This research was supported by the KBRI basic research program through Korea Brain Research Institute funded by Ministry of Science and ICT (24-BR-01-03). TMEM192-V5-APEX2 plasmids were kindly provided by Hyun-Woo Rhee (Seoul National University). TEM data were acquired at Brain Research Core Facilities in KBRI.

The step-by-step workflow for sample preparation is shown in Figure 1.
1. Cell culture and transfection
<ol>
	<li>Culture HeLa cells at a density of 1 &#215; 105 cells in 35 mm glass grid-bottomed culture dishes containing DMEM with 10% fetal bovine serum and 100 U/mL of penicillin and streptomycin at 37 &#176;C.</li>
	<li>The day after seeding the cells, co-transfect the cells at 30%-40% confluency with TMEM192-V5-APEX2 and Mito-mEosEM plasmids using the transfection reagent according to the manufacturer&#39;s instructions(1 &#181;g each of TMEM192-V5-APEX2 and Mito-mEosEM plasmid DNA with 3 &#181;L of transfection reagent). After transfection, treat the cells with DMSO or U18666A (2 &#181;g/mL), and Bafilomycin A1 (100 nM) for 18 h.</li>
</ol>
2. Confocal microscopic imaging
<ol>
	<li>After treating the cells with Bafilomycin and U18666A for 18 h, remove the culture media and immediately add 250 &#181;L of warm (30 &#176;C-37 &#176;C) fixative solution (2.5% glutaraldehyde and 2% paraformaldehyde in 0.1 M sodium cacodylate solution [pH 7.0]) by gentle pipetting. Immediately remove the fixative, replace it with 1.5 mL of fresh fixative, and incubate on ice for 30 min. Then, wash the cells for 3 x 10 min in 1 mL of ice-cold 0.1 M sodium cacodylate buffer. 
	NOTE: Aldehyde fumes are toxic. All experiments should be conducted in a ventilated fume hood.</li>
	<li>Add 1 mL of cold 20 mM glycine solution to quench the aldehyde and incubate on ice for 10 min. Wash 3 x 10 min in cold 0.1 M sodium cacodylate buffer.</li>
	<li>Prepare a fresh solution containing 300 &#181;M Amplex-Red by adding 10 mM H2O2 to a 0.1 M cacodylate solution. Add 500 &#181;L of Amplex-Red solution to the sample dish and incubate for 5 min on ice.</li>
	<li>Remove the Amplex-Red solution and rinse 3 x 10 min with 0.1 M sodium cacodylate buffer.</li>
	<li>After Amplex-Red staining, use a confocal microscope to image the area around the cells of interest in 3 x 3 tile scan mode. Capture both differential interference contrast (DIC) and fluorescence signal to identify and record the grid and letters on the grid dish using a 40x (or 60x) objective. 
	NOTE: These overview images serve as a map for identifying specific target cells in the plastic-embedded specimens.</li>
	<li>Image the target cells at high magnification using a 40x (or 60x) objective and acquire a Z-stack image. 
	NOTE: This high-magnification image is superimposed on an electron microscope image to create a correlative image.</li>
</ol>
3. Sample preparation for EM block
<ol>
	<li>After LM imaging, add 1 mL of 1% reduced osmium tetroxide (prepared by containing 3% potassium ferrocyanide in 0.3 M cacodylate buffer with an equal volume of 2% aqueous osmium tetroxide) for postfixation at 4 &#176;C for 1 h, and then rinse for 3 x 10 min with distilled water at 4 &#176;C. 
	NOTE: Osmium tetroxide fumes are toxic. All experiments should be performed in a ventilated fume hood.</li>
	<li>Rinse 3 x 10 min with distilled water at room temperature, then dehydrate in a graded ethanol series for 15 min each time (50%, 60%, 70%, 80%, 90%, 95%, 100%, and 100%) at room temperature.</li>
	<li>Mix ethanol with epoxy resin in volume ratios of 3:1, 1:1, and 1:3, preparing a total of 300 &#181;L of each mixture. Pour each epoxy resin into the dish, allowing it to incubate at room temperature for 1 h. Then, use the 100% epoxy resin for overnight impregnation.</li>
	<li>Prepare the embedding capsule or PCR tube by filling it with fresh epoxy resin, place the tube or capsule upside down on the region of interest, and place it in an oven at 60 &#176;C for 48 h to polymerize.</li>
</ol>
4. Sectioning and TEM imaging
<ol>
	<li>After the epoxy resin has completely polymerized, separate the bottom glass from the polymer block by immersing it in liquid nitrogen.</li>
	<li>Place the specimen block in the sample holder of the ultramicrotome and set it in the trimming block. Use a razor blade to trim the resin into a trapezoidal or rectangular shape around the object of interest.</li>
	<li>Place the trimmed specimen block in the sample holder and adjust the diamond blade. Ensure that the leading edge of the specimen block is perfectly parallel to the edge of the diamond blade.</li>
	<li>Using an ultramicrotome, cut ultrathin sections approximately 60-70 nm from the flat-embedded cells. Collect the serial sections on a formvar-coated slot grid and dry them on a hot plate.</li>
	<li>Stain the section with the contrast stain for 2 min and lead citrate for 1 min.</li>
	<li>Place the grid in the TEM sample holder for observation. Identify the target cell of interest based on the DIC image from the light microscope. Image the entire cell area using tiled, overlapping images at 1,700x magnification.</li>
</ol>
5. Image stitching using imageJ Fiji.
<ol>
	<li>To begin image stitching, start imageJ Fiji25 and click on File | New | TrakEM2 (blank) | Select storage folder to create a new TrakEM226 project (Figure 2A1).</li>
	<li>Open the tile image dataset by dragging and dropping the raw tile image data into the workspace (Figure 2A2).</li>
	<li>Click the right mouse button and select Align | Montage all images in this layer | elastic (non-linear block correspondences), and then click OK to accept the default values remaining parameters (Figure 2A3).</li>
	<li>Click the right mouse button and select Export | Make flat image to create a stitched image, then click File | Save As to save the image.</li>
</ol>
6. Resizing the fluorescence image
<ol>
	<li>Open the photo enlargement software (see Table of Materials). Click on File | Open and select the fluorescence image.</li>
	<li>Enter the width and height to be the same size as the electron microscope image, and select the algorithm to be applied in the resize method (Figure 2B1).</li>
	<li>Click on File | Save AS to save the resized image.</li>
</ol>
7. Landmark-based image registration with the Big Warp plugin
<ol>
	<li>Open ImageJ Fiji and drag and drop the resized fluorescence micrograph (FM image) and the stitched electron micrograph (EM image).</li>
	<li>Click on Plugins | BigData Viewer | Big Wrap to start the registration , then click on the dropdown menu to select the FM image as the moving image and the EM image as the target image (Figure 2C1). 
	NOTE: Each FM and EM image will have three new popup windows (BigWrap moving Image, BigWrap fixed image, and Landmarks) to display landmarks.</li>
	<li>Compare the FM and EM images using functions such as zoom (mouse wheel), image rotation (moving the mouse while holding down the left button), and move (moving the mouse while holding down the right button), and then press the space bar on the keyboard to switch Landmark mode on, and press the left mouse button to mark at least three landmarks identified in the FM and EM images (Figure 2C2, pink dots and red arrow).</li>
	<li>After marking all the identified landmarks, go to File | Export Moving Image | OK and a new popup window will appear with the transformed FM image (Figure 2C3). 
	NOTE: The newly created popup window is an FM image transformed based on the basis of the landmark. Review the alignment and make adjustments, if necessary, by adding or modifying landmarks and recalculating the transformation.</li>
	<li>Click on File | Save As to save the transformed FM image.</li>
</ol>
8. Overlay of CLEM images
<ol>
	<li>In ImageJ, drag and drop the transformed FM image and the stitched EM image.</li>
	<li>Click on Image | Type | 8-Bit Color to set the same image type for both the LM and EM images (Figure 2D1). 
	NOTE: The EM image must be converted twice in the following order: 8-bit -&#62; RGB Color -&#62; 8-bit Color.</li>
	<li>Click on Image | Color &#62;| Merge Channels to combine the images into a composite image by selecting the FM and EM images from the dropdown menu, respectively (Figure 2D2).</li>
	<li>Click on File | Save As to save the CLEM image.</li>
</ol>

Correlative Microscopy Reveals Stress-Induced Mitochondria-Lysosome Interactions

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The authors have no conflicts of interest to disclose.

This protocol describes dual-color CLEM to observe the interaction of mitochondria and lysosomes in fixed cells using genetically encoded mEosEM and APEX2 tags. Over the past few decades, advances in microscopy have greatly improved the ability to observe changes in organelle networks32,33, and the study of organelle interactions has become increasingly interesting. Super-resolution microscopy has been used to observe changes in highly dynamic contacts between mitochondria and lysosomes34,35,36. However, despite the use of super-resolution microscopy, it is difficult to distinguish changes in the thickness of the inner and outer mitochondrial membranes (approximately 22 nm) and cristae junctions (approximately 28 nm)37. Transmission electron microscopy has the advantage of being able to observe the structure of biological samples at very high resolution but has the disadvantage that the images can only be viewed in black and white, making it difficult to distinguish organelles in complex images. In our protocol, we used a CLEM method that combines the advantages of light and electron microscopy to monitor the interaction of two different organelles by staining them with different colors. In particular, dual-color CLEM is an invaluable tool for observing and analyzing mitophagy, the process of mitochondrial degradation within lysosomes, as shown in Figure 3.
Live cell imaging is typically performed under physiological conditions, enabling researchers to observe the dynamic cellular processes in real time. However, electron microscopy techniques often require time-consuming sample preparation procedures, including fixation and staining. The time delay between live cell imaging and electron microscopy sample preparation limits the temporal resolution of the technique and can result in potentially missed dynamic cellular events. To overcome the limitation of live cell CLEM, we have developed a dual-color CLEM method in which the mEosEM probe23 and chemical dye (Amplex-Red) are stained by the APEX2 proximity labeling enzyme38 without losing fluorescence signal even in harsh fixative environments. In a previous live cell CLEM study39, discrepancies were found between fluorescence signals and EM images due to temporal resolution issues in TEM sampling after live cell imaging and technical limitations in sample preparation. However, by acquiring fluorescence images of fixed cells for a sufficient time (z-stack imaging of whole cells at a high magnification), we can resolve the temporal resolution issue and improve the correlation between LM and EM images (Figure 3 and Figure 4).
Another key factor for successful CLEM results is how to accurately register the LM image with the EM image. To do this, we first used a combination of DIC and fluorescence imaging modes during light microscopy imaging to create a navigation map around the target cells. Using this navigation map, we could easily identify the cells of interest on the ultramicrotome and TEM. Second, by acquiring TEM tile images of whole cells at high magnification, we were able to obtain many landmarks that could be used for image registration. In our experience, the higher the number of landmarks, the better the alignment accuracy of LM and EM images in the Big Warp plugin.
In this study, we showed the interaction domains of lysosomes and mitochondria with high-resolution CLEM results. In addition, the dual-color probe allowed for precise distinction between mitochondria and lysosomes. However, this method has some limitations. First, if the reaction time during the proximal labeling of Amplex-Red by the APEX2 enzyme is too long, the Amplex-Red will diffuse and stain wider than the target protein location38. Second, the thickness (z-axis volume) of the area where the lysosome is engulfed in the mitochondria is quite small, so the fusion of the two organelle membranes cannot be determined in a 50 nm thick serial section. Furthermore, it is a significant challenge to accurately align the z-axis of an optical section by light microscope and a mechanical section by ultramicrotome. To overcome these limitations, dual-color CLEM can be used in combination with conventional electron-tomography40 or serial section volume-CLEM41.

Mitochondria and lysosomes are the principal membrane-bound organelles that are essential for the maintenance of cellular homeostasis. Mitochondria are highly dynamic cellular organelles modulated by fission and fusion events1. In general, mitochondria are referred to as the powerhouse of the cell and are known for their important role in oxidative phosphorylation, ATP production, and metabolite storage such as&#160;calcium2. In addition to their role in energy metabolism, mitochondria are involved in a number of other cellular signaling processes, including apoptosis3,4, calcium signaling5,6, and lipid metabolism7,8. Similarly, lysosomes are highly dynamic cellular organelles that play multiple roles as cellular recycling centers to maintain cellular cleanliness through protein degradation, autophagy, endocytosis, and phagocytosis9. Apart from their role in degradation, lysosomes also play roles in cellular physiology such as nutrient sensing, homeostasis10, lipid metabolism11, and cellular dedifferentiation12.
Despite the existence of numerous studies on the functions of individual cellular organelles, our recent research has concentrated on the less-understood crosstalk between mitochondria and lysosomes, which has been demonstrated to influence both cellular physiology and pathology. Contacts between mitochondria and lysosomes have been observed in various cell types, including cancer cells, neurons, and induced pluripotent stem cell (iPSC)-derived cells13, and they are mainly observed under cellular stress or in neurodegenerative diseases such as Parkinson&#39;s disease13,14 and Charcot-Marie-Tooth disease15. The contact between mitochondria and lysosomes plays an important role in maintaining metabolic homeostasis through the exchange of ions (e.g., Ca2+)16, cholesterol17, and iron18 between the two organelles. Furthermore, the contact between mitochondria and lysosomes plays a crucial role in the quality control of cellular organelles. For example, damaged mitochondria can be targeted for degradation through the process of mitophagy, in which they are engulfed by autophagosomes and delivered to lysosomes for degradation. Consequently, the removal of dysfunctional mitochondria can be facilitated before they cause further damage to the cell or to normal mitochondria. This interaction between mitochondria and lysosomes is crucial for maintaining cellular homeostasis and preventing the accumulation of dysfunctional organelles. Thus, only damaged mitochondria are selectively removed, allowing healthy mitochondria to function optimally within the cell.
Correlative light and electron microscopy (CLEM) is an imaging technique that has recently received attention as a research method by combining the advantages of light microscopy (LM) and electron microscopy (EM) to provide detailed structural information at different scales. In CLEM, a sample is first imaged using LM to identify specific regions of interest, followed by EM sample preparation steps such as fixation, dehydration, and embedding. The same sample is then observed under an electron microscope. Although live-cell CLEM methods have been reported recently19,20,21,22, it remains challenging to accurately combine optical images of highly dynamic biological samples with electron microscopy images of fixed cells into a single plane. For highly dynamic organelles like mitochondria and lysosomes, their precise localization can change as they are observed live under the light microscope, and they continue to move even during the short delay time required for fixation.
Therefore, in this protocol, two different molecular tags that remain active even within fixative solutions were employed to visualize the interaction between mitochondria and lysosomes, which play an important role in maintaining metabolic homeostasis through the exchange of ions (e.g., Ca2+)16, cholesterol17, and iron18 between these organelles. The constructs TMEM192-V5-APEX2 and Mito-mEosEM were used to observe the interactions of mitochondria and lysosomes in cultured cells at both the EM and LM levels. For mitochondria, the mEosEM tag was selected for its ability to preserve the fluorescence signal during EM sample preparation in a fixative solution and OsO423. For lysosomes, the APEX2 tag, a protein of approximately 28 kDa, was used to study the localization of a specific protein at both EM and LM levels24. In the presence of hydrogen peroxide, the APEX2&#39;s substrate, Amplex-Red, forms an insoluble polymer that can be observed by confocal microscopy at the tag-targeted location. In this protocol, dual-color CLEM was used to visualize the structure of the interactions between lysosomes and mitochondria in fixed cells.

<table><tbody><tr><td>&lt;strong&gt;Chemicals and reagents&lt;/strong&gt;</td><td></td><td></td><td></td></tr><tr><td>25% Glutaraldehyde</td><td>Electron Microscopy Sciences</td><td>16200</td><td>Aldehyde fumes are extremely toxic. Use only in fume hood.</td></tr><tr><td>30% Hydrogen peroxide solution</td><td>Merck</td><td>107210</td><td></td></tr><tr><td>4% Paraformaldehyde solution</td><td>Biosesang</td><td>PC2031-100-00</td><td>ldehyde fumes are extremely toxic. Use only in fume hood.</td></tr><tr><td>Amplex Red</td><td>ThermoFisher</td><td>A12222</td><td></td></tr><tr><td>Epon 812</td><td>Electron Microscopy Sciences</td><td>14120</td><td></td></tr><tr><td>Ethanol</td><td>Merck</td><td>100983</td><td></td></tr><tr><td>Fugene HD</td><td>Promega</td><td>E2311</td><td>Plasmid transfection</td></tr><tr><td>Glycine</td><td>SIGMA</td><td>G8898</td><td></td></tr><tr><td>Lead Citrate 3%</td><td>Electron Microscopy Sciences</td><td>22410</td><td></td></tr><tr><td>Osmium tetroxide 4 % aqueous solution</td><td>Electron Microscopy Sciences</td><td>16320</td><td>Very toxic. Use only in fume hood.</td></tr><tr><td>Potassium hexacyanoferrate(II) trihydrate&amp;nbsp;</td><td>SIGMA</td><td>P3289</td><td></td></tr><tr><td>Sodium cacodylate trihydrate</td><td>Sigma-Aldrich</td><td>C0250-500G</td><td></td></tr><tr><td>Uranyl acetate</td><td>Electron Microscopy Sciences</td><td>22400</td><td></td></tr><tr><td>UranyLess EM stain</td><td>Electron Microscopy Sciences</td><td>22409</td><td></td></tr><tr><td>&lt;strong&gt;Plasmid construction&lt;/strong&gt;</td><td></td><td></td><td></td></tr><tr><td>Mito-mEosEM</td><td>addgene</td><td>132706</td><td>DOI:&amp;nbsp;10.1016/j.chembiol.2024.02.007</td></tr><tr><td>TMEM192-V5-APEX2</td><td>Sharma N.&amp;nbsp; et al.</td><td>N/A</td><td>DOI:&amp;nbsp;10.1016/j.chembiol.2024.02.007</td></tr><tr><td>&lt;strong&gt;Tools&lt;/strong&gt;</td><td></td><td></td><td></td></tr><tr><td>0.22 um syringe filter</td><td>Sartorius</td><td>16534</td><td></td></tr><tr><td>35 mm Gridded coverslip dish</td><td>Mattek</td><td>P35G-1.5-14-CGRD</td><td></td></tr><tr><td>BEEM Embedding Capsules</td><td>Ted Pella</td><td>130</td><td></td></tr><tr><td>Formvar Supported Single slot copper grid</td><td>Ted Pella</td><td>01705</td><td></td></tr><tr><td>Diamond knife</td><td>DiATOME</td><td>DU4530</td><td></td></tr><tr><td>Ultra-microtome</td><td>Leica</td><td>ARTOS 3D</td><td></td></tr><tr><td>&lt;strong&gt;Microscopes&lt;/strong&gt;</td><td></td><td></td><td></td></tr><tr><td>Inverted confocal microscopy</td><td>Nikon</td><td>A1 Rsi/Ti-E</td><td></td></tr><tr><td>Transmission electron microscopy</td><td>FEI</td><td>Tecnai G2</td><td></td></tr><tr><td>&lt;strong&gt;Software and Algorithms&lt;/strong&gt;</td><td></td><td></td><td></td></tr><tr><td>Fiji/Image J</td><td>NIH / open source</td><td></td><td>https://imagej.nih.gov/ij/</td></tr><tr><td>ImageJ BigWarp software package</td><td>NIH / open source</td><td></td><td>https://imagej.net/plugins/bigwarp</td></tr><tr><td>Photozoom Pro 8&amp;nbsp;</td><td>BenVista</td><td></td><td>https://www.benvista.com/photozoompro&lt;br/&gt; &lt;br/&gt; &lt;strong&gt;Alternative software&lt;br/&gt; Image Resizer &lt;/strong&gt;: https://imageresizer.com/&lt;br/&gt; &lt;strong&gt;Gigapixel AI&lt;/strong&gt; : https://www.topazlabs.com/gigapixel&lt;br/&gt; &lt;strong&gt;ON1 resize &lt;/strong&gt;: https://www.on1.com/products/resize-ai/</td></tr></tbody></table>

dual-color correlative light and electron microscopy for the visualization of interactions between mitochondria and lysosomes

Cellular organelles, such as mitochondria and lysosomes, display dynamic structures. Despite the higher resolution of transmission electron microscopy for structural analysis, light microscopy is essential for the visualization of dynamic organelles by target-specific labeling. The following protocol describes a method that combines dual-color correlative light and electron microscopy (CLEM) to observe the interactions between mitochondria and lysosomes. In this study, mitochondria were labeled with mEosEM (Mito-mEosEM) and lysosomes with TMEM192-V5-APEX2. The results obtained from CLEM images enable us to observe the changes in the interactions between mitochondria and lysosomes under external stress conditions. Treatment with bafilomycin (BFA), which inhibits lysosomal function, resulted in an increase in contact between mitochondria and lysosomes, leading to the formation of fragmented mitochondria trapped inside lysosomes. Conversely, treatment with U18666A, which inhibits cholesterol export from lysosomes, caused lysosomes to be surrounded by mitochondria, indicating a distinct form of interaction. This study presents an effective method for observing the interactions between mitochondria and lysosomes in fixed cells. Furthermore, CLEM imaging with dual-color probes offers a powerful tool for future investigations of organelle dynamics and their implications for cell function and pathology.

To visualize the interactions between mitochondria and lysosomes using this dual-color CLEM protocol, we employed Mito-mEosEM and TMEM192-V5-APEX2. Fixed cells were first imaged using a light microscope, followed by EM sampling and imaging. The results of the correlated images are shown in Figure 3A,B and Figure 4A. Following the protocol described above, we checked whether we could observe the changes in the interaction between mitochondria and lysosomes under external stress.
Treatment of cells with BFA, a lysosomal V-ATPase inhibitor, inhibits autophagy, resulting in reduced mitochondrial quality and increased interactions between mitochondria and lysosomes27. In BFA-treated cells, we observed mitochondria trapped within lysosomes (Figure 3B), compared to DMSO-treated control cells (Figure 3A). Although it is difficult to distinguish fragmented electron-dense mitochondria and cellular compartments within lysosomes in electron micrographs alone, they can be easily distinguished in correlated images (Figure 3B1,B2). The mitochondrial disruption structure (Figure 3A1,A2; blue arrow) observed in the mitochondria of DMSO-treated cells is not related to lysosomes, and the structure in the fluorescence image that appears to be a lysosome trapped inside a large mitochondrion is a structure where the mitochondrion surrounds the lysosome (Figure 3A2; red arrow).
Next, we tested the effects of U18666A (2 &#181;g/mL for 18 h), a well-known inhibitor of NPC1 that blocks the movement of cholesterol out of lysosomes28. Cells treated with U18666A showed increased direct interaction between mitochondria and lysosomes (Figure 4). In addition, the contacts between mitochondria and lysosomes in U18666A-treated samples were different from those in BFA-treated samples. In HeLa cells treated with BFA, the fragmented mitochondrion is completely trapped in the lysosome, a previously reported mitophagy morphology12,29. However, in the U18666A-treated group, some of the TMEM192-positive lysosomes appear engulfed by mitochondrion (Figure 4D,E)30,31.
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/67020/67020fig01.jpg" /> 
Figure 1: Schematic overview of TEM-based CLEM by using a genetically tagged fluorescence probe. (A) HeLa cells were grown in 35 mm glass grid-bottom culture dishes. (B) Fixed cells were labeled with Amplex-Red by APEX2 protein. (C) Fluorescence images were acquired with a confocal microscope. A motorized stage and tile-scan function were used to capture a large overview image and a high-resolution image of the target cell. The large image serves as a navigation map to identify the location of the target cell in plastic-embedded specimens. (D) Proceed with post fixation and dehydration embedding. (E) Embedding was done using a PCR tube or BEEM capsule filled with resin over the target cell on the coverslip. (F) After resin polymerization, removing the coverslip left an imprinted grid pattern on the bottom surface of the block. (G) The target cell was identified by the imprinted pattern on the resin surface, and the target cell was sectioned at approximately 60 nm thickness using an ultramicrotome. (H) Serial sections were collected on a formvar-coated copper slot grid. (I) The grids were imaged using transmission electron microscopy. Abbreviations: TEM = transmission electron microscopy; CLEM = correlative light and electron microscopy; ROI = region of interest. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/67020/67020fig01large.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/67020/67020fig02.jpg" /> 
Figure 2: Image processing and generation of the CLEM image using imageJ Fiji. (A1) Main GUI of Fiji and TrakEM2. (A2) Import of the tile image dataset acquired with TEM at 1,700x magnification. (A3) Alignment and example of a stitched image using TrakeEM2&#39;s image alignment method. (B1-B3) Conversion of small light micrograph into large high-quality images using photo enlargement software. (C1-C3) Workflow for registration between light microscopy images and electron microscopy images using the Big Wrap plug-in software. (D1-D3) The process of overlaying a transformed FM image and an EM image to synthesize a correlated microscope image. Abbreviations: TEM = transmission electron microscopy; CLEM = correlative light and electron microscopy; FM = fluorescence microscopy. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/67020/67020fig02large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 3" class="xfigimg" src="/files/ftp_upload/67020/67020fig03.jpg" /> 
Figure 3: CLEM images of the mitochondrial and lysosomal contact sites treated with or without bafilomycin A1. HeLa cells were co-transfected with Mito-mEosEM (mitochondria; green) and TMEM192-APEX2 (lysosomes; red) and then stained with Amplex-Red dye. (A) Untreated HeLa cells. Molecular information from the fluorescence signal (left panel) and ultrastructural information obtained by TEM (middle panel) were superimposed (right panel) based on mitochondrial signal and morphology. (A1,A2) In untreated HeLa cells, the mitochondrial inner membrane is disrupted (blue arrow), or the mitochondria and lysosomes are very close to each other (red arrow). (B1,B2) In HeLa cells treated with bafilomycin A1 for 18 h, mitophagy, characterized by the trapping of undegraded mitochondria in lysosomes, was observed (green arrow). Scale bars = 1 &#181;m (A1,A2,B1,B2). <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/67020/67020fig03large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 4" class="xfigimg" src="/files/ftp_upload/67020/67020fig04.jpg" /> 
Figure 4: CLEM images of the mitochondrial and lysosomal contact sites treated with U18666A&#160;(A) Two-color CLEM images of Mito-mEosEM and TMEM192-V5-APEX2 (Amplex-Red-labeled) expressed HeLa cells under U18666A treatment (2 &#181;g/mL for 18 h). (A1,A2) Red arrows indicate contact sites between mitochondria and lysosomes. Scale bars = 1 &#181;m. (B1-B3,C1-C3) Red arrow indicates the appearance of inter-organelle fusion between mitochondria and lysosomes by serial images. Scale bars = 500 nm (B1-B3). Scale bars = 1 &#181;m (C1-C3). (D,E) High-magnification TEM images of the fusion sites between mitochondria and lysosomes. Pseudocolors are used to distinguish between mitochondrion (green) and lysosome (red). Scale bars = 100 nm. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/67020/67020fig04large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

Dual-color Correlative Light and Electron Microscopy for the Visualization of Interactions between Mitochondria and Lysosomes

This protocol describes a mechanism for using correlative light and electron microscopy to visualize the interaction of mitochondria and lysosomes labeled with mEosEM and APEX2, respectively.

Cellular organelles, such as mitochondria and lysosomes, display dynamic structures. Despite the higher resolution of transmission electron microscopy for ...

dual-color-correlative-light-electron-microscopy-for-visualization

Nanyang Technological University

Research

JoVE Journal

Neuroscience

524 Views.  Korea Brain Research Institute. Mitochondriale disfunctie wordt in verband gebracht met veel ziekten, met name neurodegeneratieve en metabole stoornissen. Daarom, als we kunnen begrijpen hoe mitochondriën worden gereguleerd door lysosomen, helpt dit om nieuwe therapeutische strategieën te ontwikkelen. In dit protocol hebben we een tweekleurige CLEM-methode gedemonstreerd om mitochondriaal lysosoomcontact te visualiseren.Correlatieve licht- en elektronenmicroscopie is een beeldvormi...

Video: Dual-color Correlative Light and Electron Microscopy for the Visualization of Interactions between Mitochondria and Lysosomes

Correlative Microscopy for 3D Structural Analysis of Dynamic Interactions

Cryo-electron tomography (cryoET) allows 3D visualization of cellular structures at molecular resolution in a close-to-physiological state1. However, direct visualization of individual viral complexes in their host cellular environment with cryoET is challenging2, due to the infrequent and dynamic nature of viral entry, particularly in the case of HIV-1. While time-lapse live-cell imaging has yielded a great deal of information about many aspects of the life cycle of HIV-13-7, the resolution afforded by live-cell microscopy is limited (~ 200 nm). Our work was aimed at developing a correlation method that permits direct visualization of early events of HIV-1 infection by combining live-cell fluorescent light microscopy, cryo-fluorescent microscopy, and cryoET. In this manner, live-cell and cryo-fluorescent signals can be used to accurately guide the sampling in cryoET. Furthermore, structural information obtained from cryoET can be complemented with the dynamic functional data gained through live-cell imaging of fluorescent labeled target.
In this video article, we provide detailed methods and protocols for structural investigation of HIV-1 and host-cell interactions using 3D correlative high-speed live-cell imaging and high-resolution cryoET structural analysis. HeLa cells infected with HIV-1 particles were characterized first by confocal live-cell microscopy, and the region containing the same viral particle was then analyzed by cryo-electron tomography for 3D structural details. The correlation between two sets of imaging data, optical imaging and electron imaging, was achieved using a home-built cryo-fluorescence light microscopy stage. The approach detailed here will be valuable, not only for study of virus-host cell interactions, but also for broader applications in cell biology, such as cell signaling, membrane receptor trafficking, and many other dynamic cellular processes.

We describe a correlative microscopy method that combines high-speed 3D live-cell fluorescent light microscopy and high-resolution cryo-electron tomography. We demonstrate the capability of the correlative method by imaging dynamic, small HIV-1 particles interacting with host HeLa cells.

Cryo-electron tomography (cryoET) allows 3D visualization of cellular structures at molecular resolution in a close-to-physiological state1. However, ...

Bioengineering

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

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 ...

Correlative Light and Electron Microscopy (CLEM) as a Tool to Visualize Microinjected Molecules and their Eukaryotic Sub-cellular Targets

The eukaryotic cell relies on complex, highly regulated, and functionally distinct membrane bound compartments that preserve a biochemical polarity necessary for proper cellular function. Understanding how the enzymes, proteins, and cytoskeletal components govern and maintain this biochemical segregation is therefore of paramount importance. The use of fluorescently tagged molecules to localize to and/or perturb subcellular compartments has yielded a wealth of knowledge and advanced our understanding of cellular regulation. Imaging techniques such as fluorescent and confocal microscopy make ascertaining the position of a fluorescently tagged small molecule relatively straightforward, however the resolution of very small structures is limited 1.
On the other hand, electron microscopy has revealed details of subcellular morphology at very high resolution, but its static nature makes it difficult to measure highly dynamic processes with precision 2,3. Thus, the combination of light microscopy with electron microscopy of the same sample, termed Correlative Light and Electron Microscopy (CLEM) 4,5, affords the dual advantages of ultrafast fluorescent imaging with the high-resolution of electron microscopy 6. This powerful technique has been implemented to study many aspects of cell biology 5,7. Since its inception, this procedure has increased our ability to distinguish subcellular architectures and morphologies at high resolution. 
Here, we present a streamlined method for performing rapid microinjection followed by CLEM (Fig. 1). The microinjection CLEM procedure can be used to introduce specific quantities of small molecules and/or proteins directly into the eukaryotic cell cytoplasm and study the effects from millimeter to multi-nanometer resolution (Fig. 2). The technique is based on microinjecting cells grown on laser etched glass gridded coverslips affixed to the bottom of live cell dishes and imaging with both confocal fluorescent and electron microscopy. Localization of the cell(s) of interest is facilitated by the grid pattern, which is easily transferred, along with the cells of interest, to the Epon resin used for immobilization of samples and sectioning prior to electron microscopy analysis (Fig. 3). Overlay of fluorescent and EM images allows the user to determine the subcellular localization as well as any morphological and/or ultrastructural changes induced by the microinjected molecule of interest (Fig. 4). This technique is amenable to time points ranging from &le;5 s up to several hours, depending on the nature of the microinjected sample.

Correlative Light and Electron Microscopy CLEM as a Tool to Visualize Microinjected Molecules and their Eukaryotic Sub-cellular Targets

The CLEM technique has been adapted to analyze ultrastructural morphology of membranes, organelles, and subcellular structures affected by microinjected molecules. This method combines the powerful techniques of micromanipulation/microinjection, confocal fluorescent microscopy, and electron microscopy to allow millimeter to multi-nanometer resolution. This technique is amenable to a wide variety of applications.


The eukaryotic cell relies on complex, highly regulated, and functionally distinct membrane bound compartments that preserve a biochemical polarity ...

Biology

Multicolor Flow Cytometry-based Quantification of Mitochondria and Lysosomes in T Cells

T cells utilize different metabolic programs to match their functional needs during differentiation and proliferation. Mitochondria are crucial cellular components responsible for supplying cell energy; however, excess mitochondria also produce reactive oxygen species (ROS) that could cause cell death. Therefore, the number of mitochondria must constantly be adjusted to fit the needs of the cells. This dynamic regulation is achieved in part through the function of lysosomes that remove surplus/damaged organelles and macromolecules. Hence, cellular mitochondrial and lysosomal contents are key indicators to evaluate the metabolic adjustment of cells. With the development of probes for organelles, well-characterized lysosome or mitochondria-specific dyes have become available in various formats to label cellular lysosomes and mitochondria. Multicolor flow cytometry is a common tool to profile cell phenotypes, and has the capability to be integrated with other assays. Here, we present a detailed protocol of how to combine organelle-specific dyes with surface markers staining to measure the amount of lysosomes and mitochondria in different T cell populations on a flow cytometer.

This article illustrates a powerful method to quantify mitochondria or lysosomes in living cells. The combination of lysosome- or mitochondria-specific dyes with fluorescently conjugated antibodies against surface markers allows the quantification of these organelles in mixed cell populations, like primary cells harvested from tissue samples, by using multicolor flow cytometry.

T cells utilize different metabolic programs to match their functional needs during differentiation and proliferation. Mitochondria are crucial cellular ...

Immunology and Infection

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 ...

Study of Endoplasmic Reticulum and Mitochondria Interactions by In Situ Proximity Ligation Assay in Fixed Cells

Structural interactions between the endoplasmic reticular (ER) and mitochondrial membranes, in domains known as mitochondria-associated membranes (MAM), are crucial hubs for cellular signaling and cell fate. Particularly, these inter-organelle contact sites allow the transfer of calcium from the ER to mitochondria through the voltage-dependent anion channel (VDAC)/glucose-regulated protein 75 (GRP75)/inositol 1,4,5-triphosphate receptor (IP3R) calcium channeling complex. While this subcellular compartment is under intense investigation in both physiological and pathological conditions, no simple and sensitive method exists to quantify the endogenous amount of ER-mitochondria contact in cells. Similarly, MAMs are highly dynamic structures, and there is no suitable approach to follow modifications of ER-mitochondria interactions without protein overexpression. Here, we report an optimized protocol based on the use of an in situ proximity ligation assay to visualize and quantify endogenous ER-mitochondria interactions in fixed cells by using the close proximity between proteins of the outer mitochondrial membrane (VDAC1) and of the ER membrane (IP3R1) at the MAM interface. Similar in situ proximity ligation experiments can also be performed with the GRP75/IP3R1 and cyclophilin D/IP3R1 pairs of antibodies. This assay provides several advantages over other imaging procedures, as it is highly specific, sensitive, and suitable to multiple-condition testing. Therefore, the use of this in situ proximity ligation assay should be helpful to better understand the physiological regulations of ER-mitochondria interactions, as well as their role in pathological contexts.

Study of Endoplasmic Reticulum and Mitochondria Interactions by In Situ Proximity Ligation Assay in Fixed Cells

Here, we describe a procedure to visualize and quantify with high sensitivity the endogenous interactions between the endoplasmic reticulum and mitochondria in fixed cells. The protocol features an optimized in situ proximity ligation assay targeting the inositol 1,4,5-triphosphate receptor/glucose-regulated protein 75/voltage-dependent anion channel/cyclophilin D complex at the mitochondria-associated membrane interface.

Structural interactions between the endoplasmic reticular (ER) and mitochondrial membranes, in domains known as mitochondria-associated membranes (MAM), ...

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 ...

Visualizing Mitophagy with Fluorescent Dyes for Mitochondria and Lysosome

Mitochondria, being the powerhouses of the cell, play important roles in bioenergetics, free radical generation, calcium homeostasis, and apoptosis. Mitophagy is the primary mechanism of mitochondrial quality control and is generally studied using microscopic observation, however in vivo mitophagy assays are difficult to perform. Evaluating mitophagy by imaging live organelles is an alternative and necessary method for mitochondrial research. This protocol describes the procedures for using the cell-permeant green-fluorescent mitochondria dye MitoTracker Green and the red-fluorescent lysosome dye LysoTracker Red in live cells, including the loading of the dyes, visualization of the mitochondria and the lysosome, and expected outcomes. Detailed steps for the evaluation of mitophagy in live cells, as well as technical notes about microscope software settings, are also provided. This method can help researchers observe mitophagy using live-cell fluorescent microscopy. In addition, it can be used to quantify mitochondria and lysosomes and assess mitochondrial morphology.

Mitophagy is the primary mechanism of mitochondrial quality control. However, the evaluation of mitophagy in vivo is hindered by the lack of reliable quantitative assays. Presented here is a protocol for the observation of mitophagy in living cells using a cell-permeant green-fluorescent mitochondria dye and a red-fluorescent lysosome dye.

Mitochondria, being the powerhouses of the cell, play important roles in bioenergetics, free radical generation, calcium homeostasis, and apoptosis. ...

Ultrastructural Localization of Endogenous LC3 by On-Section Correlative Light-Electron Microscopy

The visualization of autophagic organelles at the ultrastructural level by electron microscopy (EM) is essential to establish their identity and reveal details that are important for understanding the autophagic process. However, EM methods often lack molecular information, obstructing the correlation of ultrastructural information obtained by EM to fluorescence microscopy-based localization of specific autophagy proteins. Furthermore, the rarity of autophagosomes in unaltered cellular conditions hampers investigation by EM, which requires high magnification, and hence provides a limited field of view.
In answer to both challenges, an on-section correlative light-electron microscopy (CLEM) method based on fluorescent labeling was applied to correlate a common autophagosomal marker, LC3, to EM ultrastructure. The method was used to rapidly screen cells in fluorescence microscopy for LC3 labeling in combination with other relevant markers. Subsequently, the underlying ultrastructural features of selected LC3-labeled spots were identified by CLEM. The method was applied to starved cells without adding inhibitors of lysosomal acidification.
In these conditions, LC3 was found predominantly on autophagosomes and rarely in autolysosomes, in which LC3 is rapidly degraded. These data show both the feasibility and sensitivity of this approach, demonstrating that CLEM can be used to provide ultrastructural insights on LC3-mediated autophagy in native conditions-without drug treatments or genetic alterations. Overall, this method presents a valuable tool for ultrastructural localization studies of autophagy proteins and other scarce antigens by bridging light microscopy to EM data.

Here, we present a protocol for optimized on-section correlative light-electron microscopy based on endogenous, fluorescent labeling as a tool to investigate the localization of rare proteins in relation to cellular ultrastructure. The power of this approach is demonstrated by ultrastructural localization of endogenous LC3 in starved cells without Bafilomycin treatment.

The visualization of autophagic organelles at the ultrastructural level by electron microscopy (EM) is essential to establish their identity and reveal ...

Dual-color Correlative Light and Electron Microscopy for the Visualization of Interactions between Mitochondria and Lysosomes

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