Our sincere appreciation is expressed to the following people: Drs. Sam Johnson and Benjamin Carlson of the Duke Light Microscopy Core Facility for their assistance with data visualization software, and Ms. Valerie Lapham and Dr. John M. Mackenzie, Jr. of the Center for Electron Microscopy at North Carolina State University for their advice on electron microscopy. We thank Dr. Elaine B. Boh&#243;rquez for her editorial assistance. Authors contributed in the following manner: DVB, SM, and RAL designed experiments and analyzed data. DVB performed experiments and FH performed manual rendering of data. SM is director of Renovo Neural, where SBEM data was acquired. DVB wrote the manuscript and all authors reviewed and edited the final manuscript. This work was supported by NIH grants R01DK091946 and Veterans Affairs grant I01BX002230 to RAL, and F32DK094704, to DVB.

All animal care and experiments were done in accordance with a protocol approved by the Duke University Institutional Animal Care and Use Committee.
Cocem3D stemmed from a combination of previously published protocols10,11 and was applied here to study a specific enteroendocrine cell using a PyyGFP reporter mouse12. This method can readily be applied to other cells of interest using commercially available transgenic reporter mice. The method is described in three sections: Correlative confocal microscopy, serial block-face scanning electron microscopy (SBEM), and data rendering in 3D.
Correlative Confocal Microscopy
The objective of this section is to harvest a tissue segment from the distal colon of dimensions that allow correlative confocal and SBEM imaging. The protocol is as follows:
1. Harvesting the Tissue
<ol>
	<li>Prepare 10 ml&#160;of 50 &#181;g/ml heparin solution in PBS.</li>
	<li>Prepare xylazine/ketamine anesthetic stock by mixing 10 ml of Ketamine (100 mg/ml) and 1 ml of xylazine (20 mg/ml).
	<ol>
		<li>Dilute xylazine/ketamine stock 1:4 in 0.1M PBS.</li>
	</ol>
	</li>
	<li>Prepare a 100 ml of fixative solution containing 4% paraformaldehyde and 0.1% glutaraldehyde in PBS.</li>
	<li>Anesthetize a PyyGFP mouse, 6 to 10 weeks old, with a lethal dose of xylazine/ketamine anesthetic. Inject the anesthetic intraperitoneally at 0.15 ml per 20 g of mouse weight.</li>
	<li>Confirm proper mouse anesthetization by pinching the tail or toes, and use ointment on eyes to prevent dryness.</li>
	<li>Use a variable flow peristaltic pump to perfuse fixative intracardially10. Using surgical scissors (13 cm length) cut open the abdominal cavity to expose the intestines, heart, and lungs. Hold heart with narrow-pattern curved forceps (12 cm length) and insert butterfly needle (19 G) from peristaltic pump into left ventricle. Immediately, cut right atrium with spring straight scissors (length 4 mm). Perfuse at a rate of 2 ml/min first with heparin solution for 1 min, and then with ice-cold fixative solution for 15 min until mouse&#8217;s tail is completely rigid. 
	Note: It is very important to perfuse at a slow rate to prevent bursting of small vessels in the intestinal mucosa. Bursting of vessels will compromise the tissue&#8217;s ultrastructure. If perfusion is adequate, the liver should turn pale in color within 3-5 min.</li>
	<li>Using small scissors open the abdominal cavity and excise the entire colon from the junction with the cecum to the distal rectum. Place the tissue in ice-cold PBS. While submerged in PBS, cut open with small spring scissors the colon along the mesentery.</li>
</ol>
2. Dissecting and Fixing Tissue Segments
<ol>
	<li>Using a scalpel, cut about 6 small tissue segments (2 mm2) from the distal colon. Do this on a sheet of dental wax and submerged in few drops of PBS.</li>
	<li>Post-fix the tissue segments in fixative solution for 3 hr at 4 &#176;C.</li>
</ol>
3. Micro-dissecting Tissue Blocks
<ol>
	<li>Prepare 5% low-melting agarose in PBS and keep it in a water bath at 45 &#176;C.</li>
	<li>Embed the tissue segments in 5 % low-melting agarose using a small plastic container, like the standard size Tissue-Tek Cryomold.</li>
	<li>Mount the embedded sections on a vibrating blade microtome and fill the buffer tray with ice-cold PBS.</li>
	<li>Cut 300 &#181;m tissue strips at 0.8 amplitude and 0.04 mm/sec speed. Note: At this point, the tissue strips will be dislodged from the agarose.</li>
	<li>Re-embed 300 &#181;m tissue strips in agarose and mount them on microtome perpendicularly to the vibratome&#8217;s blade.</li>
	<li>Using a vibratome, cut tissue strips at a thickness of 50 &#181;m. Note: The final tissue blocks should be about 300 &#181;m wide x 50 &#181;m thick. These dimensions are optimized from pilot confocal experiments that showed that the thickness of enteroendocrine cell is between 15-20 &#181;m and the neuropod&#8217;s length 30-70 nm. Therefore, an entire cell could be contained within a block of tissue 300 &#181;m wide by 50 &#181;m thick if the cell&#8217;s orientation is parallel to the face of the tissue block. Vary these dimensions depending on the tissue and cell type of interest.</li>
	<li>Store the tissue blocks in PBS at 4 &#176;C.</li>
</ol>
4. Confocal Imaging
<ol>
	<li>Prepare a nuclear stain solution by diluting DAPI in PBS at 1:4,000.</li>
	<li>Incubate tissue blocks in DAPI nuclear stain for 5 min. Note: DAPI staining facilitates distinguishing individual cells by their nuclei, which is helpful for correlating the confocal and SBEM images.</li>
	<li>Mount tissue blocks on charged glass slide, add a few drops of PBS, and cover them with a coverslip.</li>
	<li>Using a confocal microscope, identify blocks with intact villi and PyyGFP cells of interest and imaged them to obtain z-stacks optical sections.
	<ol>
		<li>Use a 20X/0.8 Zeiss Plan Apochromat objective to obtain z-stacks of 1 &#181;m optical sections.</li>
		<li>For the z-stack, use the channels for 405 nm (DAPI) to determine the relationship to other cells, 488 nm endogenous GFP to localize the cell of interest, and differential interference contrast (DIC) to determine location of the cell relative to the lumen.</li>
		<li>Use image resolution of 1,024 pixels or higher.</li>
	</ol>
	</li>
</ol>
5.&#160; Embedding Blocks in Agarose
<ol>
	<li>Prepare 10 ml of fixative containing 4% paraformaldehyde and 2.5% glutaraldehyde in PBS.</li>
	<li>Remove the tissue blocks from the glass slide by carefully adding PBS in the edges of the cover slip to slide the coverslip away from the glass slide without damaging the tissue block. Then, use a fine art paintbrush to transfer the tissue block from the slide to a 1.5 ml microcentrifuge tube containing 4% paraformaldehyde and 2.5% glutaraldehyde fixative.</li>
	<li>Post-fix tissue blocks O/N at 4 &#176;C.</li>
	<li>Transfer tissue blocks to PBS. Then, embed blocks flat in 5% low-melting agarose by sandwiching them between two glass slides. 
	Note: Embedding the blocks in a thin layer of agarose eases posterior manipulation during staining.</li>
	<li>Cut the agarose in a square and make a notch on the upper hand side. Note: This step is necessary to maintain the orientation in subsequent steps.</li>
	<li>Store blocks in a 1.5 ml microcentrifuge tube with PBS at 4 &#176;C until further processing.</li>
</ol>
Serial Block-face Scanning Electron Microscopy (SBEM)
In this section, the tissue block is prepared and imaged with SBEM at low-magnification. The survey image of the block face is then correlated with the confocal data to identify the region containing the cell of interest. Once the region is identified, the tissue is imaged at a resolution of 7 nm/pixel and slices of 70 nm. This was sufficient to resolve and distinguish large-dense core secretory vesicles from other organelles. In enteroendocrine cells, this type of vesicles varies between 100 and 150 nm in diameter13. The protocol is as follows:
6. Staining Tissue Sections
<ol>
	<li>Remove tissue from PBS and rinse three times, 5 min each, in 0.1 M cacodylate buffer.</li>
	<li>Stain blocks for 1 hr in 0.1 % tannic acid dissolved in 0.1 M cacodylate buffer to enhance contrast of cell membranes14.</li>
	<li>Carry out subsequent staining and dehydration of tissue according to the published protocol11. Following Deerinck et al. protocol, prepare these solutions: 1) 1% (w/v) thiocarbonhydrazide (THC); 2) lead aspartate solution; 3) 1% uranyl acetate, and 4) osmium tetraoxyde/ potassium ferrocyanide (OsO4/K Ferrocyanide). Mix 4% OsO4 and 2x K Ferrocyanide stock [0.3g K Ferrocyanide and 0.86g Na Cacodylate in 10 ml H2O] at 1:1 ratio to make OsO4/K Ferrocyanide. Note: Having the tissue embedded in agarose facilitates handling during staining. Carefully remove agarose for subsequent resin infiltration.</li>
	<li>Rinse samples three times, 5 min each, in 0.1 M cacodylate buffer and then stain with OsO4/K Ferrocyanide solution for 2 hr at 4 &#176;C.</li>
	<li>Rinse in ultra pure water three times, 5 min each, and stain with TCH for 30 min at 60 &#176;C.</li>
	<li>Rinse in ultra pure water three times, 5 min each and stain with 2% OsO4 (no K Ferrocyanide) for 60 min at RT. Rinse again in ultra pure water three times, 5 min each.</li>
	<li>Stain with 1% uranyl acetate O/N at 4 &#176;C. Rinse in ultra pure water three times, 5 min each.</li>
	<li>Stain with lead aspartate for 30 min at 60 &#176;C. Rinse in ultra pure water three times, 5 min each.</li>
	<li>Dehydrate tissues by rinsing in solutions with increasing concentrations of ethanol. Rinse 2 times, 5 min each, with 50%, 75%, 85%, 95%, and absolute 100% ethanol. At the end, rinse sections with propylene oxide two times, 10 min each.</li>
</ol>
7. Infiltrating and Embedding Tissue Sections in Resin
<ol>
	<li>Infiltrate the blocks with resin using the commercially avaialble kit. Embed tissues in resin using an epon embedding kit. Prepare a resin mix of 48% Epoxy, 20% ml DDSA, 30% ml NMA, and 2% DMP30. Agitate resin mixture vigorously for 5 min and then place mix in vacuum for 30 min to allow bubbles to come to the surface.</li>
	<li>Mix resin with propylene oxide at 1:1 ratio and shake vigorously. Remove the final dehydration rinse from tissues, and add resin mixed with propylene oxide. Place vial with samples in a rotator O/N and leave vial uncapped. The next day, add freshly made epon resin and mix for 90 min in rotator.</li>
	<li>Embed blocks in resin as flat as possible by sandwiching them in between glass slides that have been cured with liquid release agent to prevent them from sticking to each other15.</li>
	<li>Once the blocks are embedded flat, cure the blocks for an addition 48 hr at 60 &#176;C.</li>
	<li>Pull slides apart to release the blocks.</li>
</ol>
8. Mounting and Trimming the Tissue Block for Imaging
<ol>
	<li>Under a dissecting scope, match the orientation of the tissue blocks embedded in resin with that of the confocal micrographs to facilitate identifying the regions containing the cells of interest before trimming the block.</li>
	<li>Trim the resin embedded block manually down to a ~500 x 500 &#181;m block-face.</li>
	<li>Mount the block flat on a pin containing conductive epoxy and dry for 30 min. Then, lay the block flat on a surface and dry O/N&#160;at 60 &#176;C.</li>
	<li>Coat the block with colloidal silver liquid. Maintain the tissue sections flat to ensure slicing of the block at a right angle to facilitate correlating the serial block-face and confocal micrographs.</li>
</ol>
9. SBEM
<ol>
	<li>Image the block using a Scanning Electron Microscope equipped with a SBEM system (e.g., 3View).</li>
	<li>Set initial thickness section at ~2 &#181;m increments and cut until the face of tissue emerges from the block.</li>
	<li>Acquire a surver image of the entire block face.</li>
	<li>Using Fiji software, take measurements of both the serial block-face and confocal micrographs to generate a common denominator and account for sample warping during fixation and staining16.</li>
	<li>Locate the area of interest on the block face by multiplying the denominator to the coordinates in the confocal images. Use also other structural features, like the position of microvilli, goblet cells, or lamina propria, as a reference.</li>
	<li>Image region of interest at a 2.25 kV and 7 nm/pixel (or 15,147X magnification) in high vacuum mode. Slice increments should be set at 70 nm or less.</li>
	<li>Collect SBEM data in raw 16-bit .dm3 format.</li>
</ol>
10. Optimizing SBEM Images for Surface Segmentation
<ol>
	<li>Convert SBEM images from raw .dm3 format to 8-bit .tiff.</li>
	<li>Filter .tiff images using a 0.8 gaussian blur filter in Fiji.</li>
	<li>Scale down the data set to 25 % of the original size and save it as a .tiff stack to minimize the amount of RAM memory necessary to handle the set.</li>
	<li>Align the stack of SBEM images using the Fiji plugin &#8220;linear stack alignment with SIFT&#8221; in translation mode and cropped using the &#8220;crop 3D&#8221; plugin. Note: Cropping helps to further reduced the amount of RAM memory necessary for segmenting and volume rendering.</li>
</ol>
Data Rendering in 3D
11. Confocal Microscopy
<ol>
	<li>Reconstruct z-stacks using the automatic surpass mode of the &#8220;surfaces&#8221; tool. Figure 1D, shows a reconstruction of each channel using the smooth option on, the surfaces area detail at 0.126 &#181;m, and thresholding as absolute intensity.</li>
</ol>
12. Serial Block-face SEM
Note: Manual data segmentation is a very time consuming procedure and depending on the features to be rendered the procedure can take several weeks. Segmentation for the videos and figures presented in Boh&#243;rquez et al. 20148 was done manually and took about 500 hr of labor. It is recommended to prioritize the essential features needing rendering before beginning the segmentation process. The procedure is as follows:
<ol>
	<li>Use the &#8220;surfaces&#8221; tool in draw mode and the &#8220;contour&#8221; option at drawing mode of 50 msec to segment manually and volume rendered the cell of interest.</li>
	<li>Trace contours on every slice of the cell for smoother rendering or every 5 slices for faster rendering.</li>
	<li>Export final images using the &#8220;snapshot&#8221; tool at a resolution of 300 dpi or more.</li>
</ol>

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Dr. Satish Medicetty is an employee of and receives 100% salary from Renovo Neural Inc; however, this does not alter the author&#8217;s adherence to JoVE policies on sharing data and materials. No conflict of interest is declared for remaining authors.

Gastrointestinal chemosensation is emerging as an exciting new field in biomedical research. This is in great part due to the discovery of functional taste receptors in enteroendocrine cells18. Subsequent studies have shown that enteroendocrine cells express specific receptors for nutrients, including carbohydrates, lipids, and aminoacids5,6,19,20. The catalytic factor for these discoveries has been the development of reporter mice, in which enteroendocrine cells are fluorescently labeled20. We developed one of these mice, the PyyGFP mouse, to study enteroendocrine cells of the distal small intestine and colon17,21. These cells are of interest because they secrete PYY and glucagon-like peptide 1, both of which are inducers of satiety22,23. At the time, a complete ultra-structural account of these cells was missing, which we believed was essential to understand their signaling mechanisms.
Here, we described a visually a method to bridge confocal microscopy with SBEM. The method is referred as Cocem3D, which allowed us to document the complete ultrastructure of enteroendocrine cells. We have reported that these cells have a neuropod that contains three-quarters of all of the secretory vesicles8. Within the neuropod, there are neurofilaments and much like neuronal axons, neuropods are nurtured by enteric glial cells8. More important, it is though these neuropods that enteroendocrine cells physically connect to neurons innervating the intestine and colon9.
One of the strengths of cocem3D is its simplicity. Reducing the tissue sample into a block that can be imaged by confocal and SBEM facilitates the identification of a specific cell within a tissue. Secretory vesicles and other organelles can be identified with ease at a resolution of 7 nm/pixel. Because the sections in SBEM are discarded, further processing of tissues to identify specific proteins is not an option at this point. However, the development of methods such as ATUM24, in which sections from the tissue block are preserved, are likely to enable the identification of specific proteins in the cell. Determining the specific location of chemosensory receptors on enteroendocrine cells is essential information for the development of drug therapies for obesity, because enteroendocrine cells are a sensory interface between food in the gut and satiety in the brain.

Life inside a cell takes place in time and space. Changes over time are often studied using time-lapse microscopy combined with fluorescence imaging techniques, like super-resolution microscopy. Space, in particular the arrangement of organelles inside a cell or cell-to-cell interactions, can only be derived by a complete account of the cell&#8217;s fine structure. A comprehensive account of a cell&#8217;s fine structure can also bring clarity to genomic function in cases where the genome is available, like the C. elegans nematode1 or the flat placozoa tricoplax adherens2. Serial sectioning electron microscopy is now a reproducible, time efficient, and less expensive task thanks to the development of automated 3D electron microscopy technologies, like serial block-face scanning electron microscopy3 (SBEM).
The need for structural information to elucidate function is very evident in certain cell types where function depends on physical cell-to-cell interactions, such as neurons, glia, or sensory epithelial cells. We are particularly interested in elucidating how sensory signals from nutrients in the lumen of the gut are transduced into an electrical signal that ultimately modulates appetitive behaviors. The circuit is complex but begins at the wall of the intestine where nutrients come into contact with sensory epithelial cells, called enteroendocrine cells. Unlike other sensory epithelial cells, such as taste cells, enteroendocrine cells are dispersed throughout the gut epithelium at a ratio of one to one thousand4-7. Consequently, they have been difficult to identify and study, and for long time they were viewed only as a source of gut hormones. But with the development of cell-specific fluorescence reporter mice, the complex sensory function of these cells is emerging. Using one of these reporter mice, a peptide YY-GFP (PyyGFP) mouse, we found that enteroendocrine cells have a prominent cytoplasmic tail that we named neuropod. The appearance of neuropods suggested a conserved function in cell-to-cell communication. Thus, we reasoned that by documenting the ultrastructure of an enteroendocrine cell, the function of neuropods could be derived.
The need to understand the structure of an appendage in a dispersed cell which is hard to identify was the main rationale for developing a method to combine confocal microscopy and SBEM. The cell of interest was identified using PyyGFP enteroendocrine cell-specific reporter mice. The method allowed us to document the entire ultrastructure of an enteroendocrine cell and its neuropod. Within neuropods, we found structural features of neuronal axons, and outside neuropods, we found a physical relationship to enteric glia8. Indeed, neuropods contain about 70% of all secretory vesicles suggesting an essential role in the secretory function of these cells. Building on the structural data, more recently we found that through these neuropods, enteroendocrine cells and neurons innervating the gut form a neuroepithelial circuit, similar to that of taste cells in the tongue8,9.
Uncovering such features of gastrointestinal chemosensory mechanisms stemmed from structural data gathered using this correlative microscopy method. We believe this method could be useful in other areas of cell biology, particularly where cells are dispersed within the tissues at a very low ratio. We referred to the method as correlative confocal and serial block face scanning electron microscopy in 3D (Cocem3D). The method is comprised of the following main steps: tissue dissection, confocal microscopy, SBEM imaging, SBEM and confocal image correlation, and manual segmentation. Compared to other correlative methods, the concept is rather simple because the correlation is physical rather than chemical.

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			<td>DAPI&#160; nuclear stain</td>
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			<td>Postively charged glass slides and coverslips</td>
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			<td>Fine art paintbrush #1</td>
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			<td>Cacodylate buffer</td>
			<td>Electron Microscopy Sciences</td>
			<td>11650</td>
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			<td>Tannic acid</td>
			<td>Electron Microscopy Sciences</td>
			<td>21700</td>
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			<td>EMbed 812 kit&#160;</td>
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			<td>14120</td>
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			<td>Liquid silver colloidal&#160;&#160;&#160;</td>
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			<td>12630</td>
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			<td>CircuitWorks conductive epoxy&#160;&#160;</td>
			<td>ITW Chemtronics</td>
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			<td>Variable flow peristaltic pump</td>
			<td>VWR</td>
			<td>70730-064</td>
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			<td>VT1200S Vibrating blade microtome</td>
			<td>Leica</td>
			<td></td>
			<td></td>
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			<td>Zeiss 780i confocal microscope</td>
			<td>Carl Zeiss&#160;</td>
			<td></td>
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			<td>Sigma VP Scanning Electron Microscope&#160;</td>
			<td>Carl Zeiss&#160;</td>
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			<td>3view system</td>
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			<td>Renovo Neural Inc (Cleveland, OH)</td>
			<td></td>
			<td>http://www.renovoneural.com</td>
			<td>Renovo provides 3d EM services</td>
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			<td>Fiji software</td>
			<td>Open access software</td>
			<td></td>
			<td></td>
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			<td>Computer station with 16GB of RAM or more</td>
			<td></td>
			<td></td>
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			<td>Data visualization software Imaris 7.5&#160;</td>
			<td>Bitplane</td>
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correlative confocal and 3d electron microscopy of a specific sensory cell

Delineation of a cell&#8217;s ultrastructure is important for understanding its function. This can be a daunting project for rare cell types diffused throughout tissues made of diverse cell types, such as enteroendocrine cells of the intestinal epithelium. These gastrointestinal sensors of food and bacteria have been difficult to study because they are dispersed among other epithelial cells at a ratio of 1:1,000. Recently, transgenic reporter mice have been generated to identify enteroendocrine cells by means of fluorescence. One of those is the peptide YY-GFP mouse. Using this mouse, we developed a method to correlate confocal and serial block-face scanning electron microscopy. We named the method cocem3D and applied it to identify a specific enteroendocrine cell in tissue and unveil the cell&#8217;s ultrastructure in 3D. The resolution of cocem3D is sufficient to identify organelles as small as secretory vesicles and to distinguish cell membranes for volume rendering. Cocem3D can be easily adapted to study the 3D ultrastructure of other specific cell types in their native tissue.

The method presented here was used to study the ultrastructure of a specific cell within a layer of epithelium. The cell of interest in this case is the elusive enteroendocrine cell. Their identification in situ has been possible only in the last few years with the development of cell-specific fluorescence reporter mice. In 2011, we developed a mouse in which the promoter of the hormone PYY drives the expression of green fluorescent protein17. In this PyyGFP mouse, enteroendocrine cells of the distal part of the small intestine and colon are easily distinguished under UV light. This mouse model was a foundation to identify a tissue block containing an enteroendocrine cell and process it for SBEM. The correlation method is referred here as cocem3D and was built on previously published protocols10,11.
Optimizing the dimensions of the tissue block is critical for the correlation. In preliminary experiments, it was determined that in a tissue block 300 &#181;m long x 50 &#181;m thick the number of SBEM images is reduced to a minimum while covering the entire body of the cell of interest. Figure 1A shows an overlook of the dissection using a vibrating blade microtrome to obtain the tissue block with the right dimensions. At least 100 blocks are obtained before sorting through them to pick those with intact cells of interest. Selected blocks are imaged by confocal microscopy and image z-stacks are optically spaced every 1 &#181;m (Figure 1B and C). Figure 1D shows a volume rendered view of an enteroendocrine cell in the ileum of the mouse. Its prominent neuropod extends underneath epithelial cells for ~60 &#181;m.
The cell of interest is found by correlating the confocal z-stacks with a SBEM image of the entire block face. An example is presented here of a block from the distal colon of a PyyGFP mouse. After obtaining confocal z-stacks (Figure 2A), the block is embedded in a thin layer of low-melting agarose (Figure 2B). This is necessary for subsequent manipulation of the block. It is important to maintain the orientation of the block as flat as possible to match the optical slices with the SBEM slices. In Figure 2C, a representative image is shown of the entire face of the tissue block. Parts of this figure were previously published in Bohorquez et al., 20148. This was subsequently correlated with the confocal image by taking into account tissue landmarks and dimensions. Figure 2D shows an overlay of the confocal and SBEM image of the block, revealing the precise location of our cell of interest.
Once the cell has been identified, SBEM imaging is done at a resolution high enough to identify secretory vesicles and other cell organelles. The data set presented here was imaged at 7 nm per pixel as images of the block were taken every 70 nm. The data set contained 643 images that span 45 &#181;m of the tissue depth. A few microns are shaved during initial imaging of the entire block face at low magnification. The raw SBEM data set was acquired in .dm3 format and transformed to .tiff for subsequent handling. The SBEM stack was cropped using the Fiji software plugin &#8220;crop (3D)&#8221; block to contain only the region of interest. This reduces the amount of RAM memory needed for volume rendering. The cell was identified by its position and secretory vesicles, and segmented using Imaris software. Figure 3 contains a rendering in 3D of the enteroendocrine cell.
<img alt="Figure 1" src="/files/ftp_upload/52918/52918fig1.jpg" />
Figure 1: Correlative confocal microscopy. (A) Workflow to dissect a piece of intestinal tissue into a tissue block 300 x 50 &#181;m thick. (B) A representative tissue block of the right dimensions. Note that a tissue block of these dimensions from the mouse small intestine spans about 3 villi or, in the case of the colon about 8 crypts. (C) A villus in the small intestine containing a cell of interest. (D) A confocal z-stack rendered in 3D using Imaris software shows an enteroendocrine cell with a prominent neuropod running underneath the intestinal epithelium. Blue = DAPI nuclear stain. Bars = 10 &#181;m. <a href="https://www.jove.com/files/ftp_upload/52918/52918fig1large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 2" src="/files/ftp_upload/52918/52918fig2.jpg" /> 
Figure 2: Serial block-face scanning electron microscopy. (A) Confocal image of a selected tissue block from the colon containing PyyGFP enteroendocrine cells (green) of interest. (B) Once the block is imaged by confocal microscopy, it is then embedded flat in low-melting agarose using glass slides. Embedding the tissue block in agarose facilitates subsequent manipulation, and the notch on the left corner helps to mount the block within the SBEM in the right orientation. (C) SBEM image of the face of the block. (D) Correlation of confocal data with SBEM to identify cell of interest (white arrow). Images in c and d of this figure are modified from Bohorquez et al., 20148. Blue = DAPI nuclear stain. Bars = 10 &#181;m. <a href="https://www.jove.com/files/ftp_upload/52918/52918fig2large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 3" src="/files/ftp_upload/52918/52918fig3.jpg" /> 
Figure 3: Data rendering in 3D. (A) Volume rendering of enteroendocrine cell using Imaris software. The cell contains a neuropod packed with secretory vesicles. For a complete description of the ultrastructure of an enteroendocrine cell, please refer to Boh&#243;rquez et al., 20148. <a href="https://www.jove.com/files/ftp_upload/52918/52918fig3large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

Watch this Scientific Journal Video about Correlative Confocal and 3D Electron Microscopy of a Specific Sensory Cell at JoVE.com

Correlative Confocal and 3D Electron Microscopy of a Specific Sensory Cell

Here, we introduce a method, cocem3D, to unveil the ultrastructure of a specific cell in its native tissue by bridging confocal and serial block-face scanning electron microscopy.

Delineation of a cell&#8217;s ultrastructure is important for understanding its function. This can be a daunting project for rare cell types diffused ...

correlative-confocal-3d-electron-microscopy-specific-sensory

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JoVE Journal

Biology

11.4K Views.  Duke University Medical Center. Here, we introduce a method, cocem3D, to unveil the ultrastructure of a specific cell in its native tissue by bridging confocal and serial block-face scanning electron microscopy. 

Video: Correlative Confocal and 3D Electron Microscopy of a Specific Sensory Cell

Fluorescence-based Measurement of Store-operated Calcium Entry in Live Cells: from Cultured Cancer Cell to Skeletal Muscle Fiber

Store operated Ca2+ entry (SOCE), earlier termed capacitative Ca2+ entry, is a tightly regulated mechanism for influx of extracellular Ca2+ into cells to replenish depleted endoplasmic reticulum (ER) or sarcoplasmic reticulum (SR) Ca2+ stores1,2. Since Ca2+ is a ubiquitous second messenger, it is not surprising to see that SOCE plays important roles in a variety of cellular processes, including proliferation, apoptosis, gene transcription and motility. Due to its wide occurrence in nearly all cell types, including epithelial cells and skeletal muscles, this pathway has received great interest3,4. However, the heterogeneity of SOCE characteristics in different cell types and the physiological function are still not clear5-7.
The functional channel properties of SOCE can be revealed by patch-clamp studies, whereas a large body of knowledge about this pathway has been gained by fluorescence-based intracellular Ca2+ measurements because of its convenience and feasibility for high-throughput screening. The objective of this report is to summarize a few fluorescence-based methods to measure the activation of SOCE in monolayer cells, suspended cells and muscle fibers5,8-10. The most commonly used of these fluorescence methods is to directly monitor the dynamics of intracellular Ca2+ using the ratio of F340nm and F380nm (510 nm for emission wavelength) of the ratiometric Ca2+ indicator Fura-2. To isolate the activity of unidirectional SOCE from intracellular Ca2+ release and Ca2+ extrusion, a Mn2+ quenching assay is frequently used. Mn2+ is known to be able to permeate into cells via SOCE while it is impervious to the surface membrane extrusion processes or to ER uptake by Ca2+ pumps due to its very high affinity with Fura-2. As a result, the quenching of Fura-2 fluorescence induced by the entry of extracellular Mn2+ into the cells represents a measurement of activity of SOCE9. Ratiometric measurement and the Mn+2 quenching assays can be performed on a cuvette-based spectrofluorometer in a cell population mode or in a microscope-based system to visualize single cells. The advantage of single cell measurements is that individual cells subjected to gene manipulations can be selected using GFP or RFP reporters, allowing studies in genetically modified or mutated cells. The spatiotemporal characteristics of SOCE in structurally specialized skeletal muscle can be achieved in skinned muscle fibers by simultaneously monitoring the fluorescence of two low affinity Ca2+ indicators targeted to specific compartments of the muscle fiber, such as Fluo-5N in the SR and Rhod-5N in the transverse tubules9,11,12.

The extent of store-operated Ca2+ entry (SOCE) can be monitored using fluorescent Ca2+ indicators. Mn2+ quenching of such indicators assays SOCE in cultured cells and skeletal muscle fibers. A technique allowing spatial and temporal resolution of SOCE by confocal imaging of mechanically skinned muscle fibers is also described.

Store operated Ca2+ entry (SOCE), earlier termed capacitative Ca2+ entry, is a tightly regulated mechanism for influx of extracellular Ca2+ into cells to ...

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 ...

Analyzing Craniofacial Morphogenesis in Zebrafish Using 4D Confocal Microscopy

Time-lapse imaging is a technique that allows for the direct observation of the process of morphogenesis, or the generation of shape. Due to their optical clarity and amenability to genetic manipulation, the zebrafish embryo has become a popular model organism with which to perform time-lapse analysis of morphogenesis in living embryos. Confocal imaging of a live zebrafish embryo requires that a tissue of interest is persistently labeled with a fluorescent marker, such as a transgene or injected dye. The process demands that the embryo is anesthetized and held in place in such a way that healthy development proceeds normally. Parameters for imaging must be set to account for three-dimensional growth and to balance the demands of resolving individual cells while getting quick snapshots of development. Our results demonstrate the ability to perform long-term in vivo imaging of fluorescence-labeled zebrafish embryos and to detect varied tissue behaviors in the cranial neural crest that cause craniofacial abnormalities. Developmental delays caused by anesthesia and mounting are minimal, and embryos are unharmed by the process. Time-lapse imaged embryos can be returned to liquid medium and subsequently imaged or fixed at later points in development. With an increasing abundance of transgenic zebrafish lines and well-characterized fate mapping and transplantation techniques, imaging any desired tissue is possible. As such, time-lapse in vivo imaging combines powerfully with zebrafish genetic methods, including analyses of mutant and microinjected embryos.

Time-lapse confocal imaging is a powerful technique useful for characterizing embryonic development. Here, we describe the methodology and characterize craniofacial morphogenesis in wild-type, as well as pdgfra, smad5, and smo mutant embryos.

Time-lapse imaging is a technique that allows for the direct observation of the process of morphogenesis, or the generation of shape. Due to their optical ...

Live Cell Imaging and 3D Analysis of Angiotensin Receptor Type 1a Trafficking in Transfected Human Embryonic Kidney Cells Using Confocal Microscopy

Live-cell imaging is used to simultaneously capture time-lapse images of angiotensin type 1a receptors (AT1aR) and intracellular compartments in transfected human embryonic kidney-293 (HEK) cells following stimulation with angiotensin II (Ang II). HEK cells are transiently transfected with plasmid DNA containing AT1aR tagged with enhanced green fluorescent protein (EGFP). Lysosomes are identified with a red fluorescent dye. Live-cell images are captured on a laser scanning confocal microscope after Ang II stimulation and analyzed by software in three dimensions (3D, voxels) over time. Live-cell imaging enables investigations into receptor trafficking and avoids confounds associated with fixation, and in particular, the loss or artefactual displacement of EGFP-tagged membrane receptors. Thus, as individual cells are tracked through time, the subcellular localization of receptors can be imaged and measured. Images must be acquired sufficiently rapidly to capture rapid vesicle movement. Yet, at faster imaging speeds, the number of photons collected is reduced. Compromises must also be made in the selection of imaging parameters like voxel size in order to gain imaging speed. Significant applications of live-cell imaging are to study protein trafficking, migration, proliferation, cell cycle, apoptosis, autophagy and protein-protein interaction and dynamics, to name but a few.

Here we present a protocol to image cells expressing green fluorescent protein-tagged angiotensin type 1a receptors during endocytosis initiated by angiotensin II treatment. This technique includes labeling lysosomes with a second fluorescent marker, and then utilizing software to analyze the co-localization of receptor and lysosome in three dimensions over time.

Live-cell imaging is used to simultaneously capture time-lapse images of angiotensin type 1a receptors (AT1aR) and intracellular compartments in ...

Flash-and-Freeze: A Novel Technique to Capture Membrane Dynamics with Electron Microscopy

Cells constantly change their membrane architecture and protein distribution, but it is extremely difficult to visualize these events at a temporal and spatial resolution on the order of ms and nm, respectively. We have developed a time-resolved electron microscopy technique, &#34;flash-and-freeze,&#34; that induces cellular events with optogenetics and visualizes the resulting membrane dynamics by freezing cells at defined time points after stimulation. To demonstrate this technique, we expressed channelrhodopsin, a light-sensitive cation channel, in mouse hippocampal neurons. A flash of light stimulates neuronal activity and induces neurotransmitter release from synaptic terminals through the fusion of synaptic vesicles. The optogenetic stimulation of neurons is coupled with high-pressure freezing to follow morphological changes during synaptic transmission. Using a commercial instrument, we captured the fusion of synaptic vesicles and the recovery of the synaptic vesicle membrane. To visualize the sequence of events, large datasets were generated and analyzed blindly, since morphological changes were followed in different cells over time. Nevertheless, flash-and-freeze allows the visualization of membrane dynamics in electron micrographs with ms temporal resolution.

We developed a novel technique in electron microscopy, &#34;flash-and-freeze,&#34; that enables the visualization of membrane dynamics with ms temporal resolution. This technique combines the optogenetic stimulation of neurons with high-pressure freezing. Here, we demonstrate the procedures and describe the protocols in detail.

Cells constantly change their membrane architecture and protein distribution, but it is extremely difficult to visualize these events at a temporal and ...

Combining Mitotic Cell Synchronization and High Resolution Confocal Microscopy to Study the Role of Multifunctional Cell Cycle Proteins During Mitosis

Study of the various regulatory events of the cell cycle in a phase-dependent manner provides a clear understanding about cell growth and division. The synchronization of cell populations at specific stages of the cell cycle has been found to be very useful in such experimental endeavors. Synchronization of cells by treatment with chemicals that are relatively less toxic can be advantageous over the use of pharmacological inhibitory drugs for the study of consequent cell cycle events and to obtain specific enrichment of selected mitotic stages. Here, we describe the protocol for synchronizing human cells at different stages of the cell cycle, including both in S phase and M phase with a double thymidine block and release procedure for studying the functionality of mitotic proteins in chromosome alignment and segregation. This protocol has been extremely useful for studying the mitotic roles of multifunctional proteins which possess established interphase functions. In our case, the mitotic role of Cdt1, a protein critical for replication origin licensing in G1 phase, can be studied effectively only when G2/M-specific Cdt1 can be depleted. We describe the detailed protocol for depletion of G2/M-specific Cdt1 using double thymidine synchronization. We also explain the protocol of cell fixation, and live cell imaging using high resolution confocal microscopy after thymidine release. The method is also useful for analyzing the function of mitotic proteins under both physiological and perturbed conditions such as for Hec1, a component of the Ndc80 complex, as it enables one to obtain large sample sizes of mitotic cells for fixed and live cell analysis as we show here.&#160;&#160;

We present a protocol for double thymidine synchronization of HeLa cells followed by analysis using high resolution confocal microscopy. This method is key to obtaining large number of cells that proceed synchronously from S phase to mitosis, enabling studies on mitotic roles of multifunctional proteins which also possess interphase functions.

Study of the various regulatory events of the cell cycle in a phase-dependent manner provides a clear understanding about cell growth and division. The ...

Biological Sample Preparation by High-pressure Freezing, Microwave-assisted Contrast Enhancement, and Minimal Resin Embedding for Volume Imaging

The described sample preparation technique is designed to combine the best quality of ultrastructural preservation with the most suitable contrast for the imaging modality in a focused ion beam scanning electron microscope (FIB-SEM), which is used to obtain stacks of sequential images for 3D reconstruction and modelling. High-pressure freezing (HPF) allows close to native structural preservation, but the subsequent freeze substitution often does not provide sufficient contrast, especially for a bigger specimen, which is needed for high-quality imaging in the SEM required for 3D reconstruction. Therefore, in this protocol, after the freeze substitution, additional contrasting steps are carried out at room temperature. Although these steps are performed in a microwave, it is also possible to follow traditional bench processing, which requires longer incubation times.The subsequent embedding in minimal amounts of resin allows for faster and more precise targeting and preparation inside the FIB-SEM. This protocol is especially useful for samples that require preparation by high-pressure freezing for a reliable ultrastructural preservation but do not gain enough contrast during the freeze substitution for volume imaging using FIB-SEM. In combination with the minimal resin embedding, this protocol provides an efficient workflow for the acquisition of high-quality volume data.

Here we present a protocol for combining two sample processing techniques, high-pressure freezing and microwave-assisted sample processing, followed by minimal resin embedding for acquiring data with a focused ion beam scanning electron microscope (FIB-SEM). This is demonstrated using a mouse tibial nerve sample and Caenorhabditis elegans.

The described sample preparation technique is designed to combine the best quality of ultrastructural preservation with the most suitable contrast for the ...

Reconstruction of Single-Cell Innate Fluorescence Signatures by Confocal Microscopy

Described here is confocal reflection microscopy-assisted single-cell innate fluorescence analysis (CRIF), a minimally invasive method for reconstructing the innate cellular fluorescence signature from each individual live cell in a population distributed in a three-dimensional (3D) space. The innate fluorescence signature of a cell is a collection of fluorescence signals emitted by various biomolecules within the cell. Previous studies established that innate fluorescence signatures reflect various cellular properties and differences in physiological status and are a rich source of information for cell characterization and identification. Innate fluorescence signatures have been traditionally analyzed at the population level, necessitating a clonal culture, but not at the single-cell level. CRIF is particularly suitable for studies that require 3D resolution and/or selective extraction of fluorescence signals from individual cells. Because the fluorescence signature is an innate property of a cell, CRIF is also suitable for tag-free prediction of the type and/or physiological status of intact and single cells. This method may be a powerful tool for streamlined cell analysis, where the phenotype of each single cell in a heterogenous population can be directly assessed by its autofluorescence signature under a microscope without cell tagging.

Here, a protocol is presented for optically extracting and cataloging innate cellular fluorescence signatures (i.e., cellular autofluorescence) from every individual live cell distributed in a three-dimensional space. This method is suitable for studying the innate fluorescence signature of diverse biological systems at a single-cell resolution, including cells from bacteria, fungi, yeasts, plants, and animals.

Described here is confocal reflection microscopy-assisted single-cell innate fluorescence analysis (CRIF), a minimally invasive method for reconstructing ...

Cryo-Electron Microscopic Grid Preparation for Time-Resolved Studies using a Novel Robotic System, Spotiton

The capture of short-lived molecular states triggered by the early encounter of two or more interacting particles continues to be an experimental challenge of great interest to the field of cryo-electron microscopy (cryo-EM). A few methodological strategies have been developed that support these &#34;time-resolved&#34; studies, one of which, Spotiton-a novel robotic system-combines the dispensing of picoliter-sized sample droplets with precise temporal and spatial control. The time-resolved Spotiton workflow offers a uniquely efficient approach to interrogate early structural rearrangements from minimal sample volume. Fired from independently controlled piezoelectric dispensers, two samples land and rapidly mix on a nanowire EM grid as it plunges toward the cryogen. Potentially hundreds of grids can be prepared in rapid succession from only a few microliters of a sample. Here, a detailed step-by-step protocol of the operation of the Spotiton system is presented with a focus on troubleshooting specific problems that arise during grid preparation.

The protocol presented here describes the use of Spotiton, a novel robotic system, to deliver two samples of interest onto a self-wicking, nanowire grid that mix for a minimum of 90 ms prior to vitrification in liquid cryogen.

The capture of short-lived molecular states triggered by the early encounter of two or more interacting particles continues to be an experimental ...

Strategies for Optimization of Cryogenic Electron Tomography Data Acquisition

Cryogenic electron tomography (cryoET) is a powerful method to study the 3D structure of biological samples in a close-to-native state. Current state-of-the-art cryoET combined with subtomogram averaging analysis enables the high-resolution structural determination of macromolecular complexes that are present in multiple copies within tomographic reconstructions. Tomographic experiments usually require a vast amount of tilt series to be acquired by means of high-end transmission electron microscopes with important operational running-costs. Although the throughput and reliability of automated data acquisition routines have constantly improved over the recent years, the process of selecting regions of interest at which a tilt series will be acquired cannot be easily automated and it still relies on the user's manual input. Therefore, the set-up of a large-scale data collection session is a time-consuming procedure that can considerably reduce the remaining microscope time available for tilt series acquisition. Here, the protocol describes the recently developed implementations based on the SerialEM package and the PyEM software that significantly improve the time-efficiency of grid screening and large-scale tilt series data collection. The presented protocol illustrates how to use SerialEM scripting functionalities to fully automate grid mapping, grid square mapping, and tilt series acquisition. Furthermore, the protocol describes how to use PyEM to select additional acquisition targets in off-line mode after automated data collection is initiated. To illustrate this protocol, its application in the context of high-end data collection of Sars-Cov-2 tilt series is described. The presented pipeline is particularly suited to maximizing the time-efficiency of tomography experiments that require a careful selection of acquisition targets and at the same time a large amount of tilt series to be collected.

The increasing demand for large-scale data collection in cryogenic electron tomography requires high-throughput image acquisition routines. Described here is a protocol that implements the recent developments of advanced acquisition strategies aimed at maximizing the time-efficiency and throughput of tomographic data collection.