The authors thank Tim Mohun for his invalubale contributions in developing HREM and Petra Heffeter for providing samples.

All procedures were performed in accordance with ethical guidelines at the Medical University of Vienna.
1. Sample Preparation
<ol>
	<li>Preparing embryos and embryo tissues (up to 5 x 5 x 5 mm3 )
	<ol>
		<li>Fix the embryos or parts of the embryos in 4% PFA/PBS or Bouin's fixative at 4 &#176;C for at least 16 - 24 h under constant rocking. 
		NOTE: Embryos of several species and of various developmental stages can be used, such as 48-h zebrafish, 19-h chick and quail, and also prenatal mouse embryos, which were used here (see Representative Results). The specimens were harvested and transferred into PBS at room temperature for staging, before they were put into fixative. If necessary, trim the specimens with micro-scissors or scalpels prior to fixation to fit into the embedding molds.</li>
		<li>Remove the fixative and wash the embryo tissue in PBS at 4 &#176;C under constant rocking for 24 h (2-3 changes).</li>
		<li>Dehydrate the samples in 70%, 80%, 90%, 96% ethanol at 4 &#176;C for 2 - 3 h each. Place tubes containing the samples on a rotator. 
		NOTE: For small samples (&#60; 2 x 2 x 2 mm3), the time a sample rests in each ethanol may be reduced to 60 min. For large samples (&#62; 5 x 5 x 5 mm3), it may be extended to 4 h.</li>
		<li>Perform the following infiltration steps under a fume hood wearing protective gloves.
		<ol>
			<li>Prepare the infiltration solution by adding 1.25 g benzoyl peroxide (plasticized catalyst) and 0.4 g eosin to 100 mL of solution A (for more details, see Table of Materials). Stir this in a beaker on a magnetic stirring plate at 4 &#176;C until eosin and catalyst are fully dissolved (2 - 3 h).</li>
			<li>Place the samples for 12 - 24 h in the infiltration solution. Keep it at 4 &#176;C under continuous rocking or rotation. Replace infiltration solution with fresh solution after 3 and 12 h. 
			NOTE: Extend infiltration time to 48 h for large embryos (&#62; 1 cm crown/rump length).</li>
		</ol>
		</li>
	</ol>
	</li>
	<li>Preparing adult tissues samples (up to 5 x 5 x 7 mm3 )
	<ol>
		<li>Fix the tissues in 2% PFA/PBS containing 4% carbolic acid at 4 &#176;C for at least 3 days. 
		NOTE: Tissue sample stemming from skin, liver, pancreas, kidney, thyroid, heart, striated muscle, brain, nerves and tumor models of several neonatal and adult humans, rodents, pigs, fruit flies, and zebra fishes can be fixed in this solution and then processed for HREM imaging (we demonstrate the fixing of the samples; see Representative Results). Excise the tissue samples with scalpels, scissors or biopsy punches and directly transfer them into fixative. For fixation of nervous tissue use 4% PFA/PBS.</li>
		<li>Remove the fixative and wash the tissue under running tap water for 3-6 h.</li>
		<li>Dehydrate the samples by subsequently placing them in 70%, 80%, 90%, 96% ethanol, mixed with 0.4 g eosin per 100 mL (2 - 3 h each). Keep the solutions with samples at 4 &#176;C under constant rocking or on a rotator. 
		NOTE: For small samples (&#60; 2 x 2 x 2 mm3), the time a sample rests in each ethanol may be reduced to 60 min. For large samples (&#62; 5 x 5 x 5 mm3), it may be extended to 4-8 h.</li>
		<li>Perform all the following infiltration steps under a fume hood wearing protective gloves.
		<ol>
			<li>Prepare the infiltration solution by adding 1.25 g benzoyl peroxide (plasticized catalyst) and 0.4 g eosin to 100 mL of Solution A (for more details, see Table of Materials). Stir in a beaker on a magnetic stirring plate at 4 &#176;C until the eosin and catalyst are fully dissolved (2 - 3 h).</li>
			<li>Place samples in infiltration solution for 24 - 36 h. Keep it at 4 &#176;C under continuous rocking or rotation. Replace infiltration solution with fresh solution after 3 and 12 h. 
			NOTE: For large samples (&#62; 5 x 5 x 5 mm3) extend infiltration time to 48 h.</li>
		</ol>
		</li>
	</ol>
	</li>
	<li>Preparing other organic material 
	NOTE: Trim skin substitutes and paper to the desired size with ordinary scissors. No fixation is needed.
	<ol>
		<li>Skip fixation, washing and dehydration steps.</li>
		<li>Perform all the following infiltration steps under a fume hood wearing protective gloves.
		<ol>
			<li>Prepare the infiltration solution by adding 1.25 g benzoyl peroxide (plasticized catalyst) and 0.4 g eosin to 100 mL of Solution A (for more details, see Table of Materials). Stir in a beaker at 4 &#176;C on a magnetic stirring plate until eosin and catalyst are fully dissolved (2 - 3 h).</li>
			<li>Place samples in infiltration solution for 3 - 12 h. Keep it at 4 &#176;C under continuous rocking or rotation. Replace infiltration solution with fresh solution after 1 and 2 - 4 h.</li>
		</ol>
		</li>
	</ol>
	</li>
</ol>
2. Embedding
NOTE: Perform all steps under a fume hood wearing protective gloves.
<ol>
	<li>Prepare fresh infiltration solution as described above (step 1.1.4.1). Add 4 mL Solution B (for details, see Table of Materials) to 100 mL infiltration solution to prepare the embedding solution. 
	NOTE: If precise orientation of the specimen in the embedding molds is required and to allow the embedding of several samples in parallel, slow the polymerization of the stock solution by putting the beaker into an ice-bath.</li>
	<li>Use molding cup trays (for details, see Table of Materials) with molds having a routine volume of 6 x 8 x 5 mm3 or 13 x 19 x 5 mm3, or customized molds with volumes of up to 8 x 10 x 15 mm3 (Figure 2).</li>
	<li>Fill the molds with embedding solution and transfer the sample into the mold using a spoon. During transfer ensure that the sample is fully covered with embedding solution to avoid air trapping. 
	NOTE: This can be done at room temperature, if started immediately after collecting the samples and the embedding solution from the refrigerator.
	<ol>
		<li>Orient the sample inside the mold using needles or forceps.</li>
		<li>As soon as embedding solution starts to become viscous, place a block holder (see Table of Materials) on top of the mold and add embedding solution through the central hole of the block holder until the base of the block holder is covered with 1 - 2 mm of embedding solution.</li>
		<li>Seal the molding cup tray by covering it with liquid paraffin wax and wait until it has hardened. Alternatively place a second molding cup tray upside-down on the molding cup tray with samples and seal it with adhesive tape to air-proof.</li>
	</ol>
	</li>
	<li>Allow blocks to finish polymerization by storing the sealed molding cup trays for 1 - 2 days at room temperature.</li>
	<li>For post-polymerization processing, place the molding cup trays with the polymerized blocks in a standard laboratory oven and bake at 70 - 80 &#176;C for a minimum of 1 - 2 days until they become dark red. Collect the trays from the oven and remove the blocks from the mold. 
	NOTE: Baking can be done under normal environmental conditions. The covering molding cup tray, which ensures airproof sealing in the first phase after embedding, can be removed to allow for checking the color of the resin after 10 h. Immediately after baking, the resin is soft and the blocks can be easily removed from the molding cup tray. Within 2 - 3 h they are hard and can be stored at room temperature for several months.</li>
</ol>
3. Data Generation
NOTE: The HREM prototype18,25 used here comprises the following items: (i) Rotary microtome with a block holder that stops after each cut at its upper turning point. (ii) Standard, non-disposable microtome knife, hard metal, profile D (for details, see Table of Materials). (iii) Head of fluorescence compound microscope with objective revolver and a GFP-filter cube (excitation 470/40; emission filter 525/50) in its optical axis. The optical axis is arranged perpendicular to the freshly cut surface of a block mounted on the microtome and is held by a device that allows up- and downward movement. (iv) Motorized cross table carrying the microtome. The table can be shifted in the direction of the optical axis and laterally. (v) Digital video camera attached to the fluorescent compound microscope. (vi) Monochrome light source (470 nm). (vii) Computer connected to the camera, with data generation software (for details, see Table of Materials).
<ol>
	<li>Procedure
	<ol>
		<li>Indicate the region of interest.
		<ol>
			<li>Use standard laboratory light source (see Table of Materials) for directing white light obliquely to the block surface.</li>
			<li>Identify the outlines of the embedded material as it projects to the block surface. Indicate the region of interest and later field of view on the block surface using a pen or razor blade. 
			NOTE: This step is mandatory to define the extent of the field of view prior to sectioning.</li>
		</ol>
		</li>
		<li>Define the field of view.
		<ol>
			<li>Mount the resin block with sample on the microtome.</li>
			<li>Move the block holder to its stopping position (upper turning point of block holder excursion).</li>
			<li>Start the digital camera and data generation software, and acquire a live image.</li>
			<li>Choose an objective lens, with an appropriate magnification to cover the region of interest indicated on the block surface. 
			NOTE: Objectives with magnifications 2.5X, 5X, 10X, 20X and numeric apertures between 0.07 and 0.40 are commonly used.</li>
			<li>Move the optic up- and downward and the microtome laterally using the motorized cross table until the region of interest matches with the field of view displayed on the computer screen.</li>
		</ol>
		</li>
		<li>Focus.
		<ol>
			<li>Perform step by step manual sectioning until first structures of the sample become visible.</li>
			<li>Arrange the block surface in the focus plane of the optic, by moving the microtome in the direction of the optical axis.</li>
			<li>Choose section thickness; section thicknesses from 0.5 to 5 &#181;m are recommended.</li>
		</ol>
		</li>
	</ol>
	</li>
	<li>Data generation and processing
	<ol>
		<li>Start software routine that orchestrates sectioning and image capturing. Follow the documented instruction of the imaging capturing software.</li>
		<li>Stop software routine and put a scaling slide in front of the block surface as soon as the sample is completely sectioned.</li>
		<li>Capture an image interactively for later calibration. 
		NOTE: This must be done every time an objective is used in a newly arranged set up.</li>
		<li>Convert the image series to 8 bit gray-scale images in.jpg format.</li>
		<li>Load image series into 3D visualization software and adjust contrasts. 
		NOTE: Image conversion and 3D visualization can be done with various commercially or freely available standard software packages, following the instructions of the respective software.</li>
	</ol>
	</li>
</ol>

The authors have nothing to disclose.

HREM is a highly robust microscopic method that is ideal for visualizing a broad spectrum of organic materials used in biomedicine and industry18,21,26,27,28,29,30,31,32,33,34,35,36,37,38,39,40. It can be employed as an exclusive imaging modality, as currently used by the Mechanisms of Developmental Disorders (DMDD) program41,42,43,44, or as an integrative part of multimodal imaging pipelines45.
A fully functioning HREM data generation apparatus can be assembled from conventional laboratory components and comprises a motorized microtome, microscope, motorized cross table, and a computer with appropriate software25. It is critical to use a microtome equipped with a block holder that reproducibly stops after each section at a defined position and GFP filter cubes inside the optical pathway. However, fully functioning all-inclusive solutions can be purchased from companies such as Indigo Scientific.
HREM faces the same limitations as all histological techniques, except that no artifacts are introduced during sectioning or section mounting. However, there are limitations, which result from the necessity to stain specimens prior to sectioning and from the characteristics of the embedding material. Penetration of eosin through the entire specimen is required for obtaining sufficient tissue contrasts; very dense material, adipose tissues and anorganic substances effectively hinder eosin penetration and this results in unstained tissues in the center of the objects. Using special fixatives helps stain skin specimens, but there is still no proper method for fully overcoming the problem. Another limitation is that resins that block higher than 2 cm tend to break during sectioning. This can be partly avoided by cutting the specimens and processing parts separately.
Correct positioning of small samples or samples with irregular surfaces in the molds during embedding is often problematic. Covering the samples with agarose and processing the agarose blocks as described in the protocol usually resolves this issue19. An alternative approach, which also helps if blocks break during sectioning, is to remove the already hardened block from its holder and embed it anew, following the described embedding procedure.
A typical HREM data set comprises 500 to 3,000 single images. Its numerical resolution is determined by the distance between the successive images (i.e., by section thickness), the characteristic of the camera target, and the properties of the utilized optics. We used section thicknesses between 1 &#181;m and 5 &#181;m and achieved good results, although the presented protocols do not entirely eliminate shining from artifacts20,46. These artifacts are caused by intensely stained tissues located deep inside the block, resulting in blurring of tissue information on the block surfaces by.
The cameras had target dimensions of 2,560 x 1,920 pixels2, 2,048 x 2,048 pixels2, and 4,096 x 4,096 pixels2 and were combined with 1.25X, 2.5X, 5X, 10X, and 20X objective lenses. This resulted in numeric pixel sizes between 0.18 x 0.18 &#181;m2 and 5.92 x 5.92 &#181;m2, which proved to be sufficient for 3D analysis of tissue architecture and cell shapes, and even for visualizing nuclei. Given the high numeric resolution, other cell organelles should be visible as well. Insufficient contrasts due to simple eosin staining, and the optical properties of the objectives dramatically lower the possibility to discriminate structures. The maximal true spatial resolution of the HREM data, which takes into account the numeric aperture, is approximately 1 x 1 x 1 &#181;m3, and therefore only permits effective discrimination of structures larger than approximately 3 x 3 x 3 &#181;m3.
A common problem to all digital imaging techniques is the tradeoff between the size of the field of view, which defines the part of the specimen that can be displayed on the camera target, and the numeric resolution of the image. The larger the field of view, the lower the maximal possible numeric resolution46. The HREM setup used here permits the generation of HREM data with a field of view between 0.74 x 0.74 mm2 (20X objective) displayed in a numeric resolution of 0.18 x 0.18 &#181;m2 and 12.12 x 12.12 mm2 (1.25X objective) displayed in a numeric resolution of 2.96 x 2.96 &#181;m2. Alternative, commercialized set-ups can provide larger fields of views, but at the cost of true resolution. Nevertheless, they provide excellent results, as obvious from the data displayed on the homepage of the DMDD program47.

Structural analysis of organic and anorganic materials is the first step in understanding their physical properties and function. The basis for such analysis often is two-dimensional (2D) information gained by careful observation of histological sections, with a variety of simple and sophisticated imaging methods that extract details of tissue architecture, cell morphology and topology, molecular composition and biomechanical properties1,2,3. However, 2D information is not suited for researching spatially complex arrangements. Hence a growing number of in vivo and ex vivo methods that permit the generation of digital volume data were established in the last decades4 and many more are under development.
The methodical principle of most volume data generation methods is the generation of virtual stacks of digital images displaying sections gained by virtual or physical sectioning of an object. If the section images are aligned properly, this creates a volume, which can be re-sectioned in virtual section planes, or used for creating 3D surface and volume rendered models. Popular techniques for visualizing humans and larger biological specimens are magnetic resonance tomography (MRT), computed tomography (CT), positron emission tomography (PET) and single-photon emission computed tomography (SPECT). Small specimens are usually visualized by using micro-magnetic resonance imaging (&#181;MRI), optical projection tomography (OPT), optical coherence tomography (OCT), photoacoustic tomography (PAT), histological sectioning based methods, confocal microscopy, and electron tomography5,6,7,8,9,10,11,12,13,14,15,16,17.
A relatively new volume data generation technique, which produces digital data of small specimens and histological tissue samples is the HREM method, which was developed in close co-operation with Tim Mohun18,19. It is a simple microscope based technique, which generates digital volume data from resin embedded material that is sectioned on a microtome. The data facilitate detailed analysis of tissue architecture and cell distributions as well as metric analysis of small features on an intermediate light microscopic level.
HREM produces stacks of inherently aligned digital images that appear as if captured from eosin stained histological sections. Tissue contrast and data resolution in respect to the field of view exceed that of data produced with &#181;CT, &#181;MRI, and OPT, but are lower than that achievable with confocal, light sheet and electron microscopy20. However, in contrast to the latter, HREM is capable of visualizing specimens with relative large volumes of up to 5 x 5 x 7 mm3 in histological quality. A number of recent studies provide detailed characterizations and comparisons of advantages and disadvantages of the single imaging techniques and, for the sake of objectivity, we refer to those for further information regarding their limitations and potential fields of applications4,21,22,23,24.
This study focuses on the HREM imaging method and aims to provide very simple protocols for generating HREM data of a broad spectrum of organic materials, as well as examples of their application. The workflow for creating HREM data is simple and applies to all materials that can embed in methacrylate resin (Figure 1). Yet there are tissue specific differences in sample preparation, that need to be considered. We therefore provide three standard protocols for preparing various samples. Embedding and data generation protocol steps are identical for all of them.

<table><tbody><tr><td>JB-4 Plus Embedding Kit</td><td>Polysciences Europe GmbH</td><td>18570-1</td><td>includes Benzoyl Peroxide, Plasticized (Catalyst) and Solution A+B</td></tr><tr><td>Polyethylene Molding Cup Trays, 6 x 8 x 5 mm hexagon (9 cavities)</td><td>Polysciences Europe GmbH</td><td>17177A-3</td><td></td></tr><tr><td>Polyethylene Molding Cup Trays, 13 x 19 x 5 mm (9 cavities)</td><td>Polysciences Europe GmbH</td><td>17177C-3</td><td></td></tr><tr><td>JB-4 Plastic Block Holders</td><td>Polysciences Europe GmbH</td><td>15899-50</td><td></td></tr><tr><td>Eosin</td><td>Waldeck GmbH &amp;amp; Co. KG, Division Chroma</td><td>1A-196</td><td></td></tr><tr><td>Microtec CUT 4060E</td><td></td><td></td><td>rotary microtome</td></tr><tr><td>Leica DM LM, fluorescence compound microscope</td><td>Leica Mikrosysteme Handelges.m.b.H</td><td></td><td></td></tr><tr><td>GFP filter set</td><td>Leica Mikrosysteme Handelges.m.b.H</td><td>11090937180000</td><td></td></tr><tr><td>Motorised cross table</td><td>Walter Uhl, technische Mikroskopie GmbH &amp;amp; CO. KG</td><td>KT5-LSMA</td><td></td></tr><tr><td>Digital video camera SPOT-FLEX</td><td>Visitron Systems GmbH.</td><td></td><td></td></tr><tr><td>precisExcite High-Power LED</td><td>Visitron Systems GmbH.</td><td></td><td>light source</td></tr><tr><td>VisiView 2.1.4</td><td>Visitron Systems GmbH.</td><td></td><td>Image capturing software</td></tr><tr><td>Hard metal knife (tungsten carbide), profile D</td><td>Leica Mikrosysteme Handelges.m.b.H</td><td></td><td></td></tr><tr><td>KL 2500 LCD</td><td>Schott AG</td><td></td><td>light source</td></tr></tbody></table>

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.

HREM generates series of inherently aligned digital images with contrasts similar to images of hematoxylin/eosin stained histological sections. As opposed to 2D section images, stacks of HREM images allow for visualization and analysis of the tissue architecture, morphology, and topology of a broad variety of organic materials in 3D. The high contrast often facilitates quick and simple 3D visualization of volume rendered computer models and semiautomatic contour findings for the generation of surface rendered models.
The size of HREM data varies according to the size of the camera target, the mode of image capturing (8, 12, 16 bit, gray scale, color) and the number of single block face images. Smaller data sets of 1,000 8-bit.jpg gray scale images of 2,048 pixels x 2,048 pixels have a size of about 900 MB. Larger data sets of 3,000 8-bit.jpg gray scale images of 4,096 pixel x 4,096 pixel have volumes of approximately 20 GB.
The protocols provided are simple and robust, and during the last decade, they were employed for generating HREM data of many different specimens. Protocol section 1.1 was optimized for processing whole embryos of biomedical model organisms with a length of up to 1 cm and embryo tissue samples with a dimension of up to 5 x 5 x 5 mm3; we used this method for producing HREM data of embryos of the following species: mouse (Mus musculus), chick (Gallus gallus), zebra fish (Danio rerio), quail (Coturnix coturnix), African clawed frog (Xenopus laevis), horse (Equus ferus caballus), milkweed bug (Oncopeltus fasciatus), crocodile (Crocodylia), and octopus (Octopus vulgaris). The data of all species were of excellent quality (e.g., Figure 3)
Protocol section 1.2 was optimized for processing juvenile and adult tissue samples with dimensions up to 5 x 5 x 7 mm3 and employed for visualizing tissue samples of the skin, liver, pancreas, kidney, thyroid, heart, striated muscle, brain, nerves and tumor models harvested from humans (Homo sapiens), mice (Mus musculus), rats (Rattus norvegicus), pigs (Sus scrofa domestica) ferret (Mustela furo), fruit fly (Drosophila melanogaster), and zebra fish (Danio rerio). While the results were excellent with most specimens (e.g., Figure 4, Animation 1), the very central parts of skin (with epidermis) and brain specimens larger than 3 x 3 x 3 mm3 often remained unstained due to insufficient penetration of eosin through these tissues.
Protocol section 1.3 was optimized for processing fibrous organic material and employed for visualizing the fiber architecture of coated paper, uncoated paper, native dermal substitute material and stem cell seeded dermal substitute material. These samples were easy and quick to process. The data of the uncoated paper and most skin substitutes were of good quality (Figure 5, Animation 2). Problems occurred in processing the coated paper, when the anorganic material hindered penetration of eosin. Another issue occurred when processing the skin substitutes on the basis of agar because they became partly digested by the infiltration solution.
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/56071/56071fig1.jpg" /> 
Figure 1: Workflow. Steps shown in the red boxes require modifications according to sample characteristics. Steps in the green boxes are those that are similar in all samples. <a href="http://cloudflare.jove.com/files/ftp_upload/56071/56071fig1large.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/56071/56071fig2.jpg" /> 
Figure 2: Customized Molds. Molds can be adapted by cutting a hole into the original mold, inserting the bulb of a Pasteur pipet and sealing it with modeling material. <a href="http://cloudflare.jove.com/files/ftp_upload/56071/56071fig2large.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/56071/56071fig3.jpg" /> 
Figure 3: Exemplarily Visualized Embryos. (A, B) Mouse embryo harvested at embryonic day, E=9.5. HREM section image is shown in (A). Volume rendered 3D model showing the surface is shown in (B). (C) Virtual sagittal section through a volume rendered model of the neck of an E15.5 mouse embryo. (D) Chick embryo at developmental Hamburger Hamilton (HH) stage 18. Surface model of the lumina of the cardiovascular components combined with volume rendering of all embryo tissues. Scale bars = 200 &#181;m. <a href="http://cloudflare.jove.com/files/ftp_upload/56071/56071fig3large.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/56071/56071fig4.jpg" /> 
Figure 4: Exemplarily Visualized Adult Tissue Samples. (A) Part of an HREM section image through a human nerve. Inlay shows a section of the image in more detail. (B) Part of an HREM section image through porcine liver. Note the nuclei. (C) Part of an HREM section image of human lymphatic tissue. (D, E) Thick skin of a human thumb pad. Volume rendered 3D model of the entire biopsy. (D) Surface rendered models of arteries, veins, and nerves in front of a virtual resection through HREM data (E). (F) Experimental tumor in adult mouse tissue. Volume rendered 3D model in front of three virtual sections through HREM data. Note the necrotic parts (arrowheads). Scale bars = 200 &#181;m. <a href="http://cloudflare.jove.com/files/ftp_upload/56071/56071fig4large.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/56071/56071fig5.jpg" /> 
Figure 5: Exemplarily Visualized Fibrous Material. (A, B) HREM section image of uncoated, brown paper. (B) shows a section of (A). Note the fibers and their lumen. (C) HREM section of coated paper. Note that the fibers remain unstained. (D) Volume rendered model of native dermal substitute material. Note the different caliber and form of the fibers. Scale bars = 100 &#181;m. <a href="http://cloudflare.jove.com/files/ftp_upload/56071/56071fig5large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Movie 1" src="/files/ftp_upload/56071/56071mov1.jpg" /> 
Animation 1: Volume Rendered Model of Thick Skin of a Human Thumb Pad. The size of the tissue block is approximately 4.2 x 2.7 x 2.7 mm3. Voxel size is 1.07 x 1.07 x 2 &#181;m3. <a href="http://cloudflare.jove.com/files/ftp_upload/56071/animation1.mp4" target="_blank">Please click here to view this video. (Right-click to download.)</a>
<img alt="Movie 2" src="/files/ftp_upload/56071/56071mov2.jpg" /> 
Animation 2: Volume Rendered Model of Native Dermal Substitute Material. The size of the sample is approximately 1.2 x 0.85 x 0.4 mm3. Voxel size is 0.54 x 0.54 x 2 &#181;m3. <a href="http://cloudflare.jove.com/files/ftp_upload/56071/animation2.mp4" target="_blank">Please click here to view this video. (Right-click to download.)</a>

Watch this Scientific Journal Video about High-resolution Episcopic Microscopy HREM - Simple and Robust Protocols for Processing and Visualizing Organic Materials at JoVE.com

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.

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 ...

high-resolution-episcopic-microscopy-hrem-simple-robust-protocols-for

Research

JoVE Journal

Developmental Biology

10.2K Views.  Medical University of Vienna.  Wij bieden eenvoudige en robuuste protocollen voor het verwerken van biopsiemateriaal van verschillende soorten, embryo's van biomedische modelorganismen en monsters van andere organische weefsels om de digitale volumegegevensopwekking mogelijk te maken met de hoge resolutie episcopische microscopie methode. 

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

High-resolution Fiber-optic Microendoscopy for in situ Cellular Imaging

Many biological and clinical studies require the longitudinal study and analysis of morphology and function with cellular level resolution. Traditionally, multiple experiments are run in parallel, with individual samples removed from the study at sequential time points for evaluation by light microscopy. Several intravital techniques have been developed, with confocal, multiphoton, and second harmonic microscopy all demonstrating their ability to be used for imaging in situ 1. With these systems, however, the required infrastructure is complex and expensive, involving scanning laser systems and complex light sources. Here we present a protocol for the design and assembly of a high-resolution microendoscope which can be built in a day using off-the-shelf components for under US$5,000. The platform offers flexibility in terms of image resolution, field-of-view, and operating wavelength, and we describe how these parameters can be easily modified to meet the specific needs of the end user.
We and others have explored the use of the high-resolution microendoscope (HRME) in in vitro cell culture 2-5, in excised 6 and living animal tissues 2,5, and in human tissues in vivo 2,7. Users have reported the use of several different fluorescent contrast agents, including proflavine 2-4, benzoporphyrin-derivative monoacid ring A (BPD-MA) 5, and fluoroscein 6,7, all of which have received full, or investigational approval from the FDA for use in human subjects. High-resolution microendoscopy, in the form described here, may appeal to a wide range of researchers working in the basic and clinical sciences. The technique offers an effective and economical approach which complements traditional benchtop microscopy, by enabling the user to perform high-resolution, longitudinal imaging in situ.

High-resolution Fiber-optic Microendoscopy for in situ Cellular Imaging

In many biological and clinical situations it is advantageous to study cellular processes as they evolve in their native microenvironment. Here we describe the assembly and use of a low-cost fiber-optic microscope which can provide real time imaging in cell culture, animal studies, and clinical patient studies.

Many biological and clinical studies require the longitudinal study and analysis of morphology and function with cellular level resolution.  ...

Bioengineering

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 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) ...

Processing Embryo, Eggshell, and Fungal Culture for Scanning Electron Microscopy

Although scanning electron microscopy (SEM) is being widely used for the ultra-structural analysis of various biological and non-biological samples, methods involved in processing different biological samples involve unique practices. All conventional practices described in the literature for processing samples still find useful applications, but subtle changes in the sample preparation can alter image quality, as well as, introduce artifacts. Hence, using a unique sample preparation technique specific to the type of tissue analyzed is required to obtain a good quality image with ultrastructural resolution. The focus of this study is to provide the optimal sample preparation protocols for imaging embryos, rigid eggshells, and fungal cultures using SEM. The following optimizations were recommended to yield good results for the three different delicate biological samples studied. Use of milder fixatives like 4% paraformaldehyde or 3% glutaraldehyde followed by dehydration with ethanol series is mandatory. Fungal mycelium on agar blocks obtained by slide cultures yields a better ultrastructural integrity compared to cultures taken directly from agar plates. Chemical drying of embryos with HMDS provides drying without introducing surface tension artifacts compared to critical point drying. HMDS prevents cracking caused by shrinkage as samples are less brittle during drying. However, for fungal culture, critical point drying provides acceptable image quality compared to chemical drying. Eggshells can be imaged with no special preparation steps except for thorough washing and air drying prior to mounting. Preparation methodologies were standardized based on acceptable image quality obtained with each trial.&#160;

Here, we present detailed processing protocols for imaging delicate tissue samples using scanning electron microscopy (SEM). Three different processing methods, namely, hexamethyl disilazana (HMDS) chemical drying, simple air drying, and critical point drying are described for preparing rigid eggshells, embryos at early developmental stages, and fungal cultures respectively.

Although scanning electron microscopy (SEM) is being widely used for the ultra-structural analysis of various biological and non-biological samples, ...

Environment

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 ...

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

Visualization of Endoplasmic Reticulum Subdomains in Cultured Cells

The lipids and proteins in eukaryotic cells are continuously exchanged between cell compartments, although these retain their distinctive composition and functions despite the intense interorganelle molecular traffic. The techniques described in this paper are powerful means of studying protein and lipid mobility and trafficking in vivo and in their physiological environment. Fluorescence recovery after photobleaching (FRAP) and fluorescence loss in photobleaching (FLIP) are widely used live-cell imaging techniques for studying intracellular trafficking through the exo-endocytic pathway, the continuity between organelles or subcompartments, the formation of protein complexes, and protein localization in lipid microdomains, all of which can be observed under physiological and pathological conditions. The limitations of these approaches are mainly due to the use of fluorescent fusion proteins, and their potential drawbacks include artifactual over-expression in cells and the possibility of differences in the folding and localization of tagged and native proteins. Finally, as the limit of resolution of optical microscopy (about 200 nm) does not allow investigation of the fine structure of the ER or the specific subcompartments that can originate in cells under stress (i.e. hypoxia, drug administration, the over-expression of transmembrane ER resident proteins) or under pathological conditions, we combine live-cell imaging of cultured transfected cells with ultrastructural analyses based on transmission electron microscopy.

We describe the imaging approaches we use to investigate the distribution and mobility of the transfected fluorescent proteins resident in the endoplasmic reticulum (ER) by means of the confocal imaging of living cells. We also ultrastructurally analyze the effect of their expression on the architecture of this subcellular compartment.

The lipids and proteins in eukaryotic cells are continuously exchanged between cell compartments, although these retain their distinctive composition and ...

Visualizing Retinal Synaptic Structures in 3D with Volume Electron Microscopy

Preparation of Retinal Samples for Volume Electron Microscopy

Volume electron microscopy (Volume EM) has emerged as a powerful tool for visualizing the 3D structure of cells and tissues with nanometer-level precision. Within the retina, various types of neurons establish synaptic connections in the inner and outer plexiform layers. While conventional EM techniques have yielded valuable insights into retinal subcellular organelles, their limitation lies in providing 2D image data, which can hinder accurate measurements. For instance, quantifying the size of three distinct synaptic vesicle pools, crucial for synaptic transmission, is challenging in 2D. Volume EM offers a solution by providing large-scale, high-resolution 3D data. It is worth noting that sample preparation is a critical step in Volume EM, significantly impacting image clarity and contrast. In this context, we outline a sample preparation protocol for the 3D reconstruction of photoreceptor axon terminals in the retina. This protocol includes three key steps: retina dissection and fixation, sample embedding processes, and selection of the area of interest.

Author Spotlight: Retinal Neuroscience Studies with Volume Electron Microscopy

This protocol describes the detailed steps for preparing retinal samples for volume electron microscopy, focusing on the structural features of retinal photoreceptor terminals.

Volume electron microscopy (Volume EM) has emerged as a powerful tool for visualizing the 3D structure of cells and tissues with nanometer-level ...

Neuroscience

Optimized Protocol for Retinal Organoid Sample Preparation for TEM

Preparing Retinal Organoid Samples for Transmission Electron Microscopy

Retinal organoids (ROs) are a three-dimensional culture system mimicking human retinal features that have differentiated from induced pluripotent stem cells (iPSCs) under specific conditions. Synapse development and maturation in ROs have been studied immunocytochemically and functionally. However, the direct evidence of the synaptic contact ultrastructure is limited, containing both special ribbon synapses and conventional chemical synapses. Transmission electron microscopy (TEM) is characterized by high resolution and a respectable history elucidating retinal development and synapse maturation in humans and various species. It is a powerful tool to explore synaptic structure in ROs and is widely used in the research field of ROs. Therefore, to better explore the structure of RO synaptic contacts at the nanoscale and obtain high-quality microscopic evidence, we developed a simple and repeatable method of RO TEM sample preparation. This paper describes the protocol, reagents used, and detailed steps, including RO fixation preparation, post fixation, embedding, and visualization.

Author Spotlight: Analyzing the Synaptic Ultrastructure in Mature Retinal Organoids Using TEM

This protocol provides an optimized and elaborate preparation procedure for retinal organoid samples for transmission electron microscopy. It is suitable for applications that involve the analysis of synapses in mature retinal organoids.

Retinal organoids (ROs) are a three-dimensional culture system mimicking human retinal features that have differentiated from induced pluripotent stem ...

Correlative Microscopy Reveals Stress-Induced Mitochondria-Lysosome Interactions

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

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

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

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

Microscopy Techniques for Interpreting Fungal Colonization in Mycoheterotrophic Plants Tissues and Symbiotic Germination of Seeds

Structural botany is an indispensable perspective to fully understand the ecology, physiology, development, and evolution of plants. When researching mycoheterotrophic plants (i.e., plants that obtain carbon from fungi), remarkable aspects of their structural adaptations, the patterns of tissue colonization by fungi, and the morphoanatomy of subterranean organs can enlighten their developmental strategies and their relationships with hyphae, the source of nutrients. Another important role of symbiotic fungi is related to the germination of orchid seeds; all Orchidaceae species are mycoheterotrophic during germination and seedling stage (initial mycoheterotrophy), even the ones that photosynthesize in adult stages. Due to the lack of nutritional reserves in orchid seeds, fungal symbionts are essential to provide substrates and enable germination. Analyzing germination stages by structural perspectives can also answer important questions regarding the fungi interaction with the seeds. Different imaging techniques can be applied to unveil fungi endophytes in plant tissues, as are proposed in this article. Freehand and thin sections of plant organs can be stained and then observed using light microscopy. A fluorochrome conjugated to wheat germ agglutinin can be applied to the fungi and co-incubated with Calcofluor White to highlight plant cell walls in confocal microscopy. In addition, the methodologies of scanning and transmission electron microscopy are detailed for mycoheterotrophic orchids, and the possibilities of applying such protocols in related plants is explored. Symbiotic germination of orchid seeds (i.e., in the presence of mycorrhizal fungi) is described in the protocol in detail, along with possibilities of preparing the structures obtained from different stages of germination for analyses with light, confocal, and electron microscopy.

This protocol aims to provide detailed procedures for collecting, fixing, and maintaining mycoheterotrophic plant samples, applying different microscopy techniques such as scanning and transmission electron microscopy, light, confocal, and fluorescence microscopy to study fungal colonization in plants tissues and seeds germinated with mycorrhizal fungi.

Structural botany is an indispensable perspective to fully understand the ecology, physiology, development, and evolution of plants. When researching ...

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

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