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 border="0" cellpadding="0" cellspacing="0" width="672">
	<tbody>
		<tr height="84">
			<td height="84" style="height:84px;width:231px;">JB-4 Plus Embedding Kit</td>
			<td style="width:115px;">Polysciences Europe GmbH</td>
			<td style="width:161px;">18570-1</td>
			<td style="width:167px;">includes Benzoyl Peroxide, Plasticized (Catalyst) and Solution A+B</td>
		</tr>
		<tr height="42">
			<td height="42" style="height:42px;width:231px;">Polyethylene Molding Cup Trays, 6x8x5mm hexagon (9 cavities)</td>
			<td style="width:115px;">Polysciences Europe GmbH</td>
			<td style="width:161px;">17177A-3</td>
			<td style="width:167px;"></td>
		</tr>
		<tr height="42">
			<td height="42" style="height:42px;width:231px;">Polyethylene Molding Cup Trays, 13x19x5mm (9 cavities)</td>
			<td style="width:115px;">Polysciences Europe GmbH</td>
			<td style="width:161px;">17177C-3</td>
			<td style="width:167px;"></td>
		</tr>
		<tr height="42">
			<td height="42" style="height:42px;">JB-4 Plastic Block Holders</td>
			<td style="width:115px;">Polysciences Europe GmbH</td>
			<td>15899-50</td>
			<td></td>
		</tr>
		<tr height="63">
			<td height="63" style="height:63px;width:231px;">Eosin</td>
			<td style="width:115px;">Waldeck GmbH &#38; Co. KG, Division Chroma</td>
			<td style="width:161px;">1A-196</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:231px;">Microtec CUT 4060E</td>
			<td></td>
			<td style="width:161px;"></td>
			<td>rotary microtome</td>
		</tr>
		<tr height="84">
			<td height="84" style="height:84px;width:231px;">Leica DM LM, fluorescence compound microscope</td>
			<td style="width:115px;">Leica Mikrosysteme Handelges.m.b.H</td>
			<td style="width:161px;"></td>
			<td></td>
		</tr>
		<tr height="84">
			<td height="84" style="height:84px;width:231px;">GFP filter set</td>
			<td style="width:115px;">Leica Mikrosysteme Handelges.m.b.H</td>
			<td style="width:161px;">11090937180000</td>
			<td></td>
		</tr>
		<tr height="84">
			<td height="84" style="height:84px;width:231px;">Motorised cross table</td>
			<td style="width:115px;">Walter Uhl, technische Mikroskopie GmbH &#38; CO. KG</td>
			<td style="width:161px;">KT5-LSMA</td>
			<td></td>
		</tr>
		<tr height="42">
			<td height="42" style="height:42px;width:231px;">Digital video camera SPOT-FLEX</td>
			<td style="width:115px;">Visitron Systems GmbH.</td>
			<td style="width:161px;"></td>
			<td></td>
		</tr>
		<tr height="42">
			<td height="42" style="height:42px;width:231px;">precisExcite High-Power LED</td>
			<td style="width:115px;">Visitron Systems GmbH.</td>
			<td style="width:161px;"></td>
			<td>light source</td>
		</tr>
		<tr height="42">
			<td height="42" style="height:42px;width:231px;">VisiView 2.1.4</td>
			<td style="width:115px;">Visitron Systems GmbH.</td>
			<td style="width:161px;"></td>
			<td>Image capturing software</td>
		</tr>
		<tr height="84">
			<td height="84" style="height:84px;width:231px;">Hard metal knife (tungsten carbide), profile D</td>
			<td style="width:115px;">Leica Mikrosysteme Handelges.m.b.H</td>
			<td style="width:161px;"></td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:231px;">KL 2500 LCD</td>
			<td style="width:115px;">Schott AG</td>
			<td style="width:161px;"></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="https://cloudflare-jove-com.remotexs.ntu.edu.sg/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="https://cloudflare-jove-com.remotexs.ntu.edu.sg/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="https://cloudflare-jove-com.remotexs.ntu.edu.sg/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="https://cloudflare-jove-com.remotexs.ntu.edu.sg/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="https://cloudflare-jove-com.remotexs.ntu.edu.sg/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="https://cloudflare-jove-com.remotexs.ntu.edu.sg/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="https://cloudflare-jove-com.remotexs.ntu.edu.sg/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

Nanyang Technological University

Research

JoVE Journal

Developmental Biology

10.1K Views.  Medical University of Vienna. 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.

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

Analysis of Zebrafish Kidney Development with Time-lapse Imaging Using a Dissecting Microscope Equipped for Optical Sectioning

In order to understand organogenesis, the spatial and temporal alterations that occur during development of tissues need to be recorded. The method described here allows time-lapse analysis of normal and impaired kidney development in zebrafish embryos by using a fluorescence dissecting microscope equipped for structured illumination and z-stack acquisition. To visualize nephrogenesis, transgenic zebrafish (Tg(wt1b:GFP)) with fluorescently labeled kidney structures were used. Renal defects were triggered by injection of an antisense morpholino oligonucleotide against the Wilms tumor gene wt1a, a factor known to be crucial for kidney development.
The advantage of the experimental setup is the combination of a zoom microscope with simple strategies for re-adjusting movements in x, y or z direction without additional equipment. To circumvent focal drift that is induced by temperature variations and mechanical vibrations, an autofocus strategy was applied instead of utilizing a usually required environmental chamber. In order to re-adjust the positional changes due to a xy-drift, imaging chambers with imprinted relocation grids were employed.
In comparison to more complex setups for time-lapse recording with optical sectioning such as confocal laser scanning&#160;or light sheet microscopes, a zoom microscope is easy to handle. Besides, it offers dissecting microscope-specific benefits such as high depth of field and an extended working distance.
The method to study organogenesis presented here can also be used with fluorescence stereo microscopes not capable of optical sectioning. Although limited for high-throughput, this technique offers an alternative to more complex equipment that is normally used for time-lapse recording of developing tissues and organ dynamics.

The method described here allows time-lapse analysis of organ development in zebrafish embryos by using a fluorescence dissecting microscope capable of performing optical sectioning and simple strategies of readjustment to correct focal and planar drift.

In order to understand organogenesis, the spatial and temporal alterations that occur during development of tissues need to be recorded. The method ...

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) ...

A Simple Chamber for Long-term Confocal Imaging of Root and Hypocotyl Development

Several aspects of plant development, such as lateral root morphogenesis, occur on time spans of several days. To study underlying cellular and subcellular processes, high resolution time-lapse microscopy strategies that preserve physiological conditions are required. Plant tissues must have adequate nutrient and water supply with sustained gaseous exchange but, when submerged and immobilized under a coverslip, they are particularly susceptible to anoxia. One strategy that has been successfully employed is the use of a perfusion system to maintain a constant supply of oxygen and nutrients. However, such arrangements can be complicated, cumbersome, and require specialized equipment. Presented here is an alternative strategy for a simple imaging system using perfluorodecalin as an immersion medium. This system is easy to set up, requires minimal equipment, and is easily mounted on a microscope stage, allowing several imaging chambers to be set up and imaged in parallel. In this system, lateral root growth rates are indistinguishable from growth rates under standard conditions on agar plates for the first two days, and lateral root growth continues at reduced rates for at least another day. Plant tissues are supplied with nutrients via an agar slab that can be used also to administer a range of pharmacological compounds. The system was established to monitor lateral root development but is readily adaptable to image other plant organs such as hypocotyls and primary roots.

Presented here is a simple technique for high-resolution confocal time-lapse imaging of root and hypocotyl development for up to 3 days using high numerical-aperture objectives and perfluorodecalin as an immersion medium.

Several aspects of plant development, such as lateral root morphogenesis, occur on time spans of several days. To study underlying cellular and ...

Protocols for Visualizing Steroidogenic Organs and Their Interactive Organs with Immunostaining in the Fruit Fly Drosophila melanogaster

In multicellular organisms, a small group of cells is endowed with a specialized function in their biogenic activity, inducing a systemic response to&#160;growth and reproduction. In insects, the larval prothoracic gland (PG) and the adult female ovary play essential roles in biosynthesizing the principal steroid hormones called ecdysteroids. These ecdysteroidogenic organs are innervated from the nervous system, through&#160;which the timing of biosynthesis is affected by environmental cues. Here we describe a protocol for visualizing ecdysteroidogenic organs and their interactive organs in larvae and adults of the fruit fly Drosophila melanogaster, which provides a suitable model system for studying steroid hormone biosynthesis and its regulatory mechanism. Skillful dissection allows us to maintain the positions of ecdysteroidogenic organs and their interactive organs including the brain, the ventral nerve cord, and other tissues. Immunostaining with antibodies against ecdysteroidogenic enzymes, along with transgenic fluorescence proteins driven by tissue-specific promoters, are available to label ecdysteroidogenic cells. Moreover, the innervations of the ecdysteroidogenic organs can also be labeled by specific antibodies or a collection of GAL4 drivers in various types of neurons. Therefore, the ecdysteroidogenic organs and their neuronal connections can be&#160;visualized simultaneously&#160;by immunostaining and transgenic techniques. Finally, we describe how to visualize germline stem cells, whose proliferation and maintenance are controlled by ecdysteroids. This method contributes to comprehensive understanding of steroid hormone biosynthesis and its neuronal regulatory mechanism.

Protocols for Visualizing Steroidogenic Organs and Their Interactive Organs with Immunostaining in the Fruit Fly Drosophila melanogaster

We describe a protocol for dissection, fixation, and immunostaining of steroidogenic organs in Drosophila larvae and adult females to study steroid hormone biosynthesis and its regulatory mechanism. In addition to steroidogenic organs, we visualize the innervation of steroidogenic organs as well as steroidogenic target cells such as germline stem cells.

In multicellular organisms, a small group of cells is endowed with a specialized function in their biogenic activity, inducing a systemic response ...

Protocol for the Differentiation of Human Induced Pluripotent Stem Cells into Mixed Cultures of Neurons and Glia for Neurotoxicity Testing

Human pluripotent stem cells can differentiate into various cell types that can be applied to human-based in vitro toxicity assays. One major advantage is that the reprogramming of somatic cells to produce human induced pluripotent stem cells (hiPSCs) avoids the ethical and legislative issues related to the use of human embryonic stem cells (hESCs). HiPSCs can be expanded and efficiently differentiated into different types of neuronal and glial cells, serving as test systems for toxicity testing and, in particular, for the assessment of different pathways involved in neurotoxicity. This work describes a protocol for the differentiation of hiPSCs into mixed cultures of neuronal and glial cells. The signaling pathways that are regulated and/or activated by neuronal differentiation are defined. This information is critical to the application of the cell model to the new toxicity testing paradigm, in which chemicals are assessed based on their ability to perturb biological pathways. As a proof of concept, rotenone, an inhibitor of mitochondrial respiratory complex I, was used to assess the activation of the Nrf2 signaling pathway, a key regulator of the antioxidant-response-element-(ARE)-driven cellular defense mechanism against oxidative stress.

Human induced pluripotent stem cells (hiPSCs) are considered a powerful tool for drug and chemical screening and for the development of new in vitro models for toxicity testing, including neurotoxicity. Here, a detailed protocol for the differentiation of hiPSCs into neurons and glia is described.

Human pluripotent stem cells can differentiate into various cell types that can be applied to human-based in vitro toxicity assays. One major advantage is ...

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

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

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

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

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 ...

Live Cell Imaging of Chromosome Segregation During Mitosis

Chromosomes must be reliably and uniformly segregated into daughter cells during mitotic cell division. Fidelity of chromosomal segregation is controlled by multiple mechanisms that include the Spindle Assembly Checkpoint (SAC). The SAC is part of a complex feedback system that is responsible for prevention of a cell progress through mitosis unless all chromosomal kinetochores have attached to spindle microtubules. Chromosomal lagging and abnormal chromosome segregation is an indicator of dysfunctional cell cycle control checkpoints and can be used to measure the genomic stability of dividing cells. Deregulation of the SAC can result in the transformation of a normal cell into a malignant cell through the accumulation of errors during chromosomal segregation. Implementation of the SAC and the formation of the kinetochore complex are tightly regulated by interactions between kinases and phosphatase such as Protein Phosphatase 2A (PP2A). This protocol describes live cell imaging of lagging chromosomes in mouse embryonic fibroblasts isolated from mice that had a knockout of the PP2A-B56&#947; regulatory subunit. This method overcomes the shortcomings of other cell cycle control imaging techniques such as flow cytometry or immunocytochemistry that only provide a snapshot of a cell cytokinesis status, instead of a dynamic spatiotemporal visualization of chromosomes during mitosis.

This protocol describes an easy and convenient method to label and visualize live chromosomes in mitotic cells using Histone2B-GFP BacMam 2.0 label and a spinning disc confocal microscopy system.

Chromosomes must be reliably and uniformly segregated into daughter cells during mitotic cell division. Fidelity of chromosomal segregation is controlled ...

Thin Film Composite Silicon Elastomers for Cell Culture and Skin Applications: Manufacturing and Characterization

In this protocol, we present methods to fabricate thin elastomer composite films for advanced cell culture applications and for the development of skin adhesives. Two different poly-(dimethyl siloxanes) (PDMS and soft skin adhesive (SSA)), have been used for in depth investigation of biological effects and adhesive characteristics. The composite films consist of a flexible backing layer and an adhesive top coating. Both layers have been manufactured by doctor blade application technique. In the present investigation, the adhesive behavior of the composite films has been investigated as a function of the layer thickness or a variation of the Young&#39;s modulus of the top layer. The Young&#39;s modulus of PDMS has been changed by varying the base to crosslinker mixing ratio. In addition, the thickness of SSA films has been varied from approx. 16 &#181;m to approx. 320 &#181;m. Scanning electron microscopy (SEM) and optical microscopy have been used for thickness measurements. The adhesive properties of elastomer films depend strongly on the film thickness, the Young&#39;s modulus of the polymers and surface characteristics. Therefore, normal adhesion of these films on glass substrates exhibiting smooth and rough surfaces has been investigated. Pull-off stress and work of separation are dependent on the mixing ratio of silicone elastomers.
Additionally, the thickness of the soft skin adhesive placed on top of a supportive backing layer has been varied in order to produce patches for skin applications. Cytotoxicity, proliferation and cellular adhesion of L929 murine fibroblasts on PDMS films (mixing ratio 10:1) and SSA films (mixing ratio 50:50) have been conducted. We have shown here, for the first time, the side by side comparison of thin composite films manufactured of both polymers and present the investigation of their biological- and adhesive properties.

A protocol for the manufacturing process of polymeric thin film composite structures possessing either different Young&#39;s moduli or thicknesses is presented. Films are produced for advanced cell culture studies or as skin adhesives.

In this protocol, we present methods to fabricate thin elastomer composite films for advanced cell culture applications and for the development of skin ...

Visualizing the Node and Notochordal Plate In Gastrulating Mouse Embryos Using Scanning Electron Microscopy and Whole Mount Immunofluorescence

The post-implantation mouse embryo undergoes major shape changes after the initiation of gastrulation and morphogenesis. A hallmark of morphogenesis is the formation of the transient organizers, the node and notochordal plate, from cells that have passed through the primitive streak. The proper formation of these signaling centers is essential for the development of the body plan and techniques to visualize them are of high interest to mouse developmental biologists. The node and notochordal plate lie on the ventral surface of gastrulating mouse embryos around embryonic day (E) 7.5 of development. The node is a cup-shaped structure whose cells possess a single slender cilium each. The proper subcellular localization and rotation of the cilia in the node pit determines left-right asymmetry. The notochordal plate cells also possess single cilia albeit shorter than those of the node cells. The notochordal plate forms the notochord which acts as an important signaling organizer for somitogenesis and neural patterning. Because the cells of the node and notochordal plate are transiently present on the surface and possess cilia, they can be visualized using scanning electron microscopy (SEM). Among other techniques used to visualize these structures at the cellular level is whole mount immunofluorescence (WMIF) using the antibodies against the proteins that are highly expressed in the node and notochordal plate. In this report, we describe our optimized protocols to perform SEM and WMIF of the node and notochordal plate in developing mouse embryos to help in the assessment of tissue shape and cellular organization in wild-type and gastrulation mutant embryos.

The node and notochordal plate are transient signaling organizers in developing mouse embryos that can be visualized using several techniques. Here, we describe in detail how to perform two of the techniques to study their structure and morphogenesis: 1) scanning electron microscopy (SEM); and 2) whole mount immunofluorescence (WMIF).

The post-implantation mouse embryo undergoes major shape changes after the initiation of gastrulation and morphogenesis. A hallmark of morphogenesis is ...