, whereas over fixation of tissues results in higher non-specific labeling. Therefore, careful consideration must be given in determining and titrating an optimal level of tissue fixation for proper and specific labeling of proteins. In this method using heart tissues, the fixation time with glutaraldehyde was titrated using 24 h and 48 h as timepoints. Based on the staining images of the sections fixed for both the timepoints, it was found that sections fixed for 24 h displayed better results. To date, QD nanocrystals are available in multiple sizes, including 525, 565, 585, 605, 655, and 705 nm11,29. Each of these QD has its own emission spectra and emits fluorescence at different wavelengths. Additionally, these commercially available QDs of different sizes display different shapes; for example, QD 525, 565, and 585 are virtually spherical with different sizes, whereas QD 605, 655, and 705 are irregular oblong shaped. Of these different QD nanocrystals, QD 525, 565, and 655 are easily distinguishable from one another11,29. These differences in emission spectra and shapes make QD a great candidate for multi-labeling of proteins and visualization by fluorescence and electron microscopy. In this study, a commercially available QD, QD 655, was used to label the Sigmar1 protein to distinguish it from any non-specific background in the stained sections.
Another counterpart of QD for protein labeling in high-resolution microscopy is the immunogold particle. The immunogold particles are traditionally used to label proteins for high-resolution microscopy. These gold particles are highly electron-dense and easily identifiable compared to QD nanocrystals. However, QD exhibits better efficiency with better penetration in tissues, higher stability and shelf life, and better retention of ultrastructural components, making them a better candidate for protein labeling4,5. QD also has a unique ability to be detected by both light and electron microscopy, which adds to its value over immunogold labeling10.
One limitation of this QD-mediated immunolabeling is the use of osmium tetroxide during processing. Osmium tetroxide is used to increase the electron density, conductivity, and contrast of otherwise less electron-dense and less contrasting biological membrane structures5,30. However, the use of osmium tetroxide instantaneously and irreversibly destroys the property of the specimen to create fluorescence when labeled with QD6. This limits the use of QD in fluorescence microscopy. An alternative approach omitting the use of osmium tetroxide will be advantageous in retaining the fluorescent properties and hence the dual application of QD-mediated immunolabeling. Some of the newer models of TEM have the option of attaching the Energy Dispersive X-Ray Analysis (EDX) system that allows identification of the elemental composition of materials. Another limitation of the study is the lack of elemental mapping of a sample and image analysis using EDX. Therefore, future studies should focus on the EDX analysis of the QD spectra to analyze the elemental composition.
QD labeling of proteins has gained a lot of attention in recent times. QD offers several applications and advantages in both biological research and medical therapeutics. QD being additionally wrapped with polydentate ligand exhibits increased stability maintaining the quantum yield. Further, encapsulating QD with these bio-favorable agents also increases its bioavailability in tissues making it a good candidate for potential application in detecting tumors, live-cell imaging, drug delivery, and tissue imaging3,31,32.

The human body is composed of many proteins responsible for numerous bodily functions. The function of proteins largely depends on their localization in the organ and cellular organelles. Several techniques, including subcellular fractionation, immunofluorescence, and detergent-mediated protein extraction, are commonly used to determine the protein&#39;s subcellular localization1,2. Microscopy using immunofluorescent dye is the most widely used method among these techniques. However, the fluorescent dyes used in this technique are less stable and prone to photobleaching3. Other techniques involved high-resolution microscopy to visualize the protein at an ultrastructural level by immunolabeling proteins with electron-dense, heavy metals (gold, ferritin) or quantum dots nanocrystals and followed by visualizing them using transmission electron microscopy (TEM)4,5.
QD is a semiconductor nanocrystal composed of semiconductor metal compounds with controllable photoluminescence properties holding a great significance in biological systems3. QD nanocrystals are made in a core-shell format where a nanocrystal is encapsulated in the formation of nanocrystals to assure their proper stability and functioning. Commonly used core-shell nanocrystals combination are CdSe/ZnS, CdSe/CdS CdSe/ZnSe, CdTe/CdS, CdTe/ZnS, and CdTe/CdS/ZnS (core/shell/shell)3. Among these nanocrystal combinations, CdSe/ZnS and CdSe/CdS are most vigorously studied and frequently used as secondary antibody conjugates3,6. These QD nanoparticles also possess fluorescent properties with different excitation and emission spectra than traditional fluorophores. QD employs the excitation of electrons from the bulk valence band to exhibit higher fluorescence quantum yields compared to traditional fluorophores. The nanocrystal arrangement of semiconductor metals makes QD-mediated labeling more stable and resistant to photobleaching6. In addition, the nanocrystal core in QD and its crystalline structure allows QD of different sizes to have a wide range of absorption spectra and very narrow emission peaks7. Moreover, these QD particles are large enough to yield high electron density, making them useful in high-resolution microscopy techniques, including transmission electron microscopy5,8,9. These QD nanocrystals are also commercially available in multiple sizes with different fluorescence emission spectra and shapes, making them a great candidate for labeling with multiple antibodies10,11.
QD technology attracted great significance in biological research due to multiple functional properties, including use in live-cell imaging, the study of transport mechanisms in the cell, membrane transport of protein&#39;s diffusion movement, functional heterogeneity of cells, and marking of intracellular organelle3,12,13,14,15,16. QD is also useful in molecular diagnostics for targeting and detecting cancer tissues, characterizing the tumor&#39;s molecular profile and immune status, and visualizing vitreous body and epiretinal membranes3,17,18. In addition, QD can also be used in medical therapies to treat tumor malignancies via photodynamic therapy and ophthalmic anomalies by delivering medicines to eyes3,17,18,19.
Using these highly useful QD nanocrystals, the present study determined the subcellular localization of a protein named Sigma 1 receptor (Sigmar1). Sigmar1 is a ubiquitously expressed multi-tasking molecular chaperone protein. Extensive studies focusing on Sigmar1&#39;s subcellular localization in different tissues and organs reported a cell and tissue-type specific subcellular localization eliciting molecular function20. In different cells (neuronal, photoreceptor, and immune cells) and tissues (liver and brain) using different biochemical approaches, studies reported the localization of Sigmar1 on endoplasmic reticulum (ER), mitochondria-associated ER membrane (MAM), nuclear envelope, plasma membrane, nucleoplasmic reticulum, nucleus, and mitochondria. Despite all these studies, the subcellular localization of Sigmar1 in the heart remained unknown20. Therefore, the subcellular localization of Sigmar1 in cardiac tissue was determined using the QD-mediated immunolabeling followed by TEM imaging.

<table><tbody><tr><td>200 Mesh copper grid</td><td>Ted Pella</td><td>G200HH</td><td></td></tr><tr><td>6-month-old male mice with FVB/N background</td><td>Jackson Laboratory, Bar Harbor, ME</td><td></td><td></td></tr><tr><td>Acetone 100%</td><td>Fisher Chemicals</td><td>A949</td><td></td></tr><tr><td>Antibody diluent</td><td>Dako</td><td>S3022</td><td></td></tr><tr><td>anti-Sigmar1 antibody</td><td>Cell Signaling</td><td>61994S</td><td></td></tr><tr><td>Biotinylated goat anti-rabbit IgG antibody</td><td>Sigma Aldrich</td><td>B7389</td><td></td></tr><tr><td>BSAc (10%)</td><td>Electron Microscopy Sciences</td><td>25557</td><td></td></tr><tr><td>Calcium chloride</td><td>Sigma Aldrich</td><td>C7902</td><td></td></tr><tr><td>Cytoseal Xyl</td><td>Thermo fisher</td><td>8312-4</td><td></td></tr><tr><td>DER 732C36AA10:C33</td><td>Electron Microscopy Sciences</td><td>13010</td><td></td></tr><tr><td>Dextrose</td><td>Sigma Aldrich</td><td>G7528</td><td></td></tr><tr><td>Diamond Knife</td><td>Diatome</td><td>Histo; Ultra 45&lt;sup&gt;0&lt;/sup&gt;</td><td></td></tr><tr><td>DMAE</td><td>Electron Microscopy Sciences</td><td>13300</td><td></td></tr><tr><td>Electron microscope</td><td>JEOL</td><td>JEOL-1400 Flash</td><td></td></tr><tr><td>ERL 4221</td><td>Electron Microscopy Sciences</td><td>15004</td><td></td></tr><tr><td>Ethanol 100%</td><td>Fisher Chemicals</td><td>A405P</td><td></td></tr><tr><td>Glutaraldehyde 3%</td><td>Electron Microscopy Sciences</td><td>16538-15</td><td></td></tr><tr><td>Glycine</td><td>Alfa Aesar</td><td>A13816</td><td></td></tr><tr><td>Hydrochloric acid</td><td>Fisher Scientific</td><td>SA56</td><td></td></tr><tr><td>Micromolds</td><td>Ted Pella</td><td>10505</td><td></td></tr><tr><td>Microtome</td><td>Leica Microsystem</td><td>EM UC7</td><td></td></tr><tr><td>Normal goat serum</td><td>Invitrogen</td><td>PCN5000</td><td></td></tr><tr><td>NSA</td><td>Electron Microscopy Sciences</td><td>19050</td><td></td></tr><tr><td>Osmium tetroxide</td><td>Electron Microscopy Sciences</td><td>19150</td><td></td></tr><tr><td>Parafilm</td><td>Genesse Scientific</td><td>16-101</td><td></td></tr><tr><td>Potassium Chloride</td><td>Sigma Aldrich</td><td>P5655</td><td></td></tr><tr><td>Potassium Phosphate monobasic</td><td>Sigma Aldrich</td><td>71640</td><td></td></tr><tr><td>Qdot 655 Streptavidin Conjugate</td><td>Invitrogen</td><td>Q10121MP</td><td></td></tr><tr><td>Sodium Acetate</td><td>Fisher Scientific</td><td>BP334</td><td></td></tr><tr><td>Sodium Cacodylate</td><td>Electron Microscopy Sciences</td><td>12300</td><td></td></tr><tr><td>Sodium Chloride</td><td>Fisher Scientific</td><td>BP358</td><td></td></tr><tr><td>Sodium metaperiodate</td><td>Sigma Aldrich</td><td>71859</td><td></td></tr><tr><td>Sodium Phosphate dibasic</td><td>Sigma Aldrich</td><td>P9541</td><td></td></tr><tr><td>Surgical blade (size 10)</td><td>Aspen surgical</td><td>371110</td><td></td></tr><tr><td>TEM&amp;nbsp; image software</td><td>AMT-V700</td><td>AMT TEM imaging systems</td><td></td></tr><tr><td>TEM imaging camera</td><td>XR80 TEM series</td><td>AMT TEM imaging systems</td><td></td></tr><tr><td>Toluidine Blue O solution (0.5%)</td><td>Fisher Scientic</td><td>S25612</td><td></td></tr><tr><td>Uranyl acetate</td><td>Polysciences</td><td>21447</td><td></td></tr></tbody></table>

visualizing subcellular localization of a protein in the heart using quantum dots-mediated immuno-labeling followed by transmission electron microscopy

The subcellular localization is critical to delineating proper function and determining the molecular mechanisms of a particular protein. Several qualitative and quantitative techniques are used to determine the subcellular localization of proteins. One of the emerging techniques in determining the subcellular localization of a protein is quantum dots (QD)-mediated immunolabeling of a protein followed by imaging them with transmission electron microscopy (TEM). QD is a semiconductor nanocrystal with a dual property of crystalline structure and high electron density, which makes them applicable to electron microscopy. This current method visualized the subcellular localization of Sigma 1 receptor (Sigmar1) protein using QD-TEM in the heart tissue at ultrastructural level. Small cubes of the heart tissue sections from a wild-type mouse were fixed in 3% glutaraldehyde, subsequently osmicated, stained with uranyl acetate, followed by sequential dehydration with ethanol and acetone. These dehydrated heart tissue sections were embedded in low-viscosity epoxy resins, cut into thin sections of 500 nm thickness, put on the grid, and subsequently subjected to antigen unmasking with 5% sodium metaperiodate, followed by quenching of the residual aldehydes with glycine. The tissues were blocked, followed by sequential incubation in primary antibody, biotinylated secondary antibody, and streptavidin-conjugated QD. These stained sections were blot dried and imaged at high magnification using TEM. The QD-TEM technique allowed the visualization of Sigmar1 protein&#39;s&#160;subcellular localization at the ultrastructural level in the heart. These techniques can be used to visualize the presence of any protein and subcellular localization in any organ system.

The present QD-TEM methodology visualized the presence of Sigmar1 and its localization on the subcellular heart compartments by performing anti-Sigmar1 targeted QD labeling on adult mouse ultra-thin heart sections. Electron-dense anti-Sigmar1 labeled QD on mitochondrial membranes (inner and outer), lysosomes, and the endoplasmic reticulum / sarcoplasmic reticulum (ER/SR) membrane-mitochondrial interface (Figure 2A,B) were illustrated by the QD-TEM images. In addition, heart sections were also labeled with rabbit IgG and QD as an Isotype control for anti-Sigmar1 rabbit primary antibody (Figure 2C,D). Therefore, QD-TEM imaging highlighted the localization of endogenous Sigmar1 and its enrichment mostly on the lysosomes and the mitochondrial membranes.
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/64085/64085fig01.jpg" /> 
Figure 1: Schematic illustration showing the sequential steps and procedures used to perform QD-TEM. <a href="https://www.jove.com/files/ftp_upload/64085/64085fig01large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 2" class="xfigimg" src="/files/ftp_upload/64085/64085fig02.jpg" /> 
Figure 2: Sigmar1 immuno-labeled QD in adult mice heart tissues. (A,B) Representative TEM images showing Sigmar1 labeled QD on mitochondria (outer- and inner-membrane), mitochondria-associated endoplasmic reticulum (ER) / sarcoplasmic reticulum (SR) membranes, and lysosomes. (C,D) TEM image of the heart sections of isotype control labeled with QDs and rabbit IgG. Scale bar: 200 nm. <a href="https://www.jove.com/files/ftp_upload/64085/64085fig02large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
Supplementary File 1: Composition of PBS and other solutions used for tissue dehydration. <a href="https://www.jove.com/files/ftp_upload/64085/64085_Supplementary File 1.docx" target="_blank">Please click here to download this File.</a>

Watch this Scientific Journal Video about Visualizing Subcellular Localization of a Protein in the Heart Using Quantum Dots-Mediated Immuno-Labeling Followed by Transmission Electron Microscopy at JoV

Visualizing Subcellular Localization of a Protein in the Heart Using Quantum Dots-Mediated Immuno-Labeling Followed by Transmission Electron Microscopy

The present protocol describes a method of immuno-labeling a protein in the heart tissue sections using quantum dots. This technique provides a useful tool to visualize any protein&#39;s subcellular localization and expression at the ultrastructural level.

The subcellular localization is critical to delineating proper function and determining the molecular mechanisms of a particular protein. Several ...

visualizing-subcellular-localization-protein-heart-using-quantum-dots

Research

JoVE Journal

Biology

2.7K Views.  Louisiana State University Health Sciences Center-Shreveport. Het bepalen van de subcellulaire lokalisatie van een eiwit is van cruciaal belang voor het afbakenen van de juiste functie en de mechanismen die daarbij betrokken zijn. Deze methode kan ons helpen de subcellulaire lokalisatie van een eiwit op ultrastructuurniveau te visualiseren. Quantum dots-gemedieerde immunolabeling is stabieler, bestand tegen fotobleaching, heeft zijn eigen emissiespectra, levert een hoge elektronendichtheid op en vertoont een hogere efficiëntie en retentie op weefsels m...

Video: Visualizing Subcellular Localization of a Protein in the Heart Using Quantum Dots-Mediated Immuno-Labeling Followed by Transmission Electron Microscopy

Mitochondria and Endoplasmic Reticulum Imaging by Correlative Light and Volume Electron Microscopy

Cellular organelles, such as mitochondria and endoplasmic reticulum (ER), create a network to perform a variety of functions. These highly curved structures are folded into various shapes to form a dynamic network depending on the cellular conditions. Visualization of this network between mitochondria and ER has been attempted using super-resolution fluorescence imaging and light microscopy; however, the limited resolution is insufficient to observe the membranes between the mitochondria and ER in detail. Transmission electron microscopy provides good membrane contrast and nanometer-scale resolution for the observation of cellular organelles; however, it is exceptionally time-consuming when assessing the three-dimensional (3D) structure of highly curved organelles. Therefore, we observed the morphology of mitochondria and ER via correlative light-electron microscopy (CLEM) and volume electron microscopy techniques using enhanced ascorbate peroxidase 2 and horseradish peroxidase staining. An en bloc staining method, ultrathin serial sectioning (array tomography), and volume electron microscopy were applied to observe the 3D structure. In this protocol, we suggest a combination of CLEM and 3D electron microscopy to perform detailed structural studies of mitochondria and ER.

We present a protocol to study the distribution of mitochondria and endoplasmic reticulum in whole cells after genetic modification using correlative light and volume electron microscopy including ascorbate peroxidase 2 and horseradish peroxidase staining, serial sectioning of cells with and without the target gene in the same section, and serial imaging via electron microscopy.

Cellular organelles, such as mitochondria and endoplasmic reticulum (ER), create a network to perform a variety of functions. These highly curved ...

Biochemistry

Label-Free Imaging of Single Proteins Secreted from Living Cells via iSCAT Microscopy

We demonstrate interferometric scattering (iSCAT) microscopy, a method capable of detecting single unlabeled proteins secreted from individual living cells in real time. In this protocol, we cover the fundamental steps to realize an iSCAT microscope and complement it with additional imaging channels to monitor the viability of a cell under study. Following this, we use the method for real-time detection of single proteins as they are secreted from a living cell which we demonstrate with an immortalized B-cell line (Laz388). Necessary steps concerning the preparation of microscope and sample as well as the analysis of the recorded data are discussed. The video protocol demonstrates that iSCAT microscopy offers a straightforward method to study secretion at the single-molecule level.

We present a protocol for the real-time optical detection of single unlabeled proteins as they are secreted from living cells. This is based on interferometric scattering (iSCAT) microscopy, which can be applied to a variety of different biological systems and configurations.

We demonstrate interferometric scattering (iSCAT) microscopy, a method capable of detecting single unlabeled proteins secreted from individual living ...

Engineering

Isolation and Genetic Manipulation of Adult Cardiac Myocytes for Confocal Imaging

Cardiac myocytes isolated from adult hearts are widely accepted as a model somewhere half way between embryonic and neonatal muscle cells on one side and a working heart on the other. Thus, cardiomyocytes serve as good models for cardiac cellular physiology and pathophysiology, for pharmaceutical investigations as well as for the exploration of transgenic animal models. Here we describe a method of isolating the cells from the heart. Furthermore we show how a genetic manipulation on cardiac myocytes can be performed without breeding a transgenic animal: This is the combination of long term culture (1 week) and adenoviral gene transfer. The latter one is described from the construction of the virus to the transduction of the cells. It can be used for the expression of genetically encoded biosensors (GEBs), fluorescent fusion proteins, but also for protein over expression and down regulation, e.g. using RNAi. Here we provide an example for the expression of a fusion protein staining a subcellular structure (Golgi). Such protein expression can be visualized by confocal imaging of z-stacks for a 3D-reconstruction of subcellular structures. The protocol comprises state-of-the-art in cell culture, molecular biology and biophysics and thus provides an approach for exploring new horizons in cellular cardiology.

Adult cardiac myocytes are primary cells that can be isolated from animal hearts and cultured for several days. Within this culture period adenoviral gene transfer can be used to express genetically encoded biosensors (GEBs) or fluorescent fusion proteins. Both approaches allow cellular investigations by means of confocal microscopy.

Cardiac myocytes isolated from adult hearts are widely accepted as a model somewhere half way between embryonic and neonatal muscle cells on one side and ...

Correlative Light- and Electron Microscopy Using Quantum Dot Nanoparticles

A method is described whereby quantum dot (QD) nanoparticles can be used for correlative immunocytochemical studies of human pathology tissue using widefield fluorescence light microscopy and transmission electron microscopy (TEM). To demonstrate the protocol we have immunolabeled ultrathin epoxy sections of human somatostatinoma tumor using a primary antibody to somatostatin, followed by a biotinylated secondary antibody and visualization with streptavidin conjugated 585 nm cadmium-selenium (CdSe) quantum dots (QDs). The sections are mounted on a TEM specimen grid then placed on a glass slide for observation by widefield fluorescence light microscopy. Light microscopy reveals 585 nm QD labeling as bright orange fluorescence forming a granular pattern within the tumor cell cytoplasm. At low to mid-range magnification by light microscopy the labeling pattern can be easily recognized and the level of non-specific or background labeling assessed. This is a critical step for subsequent interpretation of the immunolabeling pattern by TEM and evaluation of the morphological context. The same section is then blotted dry and viewed by TEM. QD probes are seen to be attached to amorphous material contained in individual secretory granules. Images are acquired from the same region of interest (ROI) seen by light microscopy for correlative analysis. Corresponding images from each modality may then be blended to overlay fluorescence data on TEM ultrastructure of the corresponding region.

A method is described whereby quantum dot (QD) nanoparticles can be used for correlative immunocytochemical studies of epoxy embedded human pathology tissue. We employ commercial antibody fragment conjugated QDs that are visualized by widefield fluorescence light microscopy and transmission electron microscopy.

A method is described whereby quantum dot (QD) nanoparticles can be used for correlative immunocytochemical studies of human pathology tissue using ...

Bioengineering

Mapping Molecular Diffusion in the Plasma Membrane by Multiple-Target Tracing (MTT)

Our goal is to obtain a comprehensive description of molecular processes occurring at cellular membranes in different biological functions. We aim at characterizing the complex organization and dynamics of the plasma membrane at single-molecule level, by developing analytic tools dedicated to Single-Particle Tracking (SPT) at high density: Multiple-Target Tracing (MTT)1. Single-molecule videomicroscopy, offering millisecond and nanometric resolution1-11, allows a detailed representation of membrane organization12-14 by accurately mapping descriptors such as cell receptors localization, mobility, confinement or interactions.
We revisited SPT, both experimentally and algorithmically. Experimental aspects included optimizing setup and cell labeling, with a particular emphasis on reaching the highest possible labeling density, in order to provide a dynamic snapshot of molecular dynamics as it occurs within the membrane. Algorithmic issues concerned each step used for rebuilding trajectories: peaks detection, estimation and reconnection, addressed by specific tools from image analysis15,16. Implementing deflation after detection allows rescuing peaks initially hidden by neighboring, stronger peaks. Of note, improving detection directly impacts reconnection, by reducing gaps within trajectories. Performances have been evaluated using Monte-Carlo simulations for various labeling density and noise values, which typically represent the two major limitations for parallel measurements at high spatiotemporal resolution.
The nanometric accuracy17 obtained for single molecules, using either successive on/off photoswitching or non-linear optics, can deliver exhaustive observations. This is the basis of nanoscopy methods17 such as STORM18, PALM19,20, RESOLFT21 or STED22,23, which may often require imaging fixed samples. The central task is the detection and estimation of diffraction-limited peaks emanating from single-molecules. Hence, providing adequate assumptions such as handling a constant positional accuracy instead of Brownian motion, MTT is straightforwardly suited for nanoscopic analyses. Furthermore, MTT can fundamentally be used at any scale: not only for molecules, but also for cells or animals, for instance. Hence, MTT is a powerful tracking algorithm that finds applications at molecular and cellular scales.

Mapping Molecular Diffusion in the Plasma Membrane by Multiple-Target Tracing MTT

Multiple-Target Tracing is a homemade algorithm developed for tracking individually labeled molecules within the plasma membrane of living cells. Efficiently detecting, estimating and tracing molecules over time at high-density provide a user-friendly, comprehensive tool to investigate nanoscale membrane dynamics.

Our goal is to obtain a comprehensive description of molecular processes occurring at cellular membranes in different biological functions. We aim at ...

Visualization of miniSOG Tagged DNA Repair Proteins in Combination with Electron Spectroscopic Imaging (ESI)

The limits to optical resolution and the challenge of identifying specific protein populations in transmission electron microscopy have been obstacles in cell biology. Many phenomena cannot be explained by in vitro analysis in simplified systems and need additional structural information in situ, particularly in the range between 1 nm and 0.1 &#181;m, in order to be fully understood. Here, electron spectroscopic imaging, a transmission electron microscopy technique that allows simultaneous mapping of the distribution of proteins and nucleic acids, and an expression tag, miniSOG, are combined to study the structure and organization of DNA double-strand break repair foci.

Visualization of miniSOG Tagged DNA Repair Proteins in Combination with Electron Spectroscopic Imaging ESI

Electron spectroscopic imaging can image and distinguish nucleic acid from protein at nanometer resolution. It can be combined with the miniSOG system, which is able to specifically label tagged proteins in transmission electron microscopy samples. We illustrate the use of these technologies using double-strand break repair foci as an example.

The limits to optical resolution and the challenge of identifying specific protein populations in transmission electron microscopy have been obstacles in ...

Unveiling Mitochondrial Ultrastructure in Neurons Using STED Microscopy

Using Live Cell STED Imaging to Visualize Mitochondrial Inner Membrane Ultrastructure in Neuronal Cell Models

Mitochondria play many essential roles in the cell, including energy production, regulation of Ca2+ homeostasis, lipid biosynthesis, and production of reactive oxygen species (ROS). These mitochondria-mediated processes take on specialized roles in neurons, coordinating aerobic metabolism to meet the high energy demands of these cells, modulating Ca2+ signaling, providing lipids for axon growth and regeneration, and tuning ROS production for neuronal development and function. Mitochondrial dysfunction is therefore a central driver in neurodegenerative diseases. Mitochondrial structure and function are inextricably linked. The morphologically complex inner membrane with structural infolds called cristae harbors many molecular systems that perform the signature processes of the mitochondrion. The architectural features of the inner membrane are ultrastructural and therefore, too small to be visualized by traditional diffraction-limited resolved microscopy. Thus, most insights on mitochondrial ultrastructure have come from electron microscopy on fixed samples. However, emerging technologies in super-resolution fluorescence microscopy now provide resolution down to tens of nanometers, allowing visualization of ultrastructural features in live cells. Super-resolution imaging therefore offers an unprecedented ability to directly image fine details of mitochondrial structure, nanoscale protein distributions, and cristae dynamics, providing fundamental new insights that link mitochondria to human health and disease. This protocol presents the use of stimulated emission depletion (STED) super-resolution microscopy to visualize the mitochondrial ultrastructure of live human neuroblastoma cells and primary rat neurons. This procedure is organized into five sections: (1) growth and differentiation of the SH-SY5Y cell line, (2) isolation, plating, and growth of primary rat hippocampal neurons, (3) procedures for staining cells for live STED imaging, (4) procedures for live cell STED experiments using a STED microscope&#160;for reference, and (5) guidance for segmentation and image processing using examples to measure and quantify morphological features of the inner membrane.

Author Spotlight: Decoding Mitochondrial Aging 

This protocol presents a workflow for the propagation, differentiation, and staining of cultured SH-SY5Y cells and primary rat hippocampal neurons for mitochondrial ultrastructure visualization and analysis using stimulated emission depletion (STED) microscopy.

Mitochondria play many essential roles in the cell, including energy production, regulation of Ca2+ homeostasis, lipid biosynthesis, and production of ...

Neuroscience

Local Field Fluorescence Microscopy: Imaging Cellular Signals in Intact Hearts

In the heart, molecular signaling studies are usually performed in isolated myocytes. However, many pathological situations such as ischemia and arrhythmias can only be fully understood at the whole organ level. Here, we present the spectroscopic technique of local field fluorescence microscopy (LFFM) that allows the measurement of cellular signals in the intact heart. The technique is based on a combination of a Langendorff perfused heart and optical fibers to record fluorescent signals. LFFM has various applications in the field of cardiovascular physiology to study the heart under normal and pathological conditions. Multiple cardiac variables can be monitored using different fluorescent indicators. These include cytosolic [Ca2+], intra-sarcoplasmic reticulum [Ca2+] and membrane potentials. The exogenous fluorescent probes are excited and the emitted fluorescence detected with three different arrangements of LFFM epifluorescence techniques presented in this paper. The central differences among these techniques are the type of light source used for excitation and on the way the excitation light is modulated. The pulsed LFFM (PLFFM) uses laser light pulses while continuous wave LFFM (CLFFM) uses continuous laser light for excitation. Finally, light-emitting diodes (LEDs) were used as a third light source. This non-coherent arrangement is called pulsed LED fluorescence microscopy (PLEDFM).

In the heart, molecular events coordinate the electrical and contractile function of the organ. A set of local field fluorescence microscopy techniques presented here enables the recording of cellular variables in intact hearts. Identifying mechanisms defining the cardiac function is critical in understanding how the heart works under pathological situations.

In the heart, molecular signaling studies are usually performed in isolated myocytes. However, many pathological situations such as ischemia and ...

Multi-color Localization Microscopy of Single Membrane Proteins in Organelles of Live Mammalian Cells

Knowledge about the localization of proteins in cellular subcompartments is crucial to understand their specific function. Here, we present a super-resolution technique that allows for the determination of the microcompartments that are accessible for proteins by generating localization and tracking maps of these proteins. Moreover, by multi-color localization microscopy, the localization and tracking profiles of proteins in different subcompartments are obtained simultaneously. The technique is specific for live cells and is based on the repetitive imaging of single mobile membrane proteins. Proteins of interest are genetically fused with specific, so-called self-labeling tags. These tags are enzymes that react with a substrate in a covalent manner. Conjugated to these substrates are fluorescent dyes. Reaction of the enzyme-tagged proteins with the fluorescence labeled substrates results in labeled proteins. Here, Tetramethylrhodamine (TMR) and Silicon Rhodamine (SiR) are used as fluorescent dyes attached to the substrates of the enzymes. By using substrate concentrations in the pM to nM range, sub-stoichiometric labeling is achieved that results in distinct signals. These signals are localized with ~15&#8211;27 nm precision. The technique allows for multi-color imaging of single molecules, whereby the number of colors is limited by the available membrane-permeable dyes and the repertoire of self-labeling enzymes. We show the feasibility of the technique by determining the localization of the quality control enzyme (Pten)-induced kinase 1 (PINK1) in different mitochondrial compartments during its processing in relation to other membrane proteins. The test for true physical interactions between differently labeled single proteins by single molecule FRET or co-tracking is restricted, though, because the low labeling degrees decrease the probability for having two adjacent proteins labeled at the same time. While the technique is strong for imaging proteins in membrane compartments, in most cases it is not appropriate to determine the localization of highly mobile soluble proteins.

Here, we present a protocol for multi-color localization of single membrane proteins in organelles of live cells. To attach fluorophores, self-labeling proteins are used. Proteins, located in different membranes compartments of the same organelle, can be localized with a precision of ~18 nm.

Knowledge about the localization of proteins in cellular subcompartments is crucial to understand their specific function. Here, we present a ...

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

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

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

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

Visualizing Subcellular Localization of a Protein in the Heart Using Quantum Dots-Mediated Immuno-Labeling Followed by Transmission Electron Microscopy

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