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.

<table><tbody><tr><td>Phosphate buffered saline</td><td>Life technologies</td><td>10010023</td><td></td></tr><tr><td>Heparin sodium salt</td><td>Sigma</td><td>H4784</td><td></td></tr><tr><td>Paraformaldehyde&amp;nbsp;</td><td>Sigma</td><td>158127</td><td>4%, freshly made in PBS, final pH 7.4</td></tr><tr><td>Glutaraldehyde</td><td>Sigma</td><td>G5882</td><td></td></tr><tr><td>Dental wax</td><td>Electron Microscopy Sciences</td><td>72660</td><td></td></tr><tr><td>Low-melting agarose</td><td>Life technologies</td><td>16520-100</td><td>5%, freshly made in PBS</td></tr><tr><td>Standard Tissue-Tek Cryomold</td><td>Electron Microscopy Sciences</td><td>62534-25</td><td></td></tr><tr><td>DAPI&amp;nbsp; nuclear stain</td><td>Life technologies</td><td>D1306</td><td></td></tr><tr><td>Postively charged glass slides and coverslips</td><td></td><td></td><td></td></tr><tr><td>Fine art paintbrush #1</td><td></td><td></td><td></td></tr><tr><td>Cacodylate buffer</td><td>Electron Microscopy Sciences</td><td>11650</td><td></td></tr><tr><td>Tannic acid</td><td>Electron Microscopy Sciences</td><td>21700</td><td></td></tr><tr><td>EMbed 812 kit&amp;nbsp;</td><td>Electron Microscopy Sciences</td><td>14120</td><td></td></tr><tr><td>Liquid releasing agent&amp;nbsp;</td><td>Electron Microscopy Sciences</td><td>70880</td><td></td></tr><tr><td>Liquid silver colloidal&amp;nbsp;&amp;nbsp;&amp;nbsp;</td><td>Electron Microscopy Sciences</td><td>12630</td><td></td></tr><tr><td>CircuitWorks conductive epoxy&amp;nbsp;&amp;nbsp;</td><td>ITW Chemtronics</td><td>CW2400</td><td></td></tr><tr><td>Variable flow peristaltic pump</td><td>VWR</td><td>70730-064</td><td></td></tr><tr><td>VT1200S Vibrating blade microtome</td><td>Leica</td><td></td><td></td></tr><tr><td>Zeiss 780i confocal microscope</td><td>Carl Zeiss&amp;nbsp;</td><td></td><td></td></tr><tr><td>Sigma VP Scanning Electron Microscope&amp;nbsp;</td><td>Carl Zeiss&amp;nbsp;</td><td></td><td></td></tr><tr><td>3view system</td><td>Gatan</td><td></td><td></td></tr><tr><td>Renovo Neural Inc (Cleveland, OH)</td><td></td><td>http://www.renovoneural.com</td><td>Renovo provides 3d EM services</td></tr><tr><td>Fiji software</td><td>Open access software</td><td></td><td></td></tr><tr><td>Computer station with 16 GB of RAM or more</td><td></td><td></td><td></td></tr><tr><td>Data visualization software Imaris 7.5&amp;nbsp;</td><td>Bitplane</td><td></td><td></td></tr></tbody></table>

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

Research

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

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

Correlative Super-resolution and Electron Microscopy to Resolve Protein Localization in Zebrafish Retina

We present a method to investigate the subcellular protein localization in the larval zebrafish retina by combining super-resolution light microscopy and scanning electron microscopy. The sub-diffraction limit resolution capabilities of super-resolution light microscopes allow improving the accuracy of the correlated data. Briefly, 110 nanometer thick cryo-sections are transferred to a silicon wafer and, after immunofluorescence staining, are imaged by super-resolution light microscopy. Subsequently, the sections are preserved in methylcellulose and platinum shadowed prior to imaging in a scanning electron microscope (SEM). The images from these two microscopy modalities are easily merged using tissue landmarks with open source software. Here we describe the adapted method for the larval zebrafish retina. However, this method is also applicable to other types of tissues and organisms. We demonstrate that the complementary information obtained by this correlation is able to resolve the expression of mitochondrial proteins in relation with the membranes and cristae of mitochondria as well as to other compartments of the cell.

This protocol describes the necessary steps to obtain subcellular protein localization results on zebrafish retina by correlating super-resolution light microscopy and scanning electron microscopy images.

We present a method to investigate the subcellular protein localization in the larval zebrafish retina by combining super-resolution light microscopy and ...

Developmental Biology

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 ...

Visualization of Endosome Dynamics in Living Nerve Terminals with Four-dimensional Fluorescence Imaging

Four-dimensional (4D) light imaging has been used to study behavior of small structures within motor nerve terminals of the thin transversus abdominis muscle of the garter snake. Raw data comprises time-lapse sequences of 3D z-stacks. Each stack contains 4-20 images acquired with epifluorescence optics at focal planes separated by 400-1,500 nm. Steps in the acquisition of image stacks, such as adjustment of focus, switching of excitation wavelengths, and operation of the digital camera, are automated as much as possible to maximize image rate and minimize tissue damage from light exposure. After acquisition, a set of image stacks is deconvolved to improve spatial resolution, converted to the desired 3D format, and used to create a 4D "movie" that is suitable for variety of computer-based analyses, depending upon the experimental data sought. One application is study of the dynamic behavior of two classes of endosomes found in nerve terminals-macroendosomes (MEs) and acidic endosomes (AEs)-whose sizes (200-800 nm for both types) are at or near the diffraction limit. Access to 3D information at each time point provides several advantages over conventional time-lapse imaging. In particular, size and velocity of movement of structures can be quantified over time without loss of sharp focus. Examples of data from 4D imaging reveal that MEs approach the plasma membrane and disappear, suggesting that they are exocytosed rather than simply moving vertically away from a single plane of focus. Also revealed is putative fusion of MEs and AEs, by visualization of overlap between the two dye-containing structures as viewed in each three orthogonal projections.

Four-dimensional (4D) imaging is utilized to study the behavior and interactions among two types of endosomes in living vertebrate nerve terminals. Movement of these small structures is characterized in three dimensions, permitting confirmation of events such as endosome fusion and exocytosis.

Four-dimensional (4D) light imaging has been used to study behavior of small structures within motor nerve terminals of the thin transversus abdominis ...

Neuroscience

Correlative Light Electron Microscopy (CLEM) for Tracking and Imaging Viral Protein Associated Structures in Cryo-immobilized Cells

Due to its high resolution, electron microscopy (EM) is an indispensable tool for virologists. However, one of the main difficulties when analyzing virus-infected or transfected cells via EM are the low efficiencies of infection or transfection, hindering the examination of these cells. In order to overcome this difficulty, light microscopy (LM) can be performed first to allocate the subpopulation of infected or transfected cells. Thus, taking advantage of the use of fluorescent proteins (FPs) fused to viral proteins, LM is used here to record the positions of the "positive-transfected" cells, expressing a FP and growing on a support with an alphanumeric pattern. Subsequently, cells are further processed for EM via high pressure freezing (HPF), freeze substitution (FS) and resin embedding. The ultra-rapid freezing step ensures excellent membrane preservation of the selected cells that can then be analyzed at the ultrastructural level by transmission electron microscopy (TEM). Here, a step-by-step correlative light electron microscopy (CLEM) workflow is provided, describing sample preparation, imaging and correlation in detail. The experimental design can be also applied to address many cell biology questions.

Correlative Light Electron Microscopy CLEM for Tracking and Imaging Viral Protein Associated Structures in Cryo-immobilized Cells

A correlative light electron microscopy (CLEM) method is applied to image virus-induced intracellular structures via electron microscopy (EM) in cells that are previously selected by light microscopy (LM). LM and EM are combined as a hybrid imaging approach to achieve an integrated view of virus-host interactions.

Due to its high resolution, electron microscopy (EM) is an indispensable tool for virologists. However, one of the main difficulties when analyzing ...

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 ...

Large-scale Three-dimensional Imaging of Cellular Organization in the Mouse Neocortex

The mammalian neocortex is composed of many types of excitatory and inhibitory neurons, each with specific electrophysiological and biochemical properties, synaptic connections, and in vivo functions, but their basic functional and anatomical organization from cellular to network scale is poorly understood. Here we describe a method for the three-dimensional imaging of fluorescently-labeled neurons across large areas of the brain for the investigation of the cortical cellular organization. Specific types of neurons are labeled by the injection of fluorescent retrograde neuronal tracers or expression of fluorescent proteins in transgenic mice. Block brain samples, e.g., a hemisphere, are prepared after fixation, made transparent with tissue clearing methods, and subjected to fluorescent immunolabeling of the specific cell types. Large areas are scanned using confocal or two-photon microscopes equipped with large working distance objectives and motorized stages. This method can resolve the periodic organization of the cell type-specific microcolumn functional modules in the mouse neocortex. The procedure can be useful for the study of three-dimensional cellular architecture in the diverse brain areas and other complex tissues.

Here we describe a procedure for tissue clearing, fluorescent labeling, and large-scale imaging of mouse brain tissue which, thereby, enables visualization of the three-dimensional organization of cell types in the neocortex.

The mammalian neocortex is composed of many types of excitatory and inhibitory neurons, each with specific electrophysiological and biochemical ...

Three-dimensional Imaging of Bacterial Cells for Accurate Cellular Representations and Precise Protein Localization

The shape of a bacterium is important for its physiology. Many aspects of cell physiology such as cell motility, predation, and biofilm production can be affected by cell shape. Bacterial cells are three-dimensional (3D) objects, although they are rarely treated as such. Most microscopy techniques result in two-dimensional (2D) images leading to the loss of data pertaining to the actual 3D cell shape and localization of proteins. Certain shape parameters, such as Gaussian curvature (the product of the two principal curvatures), can only be measured in 3D because 2D images do not measure both principal curvatures. Additionally, not all cells lie flat when mounting and 2D imaging of curved cells may not accurately represent the shapes of these cells. Accurately measuring protein localization in 3D can help determine the spatial regulation and function of proteins. A forward convolution technique has been developed that uses the blurring function of the microscope to reconstruct 3D cell shapes and to accurately localize proteins. Here, a protocol for preparing and mounting samples for live cell imaging of bacteria in 3D both to reconstruct an accurate cell shape and to localize proteins is described. The method is based on simple sample preparation, fluorescent image acquisition, and MATLAB-based image processing. Many high-quality fluorescent microscopes can be simply modified to take these measurements. These cell reconstructions are computationally intensive and access to high-throughput computational resources is recommended, although not necessary. This method has been successfully applied to multiple bacterial species and mutants, fluorescent imaging modalities, and microscope manufacturers.

This protocol explains how to prepare and mount bacterial samples for live three-dimensional imaging and how to reconstruct the three-dimensional shape of E. coli from those images.

The shape of a bacterium is important for its physiology. Many aspects of cell physiology such as cell motility, predation, and biofilm production can be ...

Immunology and Infection

Confocal Microscopy Reveals Cell Surface Receptor Aggregation Through Image Correlation Spectroscopy

Confocal microscopy provides an accessible methodology to capture sub-cellular interactions critical for the characterization and further development of pre-clinical agents labeled with fluorescent probes. With recent advancements in antibody based cytotoxic drug delivery systems, understanding the alterations induced by these agents within the realm of receptor aggregation and internalization is of critical importance. This protocol leverages the well-established methodology of fluorescent immunocytochemistry and the open source FIJI distribution of ImageJ, with its inbuilt autocorrelation and image mathematical functions, to perform spatial image correlation spectroscopy (ICS). This protocol quantitates the fluorescent intensity of labeled receptors as a function of the beam area of the confocal microscope. This provides a quantitative measure of the state of target molecule aggregation on the cell surface. This methodology is focused on the characterization of static cells with potential to expand into temporal investigations of receptor aggregation. This protocol presents an accessible methodology to provide quantification of clustering events occurring at the cell surface, utilizing well established techniques and non-specialized imaging apparatus.

Antibodies that bind to target receptors on the cell surface can confer conformation and clustering alterations. These dynamic changes have implications for characterizing drug development in target cells. This protocol utilizes confocal microscopy and image correlation spectroscopy through ImageJ/FIJI to quantify the extent of receptor clustering on the cell surface.

Confocal microscopy provides an accessible methodology to capture sub-cellular interactions critical for the characterization and further development of ...

Confocal Fluorescence Microscopy: A Technique to Determine the Localization of Proteins in Mouse Fibroblasts

Source: Dominique R. Bollino1, Eric A. Legenzov2, Tonya J. Webb1 
1 Department of Microbiology and Immunology, University of Maryland School of Medicine and the Marlene and Stewart Greenebaum Comprehensive Cancer Center, Baltimore, Maryland 21201 
2 Center for Biomedical Engineering and Technology, University of Maryland School of Medicine, Baltimore, Maryland 21201
Confocal fluorescence microscopy is an imaging technique that enables increased optical resolution as compared to conventional &#34;wide-field&#34; epifluorescence microscopy. Confocal microscopes are able to achieve improved x-y optical resolution through &#34;laser scanning&#34;- typically a set of voltage-controlled mirrors (galvanometer or &#34;galvo&#34; mirrors) that direct laser-illumination to each pixel of the specimen at a time. More importantly, confocal microscopes achieve superior z-axial resolution by using a pinhole to remove out of focus light originating from locations that are not in the z-plane being scanned, thus enabling the detector to collect data from a specified z-plane. Because of the high z-resolution achievable in confocal microscopy, it is possible to collect images from a series of z-planes (also-called z-stack) and construct a 3D image through software.
Before discussing the mechanism of a confocal microscope, it is important to consider how a sample interacts with light. Light is composed of photons, packets of electromagnetic energy. A photon impinging on a biological sample can interact with the molecules comprising the sample in one of four ways: 1) the photon does not interact and passes through the sample; 2) the photon is reflected/scattered; 3) the photon is absorbed by a molecule and the absorbed energy is released as heat through processes collectively known as nonradiative decay; and 4) the photon is absorbed and the energy is then rapidly reemitted as a secondary photon through the process known as fluorescence. A molecule whose structure permits fluorescence emission is called a fluorophore. Most biological samples contain negligible endogenous fluorophores; therefore exogenous fluorophores must be used to highlight features of interest in the sample. During fluorescence microscopy, the sample is illuminated with light of the appropriate wavelength for absorption by the fluorophore. Upon absorbing a photon, a fluorophore is said to be &#34;excited&#34; and the process of absorption is referred to as &#34;excitation&#34;. When a fluorophore gives up energy in the form of a photon, the process is known as &#34;emission&#34;, and the emitted photon is called fluorescence.
The light beam used to excite a fluorophore is focused by the objective lens of a microscope and converges at a &#34;focal point&#34; where it is maximally focused. Beyond the focal point the light again diverges. The entering and exiting beams may be visualized as a pair of cones touching at the focal point (see Figure 1, left panel). The phenomenon of diffraction imposes a limit on how tightly a beam of light can be focused - the beam actually focuses to a spot of finite size. Two factors determine the size of the focal spot: 1) the wavelength of the light, and 2) the light-gathering ability of the objective lens, which is characterized by its numerical aperture (NA). The focal &#34;spot&#34; extends not only in the x-y plane, but also in the z direction, and is in reality a focal volume. The dimensions of this focal volume define the maximum resolution achievable by optical imaging. Although the number of photons is greatest within the focal volume, the conical light paths above and below the focus also contain a lower density of photons. Any fluorophore in the light path can thus be excited. In conventional (wide-field) epifluorescence microscopy, emission from fluorophores above and below the focal plane contribute out-of-focus fluorescence (a &#34;hazy background&#34;), which reduces image resolution and contrast, as demonstrated in Figure 1, with the red cube representing fluorophore emission above the focal plane (red star) that results in out-of-focus fluorescence (top right). This problem is ameliorated in confocal microscopy, due to the utilization of a pinhole. (Figure 2, bottom right). As depicted in Figure 3, the pinhole allows emissions originating from the focal place to reach the detector (left), while blocking the out-of-focus fluorescence (right) from reaching the detector, thus improving both resolution and contrast.
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/10501/10501fig1.jpg" /> 
Figure 1. Optical resolution of epifluorescence versus confocal microscopy. <a href="https://www.jove.com/files/ftp_upload/10501/10501fig1large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
The light beam used to excite a fluorophore is focused by the objective lens of a microscope and converges at a focal volume and then diverges (left). The red star represents the focal plane of a sample that is being imaged while the red square represents fluorophore emission above the focal plane. When capturing an image of this sample using an epifluorescent microscope, the emission from the out-of-focus red square will visible and contribute to a &#34;hazy background&#34; (top right). Confocal microscopes have a pinhole which prevents the detection of light emitted outside of the focal plane, eliminating the &#34;hazy background&#34; (bottom right).
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/10501/10501fig2.jpg" /> 
Figure 2. Pinhole effect in confocal microscopy. <a href="https://www.jove.com/files/ftp_upload/10501/10501fig2large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
Although the highest intensity of the excitation light is at the focal point of the lens (left, red oval), other parts of the sample not in the focal point (right, red star) will get light and fluoresce. To prevent light emitted from these out-of-focus regions to reach the detector, a screen with a pinhole is present in front of the detector. Only the in-focus light (left) emitting from the focal plane is able to travel through the pinhole and reach the detector. The out-of-focus light (right) is blocked with the pinhole and fails to reach the detector.
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/10501/10501fig3.jpg" /> 
Figure 3. Principal components of a confocal laser scanning microscope. <a href="https://www.jove.com/files/ftp_upload/10501/10501fig3large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
For the sake of simplicity, the mechanistic description of a confocal microscope will be limited to that of the Nikon Eclipse Ti A1R. Although there may be minor technical differences between different confocal microscopes, the A1R serves well as a good model for describing confocal microscope function. The excitation light beam, produced by an array of diode lasers, is reflected by the primary dichroic mirror into the objective, which focuses the light onto the specimen being imaged. The primary dichroic mirror selectively reflects the excitation light while allowing light at other wavelengths to pass through. The light then encounters the scanning mirrors that sweep the light beam across the specimen in an x-y manner, illuminating a single (x,y) pixel at a time. Fluorescence emitted by fluorophores at the illuminated pixel is collected by the objective lens and passes through the primary dichroic mirror to reach an array of photomultiplier tubes (PMTs). Secondary dichroic mirrors direct the emission light to the appropriate PMT. Excitation light scattered by the sample back into the objective is reflected by the primary dichroic mirror back towards the specimen, and thus prevented from entering the detection light path and reaching the PMTs (see Figure 3). This allows the relatively feeble fluorescence to be quantified without contamination by light scattered from the excitation light beam, which is typically orders of magnitude more intense than the fluorescence. Because the pinhole blocks light from outside of the focal volume, the light arriving at the detector comes from a narrow, selected z-plane. Therefore, images can be collected from a series of adjacent z-planes; this series of images is often referred to as a &#39;z-stack&#39;. By using the appropriate software, a z-stack can be processed to generate a 3D image of the specimen. A particular advantage of confocal microscopy is the ability to distinguish the subcellular localization of staining. For example, the differentiation between membrane staining from intracellular staining, which is very challenging with conventional epifluorescence microscopy (1, 2, 3).
Sample preparation is an important facet of confocal imaging. A strength of optical microscopy techniques is the flexibility to image live or fixed cells. When attempting to produce 3D images, because of the number of images that must be acquired for a z-stack, the difficulty of maintaining cell health, and the movement of live cells and their organelles, the use of fixed cells is typical. The procedure for fixing and staining cells for confocal fluorescence is similar to that conventionally used in immunofluorescence. After culture in chamber slides or on coverslips, cells are fixed using paraformaldehyde to preserve cellular morphology. Non-specific antibody binding is blocked using bovine serum albumin, milk, or normal serum. In order to maintain the specificity of the secondary antibodies, the solution used should not originate from the same species in which the primary antibodies were generated. The cells are incubated with primary antibodies that bind the antigen of interest. When labeling several cellular targets, the primary antibodies must each be derived from a different species. Antibodies tagging an antigen are then bound by fluorophore-conjugated secondary antibodies. Fluorophore-conjugated secondary antibodies should be selected so that they are compatible with the wavelengths of laser excitation available in the confocal microscope. When visualizing multiple antigens, the excitation/emission spectra of the fluorophores should differ enough so that their signals can be discriminated by microscopic analysis. The stained specimen is then mounted on a slide for imaging. A mounting medium is used to prevent photobleaching and specimen dehydration. If desired, a mounting medium containing a nuclear counterstain (e.g. DAPI or Hoechst) can be used (4).
In the following protocol, mouse fibroblasts transfected to express CD1d (LCD1) were stained with antibodies recognizing CD1d and CD107a (LAMP-1). CD1d is a major histocompatibility complex 1 (MHC 1)-like receptor present on the surface of antigen presenting cells that presents lipid antigens. LAMP-1 (lysosomal associated membrane protein-1) is a transmembrane protein primarily present in lysosomal membranes. For proper antigen presentation, CD1d is trafficked through the low pH lysosomal compartment, so LAMP-1 is being used as a marker of the lysosomal compartment for this protocol. By probing the LCD1 cells with anti-CD1d and anti- LAMP-1 that were produced in different species, secondary antibodies with unique fluorophores can be used to determine the localization of each protein in the cell and whether CD1d is present in the LAMP-1 positive lysosomal compartments.

Confocal Fluorescence Microscopy and Localization of Proteins

Source: Dominique R. Bollino1, Eric A. Legenzov2, Tonya J. Webb1
1 Department of Microbiology and Immunology, University of Maryland School of Medicine ...