The authors would like to thank Roland Myers for kindly providing the original TEM protocols. In addition, we would like to thank Dr. John Colbourne for providing the Daphnia specimen, Dr. Santhosh Girirajan for providing the Drosophila specimen, and Dr. Fadia Kamal for providing the mouse embryo. The investigators acknowledge funding support from the NIH (1R24OD18559-01-A2, PI: KCC), the Jake Gittlen Laboratories for Cancer Research, and from pilot award funding from the PSU Huck Institutes of the Life Sciences and the Institute for CyberScience.

All procedures on live animals were approved by the Institutional Animal Care and Use Committee (IACUC) at the Pennsylvania State University.
1. Day 1: Fixation
<ol>
	<li>To improve the fixation and reduce the volume of gut contents, starve the juvenile fish for at least 24 h for a 10 mm specimen, or longer for larger specimens (e.g., 72 h for a 14 mm specimen).</li>
	<li>Pre-chill 10% Neutral Buffered Formalin (NBF) and 2x Tricaine-S (MS-222, 400 mg/L) on ice to 4 &#176;C.</li>
	<li>Double the fish water volume with chilled 2x MS-222 for rapid and humane euthanasia.</li>
	<li>Wait for 30 s (larvae) to 60 s (juveniles) after the fish stops moving.</li>
	<li>Immerse the fish in chilled 10% NBF for 10 min.</li>
	<li>Fix the fish in chilled 10% NBF solution overnight in flat-bottomed containers at room temperature. 
	NOTE: Flat-bottomed containers are required to minimize the bending of the specimen. The volume of fixative and solutions of all subsequent steps should be at least 20 times the sample volume unless otherwise specified. For 1-5 larval zebrafish, the recommended solution volume is at least 1 mL. Insufficient fixative results in poor fixation, and a surplus of reagent volume does not negatively impact the outcome of the described procedure. For complete fixation of larger fish, slit the belly starting slightly anterior to the anal pore. Use minimal penetration of scissors or knife and apply gentle force away from the fish while cutting to minimize damage to internal organs. Fixation protocols have been described by others31,32,33.</li>
</ol>
2. Day 2: Dehydration &#38; Staining
<ol>
	<li>Rinse fixed specimens 3 times in 1x phosphate buffered saline (PBS) pH 7.4 for 10 min.</li>
	<li>Submerge the samples in 35% ethyl alcohol (EtOH) for 20 min at room temperature with gentle agitation. 
	NOTE: For all gentle agitation steps, use a tabletop shaker and set for a mixing effect while carefully avoiding bumping and rubbing of the sample against the container surfaces.</li>
	<li>Submerge the samples in 50% EtOH for 20 min at room temperature with gentle agitation.</li>
	<li>Dissolve phosphotungstic acid (PTA) powder in water to prepare a 1% w/v stock solution.</li>
	<li>Dilute the 1% PTA stock solution 3:7 in 100% EtOH to obtain a 0.3% PTA staining solution.</li>
	<li>Stain the samples overnight in the 0.3% PTA solution at room temperature with gentle agitation.</li>
</ol>
3. Day 3: Infiltration
<ol>
	<li>Prepare 1:1 v/v mixture of 100% EtOH and LR White acrylic resin and set aside.</li>
	<li>Rinse 3 times in 70% EtOH for 10 min.</li>
	<li>Submerge the samples in 90% EtOH for 30 min at room temperature with gentle agitation.</li>
	<li>Submerge the samples in 95% EtOH for 30 min at room temperature with gentle agitation.</li>
	<li>Submerge the samples twice in 100% EtOH for 30 min at room temperature with gentle agitation.</li>
	<li>Submerge the samples in the 1:1 EtOH and LR White acrylic resin mixture overnight at room temperature with gentle agitation.</li>
</ol>
4. Day 4: Embedding
<ol>
	<li>Submerge the samples in 100% LR White resin for 2 h at room temperature with gentle agitation.</li>
	<li>Replace the resin in step 4.1 with fresh 100% LR White resin and incubate for 1 h at room temperature with gentle agitation.</li>
	<li>Assemble the embedding apparatus (Figure 1).</li>
	<li>Cut polyimide tubing of appropriate diameter and length. 
	NOTE: The tubing with an inner diameter that is at least 0.1 mm larger than the diameter of the specimen is recommended. For typical specimens, a standard length of 30 mm of tubing is used for each sample. When desired, elongated specimens can be accommodated by using longer tubing.
	<ol>
		<li>Attach a P1000 micropipette to the wide end of the embedding adapter and insert the polyimide tubing to the narrow end (Figure 1C). If the embedding adapter is not available, go to Step 4.3.3. Otherwise, continue to Step 4.4.</li>
		<li>Clip off the end of a micropipette tip straight across such that the polyimide tubing fits snugly. Cut at 44 mm from the wide end of a 200 &#181;L yellow micropipette tip for the 0.0403&#34; tubing and 59 mm from bottom of 1 mL blue tip for the 0.105&#34; tubing. Trim as needed.</li>
		<li>Attach the cut micropipette tip to a micropipette.</li>
	</ol>
	</li>
	<li>Transfer the samples to a small weigh boat or V-shaped solution basin and fully submerge the samples in 100% LR White resin.</li>
	<li>With the embedding apparatus from Step 4.3, aim the tubing at the wide end of the fixed sample (or towards the head in the case of animal specimens) and pipette the specimen slowly into the tubing. Position the sample in the middle of the tubing and ensure the tubing above and below the specimen is filled with resin.
	<ol>
		<li>Pipette the sample into the tubing such that the sample is moving in the natural, forward direction. 
		NOTE: Slow pipetting avoids air bubbles and specimen damage caused by abrasion against the tubing edge. Backward movement into the tubing can easily damage extremities (limbs, wings, fins, etc.). It is recommended to allow some resin overflow in the embedding apparatus so that, later, air does not enter the tubing during removal.</li>
	</ol>
	</li>
	<li>Immediately seal the open end with oil-based soft modeling clay.
	<ol>
		<li>Flatten the clay into a 1 mm-thick sheet.</li>
		<li>Stabilize the tubing between index and middle fingers.</li>
		<li>Slowly press the clay against the end of the tubing with thumb.</li>
		<li>Remove excess clay.</li>
	</ol>
	</li>
	<li>Remove the embedding apparatus from the micropipette.</li>
	<li>Pull out the tubing by gentle rotation and seal the other end with the clay.
	<ol>
		<li>Hold a finger against the sealed end to prevent the ejection of the clay.</li>
		<li>Slowly push the unsealed end of the tubing into the clay.</li>
	</ol>
	</li>
	<li>Place the tubing horizontally and polymerize the resin for 24 h at 65 &#176;C. 
	NOTE: Horizontal placement avoids the sample movement during polymerization. If a small amount of air was trapped in Step 4.8, the end with the air bubble may be elevated slightly to prevent air bubble movement toward the specimen. Importantly, too much elevation during polymerization may cause the specimen to fall towards the low end due to gravity. A properly embedded sample is shown compared to one with an air bubble, which should be avoided because the refraction from air/resin interfaces can degrade image quality (Figure 2).</li>
</ol>
5. Day 5: Collection
<ol>
	<li>Collect the samples for image acquisition at the end of Day 5. 
	NOTE: The removal of polyimide tubing is possible but not necessary for imaging. Synchrotron-based micro-CT imaging for the zebrafish sample was performed at Beamline 2-BM at Advanced Photon Source (APS) in Argonne National Laboratory (Lemont, IL, USA). For PTA-stained zebrafish, an optimal X-ray energy range of 5-30 keV was determined, and an X-ray energy of 13.8 keV was used for imaging. 1501 projections were obtained at 13.8 keV over 180 degrees (1 projection every 0.12 degrees) with a 2048-by-2048 pixel camera. Additionally, two flat-field (gain) images (one at the beginning and one at the end of the acquisition) and one dark-field image were also acquired. Flat-field and dark-field corrections, ring artifact reduction, and image reconstruction were done using the open source TomoPy toolkit34. PTA-stained Daphnia and Drosophila samples were imaged on an X-ray microscope12,35. Specific micro-CT settings are dependent on the object and the quality of the stain. For example, the Drosophila specimen was scanned at 40 kVp, 74 mA, and 15 s at a voxel resolution of 3.10&#160;&#215; 3.10&#160;&#215; 3.10 &#181;m3.</li>
</ol>

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All of the authors declare that there are no conflicts of interests.

In this manuscript, we present a detailed protocol for rigid immobilization of fixed and stained millimeter-scale (0.5 to 2.5 mm diameter) specimens for micro-CT imaging. Given the rapid advance of micro-CT technology with regard to resolution and speed, a method of permanent sample preservation with re-imaging capabilities is highly desirable. Traditionally, dense embedding resin are commonly used in electron microscopy to provide structural support to cut ultrathin sections and minimize the damage to the specimen. Our method adapted its use for rigid immobilization during non-destructive imaging. To minimize the imaging artifacts from the presence of excess resin, we have implemented the use of X-ray transparent polyimide tubing to create round samples without edges, and to restrict the volume of resin around the sample.
Optimal outcome of the embedding procedure is contingent upon careful execution of several critical steps and attention to sample integrity throughout the procedure. Indeed, damaging the specimen will result in a compromised final image regardless of the success of execution of imaging. Since chilling contributes to a diminution of pain responses, and unfixed tissue degrades more rapidly at higher temperatures, we use pre-chilled reagents. Large specimens may need to be cut to allow the entry of fixative into the inside of the specimen so that internal organs such as liver, pancreas, and gut are completely fixed. Unfixed tissues deteriorate and lose structural integrity, destroying biological structure. Another key step is to maintain the specimen in its natural alignment. Therefore, we advocate for the use of flat-bottomed containers, which we use throughout the procedure and are particularly important during fixation. Fixation in polypropylene tubes or conical tubes that usually have a V-shaped bottom should be avoided because they can cause elongated samples to bend, which distorts the natural morphology of the specimen. In addition, to minimize the use of resin, the inner diameter of the polyimide tubing is only slightly larger than the width of the specimen. Bending of the specimen can also result in damage to the specimen as it enters the tubing. It is also recommended to transfer the specimen into the tubing in a natural forward manner to avoid damaging extremities. Proper dehydration is another critical step. This is accomplished with small increments of increasing EtOH concentrations to slowly replace water with EtOH, which is more miscible with the resin. Large increases in EtOH concentration are associated with tissue shrinkage and can be ameliorated by additional increments of EtOH incubation to make the transition more gradual. Finally, to minimize the movement of the specimen or air pockets, if there are any, the samples are placed horizontally during the resin polymerization (end of Day 4). Air pockets or sealant next to the specimen causes optical artifacts with X-ray imaging associated with edge diffraction, reducing image quality.
With respect to possible modifications to the protocol, other embedding resins and metal stains can be used in place of LR White acrylic and PTA, respectively. Embed812, an epoxy resin commonly used in electron microscopy, is highly viscous, more difficult to transfer, may cause the distortion of the specimens, and is associated with higher background than acrylic or glycol methacrylate resins. While JB4 Plus, a glycol methacrylate, is less viscous than EMbed812 and has lower background, random formation of air pockets frequently appear in the vicinity of the specimen. As mentioned, air pockets degrade the image quality and are particularly problematic for precious samples. It is worth noting that Technovit 7100, another glycol methacrylate, interferes with PTA staining, resulting in poor contrast in the final image. Comparatively, LR White acrylic has water-like viscosity, low background, and does not interfere with PTA staining. In our hands, the success rate of embedding in LR White without sample damage or air bubbles is greater than 95%. For these reasons, we advocate its use as the embedding resin for tissue micro-CT. With respect to stains, the outcome of various metal stains such as osmium tetroxide, iodine, and PTA in micro-CT imaging has been compared and discussed elsewhere15,16,17 and is outside the scope of this manuscript.
The sample size is a major limitation to our current protocol. Since we employ suction to transfer specimens into the tubing, the ability to see the specimen is critical for orientation and ensuring that it is indeed in the tubing. As a result, sub-millimeter samples such as Tardigrade (i.e., water bears) are extremely difficult to embed because they require the use a microscope to be visualized. Once suction is applied, microscopic specimens are easily lost from the focal plane and become challenging to rediscover, making it difficult to determine if they passed too far through the attached end of the tubing. Larger samples, such as mouse embryos, (3 - 10 mm) can be embedded with slight modifications to the embedding protocol (Figure 4D). In particular, the polyimide tubing was filled to 1/3 with liquid resin, which was polymerized prior to embedding. The fixed and stained sample was placed on top of the pre-polymerized resin. The tubing was then fully filled with un-polymerized resin and followed by a second polymerization.
The significance of our embedding protocol is manifold. Rigidly immobilized whole animal or tissue specimens are desirable for reasons including: (1) creating long-term repositories of samples that are difficult to generate or prepare; (2) re-acquisition of data; (3) enabling serial imaging using multiple imaging modalities; and (4) providing standards for calibration and technology development. Samples prepared with our embedding procedure are encased in solid resin that is resilient against damage and can therefore be easily stored perhaps indefinitely. Long-term storage is particularly useful for rare or laboriously generated specimens such as those associated with phenome projects. Further, in the event that the digital data are lost, the data can be generated from the original sample. The capability of re-imaging over a long period of time potentially allows the same sample to be interrogated with more advanced imaging modalities in the future. Since the samples are physically stable between imaging instances, images are acquired from the same specimen in the same orientation, facilitating registration between earlier and later data sets. For example, a low-resolution image may only allow segmentation of tissues in a zebrafish larva. The same larva can later be re-imaged at higher-resolution so that more detailed computational analyses at cellular resolution. This capability for direct comparison between registered scans suggests resin-embedded samples as a potential standard for technology development of micro-CT. The effects of any alteration to the imaging method can be assessed by imaging of the same sample so that all variables in the sample preparation step are controlled. Finally, a resin-embedded standard sample can be used for instrument calibration to test for consistency of imaging.
Our previous work examining mutant and diseased fish histology showed that cellular resolution allows detection of subtle abnormalities in tissue structure that are overlooked in gross examination using the dissecting microscope36, or a low-power stereo microscope. We wish to expand our high-resolution analyses to human samples. Zebrafish larvae, which comprise the primary subject of our development work, are similar to human needle biopsies in their fragility, cellular and intercellular tissue heterogeneity, size (1-3 mm diameter), and elongated shape. Based on our extensive experience with zebrafish, and other work showing that micro-CT successfully stained and scanned human tissue, albeit at lower resolution37, we expect our approaches to bring added value to imaging and analysis of needle biopsies. We have estimated that the assembly of a kit sufficient for one preparation of up to 20 samples of the same condition to be potentially less than 30 USD. The samples prepared by our embedding method are resistant to physical damage and therefore easy to transport. The low cost and ease of transport suggest the possibility of collection of samples from across the world, particularly areas in which cellular resolution imaging with micro-CT is not readily accessible. Our vision is for high resolution digital files of needle biopsies (ranging from 0.1 to several terabytes) to enable the creation of a digital atlas of human tissues, and at the same time allow the researchers to refine and enhance current analysis pipelines for localization and quantitative characterization of cellular and tissue architecture within the full 3D context of the tissues scanned.

Phenotypes are observable traits of an organism that represent the consequences of unique interactions between its genetic background and the environment. These characteristics can include behavioral, biochemical, morphological, developmental, and/or physiological properties. Importantly, differences in traits between wild-type organisms and genetic mutants can provide key insight into the mechanisms and functions of affected genes. With respect to morphology, histopathology is the gold standard for assessing phenotypes at the cellular level but suffers from mechanical artifacts and does not allow accurate quantitative volumetric analysis1. Our laboratory is motivated to overcome the barriers to the application of the diagnostic power of histology to full tissue volumes at submicron resolution.
A survey of available technologies suggests that imaging by X-ray micro-computed tomography (micro-CT) may provide ideal capabilities needed for millimeter-scale whole-animal 3-dimensional (3D) histology. Micro-CT enables non-destructive, isotropic, 3D visualization and the ability to conduct quantitative analysis of tissue architecture1,2. In recent years, 3D imaging of unstained and metal-stained zebrafish by micro-CT has gained increasing traction and has been utilized for volumetric analysis of several tissues, including muscle, teeth, bone, and adipose tissues3,4,5,6,7,8,9,10. Other model organisms and tissue specimens are also amenable to micro-CT imaging. For instance, a pipeline has been introduced for quantifying dense mesoscale neuroanatomy of mouse brains imaged via synchrotron micro-CT11. Similarly, an eosin-based staining protocol has been shown to be suitable for soft tissue micro-CT imaging of whole mouse organs12. Morphometric analysis of unstained human L3 vertebrae and the visualization of silver-stained human lungs using micro-CT have demonstrated the utility of this technology for human samples13,14.
In order to actualize the great promise of micro-CT for complete morphological phenotyping of intact small animals and tissue specimens, a number of hurdles need to be overcome in relation to throughput, resolution, field-of-view, comprehensive cell staining, and long-term preservation. While each of these aspects is critically important, the focus of the present manuscript is the optimization of embedding procedures, with additional notes on sample fixation and staining. Heavy metal staining is useful because the inherent contrast between different soft tissues in micro-CT images is low. The potency of various metal stains, such as osmium tetroxide, iodine, phosphotungstic acid (PTA), and gallocyanin-chromalum, to enhance contrast for micro-CT imaging has been studied15,16,17. Uranyl acetate has also been used as a contrasting agent for micro-CT imaging of bone and cartilage18,19. PTA as used in our protocol leads to consistent staining of nearly all tissues and cells in whole zebrafish specimens, yielding images that are potentially compatible with histology-like studies through full volumes of tissue.
During micro-CT image acquisition, a 2-dimensional (2D) projection is taken as the sample is rotated by a fraction of a degree and repeated until the sample has completed a 180&#176; or 360&#176; rotation, generating a series of thousands of 2D projections used for the reconstruction of the 3D volume20. In this process, any perturbation of the specimen will cause the corresponding 2D projections to be out of alignment, resulting in poorly reconstructed 3D volumes. Sample immobilization is a way to correct for this issue and some current strategies involve submerging the sample in alcohol or embedding in agarose in polypropylene tubes or micropipette tips3,5,15,16,21,22. Liquid immersion methods are not ideal because sample movement during image acquisition can occur between imaging sets, resulting in shifted reconstructions of 3D volumes23. Further, it is well known that the physical stability of liquid-stored tissue samples is poor in time scales of months to years, as a result of chemical instability and surface abrasion associated with movement of contact points between sample and the container wall (personal observations with fixed animal and human tissue samples).
To reduce the potential for sample movement during imaging, samples can be embedded in agarose24,25,26, but this practice is associated with the risk of the stain diffusing into agarose thus reducing image contrast26. Furthermore, liquid or agarose preparations are prone to physical damage and degrade over time, making them unsuitable for long-term sample storage. We reasoned that a potentially successful and straightforward sample immobilization method could be adapted from that used for electron microscopy specimens. Polymerized resins such as EPON 812 (discontinued and replaced by EMbed 812) are commonly used to provide the hardness required to generate ultrathin sections for electron microscopy27,28. Indeed, several comparative studies between X-ray imaging and electron microscopy have demonstrated that the samples embedded in resin can be imaged by X-ray microscopy29,30. However, some of the standard practices of electron microscopy do not translate directly to micro-CT imaging. For example, micro-CT imaging of samples with squared resin edges of typical resin blocks and samples cut from those blocks is associated with edge diffraction artifacts that can interfere with imaging. Smoothing resin edges, while possible, is laborious and time-consuming.
While a variety of plastic embedding resins are employed in electron microscopy, the high background associated with harder resins such as Embed 812 led us to test others. We chose LR White due to its low viscosity, low shrinkage, low tendency to form bubbles during polymerization, and lower background. To create edge-free samples, and to minimize the quantity of resin surrounding the sample, we developed the protocols to draw specimens in liquid resin into polyimide tubing prior to the polymerization. Polyimide was chosen for its high thermal stability and high X-ray transmittance so that the removal of the tubing is not necessary for imaging. Finally, we designed a custom micropipette adapter for our sample embedding technique in order to reliably hold the tubing and prevent the contamination of micropipettes. The adapter can be 3D printed from a CAD image file. Taken together, the goal of the sample preparation methods presented here is to make embedding of small samples more straightforward, achieve enhanced staining contrast, rigid immobilization, and long-term storage of intact millimeter-scale specimens.

<table><tbody><tr><td>10X Neutral buffered formalin</td><td>Fisher Scientific</td><td>SF100-4</td><td></td></tr><tr><td>Phosphotunstic Acid</td><td>VWR</td><td>MK282402</td><td></td></tr><tr><td>Tricaine-S (MS-222)</td><td>Western Chemical</td><td>MS-222</td><td></td></tr><tr><td>LR White</td><td>Electron Microscopy Sciences</td><td>14380</td><td></td></tr><tr><td>Polyimide tubing (ID 0.04&quot;)</td><td>Nordson Medical</td><td>141-0065</td><td>The exact polyimide tubings described in the manuscript are no longer available. A close match is provided here. Tubing with custom diameter can be manufactured upon request.</td></tr><tr><td>Ethyl Alcohol 200 Proof</td><td>Pharmco-Aaper</td><td>111000200</td><td></td></tr><tr><td>Oil-based soft modeling clay</td><td>Sculpey</td><td>S302 001</td><td></td></tr><tr><td>Micropipette P200</td><td>Gilson</td><td>F123601</td><td></td></tr><tr><td>Micropipette P1000</td><td>Gilson</td><td>F123502</td><td></td></tr><tr><td>Glass vials</td><td>VWR</td><td>66015-042</td><td></td></tr><tr><td>V-shaped basin</td><td>VWR</td><td>89094-676</td><td></td></tr><tr><td>Weigh boats</td><td>VWR</td><td>10803-148</td><td></td></tr><tr><td>200 &amp;mu;L yellow micropipette tip</td><td>Fisher Scientific</td><td>02-707-500</td><td></td></tr><tr><td>1 mL blue micropipette tip</td><td>Fisher Scientific</td><td>02-681-163</td><td></td></tr><tr><td>Tabletop shaker</td><td>Thermolyne</td><td>M71735</td><td></td></tr><tr><td>Camera</td><td>Photometrics</td><td>CoolSNAP HQ2 CCD</td><td></td></tr><tr><td>X-ray microscope</td><td>Zeiss</td><td>Xradia 520 Versa</td><td></td></tr></tbody></table>

rigid embedding of fixed and stained, whole, millimeter-scale specimens for section-free 3d histology by micro-computed tomography

For over a hundred years, the histological study of tissues has been the gold standard for medical diagnosis because histology allows all cell types in every tissue to be identified and characterized. Our laboratory is actively working to make technological advances in X-ray micro-computed tomography (micro-CT) that will bring the diagnostic power of histology to the study of full tissue volumes at cellular resolution (i.e., an X-ray Histo-tomography modality). Toward this end, we have made targeted improvements to the sample preparation pipeline. One key optimization, and the focus of the present work, is a straightforward method for rigid embedding of fixed and stained millimeter-scale samples. Many of the published methods for sample immobilization and correlative micro-CT imaging rely on placing the samples in paraffin wax, agarose, or liquids such as alcohol. Our approach extends this work with custom procedures and the design of a 3-dimensional printable apparatus to embed the samples in an acrylic resin directly into polyimide tubing, which is relatively transparent to X-rays. Herein, sample preparation procedures are described for the samples from 0.5 to 10 mm in diameter, which would be suitable for whole zebrafish larvae and juveniles, or other animals and tissue samples of similar dimensions. As proof of concept, we have embedded the specimens from Danio, Drosophila, Daphnia, and a mouse embryo; representative images from 3-dimensional scans for three of these samples are shown. Importantly, our methodology leads to multiple benefits including rigid immobilization, long-term preservation of laboriously-created resources, and the ability to re-interrogate samples.

The protocol described above details the sample preparation procedure for whole zebrafish larvae and juveniles for micro-CT imaging. Notably, this method is readily adapted to other samples types (e.g., Drosophila, Daphnia, Arabidopsis, mouse organs). Incubation times in various steps are appropriate for zebrafish larvae and samples of similar size; larger samples may require longer incubation. To assist with the sample embedding steps, we designed an adapter that attaches to a 1 mL micropipette, and can be customized for various sizes of polyimide tubing (Figure 1). This custom adapter was 3D printed from a CAD file that is associated with this manuscript and available for download from http://publications.chenglab.com/LinEtAlJOVE/. Since all readers may not have access to a 3D printer, assembly of a homemade embedding apparatus using micropipette tips is described in the protocol (Step 4.3). A successfully embedded zebrafish larva is shown compared to a specimen with an air bubble (Figure 2), which will likely degrade the image quality. To demonstrate the versatility of the embedding procedure, we show the specimens from various model organisms (i.e., Danio, Drosophila, Daphnia, mouse embryo) that have gone through the sample preparation pipeline (Figure 3). Reconstructed 3D volumes of whole Danio, Daphnia, and Drosophila specimens imaged by micro-CT are shown (Figure 4). Importantly, as demonstrated by the Danio sample, these specimens can be stored for an extended period of time and reused for multiple imaging sessions (Figure 5).
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/58293/58293fig1.jpg" /> 
Figure 1. Embedding apparatus. (A) An adapter was designed to facilitate the embedding using common laboratory tools. (B) A cross-sectional illustration of the adapter. (C) The embedding apparatus after the insertion of the polyimide tubing at adapter point (a) and attaching the micropipette at adapter point (c). Adapter segment (b) is an overflow chamber designed to accommodate excess resin and protect the micropipette from contamination. <a href="https://www.jove.com/files/ftp_upload/58293/58293fig1large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 2" class="xfigimg" src="/files/ftp_upload/58293/58293fig2.jpg" /> 
Figure 2. Successfully embedded specimens are devoid of air bubbles. (A) An embedded 3 day post-fertilization (dpf) larval zebrafish without air bubble. (B) An embedded 3 dpf zebrafish with an air bubble. Air trapped during the embedding process can move toward the specimen if the sample is not placed horizontally during the polymerization. Scale bars = 1 mm. <a href="https://www.jove.com/files/ftp_upload/58293/58293fig2large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 3" class="xfigimg" src="/files/ftp_upload/58293/58293fig3.jpg" /> 
Figure 3. A wide variety of specimens can be embedded with our protocol. (A) 7 dpf zebrafish larva. (B) Adult Drosophila. (C) Adult Daphnia. (D) Mouse embryo. Larger samples such as the mouse embryo are accommodated by a few modifications to the protocol. Briefly, the polyimide tubing was filled to 1/3 of its length with resin and polymerized prior to embedding. Fixed and stained sample was placed on top of the pre-polymerized resin. The tubing was then filled with un-polymerized resin followed by a second polymerization. The polyimide tubing was removed prior to photography to allow better visualization of the sample in this figure. The removal of the tubing is not necessary for successful image acquisition by micro-CT. Scale bars: specimen images, 5 mm; enlarged insets, 1 mm. <a href="https://www.jove.com/files/ftp_upload/58293/58293fig3large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 4" class="xfigimg" src="/files/ftp_upload/58293/58293fig4.jpg" /> 
Figure 4. 3-dimensional renderings of embedded specimens. (A) Reconstructed 3 dpf zebrafish larva at 0.743&#160;&#215; 0.743&#160;&#215; 0.743 &#181;m3 voxel resolution. (B) Reconstructed adult Daphnia (3&#160;&#215; 3&#160;&#215; 3 &#181;m3 voxel resolution). (C) Reconstructed adult Drosophila (3.1&#160;&#215; 3.1&#160;&#215; 3.1 &#181;m3 voxel resolution). Digital cross-sections can be generated at any angle as shown on the left column. Surface renderings are shown on the right, rendered using commercial software. Embedding resin can be seen in all scans outside of the sample (noted by *), but does not interfere with the sample itself. The zebrafish larva was imaged at Argonne National Laboratory at the Advanced Photon Source that is synchrotron-based. The Daphnia or Drosophila specimens were imaged with a commercial X-ray microscope. <a href="https://www.jove.com/files/ftp_upload/58293/58293fig4large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 5" class="xfigimg" src="/files/ftp_upload/58293/58293fig5.jpg" /> 
Figure 5. Samples embedded with our protocol can be re-imaged with micro-CT. The same 5 dpf zebrafish larva was imaged in (A) 2011 and (B) 2013, presented as average intensity projections of the sagittal view. Image comparison between scans after the storage shows the preservation of anatomical features. (C) A close-up of muscle from both scans is presented, with segmented lines indicating corresponding regions used for (D) intensity profile generation. Intensity profiles normalized to their corresponding averages along both lines show spatially matching local peaks in both scans. (E) Average intensity values for selected regions in A and B belonging to background (Bg, purple), neuronal nuclei (N, green), white matter (W, blue), and the muscle (M, orange) were divided by an average for background (white) and used to generate signal-to-noise ratios (SNR), which corresponds well between scans. Scale bars = 100 &#181;m. <a href="https://www.jove.com/files/ftp_upload/58293/58293fig5large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

Watch this Scientific Journal Video about Rigid Embedding of Fixed and Stained, Whole, Millimeter-Scale Specimens for Section-free 3D Histology by Micro-Computed Tomography at JoVE.com

Rigid Embedding of Fixed and Stained, Whole, Millimeter-Scale Specimens for Section-free 3D Histology by Micro-Computed Tomography

We developed protocols and designed a custom apparatus to enable embedding of millimeter-scale specimens. We present sample preparation procedures with an emphasis on embedding in acrylic resin and polyimide tubing to achieve rigid immobilization and long-term storage of specimens for the interrogation of tissue architecture and cell morphology by micro-CT.

For over a hundred years, the histological study of tissues has been the gold standard for medical diagnosis because histology allows all cell types in ...

rigid-embedding-fixed-stained-whole-millimeter-scale-specimens-for

Research

JoVE Journal

Bioengineering

8.8K Views.  Penn State College of Medicine. Deze methode stelt u in staat om driedimensionale histologische beelden te genereren uit kleine millimetermonsters, waaronder kleine modelorganismen. Het belangrijkste voordeel van deze techniek is dat de ingebedde monsters zijn zeer stabiel, zodat u de monsters te houden voor langere tijd en om ze beeld meerdere keren, zeggen met verschillende beeldvormende methoden over jaren. De output van deze methode zal zeer nuttig zijn voor hoge doorvoer model systeem fenomics, toxicologie, en potentie...

Video: Rigid Embedding of Fixed and Stained, Whole, Millimeter-Scale Specimens for Section-free 3D Histology by Micro-Computed Tomography

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 ...

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 ...

Developmental Biology

A Method for Obtaining Serial Ultrathin Sections of Microorganisms in Transmission Electron Microscopy

Observing cells and cell components in three dimensions at high magnification in transmission electron microscopy requires preparing serial ultrathin sections of the specimen. Although preparing serial ultrathin sections is considered to be very difficult, it is rather easy if the proper method is used. In this paper, we show a step-by-step procedure for safely obtaining serial ultrathin sections of microorganisms. The key points of this method are: 1) to use the large part of the specimen and adjust the specimen surface and knife edge so that they are parallel to each other; 2) to cut serial sections in groups and avoid difficulty in separating sections using a pair of hair strands when retrieving a group of serial sections onto the slit grids; 3) to use a 'Section-holding loop' and avoid mixing up the order of the section groups; 4) to use a 'Water-surface-raising loop' and make sure the sections are positioned on the apex of the water and that they touch the grid first, in order to place them in the desired position on the grids; 5) to use the support film on an aluminum rack and make it easier to recover the sections on the grids and to avoid wrinkling of the support film; and 6) to use a staining tube and avoid accidentally breaking the support films with tweezers. This new method enables obtaining serial ultrathin sections without difficulty. The method makes it possible to analyze cell structures of microorganisms at high resolution in 3D, which cannot be achieved by using the automatic tape-collecting ultramicrotome method and serial block-face or focused ion beam scanning electron microscopy.

This study presents reliable and easy procedures for obtaining serial ultrathin sections of a microorganism without expensive equipment in transmission electron microscopy.

Observing cells and cell components in three dimensions at high magnification in transmission electron microscopy requires preparing serial ultrathin ...

Engineering

Multimodal Hierarchical Imaging of Serial Sections for Finding Specific Cellular Targets within Large Volumes

Targeting specific cells at ultrastructural resolution within a mixed cell population or a tissue can be achieved by hierarchical imaging using a combination of light and electron microscopy. Samples embedded in resin are sectioned into arrays consisting of ribbons of hundreds of ultrathin sections and deposited on pieces of silicon wafer or conductively coated coverslips. Arrays are imaged at low resolution using a digital consumer like smartphone camera or light microscope (LM) for a rapid large area overview, or a wide field fluorescence microscope (fluorescence light microscopy (FLM)) after labeling with fluorophores. After post-staining with heavy metals, arrays are imaged in a scanning electron microscope (SEM). Selection of targets is possible from 3D reconstructions generated by FLM or from 3D reconstructions made from the SEM image stacks at intermediate resolution if no fluorescent markers are available. For ultrastructural analysis, selected targets are finally recorded in the SEM at high-resolution (a few nanometer image pixels). A ribbon-handling tool that can be retrofitted to any ultramicrotome is demonstrated. It helps with array production and substrate removal from the sectioning knife boat. A software platform that allows automated imaging of arrays in the SEM is discussed. Compared to other methods generating large volume EM data, such as serial block-face SEM (SBF-SEM) or focused ion beam SEM (FIB-SEM), this approach has two major advantages: (1) The resin-embedded sample is conserved, albeit in a sliced-up version. It can be stained in different ways and imaged with different resolutions. (2) As the sections can be post-stained, it is not necessary to use samples strongly block-stained with heavy metals to introduce contrast for SEM imaging or render the tissue blocks conductive. This makes the method applicable to a wide variety of materials and biological questions. Particularly prefixed materials e.g., from biopsy banks and pathology labs, can directly be embedded and reconstructed in 3D.

This protocol targets specific cells in tissue for imaging at nanoscale resolution using a scanning electron microscope (SEM). Large numbers of serial sections from resin-embedded biological material are first imaged in a light microscope to identify the target and then in a hierarchical manner in the SEM.

Targeting specific cells at ultrastructural resolution within a mixed cell population or a tissue can be achieved by hierarchical imaging using a ...

Preparing Lamellae from Vitreous Biological Samples Using a Dual-Beam Scanning Electron Microscope for Cryo-Electron Tomography

Presented here is a protocol for preparing cryo-lamellae from plunge-frozen grids of Plasmodium falciparum-infected&#160;human erythrocytes, which could easily be adapted for other biological samples. The basic principles for preparing samples, milling, and viewing lamellae are common to all instruments and the protocol can be followed as a general guide to on-grid cryo-lamella preparation for cryo-electron microscopy (cryoEM) and cryo-electron tomography (cryoET). Electron microscopy grids supporting the cells are plunge-frozen into liquid nitrogen-cooled liquid ethane using a manual or automated plunge freezer, then screened on a light microscope equipped with a cryo-stage. Frozen grids are transferred into a cryo-scanning electron microscope equipped with a focused ion beam (cryoFIB-SEM). Grids are routinely sputter coated prior to milling, which aids dispersal of charge build-up during milling. Alternatively, an e-beam rotary coater can be used to apply a layer of carbon-platinum to the grids, the exact thickness of which can be more precisely controlled. Once inside the cryoFIB-SEM an additional coating of an organoplatinum compound is applied to the surface of the grid via a gas injection system (GIS). This layer protects the front edge of the lamella as it is milled, the integrity of which is critical for achieving uniformly thin lamellae. Regions of interest are identified via SEM and milling is carried out in a step-wise fashion, reducing the current of the ion beam as the lamella reaches electron transparency, in order to avoid excessive heat generation. A grid with multiple lamellae is then transferred to a transmission electron microscope (TEM) under cryogenic conditions for tilt-series acquisition. A robust and contamination-free workflow for lamella preparation is an essential step for downstream techniques, including cellular cryoEM, cryoET, and sub-tomogram averaging. Development of these techniques, especially for lift-out and milling of high-pressure frozen samples, is of high-priority in the field.

Using focused ion beam milling to produce vitreous on-grid lamellae from plunge frozen biological samples for cryo-electron tomography.

Presented here is a protocol for preparing cryo-lamellae from plunge-frozen grids of Plasmodium falciparum-infected&#160;human erythrocytes, which could ...

Biochemistry

Utilizing 3D Printing Technology to Merge MRI with Histology: A Protocol for Brain Sectioning

Magnetic resonance imaging (MRI) allows for the delineation between normal and abnormal tissue on a macroscopic scale, sampling an entire tissue volume three-dimensionally. While MRI is an extremely sensitive tool for detecting tissue abnormalities, association of signal changes with an underlying pathological process is usually not straightforward. In the central nervous system, for example, inflammation, demyelination, axonal damage, gliosis, and neuronal death may all induce similar findings on MRI. As such, interpretation of MRI scans depends on the context, and radiological-histopathological correlation is therefore of the utmost importance. Unfortunately, traditional pathological sectioning of brain tissue is often imprecise and inconsistent, thus complicating the comparison between histology sections and MRI. This article presents novel methodology for accurately sectioning primate brain tissues and thus allowing precise matching between histology and MRI. The detailed protocol described in this article will assist investigators in applying this method, which relies on the creation of 3D printed brain slicers. Slightly modified, it can be easily implemented for brains of other species, including humans.

The overall goal of this protocol is to accurately align magnetic resonance imaging (MRI) image volumes with histology sections via the creation of customized 3D-printed brain holders and slicer boxes.

Magnetic resonance imaging (MRI) allows for the delineation between normal and abnormal tissue on a macroscopic scale, sampling an entire tissue volume ...

Neuroscience

3D Imaging of Soft-Tissue Samples using an X-ray Specific Staining Method and Nanoscopic Computed Tomography

We demonstrate a laboratory-based method combining X-ray microCT and nanoCT with a specific X-ray stain, which targets the cell cytoplasm. The described protocol is easy to apply, fast and suitable for larger soft-tissue samples. The presented methodology enables the characterization of crucial tissue structures in three dimensions and is demonstrated on a whole mouse kidney. The multiscale approach allows to image the entire mouse kidney and supports the selection of further volumes of interest, which are acquired with higher resolutions ranging into the nanometer range. Thereby, soft-tissue morphology with a similar detail level as the corresponding histological light microscopy images is reproduced. Deeper insights into the 3D configuration of tissue structures are achieved without impeding further investigations through histological methods.

A protocol for 3D visualization of microscopic tissue structures by using an X-ray specific staining method designed for X-ray computed tomography is presented.

We demonstrate a laboratory-based method combining X-ray microCT and nanoCT with a specific X-ray stain, which targets the cell cytoplasm. The described ...

Reconstruction of 3-Dimensional Histology Volume and its Application to Study Mouse Mammary Glands

Histology volume reconstruction facilitates the study of 3D shape and volume change of an organ at the level of macrostructures made up of cells. It can also be used to investigate and validate novel techniques and algorithms in volumetric medical imaging and therapies. Creating 3D high-resolution atlases of different organs1,2,3 is another application of histology volume reconstruction. This provides a resource for investigating tissue structures and the spatial relationship between various cellular features. We present an image registration approach for histology volume reconstruction, which uses a set of optical blockface images. The reconstructed histology volume represents a reliable shape of the processed specimen with no propagated post-processing registration error. The Hematoxylin and Eosin (H&#38;E) stained sections of two mouse mammary glands were registered to their corresponding blockface images using boundary points extracted from the edges of the specimen in histology and blockface images. The accuracy of the registration was visually evaluated. The alignment of the macrostructures of the mammary glands was also visually assessed at high resolution.
This study delineates the different steps of this image registration pipeline, ranging from excision of the mammary gland through to 3D histology volume reconstruction. While 2D histology images reveal the structural differences between pairs of sections, 3D histology volume provides the ability to visualize the differences in shape and volume of the mammary glands.

We present an image registration approach for 3-dimensional (3D) histology volume reconstruction, which facilitates the study of the changes of an organ at the level of macrostructures made up of cells . Using this approach, we studied the 3D changes between wild-type and Igfbp7-null mammary glands.

Histology volume reconstruction facilitates the study of 3D shape and volume change of an organ at the level of macrostructures made up of cells. It can ...

Biology

Whole&#45;Organ Pathogen Tracking via Fluorescent Reporter Quantification

High&#45;Resolution Three&#45;Dimensional Whole&#45;Organ Tomography of Microbial Infections

Most infections take place within three-dimensional host tissues with intricate anatomy and locally varying host physiology. The positioning of pathogen cells within this diverse environment significantly affects their stress levels, responses, fate, and contribution to the overall progression of the disease and treatment failure. However, due to the technical difficulties in locating &#181;m-sized pathogen cells within cm-sized host organs, this area of research has been relatively unexplored. Here, we present a method for addressing this challenge. We employ serial two-photon tomography and AI-enhanced image analysis to locate individual Salmonella cells throughout the entire spleen, liver lobes, and whole lymph nodes of infected mice. Using fluorescent reporters and in vivo antibody administration, the replication rate of single Salmonella cells, their local interaction with specific immune cells, and bacterial responses to antibiotics can be determined. These methodologies open avenues for a comprehensive examination of infections, their prevention, and treatment within the three-dimensional tissue context.

Author Spotlight: Unraveling Bacterial Responses to Antibiotics and Immune System in Tissues

Here, we describe a procedure that enables organ-wide detection of pathogenic bacteria during infection and quantification of fluorescent reporter activities.

Most infections take place within three-dimensional host tissues with intricate anatomy and locally varying host physiology. The positioning of pathogen ...

Immunology and Infection

Staining and High-Resolution Imaging of Three-Dimensional Organoid and Spheroid Models

In vitro three-dimensional (3D) cell culture models, such as organoids and spheroids, are valuable tools for many applications including development and disease modeling, drug discovery, and regenerative medicine. To fully exploit these models, it is crucial to study them at cellular and subcellular levels. However, characterizing such in vitro 3D cell culture models can be technically challenging and requires specific expertise to perform effective analyses. Here, this paper provides detailed, robust, and complementary protocols to perform staining and subcellular resolution imaging of fixed in vitro 3D cell culture models ranging from 100 &#181;m to several millimeters. These protocols are applicable to a wide variety of organoids and spheroids that differ in their cell-of-origin, morphology, and culture conditions. From 3D structure harvesting to image analysis, these protocols can be completed within 4-5 days. Briefly, 3D structures are collected, fixed, and can then be processed either through paraffin-embedding and histological/immunohistochemical staining, or directly immunolabeled and prepared for optical clearing and 3D reconstruction (200 &#181;m depth) by confocal microscopy.

Here, we provide detailed, robust, and complementary protocols to perform staining and subcellular resolution imaging of fixed three-dimensional cell culture models ranging from 100 &#181;m to several millimeters, thus enabling the visualization of their morphology, cell-type composition, and interactions.

In vitro three-dimensional (3D) cell culture models, such as organoids and spheroids, are valuable tools for many applications including development and ...

Rigid Embedding of Fixed and Stained, Whole, Millimeter-Scale Specimens for Section-free 3D Histology by Micro-Computed Tomography

Samenvatting

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