We thank Drs. Yichen Huang, Chunming Guo and Zhenfang Zhou for their technical support. This research was funded by NIH/NIDDK (1R01DK091645-01A1, XL) and American Heart Association (AHA, 13GRNT16950006, XL).

All animal uses are approved by the Institutional Animal Care and Use Committee at Boston Children&#39;s Hospital.
1. Sample Preparation
<ol>
	<li>Timed mating
	<ol>
		<li>Place a C57Bl6 wild type male and a female together in the same cage.</li>
		<li>Check for the formation of a mating plug early next morning. The mating plug (aka, vaginal or copulation plug) is a hardened gelatinous deposition that blocks the vagina.</li>
		<li>Set the plug date as embryonic day 0.5 (e0.5).</li>
	</ol>
	</li>
	<li>Isolation of mouse embryos
	<ol>
		<li>Euthanize pregnant females by CO2 asphyxiation.</li>
		<li>Put mice supine on an absorbent pad and spray abdominal region with 70% ethanol.</li>
		<li>Lift the skin and make an initial 5 mm incision at the caudal abdominal region using surgical scissors. Cut and remove the abdominal skin to fully expose the internal organs</li>
		<li>Cut near vagina to remove the entire uterus.</li>
		<li>Cut between implantation sites along the uterine horn. Each segment or conceptus should have only one embryo inside.</li>
		<li>Keep the uterine segments in 1x ice-cold phosphate buffered saline (PBS).</li>
	</ol>
	</li>
	<li>Embryo dissection
	<ol>
		<li>Place each conceptus in a separate dish with ice-cold PBS under a stereoscope.</li>
		<li>Remove uterine tissues, placenta and then fetal membranes sequentially using fine dissecting forceps.</li>
		<li>Transfer the dissected embryo to a 50 ml tube with 1x ice cold PBS using a large transfer pipette. Avoid picking up embryo directly with forceps to minimize tissue damage.</li>
	</ol>
	</li>
	<li>Fixation
	<ol>
		<li>Fix the dissected embryos in 10% neutral buffered Formalin solution at 4 &#176;C for more than 24 hr with at least 10 sample volumes of fixative.</li>
	</ol>
	</li>
	<li>Pretreatment
	<ol>
		<li>Prepare the Clearing Solution: 4 M urea, 10% glycerol, 4% Sodium Dodecyl Sulfate (SDS) in double distilled water.</li>
		<li>Replace the fixative with the Clearing Solution.</li>
		<li>Gently rotate the specimen on a rocker at room temperature for 1 - 2 weeks. Exchange the Clearing Solution once on the second and the third day. The specimen slowly becomes clear and translucent.</li>
		<li>Replace the Clearing Solution with 10% formalin and leave on a rocker for 2 days.</li>
	</ol>
	</li>
	<li>Dehydration
	<ol>
		<li>Wash the pretreated samples with 1x PBS 3 times for 5 min each.</li>
		<li>Dehydrate samples in an ascending ethanol series at room temperature on a gentle rocker. For E15.5 embryos, use an ascending ethanol series including 50%, 60%, 70%, 80%, 85%, 90%, 95% and 100% ethanol, 2 hr each step. For older embryos, the incubation time needs to be increased.</li>
	</ol>
	</li>
	<li>Staining
	<ol>
		<li>Prepare Staining Solution: add 0.1375 grams of Eosin Y Disodium Salt and 0.0275 grams of Acridine Orange hemi (zinc chloride) salt to 50 ml of 100% ethanol. Stir for 2 hr. Filter with a filter paper. Store at room temperature and avoid light by covering with aluminum foil.</li>
		<li>Transfer specimen to the Staining Solution for 1 hr.</li>
		<li>Replace with fresh Staining Solution and stain overnight.</li>
	</ol>
	</li>
	<li>Infiltration
	<ol>
		<li>Prepare Infiltration Solution: combine 100 ml of Solution A of JB-4 embedding kit, 1.25 g of Catalyst C, 0.275 g of Eosin Y Disodium Salt and 0.055 g of Acridine Orange hemi (zinc chloride) salt. Stir to mix (200 rpm) in an ice bucket for 2 hr, and then filter with the filter paper.</li>
		<li>Store the Infiltration Solution at 4 &#176;C and avoid light by covering with aluminum foil.</li>
		<li>Transfer the embryos to Infiltration Solution:Staining Solution (1:1), and incubate for at least 3 hr on a rocker at 4 &#176;C.</li>
		<li>Transfer the embryos to Infiltration Solution and incubate for 3 hr on a rocker in a cold room.</li>
		<li>Replace Infiltration Solution once every day, and leave in a cold room on a gentle rocker for three more days.</li>
	</ol>
	</li>
</ol>
2. Embedding
<ol>
	<li>Keep Solution B and Infiltration Solution on ice.</li>
	<li>Make Embedding Solution (1:25 of Solution B and Infiltration Solution) immediately before embedding.</li>
	<li>Orient the specimen in a embedding mold, then pour cold Embedding Solution into the embedding mold gently. Re-orient with a pin if needed.</li>
	<li>Use a pin to examine whether the Embedding Solution is solidified. Push out the specimen block, and trim it if needed.</li>
	<li>Reorient the specimen block using the separate mold to obtain a cross orientation.</li>
	<li>Put a block holder on top of the embedding mold. Gently pour Embedding Solution into the mold until the mold and the block holder are submerged by Embedding Solution.</li>
	<li>Keep the specimen in a secondary container and leave it on ice for 3 - 4 hr in a fume hood.</li>
	<li>Store the solidified specimens in a 4 &#176;C.</li>
	<li>Peel away the embedding mold. Put the block under the stereomicroscope with oblique illumination to inspect position of the specimen in the block. On the block face, mark grid lines around the specimen.</li>
	<li>Trim the block to remove access amount of embedding material using the marked grid lines as references.</li>
</ol>
3. Image Acquisition
<ol>
	<li>Clamp the specimen block to the microtome and obtain a fresh cut surface. Set section thickness (e.g., e9.5-10.5, 1.5 &#181;m; e11.5-12.5, 2.0 &#181;m; e13.5-15.5, 2.5 &#181;m; e16.5-older, 3 &#181;m). Keep the specimen/block at the home position of microtome.</li>
	<li>Mount the stereo zoom microscope horizontally facing the block surface of specimen.</li>
	<li>Turn on the computer and microscope, and start the image acquisition software.</li>
	<li>Visualize and focus the optics to the block-face under the &#34;live view&#34; mode of imaging software.</li>
	<li>Align microscope and microtome if needed so that the block-face is at the center of viewfinder.</li>
	<li>Adjust zoom so that the entire block-face is within the viewfinder.</li>
	<li>Select green fluorescent protein (GFP) filter (excitation 470 &#177; 20 nm, dichroic 495 nm, emission 525 &#177; 25 nm) and refocus if needed.</li>
	<li>Measure and set the exposure time manually (e.g., 80 - 400 mini sec).</li>
	<li>Acquire images of the block-face after each fresh cut and save images automatically if possible.</li>
	<li>Record important parameters including resolution, section thickness and zoom.</li>
	<li>After finishing the entire specimen, export and convert all image files to JPEG format if necessary.</li>
	<li>Invert all original images using an imaging processing software, and adjust brightness and contrast.</li>
	<li>Save all the processed images in a separate folder.</li>
</ol>
4. 3D Visualization
<ol>
	<li>Upload all the processed images to a 3D visualization software following the instructions.</li>
	<li>Input resolution and section thickness to convert all images to volumetric data (i.e., voxel size) and save the 3D file.</li>
	<li>Align all images using the automatic mode. Adjust individual images manually if needed.</li>
	<li>Save the aligned image to a new file name.</li>
	<li>Analyze morphometric features of the specimen using the 3D visualization software.</li>
</ol>

<ol>
	<li>Gregg, C. L., Butcher, J. T. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Quantitative+in+vivo+imaging+of+embryonic+development:+opportunities+and+challenges.">Quantitative in vivo imaging of embryonic development: opportunities and challenges.</a> Differentiation. 84, 149-162 (2012).</li><li>Liu, X., Tobita, K., Francis, R. J., Lo, C. W. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Imaging+techniques+for+visualizing+and+phenotyping+congenital+heart+defects+in+murine+models.">Imaging techniques for visualizing and phenotyping congenital heart defects in murine models.</a> Birth defects Res C. 99, 93-105 (2013).</li><li>Norris, F. C., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+coming+of+age:+advanced+imaging+technologies+for+characterising+the+developing+mouse.">A coming of age: advanced imaging technologies for characterising the developing mouse.</a> Trends Genet. 29, 700-711 (2013).</li><li>Weninger, W. J., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Phenotyping+structural+abnormalities+in+mouse+embryos+using+high-resolution+episcopic+microscopy.">Phenotyping structural abnormalities in mouse embryos using high-resolution episcopic microscopy.</a> Dis model mech. 7, 1143-1152 (2014).</li><li>Mohun, T. J., Weninger, W. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Episcopic+three-dimensional+imaging+of+embryos.">Episcopic three-dimensional imaging of embryos.</a> Cold Spring Harb protoc. , 641-646 (2012).</li><li>Sizarov, A., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Three-dimensional+and+molecular+analysis+of+the+arterial+pole+of+the+developing+human+heart.">Three-dimensional and molecular analysis of the arterial pole of the developing human heart.</a> J Anat. 220, 336-349 (2012).</li><li>Anderson, R. H., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Normal+and+abnormal+development+of+the+intrapericardial+arterial+trunks+in+humans+and+mice.">Normal and abnormal development of the intrapericardial arterial trunks in humans and mice.</a> Cardiovasc Res. 95, 108-115 (2012).</li><li>Huang, Y. C., Chen, F., Li, X. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Clarification+of+mammalian+cloacal+morphogenesis+using+high-resolution+episcopic+microscopy.">Clarification of mammalian cloacal morphogenesis using high-resolution episcopic microscopy.</a> Dev Biol. 409, 106-113 (2016).</li><li>Mohun, T. J., Weninger, W. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Embedding+embryos+for+episcopic+fluorescence+image+capturing+(EFIC).">Embedding embryos for episcopic fluorescence image capturing (EFIC).</a> Cold Spring Harb protoc. , 675-677 (2012).</li><li>Weninger, W. J., Mohun, T. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Three-dimensional+analysis+of+molecular+signals+with+episcopic+imaging+techniques.">Three-dimensional analysis of molecular signals with episcopic imaging techniques.</a> Methods mol biol. 411, 35-46 (2007).</li><li>Richardson, D. S., Lichtman, J. W. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Clarifying+Tissue+Clearing.">Clarifying Tissue Clearing.</a> Cell. 162, 246-257 (2015).</li><li>Hama, H., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Scale:+a+chemical+approach+for+fluorescence+imaging+and+reconstruction+of+transparent+mouse+brain.">Scale: a chemical approach for fluorescence imaging and reconstruction of transparent mouse brain.</a> Nat Neurosci. 14, 1481-1488 (2011).</li><li>Susaki, E. A., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Whole-brain+imaging+with+single-cell+resolution+using+chemical+cocktails+and+computational+analysis.">Whole-brain imaging with single-cell resolution using chemical cocktails and computational analysis.</a> Cell. 157, 726-739 (2014).</li><li>Mohun, T. J., Weninger, W. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Generation+of+volume+data+by+episcopic+three-dimensional+imaging+of+embryos.">Generation of volume data by episcopic three-dimensional imaging of embryos.</a> Cold Spring Harb protoc. , 681-682 (2012).</li></ol>

The authors have nothing to disclose.

Here, we present a modified protocol using routine laboratory equipment to acquire serial HREM images that are compatible for rapid 3D visualization and morphometric analysis of complex structures. Because the high-resolution images are taken directly from the block face instead of individual sections, the fine morphological features are preserved and rapidly reconstructed digitally in a 3D visualization program.
3D imaging offers significant advantages in visualizing and understanding complex...

The advent of 3D imaging technologies has opened the possibility of systemic analysis of detailed morphological features during fetal development. A variety of 3D imaging modalities is now available include micro-magnetic resonance imaging (micro-MRI), micro computed tomography, high frequency ultrasound, optical coherence tomography, optical project tomography and episcopic microscopy1-4. Each modality has its own unique features in resolution, contrast, speed, cost, in utero capability and availability.
Episcopic fluorescence image capture (EFIC) and high-resolution episcopic microscopy (HREM) are episcopic microscopy imaging methods4. Here, high-resolution serial images are captured continuously from the block face instead of tissue sections. The resulting images faithfully reflect detailed morphological features with minimal or no tissue distortion. The acquired volumetric data is readily converted to high-resolution 3-D images suitable for accurate morphometric analysis. Because of relative low cost and high resolution, HREM and EPIC have become the superior alternatives for systemic assessment of mouse models of human birth defects2,4-8.
Both EFIC and HREM methods require specimens to be embedded in the suitable medium. EFIC detects autofluorescence emitted from the embedded tissue. Because of this, it is limited to specimens emitting high levels of autofluorescence but not those with relatively weak autofluorescence such as early stage embryos9. To overcome the dependency of autofluorescence, specimens are stained with a fluorescent dye such as eosin for HREM imaging. The choices of dyes and staining methods also make HREM more compatible to detect molecular signals in the context of tissue architecture and morphology4,10.
In this article, we present an improved HREM protocol suitable for whole body assessment of the late stage mouse embryos. To facilitate staining, we include a pretreatment step to increase tissue penetration, thereby enabling HREM imaging of older mouse embryos.

<table><tbody><tr><td>Mice</td><td>The Jackson Laboratory</td><td>C57BL6</td><td></td></tr><tr><td>Ethyl Alcohol</td><td>Pharmco-Aaper</td><td>111000200</td><td>200 Proof, Absolute, ACS/USP/Kosher Grade</td></tr><tr><td>Formalin</td><td>Sigma</td><td>HT501128-4L</td><td></td></tr><tr><td>Urea</td><td>Sigma</td><td>U5128</td><td>Clearing Solution</td></tr><tr><td>Glycerol</td><td>Fisher scientific</td><td>G33-4</td><td>Clearing Solution</td></tr><tr><td>Sodium Dodecyl Sulfate</td><td>Invitrogen</td><td>15525-017</td><td>Clearing Solution</td></tr><tr><td>Eosin Y Disodium Salt</td><td>Fisher scientific</td><td>BP241925</td><td>Infiltration Solution</td></tr><tr><td>Acridine Orange hemi (zinc chloride) salt</td><td>Sigma</td><td>A6014</td><td>Infiltration Solution</td></tr><tr><td>Whatman paper</td><td>GE</td><td>3030-153</td><td></td></tr><tr><td>JB-4 Embedding Kit - Solution A</td><td>Polysciences, Inc.</td><td>0226A-800</td><td>Infiltration Solution</td></tr><tr><td>JB-4 Embedding Kit - Solution B</td><td>Polysciences, Inc.</td><td>0226B-30</td><td>Embedding Solution</td></tr><tr><td>JB-4&amp;nbsp;Embedding Kit - Catalyst</td><td>Polysciences, Inc.</td><td>02618-12</td><td>Infiltration Solution</td></tr><tr><td>Embedding Molds</td><td>Polysciences, Inc.</td><td>23185-1</td><td>16 x 8 mm</td></tr><tr><td>Peel-A-Way Sharp Embedding Molds</td><td>Polysciences, Inc.</td><td>18986-1</td><td>22 mm x 22 mm square, truncated to 12 mm x 12 mm</td></tr><tr><td>JB-4&amp;nbsp;Plastic Block Holders</td><td>Polysciences, Inc.</td><td>15899</td><td></td></tr><tr><td>Disposable Graduated Transfer Pipes</td><td>VWR</td><td>414004-014</td><td></td></tr><tr><td>Petrl Dish</td><td>VWR</td><td>25384-088</td><td>Embryo dissection</td></tr><tr><td>Forceps Inox Tip</td><td>&amp;nbsp;ROBOZ</td><td>RS-5015</td><td>Embryo dissection</td></tr><tr><td>Graefe Tissue Forcep</td><td>&amp;nbsp;ROBOZ</td><td>RS-5153</td><td>Embryo dissection</td></tr><tr><td>Delicate Operating Scissor</td><td>&amp;nbsp;ROBOZ</td><td>RS-6700</td><td>Embryo dissection</td></tr><tr><td>14 ml Polypropylene Round-Bottom Tubes</td><td>Falcon</td><td>352059</td><td></td></tr><tr><td>50 ml Polypropylene Conical Tubes</td><td>Falcon</td><td>352098</td><td></td></tr><tr><td>Ice Bucket</td><td>Magic Touch Iceware International Corp.</td><td>R19-343</td><td></td></tr><tr><td>100 ml (3.4 oz.) Antistatic Polystyrene Weigh Boats</td><td>VWR</td><td>89106-766</td><td></td></tr><tr><td>330 ml (11.2 oz.) Antistatic Polystyrene Weigh Boats</td><td>VWR</td><td>89106-770</td><td></td></tr><tr><td>Sodium Chloride</td><td>MP Biomedicals</td><td>102892</td><td>PBS Solution</td></tr><tr><td>Potassium Chloride</td><td>Sigma</td><td>P0662</td><td>PBS Solution</td></tr><tr><td>Potassium phosphate monobasic</td><td>Sigma</td><td>P5405</td><td>PBS Solution</td></tr><tr><td>Sodium phosphate dibasic heptahydrate</td><td>Sigma-Aldrich</td><td>S9390</td><td>PBS Solution</td></tr><tr><td>Digital Camera</td><td>Hamamatsu Photonics</td><td>C11440-10C</td><td></td></tr><tr><td>Microtome</td><td>Leica</td><td>RM2265</td><td></td></tr><tr><td>Illuminator</td><td>Carl Zeiss</td><td>HXP 200C</td><td></td></tr><tr><td>Fluorescence Stereo Zoom Microscope</td><td>Carl Zeiss</td><td>Axio Zoom.V16</td><td></td></tr><tr><td>Stereo Microscope</td><td>Olympus</td><td>SZX16</td><td></td></tr><tr><td>Fiber Optic Light Source</td><td>Leica</td><td>KL 1500 LCD</td><td></td></tr><tr><td>Disposable Microtome Blade</td><td>VWR</td><td>95057-832</td><td></td></tr><tr><td>Single Edge Dispenser</td><td>Personna</td><td>62-0330-0000</td><td></td></tr><tr><td>ZEN</td><td>Carl Zeiss</td><td>pro 2012</td><td></td></tr><tr><td>Photoshop</td><td>Adobe</td><td>CS6 v13.0</td><td></td></tr><tr><td>Amira 3D Software</td><td>Visage Imaging</td><td>5.4</td><td></td></tr><tr><td>Computer</td><td>Dell T7600, 2x900GB SAS hard drive and 64 GB DDR3 RDIMM memory</td><td></td><td></td></tr><tr><td>Rocking Platform Shakers</td><td>VWR</td><td>40000-304</td><td></td></tr><tr><td>Standard Hot Plate Stirrer</td><td>VWR</td><td>12365-382</td><td></td></tr><tr><td>Analytical Balance</td><td>Mettler Toledo</td><td>AB54-S</td><td></td></tr></tbody></table>

rapid acquisition of 3d images using high-resolution episcopic microscopy

High-resolution episcopic microscopic (HREM) technology enables rapid acquisition of high-resolution digital volumetric and three-dimensional (3D) morphometric data. Here, we describe the detailed protocol to image the entire mouse embryo. The protocol consists of four major sections: sample preparation, embedding, image acquisition and finally, 3D visualization. The technology requires specimens to be stained with a fluorescent dye, which can be problematic for large or dense specimens. To overcome this limitation, we have improved the existing protocol to enhance tissue penetration of the dye by pretreating the specimen with a solution containing urea and sodium dodecyl sulfate. The protocol uses only routine laboratory equipment and reagents for easy adaptation in standard laboratory settings. We show that the resulting high-resolution 3D images faithfully recapitulate the detailed morphologic features of the internal organs of mouse embryos, thereby permitting morphometric analyses. Together, we present a detailed and improved protocol using standard laboratory equipment to acquire high-resolution 3D images of small and large sized specimens.

We have reported high quality HREM images of mouse embryos between e9.5 and e13.58. Image quality of the late staged embryos, however, is significantly compromised because of the limited tissue penetration of the fluorescent dye eosin. To increase the staining efficiency, we have tested several pretreatment methods that are compatible to fluorescent imaging11. Specifically, the e15.5 mouse embryos were treated with either Sca/e A212 or Whole-Body CUBIC<sup...

Watch this Scientific Journal Video about Rapid Acquisition of 3D Images Using High-resolution Episcopic Microscopy at JoVE.com

Rapid Acquisition of 3D Images Using High-resolution Episcopic Microscopy

We describe a detailed protocol using high-resolution episcopic microscopy to acquire three-dimensional (3D) images of mouse embryos. This improved protocol utilizes a modified tissue preparation method to enhance penetration of the fluorescent dye, thereby permitting morphometric analysis of both small and large-sized specimens.

High-resolution episcopic microscopic (HREM) technology enables rapid acquisition of high-resolution digital volumetric and three-dimensional (3D) ...

rapid-acquisition-3d-images-using-high-resolution-episcopic

Research

JoVE Journal

Developmental Biology

7.6K Views.  Wenzhou Medical University. We describe a detailed protocol using high-resolution episcopic microscopy to acquire three-dimensional (3D) images of mouse embryos. This improved protocol utilizes a modified tissue preparation method to enhance penetration of the fluorescent dye, thereby permitting morphometric analysis of both small and large-sized specimens.

Video: Rapid Acquisition of 3D Images Using High-resolution Episcopic Microscopy

High Resolution 3D Imaging of Ex-Vivo Biological Samples by Micro CT

Non-destructive volume visualization can be achieved only by tomographic techniques, of which the most efficient is the x-ray micro computerized tomography (&mu;CT).
High resolution &mu;CT is a very versatile yet accurate (1-2 microns of resolution) technique for 3D examination of ex-vivo biological samples1, 2. As opposed to electron tomography, the &mu;CT allows the examination of up to 4 cm thick samples. This technique requires only few hours of measurement as compared to weeks in histology. In addition, &mu;CT does not rely on 2D stereologic models, thus it may complement and in some cases can even replace histological methods3, 4, which are both time consuming and destructive. Sample conditioning and positioning in &mu;CT is straightforward and does not require high vacuum or low temperatures, which may adversely affect the structure. The sample is positioned and rotated 180&deg; or 360&deg;between a microfocused x-ray source and a detector, which includes a scintillator and an accurate CCD camera, For each angle a 2D image is taken, and then the entire volume is reconstructed using one of the different available algorithms5-7. The 3D resolution increases with the decrease of the rotation step. The present video protocol shows the main steps in preparation, immobilization and positioning of the sample followed by imaging at high resolution.

High Resolution 3D Imaging of Ex-Vivo Biological Samples by Micro CT

Non-destructive volume visualization can be achieved only by tomographic techniques, of which the most efficient is the x-ray micro computerized tomography ( CT).

Non-destructive volume visualization can be achieved only by tomographic techniques, of which the most efficient is the x-ray micro computerized ...

Bioengineering

High-resolution Episcopic Microscopy (HREM) - Simple and Robust Protocols for Processing and Visualizing Organic Materials

We provide simple protocols for generating digital volume data with the high-resolution episcopic microscopy (HREM) method. HREM is capable of imaging organic materials with volumes up to 5 x 5 x 7 mm3 in typical numeric resolutions between 1 x 1 x 1 and 5 x 5 x 5 &#181;m3. Specimens are embedded in methacrylate resin and sectioned on a microtome. After each section an image of the block surface is captured with a digital video camera that sits on the phototube connected to the compound microscope head. The optical axis passes through a green fluorescent protein (GFP) filter cube and is aligned with a position, at which the bock holder arm comes to rest after each section. In this way, a series of inherently aligned digital images, displaying subsequent block surfaces are produced. Loading such an image series in three-dimensional (3D) visualization software facilitates the immediate conversion to digital volume data, which permit virtual sectioning in various orthogonal and oblique planes and the creation of volume and surface rendered computer models. We present three simple, tissue specific protocols for processing various groups of organic specimens, including mouse, chick, quail, frog and zebra fish embryos, human biopsy material, uncoated paper and skin replacement material.

High-resolution Episcopic Microscopy HREM - Simple and Robust Protocols for Processing and Visualizing Organic Materials

We provide simple and robust protocols for processing biopsy material of various species, embryos of biomedical model organisms and samples of other organic tissues in order to permit digital volume data generation with the high-resolution episcopic microscopy method.

We provide simple protocols for generating digital volume data with the high-resolution episcopic microscopy (HREM) method. HREM is capable of imaging ...

Highly Resolved Intravital Striped-illumination Microscopy of Germinal Centers

Monitoring cellular communication by intravital deep-tissue multi-photon microscopy is the key for understanding the fate of immune cells within thick tissue samples and organs in health and disease. By controlling the scanning pattern in multi-photon microscopy and applying appropriate numerical algorithms, we developed a striped-illumination approach, which enabled us to achieve 3-fold better axial resolution and improved signal-to-noise ratio, i.e. contrast, in more than 100 &#181;m tissue depth within highly scattering tissue of lymphoid organs as compared to standard multi-photon microscopy. The acquisition speed as well as photobleaching and photodamage effects were similar to standard photo-multiplier-based technique, whereas the imaging depth was slightly lower due to the use of field detectors. By using the striped-illumination approach, we are able to observe the dynamics of immune complex deposits on secondary follicular dendritic cells &#8211; on the level of a few protein molecules in germinal centers.

High-resolution intravital imaging with enhanced contrast up to 120 &#181;m depth in lymph nodes of adult mice is achieved by spatially modulating the excitation pattern of a multi-focal two-photon microscope. In 100 &#181;m depth we measured resolutions of 487 nm (lateral) and 551 nm (axial), thus circumventing scattering and diffraction limits.

Monitoring cellular communication by intravital deep-tissue multi-photon microscopy is the key for understanding the fate of immune cells within thick ...

Immunology and Infection

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

Three and Four-Dimensional Visualization and Analysis Approaches to Study Vertebrate Axial Elongation and Segmentation

Somitogenesis is a hallmark of vertebrate embryonic development. For years, researchers have been studying this process in a variety of organisms using a wide range of techniques encompassing ex vivo and in vitro approaches. However, most studies still rely on the analysis of two-dimensional (2D) imaging data, which limits proper evaluation of a developmental process like axial extension and somitogenesis involving highly dynamic interactions in a complex 3D space. Here we describe techniques that allow mouse live imaging acquisition, dataset processing, visualization and analysis in 3D and 4D to study the cells (e.g., neuromesodermal progenitors) involved in these developmental processes. We also provide a step-by-step protocol for optical projection tomography and whole-mount immunofluorescence microscopy in mouse embryos (from sample preparation to image acquisition) and show a pipeline that we developed to process and visualize 3D image data. We extend the use of some of these techniques and highlight specific features of different available software (e.g., Fiji/ImageJ, Drishti, Amira and Imaris) that can be used to improve our current understanding of axial extension and somite formation (e.g., 3D reconstructions). Altogether, the techniques here described emphasize the importance of 3D data visualization and analysis in developmental biology, and might help other researchers to better address 3D and 4D image data in the context of vertebrate axial extension and segmentation. Finally, the work also employs novel tools to facilitate teaching vertebrate embryonic development.

Here, we describe computational tools and methods that allow visualization and analysis of three and four-dimensional image data of mouse embryos in the context of axial elongation and segmentation, obtained by in toto optical projection tomography, and by live imaging and whole-mount immunofluorescence staining using multiphoton microscopy.

Somitogenesis is a hallmark of vertebrate embryonic development. For years, researchers have been studying this process in a variety of organisms using a ...

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 ...

Quantitative Analysis of Autophagy using Advanced 3D Fluorescence Microscopy

Prostate cancer is the leading form of malignancies among men in the U.S. While surgery carries a significant risk of impotence and incontinence, traditional chemotherapeutic approaches have been largely unsuccessful. Hormone therapy is effective at early stage, but often fails with the eventual development of hormone-refractory tumors. We have been interested in developing therapeutics targeting specific metabolic deficiency of tumor cells. We recently showed that prostate tumor cells specifically lack an enzyme (argininosuccinate synthase, or ASS) involved in the synthesis of the amino acid arginine1. This condition causes the tumor cells to become dependent on exogenous arginine, and they undergo metabolic stress when free arginine is depleted by arginine deiminase (ADI)1,10. Indeed, we have shown that human prostate cancer cells CWR22Rv1 are effectively killed by ADI with caspase-independent apoptosis and aggressive autophagy (or macroautophagy)1,2,3. Autophagy is an evolutionarily-conserved process that allows cells to metabolize unwanted proteins by lysosomal breakdown during nutritional starvation4,5. Although the essential components of this pathway are well-characterized6,7,8,9, many aspects of the molecular mechanism are still unclear - in particular, what is the role of autophagy in the death-response of prostate cancer cells after ADI treatment? In order to address this question, we required an experimental method to measure the level and extent of autophagic response in cells - and since there are no known molecular markers that can accurately track this process, we chose to develop an imaging-based approach, using quantitative 3D fluorescence microscopy11,12.
Using CWR22Rv1 cells specifically-labeled with fluorescent probes for autophagosomes and lysosomes, we show that 3D image stacks acquired with either widefield deconvolution microscopy (and later, with super-resolution, structured-illumination microscopy) can clearly capture the early stages of autophagy induction. With commercially available digital image analysis applications, we can readily obtain statistical information about autophagosome and lysosome number, size, distribution, and degree of colocalization from any imaged cell. This information allows us to precisely track the progress of autophagy in living cells and enables our continued investigation into the role of autophagy in cancer chemotherapy.

Autophagy is a ubiquitous process that enables cells to degrade and recycle proteins and organelles. We apply advanced fluorescence microscopy to visualize and quantify the small, but essential, physical changes associated with the induction of autophagy, including the formation and distribution of autophagosomes and lysosomes, and their fusion into autolysosomes.

Prostate cancer is the leading form of malignancies among men in the U.S. While surgery carries a significant risk of impotence and incontinence, ...

Biology

Super-resolution Imaging of the Cytokinetic Z Ring in Live Bacteria Using Fast 3D-Structured Illumination Microscopy (f3D-SIM)

Imaging of biological samples using fluorescence microscopy has advanced substantially with new technologies to overcome the resolution barrier of the diffraction of light allowing super-resolution of live samples. There are currently three main types of super-resolution techniques &#8211; stimulated emission depletion (STED), single-molecule localization microscopy (including techniques such as PALM, STORM, and GDSIM), and structured illumination microscopy (SIM). While STED and single-molecule localization techniques show the largest increases in resolution, they have been slower to offer increased speeds of image acquisition. Three-dimensional SIM (3D-SIM) is a wide-field fluorescence microscopy technique that offers a number of advantages over both single-molecule localization and STED. Resolution is improved, with typical lateral and axial resolutions of 110 and 280 nm, respectively and depth of sampling of up to 30 &#181;m from the coverslip, allowing for imaging of whole cells. Recent advancements (fast 3D-SIM) in the technology increasing the capture rate of raw images allows for fast capture of biological processes occurring in seconds, while significantly reducing photo-toxicity and photobleaching. Here we describe the use of one such method to image bacterial cells harboring the fluorescently-labelled cytokinetic FtsZ protein to show how cells are analyzed and the type of unique information that this technique can provide.

Super-resolution Imaging of the Cytokinetic Z Ring in Live Bacteria Using Fast 3D-Structured Illumination Microscopy f3D-SIM

Spatiotemporal information about dynamic proteins inside live cells is crucial for understanding biology. A type of super-resolution microscopy called fast 3D-structured illumination microscopy (f3D-SIM) reveals unique information about the cytokinetic Z ring in bacteria: both its bead-like appearance and the rapid dynamics of FtsZ within the ring.

Imaging of biological samples using fluorescence microscopy has advanced substantially with new technologies to overcome the resolution barrier of the ...

Three-dimensional Super Resolution Microscopy of F-actin Filaments by Interferometric PhotoActivated Localization Microscopy (iPALM)

Fluorescence microscopy enables direct visualization of specific biomolecules within cells. However, for conventional fluorescence microscopy, the spatial resolution is restricted by diffraction to ~ 200 nm within the image plane and &#62; 500 nm along the optical axis. As a result, fluorescence microscopy has long been severely limited in the observation of ultrastructural features within cells. The recent development of super resolution microscopy methods has overcome this limitation. In particular, the advent of photoswitchable fluorophores enables localization-based super resolution microscopy, which provides resolving power approaching the molecular-length scale. Here, we describe the application of a three-dimensional super resolution microscopy method based on single-molecule localization microscopy and multiphase interferometry, called interferometric PhotoActivated Localization Microscopy (iPALM). This method provides nearly isotropic resolution on the order of 20 nm in all three dimensions. Protocols for visualizing the filamentous actin cytoskeleton, including specimen preparation and operation of the iPALM instrument, are described here. These protocols are also readily adaptable and instructive for the study of other ultrastructural features in cells.

Three-dimensional Super Resolution Microscopy of F-actin Filaments by Interferometric PhotoActivated Localization Microscopy iPALM

We present a protocol for the application of interferometric PhotoActivated Localization Microscopy (iPALM), a 3-dimensional single-molecule localization super resolution microscopy method, to the imaging of the actin cytoskeleton in adherent mammalian cells. This approach allows light-based visualization of nanoscale structural features that would otherwise remain unresolved by conventional diffraction-limited optical microscopy.

Fluorescence microscopy enables direct visualization of specific biomolecules within cells. However, for conventional fluorescence microscopy, the spatial ...

Multilayer Imaging with Microprism in Head-Fixed and Freely Moving Animals

Long-Term Imaging of Identified Neural Populations using Microprisms in Freely Moving and Head-Fixed Animals

With the advancement of multi-photon microscopy and molecular technologies, fluorescence imaging is rapidly growing to become a powerful approach for studying the structure, function, and plasticity of living brain tissues. In comparison to conventional electrophysiology, fluorescence microscopy can capture the neural activity as well as the morphology of the cells, enabling long-term recordings of the identified neuron populations at single-cell or subcellular resolution. However, high-resolution imaging typically requires a stable, head-fixed setup that restricts the movement of the animal, and the preparation of a flat surface of transparent glass allows visualization of neurons at one or more horizontal planes but is limited in studying the vertical processes running across different depths. Here, we describe a procedure to combine a head plate fixation and a microprism that gives multilayer and multimodal imaging. This surgical preparation not only gives access to the entire column of the mouse visual cortex but allows two-photon imaging in a head-fixed position and one-photon imaging in a freely moving paradigm. Using this approach, one can sample identified cell populations across different cortical layers, register their responses under head-fixed and freely moving states, and track the long-term changes over months. Thus, this method provides a comprehensive assay of the microcircuits, enabling direct comparison of neural activities evoked by well-controlled stimuli and under a natural behavioral paradigm.

Author Spotlight: Comparative Imaging of Neural Activity in Awake and Freely Moving States

When integrated with a head-plate and an optical design compatible with both single- and two-photon microscopes, the microprism lens presents a significant advantage in measuring neural responses in a vertical column under diverse conditions, including well-controlled experiments in head-fixed states or natural behavioral tasks in freely moving animals.

With the advancement of multi-photon microscopy and molecular technologies, fluorescence imaging is rapidly growing to become a powerful approach for ...

Rapid Acquisition of 3D Images Using High-resolution Episcopic Microscopy

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