This work was supported by grant FKZ 13GW0044 from the German Federal Ministry for Education and Research, project MorphiQuant-3D. We thank Carolin Bartels for technical support.

NOTE: Sample blocks should be polymerized and contain some heavy metal. Fixation and embedding protocols for the two samples shown in Figure 1A-B have been described elsewhere11. In short, the sample shown in Figure 1A was chemically fixed, stained en bloc first with 1% OsO4, then with 1% uranyl acetate, and embedded in Spurr&#39;s resin. The sample shown in Figure 1B was high pressure frozen, freeze substituted with 0.4% uranyl acetate in acetone, and embedded in Lowicryl HM20 resin. Use powder free gloves for the next preparation steps.
1. Creating Arrays
<ol>
	<li>Trimming of the sample block 
	NOTE: Always tighten screws well when inserting parts.
	<ol>
		<li>Insert the sample block into the sample holder of the ultramicrotome, place the holder into the trimming block, and slide it into the lower stage of the ultramicrotome.</li>
		<li>Trim away resin around the embedded tissue with a razor blade leaving only a small rim of resin around the sample. Trim from the top until the target is reached. 
		NOTE: The shape of the block-face may be trapezoid or rectangular (we have successfully used both shapes). It is important to use the longer sides of the rectangle as the leading and trailing edge to ensure that a large area is covered by the glue mixture stabilizing the ribbons.</li>
		<li>Insert the sample holder into the arm of the ultramicrotome and replace the trimming block in the lower stage by the knife holder. Insert a diamond trim knife (normally 45&#176;), into the knife holder. Align the block-face exactly parallel to the trim knife edge. 
		NOTE: To produce exactly parallel leading (bottom) and trailing (top) edges, turn or move only the knife to trim opposite sides. For large volumes (more than several hundred sections), a 90&#176; trim knife is advantageous.</li>
		<li>Smooth all four sides, then rotate the sample holder so that the leading and trailing edge are now in the horizontal position.</li>
		<li>Carefully coat the leading and trailing sides of the block with adhesive mixture. Use a tiny brush formed from a few hairs fixed to a toothpick13. Perform this step quickly because the solvent of this mixture evaporates within seconds. Do not contaminate the block-face with this mixture. Let the coated sample block dry for 5-10 min. 
		NOTE: For larger numbers of sections (&#62;200), it may be better to coat the leading edge only, because with time, a bulge of glue might build up at the trailing edge, and potentially pull back sections over the knife-edge.</li>
	</ol>
	</li>
	<li>Preparation of the substrate
	<ol>
		<li>Cut pieces of silicon wafers to a size that fits into the knife boat (roughly 2 x 2.5 cm2 for the Jumbo knife). If needed, mark the wafer pieces (numbers or letters) with a diamond scriber before cleaning, or with permanent marker after cleaning. Clean the silicon wafer manually with isopropanol and lint-free tissue. 
		NOTE: For correlative imaging use indium-tin-oxide (ITO)-coated glass coverslips. They must be handled very carefully, and extra cleaning is not necessary.</li>
		<li>Fix the substrate to one end of the carrier plate using a removable adhesive. 
		NOTE: Alternatively, the substrate edges may be pinned down parallel to the knife-edge using two stripes of adhesive tape.</li>
		<li>Plasma activate (glow discharge) the substrate with air to obtain a hydrophilic surface. This should be done in such a way that a drop of water placed onto the substrate spreads to a very thin film (low contact angle, Figure 2C, D). Hydrophilization parameters depend on the plasma device used; for the parameters used here, see Table of Materials. 
		NOTE: Plasma activation is very volatile, so perform this immediately before substrate usage.</li>
		<li>Insert the carrier into the clamp of a substrate holder, with the mounted substrate closer to the knife-edge to achieve optimal wetting of the substrate in the knife boat. 
		NOTE: A detailed description of the substrate holder, including the computer-aided design (CAD) drawings, is given in Wacker et al.11</li>
	</ol>
	</li>
	<li>Sectioning Preparation
	<ol>
		<li>Insert a Jumbo diamond knife into the knife holder, set the clearance angle (0&#176; for Jumbo knife), and fill the knife boat with distilled water. Approach the knife to a distance of 1&#8211;2 mm to the sample.</li>
		<li>Lower the substrate into the water using screws 1&#8211;3 (Figure 2A) of the substrate holder. Check that the waterline is located in the upper third of the substrate. 
		NOTE: Check that the glue between the substrate and carrier plate is well cured by nudging the substrate with clean forceps. It should not move.</li>
		<li>Because it is hard to see the ground clearance when using a silicon wafer, lower the substrate until you feel it touch the floor. Now raise the substrate a small amount. Make sure that neither the substrate nor the carrier touches the knife boat while cutting.</li>
		<li>Use a syringe or pipette to adjust the water level in the boat. While watching through the binocular, add or remove water until the full area of the water surface shows a homogenous reflection of the top light illumination of the ultramicrotome.</li>
		<li>Switch on the bottom light of the ultramicrotome. Ensure that the arm is in the middle position and not in retraction using the hand wheel of the ultramicrotome. Approach the knife to the sample until the reflection of the knife-edge is visible on the block face.</li>
		<li>Use the adjustment options of the ultramicrotome to align the sample to the knife. First rotate the knife, then rotate and tilt the sample. 
		NOTE: It is properly aligned if the light stripe, which can be seen in the gap between the sample and the knife, shows parallel edges (straight parallel lines and not wedge shaped).</li>
		<li>Check the inclination of the sample: Make sure that the light stripe does not become thicker or thinner while moving the sample up and down. If needed, use the arc adjustment screw to correct this. Move the knife closer to the sample until it is just above the block-face (but does not touch it).</li>
		<li>Set section thickness (feed), cutting speed, and cutting window at the control unit.</li>
		<li>Start sectioning. If necessary, wait until the first full section is being cut. Cut some sections to be sure that they stick together to form ribbons (otherwise the glue has to be applied again). Start with a high feed value (maximum, 200 nm for the Ultra knife) until the first full section is being cut. Then, set the desired feed value. For adequate ribbon stability, a section thickness of 100 nm and a cutting speed of 1 mm/s is a good starting point. The lowest achievable section thickness is around 60 nm, depending on sample quality.</li>
		<li>Stop sectioning. Remove all unneeded (partially cut) sections from the knife-edge and boat using an eyelash/cat&#39;s hair. If there is a lot of small debris floating around, remove the water completely with a pipette and fill the boat with fresh water. Now the process is ready for the first productive ribbon.</li>
	</ol>
	</li>
	<li>Sectioning
	<ol>
		<li>Start sectioning. Once a number of sections (the actual number depends on the size of sections and substrate) has been cut, stop the sectioning process and release the ribbon from the knife-edge by gently stroking over the knife-edge with an eyelash14 or better yet, with a very soft hair from a cat&#39;s fur.</li>
		<li>Manipulate (push/pull) the ribbon with the eyelash towards the substrate and attach the first section to the substrate. 
		NOTE: It is necessary to gently push the ribbon until it sticks to the dry part of the substrate.</li>
		<li>Continue sectioning and attaching the ribbons to the substrate. Start on one side and move gradually over to the other with each new ribbon. 
		NOTE: Avoid massive water movements to avoid loosening the already attached ribbons. The same is true for air currents. Use the breath shield delivered with the ultramicrotome. In unfavorable environmental conditions, an enclosure for the ultramicrotome is recommended15.</li>
		<li>When the substrate is completely covered with ribbons (usually 4&#8211;5 ribbons are feasible), gently lift-out the substrate from the knife boat using the micromanipulator screws of the substrate holder. 
		NOTE: Suitable movements are: lift up vertically (screw 1) and rotating/tilting out (screw 3), or a combination of both.</li>
		<li>Let the ribbon array dry before storing it in a dust-free environment. After drying, remove the adhesive mounted substrate as soon as possible from the carrier (on the same day, otherwise the substrate might be too hard to remove or may even break during demounting).</li>
	</ol>
	</li>
</ol>
2. Staining for the LM Imaging
NOTE: Different staining/labeling methods are possible, including immunofluorescence protocols. Here a direct, rather unspecific stain is chosen to outline the cell walls.
<ol>
	<li>Propidium iodide staining
	<ol>
		<li>Cover the bottom of a large glass Petri dish (30 cm diameter) with parafilm and line the edge of the dish with wet tissue to build a humid chamber.</li>
		<li>Use approximately 300-500 &#181;L of solution per coverslip. Place one drop for each coverslip on the parafilm and put the glass upside down onto the drop, so that the sections are in contact with the staining fluid. Cover the dish and wrap it with aluminum foil to protect the samples from light. Incubate the samples for 16 h at 4 &#176;C.</li>
		<li>Remove the coverslip with forceps and wash it by moving it up and down in a 100-mL beaker filled with 80 mL of distilled water. Repeat this step in another beaker with fresh water. Dry the coverslip carefully with compressed air.</li>
	</ol>
	</li>
</ol>
3. Recording the Image Stack in the FLM
<ol>
	<li>Place the coverslip on the stage of a common wide-field FLM.</li>
	<li>Choose the appropriate filter set (Table of Materials) for the fluorescence to be observed.</li>
	<li>With a suitable objective lens, take an image of the object in each section: try to fill the field of view and keep the orientation constant. For the root tip, a 40X air objective was used. 
	NOTE: If the ribbons are not perfectly straight, a rotating stage can help to reorient the sections while taking images. If possible, center the image on a specific feature or keep a feature such as the edge of the section at an equal distance to the edge of the recorded image.</li>
	<li>If possible, use 16 bit to limit saturated pixels and keep the exposure time constant.</li>
</ol>
4. Registration of the FLM Image Stack
<ol>
	<li>Import the image series into Fiji16 as a virtual stack.</li>
	<li>Open a new TrakEM17 (blank) from the file menu.</li>
	<li>Right click into the image field, and import the stack into TrakEM as &#34;One slice per layer&#34;.</li>
	<li>Align layers (right click into image field), set the range (first image to last), and choose none as the reference. For all settings, use the default values, and choose rigid as the desired transformation.</li>
	<li>When the registration is finished and satisfactory, save the aligned dataset by right click and choose export. Make flat image, set the range from first to last image, and let the software show the resulting stack. Save the stack in tif-format.</li>
</ol>
5. Staining and Mounting for the SEM Imaging
NOTE: For preparing staining solutions see Table of Materials. Solutions may be stored at 4 &#176;C for up to 12 months, protected from light and air.
Caution: Lead citrate and uranyl acetate contain heavy metals which are toxic. Wear gloves and dispose the waste according to the local authorities&#39; instructions.
<ol>
	<li>Cover the bottom of a large glass Petri dish (30 cm diameter) with parafilm and line the edge of the dish with wet tissue to build a humid chamber. 
	NOTE: It is important that several pellets of NaOH are positioned within the Petri dish near the staining drops to prevent excessive precipitation of lead citrate.</li>
	<li>Uranyl acetate staining
	<ol>
		<li>Centrifuge the uranyl acetate solution at 2,680 x g for a few seconds to sediment small particles.</li>
		<li>Use approximately 300&#8211;500 &#181;L of solution per coverslip. Place one drop for each coverslip on the parafilm and put the glass upside down onto the drop, so that the sections are in contact with the staining fluid.</li>
		<li>Incubate for 10 min at room temperature, and cover the dish during staining.</li>
		<li>Remove the coverslip with forceps and wash by moving it up and down in a beaker filled with distilled water (see step 2.1.3).</li>
	</ol>
	</li>
	<li>Lead citrate staining
	<ol>
		<li>During the uranyl acetate incubation, prepare the lead citrate solution. 
		NOTE: Lead citrate should always be filtered immediately before use to remove any precipitates. Also centrifuge the lead citrate solution at 2,680 x g for a few seconds as in step 5.2.1.</li>
		<li>Use approximately 300&#8211;500 &#181;L of solution per coverslip. Place one drop for each coverslip on the parafilm immediately before washing the coverslips after the uranyl acetate staining, and put the glass upside down onto the drop, so that the sections are in contact with the staining fluid. Place the drops (300&#8211;500 &#181;L) on the parafilm immediately before washing the coverslips after the uranyl acetate staining. 
		NOTE: To avoid formation of precipitates, do not breathe onto the lead citrate droplets.</li>
		<li>Place the washed coverslip upside down onto the drop (there is no need to dry it).</li>
		<li>Incubate for 5 min at room temperature, and cover the dish during the staining.</li>
		<li>Remove the coverslip with forceps and wash it as described above in a beaker with fresh water (step 5.2.4).</li>
	</ol>
	</li>
	<li>Dry the coverslip carefully with compressed air.</li>
	<li>Mounting samples for the SEM imaging
	<ol>
		<li>Mount the silicon wafers on aluminum stubs with a sticky carbon pad. 
		NOTE: ITO-coated coverslips may either be mounted using silver paint and Cu-tape &#8212; make sure the conductive surface is connected to the stub &#8212; or with carbon pads as above. In that case, a conductive connection from the ITO-surface to the stub can be made with a drop of silver paint.</li>
	</ol>
	</li>
</ol>
6. Hierarchical Imaging in the SEM
NOTE: In a field emission SEM, choose a low primary energy (3 kV or lower), a beam current in a range from 50 to 800 pA to avoid charging, and a suitable working distance for efficient collection of secondary and/or back-scattered electrons. Selection of the beam current depends on the sample properties (e.g., embedding resin); the electron dose will also be a compromise between a small current (less harmful to the sample) and a high current, which is beneficial for imaging speed and therefore lowers the total image acquisition time. Dedicated detectors for back-scattered electrons provide good contrast, are less sensitive to charging of the sample, and show less of the sample&#39;s surface artifacts (folds, knife marks). Contrast and brightness should be adjusted such that the histogram is centered.
<ol>
	<li>SEM imaging
	<ol>
		<li>First define the four corners of the array by grabbing an image of each corner at low magnification, about 100x. Create a region of interest (ROI) enclosing the whole array. Assign an imaging protocol with the following parameters: Use a secondary electron (SE) detector, which allows for high speed imaging at a large image pixel size (for example 1,000 nm), and a short dwell time (e.g., 0.2 &#181;s). 
		NOTE: To overcome electron optical limitations at a large scanning fields of view (FOV), which might result in distortions at the periphery of the images, use dedicated low magnification modes (provided by most SEM manufacturers) or use medium, 1 to 2 k scan fields for single images.</li>
		<li>Generate a section set by creating a ROI outlining just the tissue in the first section. Clone it to all subsequent sections using the stamp tool. Rotate the ROIs when needed to accommodate bent ribbons.</li>
		<li>Record the image series using an intermediate pixel size (around 50 nm) and a dwell time long enough to identify and recognize the target structure. Use a FOV for single images in a range of 6-10 k pixels. 
		NOTE: The Atlas 5 software can automatically collect mosaics composed of adjacent images to cover large ROI/section areas across the serial sections.</li>
		<li>Create a site set within this section set, containing the target structure for higher resolution SEM imaging. Make the ROI large enough to account for stage precision. Check and adjust the positions of the sites. 
		NOTE: It is important to place the ROIs in such a way that the center, where the autofocus and autostigmation will be performed, does not sit on &#34;empty&#34; material with no structural detail, e.g., vacuoles.</li>
		<li>Define the autofocus settings and check the performance over at least the length of a ribbon (i.e., the longest distance that the stage has to travel) on a small ROI close to the site that will be imaged.</li>
		<li>Define an imaging protocol for the high-resolution SEM acquisition. To see membrane compartments, choose 3&#8211;5 nm image pixel size. Select a dwell time depending on the detector so that the image is not too noisy.</li>
		<li>Before starting the acquisition, define the focus values on at least the first section of each ribbon using the check protocol option.</li>
		<li>Start the automated SEM imaging over the whole series of target ROIs.</li>
		<li>Export the acquired data as image series, preferably in tif-format.</li>
	</ol>
	</li>
</ol>
7. Registration of the SEM Image Stack
<ol>
	<li>Import image series into Fiji as a virtual stack. 
	NOTE: These will be large data files in the range of a few GB depending on the number of sections and the size of the ROI.</li>
	<li>Crop the SEM image stack for further processing to an area as close to the structure of interest as possible, and adjust the brightness and contrast.</li>
	<li>Open a new TrakEM17 (blank) from the file menu.</li>
	<li>Right click into the image field, and import the stack into TrakEM as &#34;One slice per layer&#34;.</li>
	<li>Align the layers (right click into the image field), choose least squares as mode, set the range (first image to last), and choose none as the reference. For the settings, use the default values and choose rigid as the desired transformation.</li>
	<li>When the registration is completed and satisfactory, save the aligned dataset by right click and choose export. Make a flat image, set the range from first to last image, and let the software show the resulting stack. Save the stack in tif-format.</li>
</ol>

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KIT has received reimbursements by Boeckeler Instruments for supply of a functional model of the substrate holder. Marlene Thaler is employee of ZEISS Microscopy GmbH, manufacturer of the microscope systems mentioned in this article. In addition, ZEISS offers certain solutions such as ZEISS Atlas solution packages for a wide range of applications in large-area, 3D imaging for SEM and FIB-SEM instruments. All other authors have nothing to disclose.

A workflow for targeting specific cells within a tissue by multi-modal hierarchical AT was demonstrated: A resin-embedded sample is sliced up into arrays of serial sections, which are placed on a conductive substrate using a custom-designed substrate holder. After labeling with a fluorophore and imaging in the FLM, the reconstructed volume is used for selecting the target cells. After additional staining rounds with heavy metals to introduce contrast, these targets are imaged over several hundred sections at nanoscale resolution in an SEM using an automated software platform.
For producing densely packed arrays with several long ribbons, a substrate holder similar to the one described here is necessary. A skilled and patient person may be able to attach several ribbons to a silicon substrate, semi-immersed in the knife boat, and retrieve the array by gradually lowering the water level until the ribbons are sitting on the substrate. However, in the our experience, there is a tendency to shatter formation when the substrate is touching any part of the knife boat (cf. note in 1.3.2 in protocol). In addition, this procedure is much more difficult with ITO-coated substrates: (1) due to the transparency of the ITO-glass, it is difficult to see the edge of the water where the ends of the ribbons have to be attached; and (2) because the ITO-coated surface is much rougher than the highly polished silicon wafer, the ribbons tend to break during lift-out and smaller fragments consisting of a few sections may float, thus destroying the order of the sections.&#160;&#160;
The whole workflow is also feasible without correlation to FLM data. In this case, data collection in the SEM may have to be performed in several sessions. An initial 3D reconstruction or at least evaluation of low or medium resolution data may be necessary to identify targets. In addition, conventional histological stains for brightfield LM (not requiring FLM) may be applied. Of course, other options6,7,8 are antibody labeling on the arrays, as already demonstrated in the initial paper on AT18, or genetically encoded fluorescent proteins (XFPs) or pre-embedding labeling with preservation of fluorescence during sample preparation.
A general limitation of the discussed method is the use of sections of a certain thickness and the resulting discrete sampling of the 3D volume: Resolution in Z can only be as good as the thickness of the sections since the SEM collects only data from the section surface (depending on the primary energy/landing energy selected). This means that the resulting 3D volume has anisotropic voxels, e.g., 5 x 5 x 100 nm3 if 100 nm sections and an image pixel size of 5 nm are used. For very small entities in a size range below 1 &#181;m, this may not be sufficient for a true ultrastructural description. A more technical limitation is the accuracy of the stage used in the SEM for automated imaging. Due to this, it is necessary to choose an ROI larger than the specifications of the stage accuracy to guarantee that the full target area is imaged.
Compared to SBF-SEM and FIB-SEM as block-face imaging methods, correlative AT has the definitive disadvantage of anisotropic voxels, as described above. With FIB-SEM, isotropic voxels of 5 x 5 x 5 nm3 can be obtained when a proper drift correction is in place.
Gaps in the reconstructed volume due to loss of sections during preparation of arrays might also be a concern that is not encountered with SBF-SEM or FIB-SEM. With good ribbon stabilization by glue, this usually is only an issue for the last section of a ribbon: It might be damaged when releasing it from the knife-edge using the eyelash. However, in our experience, the loss of one section in every 20&#8211;50 sections does not influence image registration.
On the other hand, the possibility to post-stain arrays confers good signal and contrast for SEM imaging, even on weakly metalized samples such as the high pressure frozen root tips shown here. Therefore, it is not necessary to compromise optimal ultrastructural preservation by numerous chemical fixation and metallization steps. Also, routine samples from the pathology lab with intermediate degrees of metallization deliver excellent data10. Such a post-embedding contrast enhancement is not possible for SBF-SEM and FIB-SEM in general. Furthermore, since these methods are destructive, i.e., consuming the sample during imaging, hierarchical imaging at different resolutions and sites or repeated imaging at later points in time is impossible. In principle, unlimited volumes, consisting of large FOVs (e.g., up to several millimeters for whole mouse brains in connectomics) created by stitching mosaics, and huge numbers of sections can be acquired by AT, while in FIB-SEM, FOVs beyond 100 &#181;m x 100 &#181;m are difficult to achieve with routine instruments.
Further automation of the described AT-workflow would be a definite advantage, since the above-mentioned methods SBF-SEM and FIB-SEM perform both sectioning and imaging within the same instrument in a fully automated manner. One kind of automation of sectioning exists: The ATUMtome12 can generate and collect thousands of sections, but the use of Kapton tape as a substrate makes such arrays difficult to image in a FLM. On the ITO-coated coverslips used here, even super-resolution imaging should be possible. A further, very desirable target for automation would be the recording of the FLM data stacks. On the other hand, automation can be expensive and except for the substrate holder, the workflow presented here relies (in terms of hardware) only on instrumentation usually available in a routine EM laboratory or core facility, making it low level access.

For reconstructing large volumes of tissue at ultrastructural resolution a number of different imaging approaches based on SEM have been used1: Comprehensive reviews are available e.g., for SBF-SEM2, FIB-SEM3, and Array Tomography (AT)4. While for the latter method the sample material is preserved as an array of serial sections on a substrate, SBF-SEM and FIB-SEM are destructive methods, working on the sample block and consuming it during imaging. Due to the charging of the resin in the SEM, they also depend on strongly metalized sample blocks5.
On the other hand, identifying certain cells or structures of interest within a tissue sample can profit particularly from correlative light and electron microscopy (CLEM)6,7,8. Using FLM for targeting precludes the application of large amounts of heavy metal since this would quench the fluorescence signal9. For such only slightly metalized samples, AT is the method of choice since arrays may easily be post-stained with heavy metal after LM imaging. Moreover, almost any sample type may be used for AT, even routine samples from the pathologist&#39;s treasure chest10.
Another big advantage of AT is the potential for hierarchical11 or multi-resolution imaging12: It is not necessary to image everything at high-resolution, as targets may be selected in a different modality (e.g., FLM) or in low-resolution SEM images. Imaging only the interesting regions of a tissue or cell population at high-resolution saves digital data storage space and produces smaller image data sets, which are easier to handle. Here, the AT workflow is demonstrated using a rather weakly metalized sample: high pressure frozen plant roots (Arabidopsis thaliana) embedded in hydrophilic resin.
How the arrays are prepared, stained, and imaged in both FLM and SEM, and how the image stacks are registered are explained. Also, how the 3D reconstruction of the FLM volume can be used to select specific cells for imaging in the SEM at nanoscale resolution is illustrated.

<table><tbody><tr><td>&lt;strong&gt;Instrumentation&lt;/strong&gt;</td><td></td><td></td><td></td></tr><tr><td>Ultramicrotome</td><td>RMC</td><td>PT-PC</td><td>Alternative: Leica UC7</td></tr><tr><td>Substrate holder</td><td>RMC</td><td>ASH-100</td><td>Alternative: home built</td></tr><tr><td>Plasma cleaner</td><td>Diener</td><td>Zepto 40kHz</td><td>Alternatives: Ted Pella Pelco or other benchtop plasma cleaner&lt;br/&gt; Example Parameters for Diener Zepto with 40kHz generator (0-100W); 0.5 mbar, 5 sccm (Air), 10% performance&amp;nbsp;</td></tr><tr><td>Widefield fluorescence light microscope</td><td>Zeiss</td><td>Axio Observer.Z1</td><td>Alternatives:&lt;br/&gt; Leica, Nikon, Olympus</td></tr><tr><td>Fluorescence filter set</td><td>Zeiss</td><td>43 HE (Cy3/DsRed)</td><td></td></tr><tr><td>Objective lens</td><td>Zeiss</td><td>Zeiss Neofluar 40x</td><td>&amp;nbsp;0.75 NA</td></tr><tr><td>Decent workstation able to handle GB-sized image data</td><td></td><td></td><td></td></tr><tr><td>FESEM</td><td>Zeiss</td><td>Ultra 55</td><td>Alternatives: FEI, Jeol, Hitachi, TESCAN</td></tr><tr><td>&lt;strong&gt;Name&lt;/strong&gt;</td><td>&lt;strong&gt;Company&lt;/strong&gt;</td><td>&lt;strong&gt;Catalog Number&lt;/strong&gt;</td><td>&lt;strong&gt;Comments&lt;/strong&gt;</td></tr><tr><td>&lt;strong&gt;Sectioning&lt;/strong&gt;</td><td></td><td></td><td></td></tr><tr><td>Razor blades</td><td>Plano</td><td>T585-V</td><td></td></tr><tr><td>Diamond knife for trimming 45&amp;deg;</td><td>Diatome</td><td>DTB45</td><td></td></tr><tr><td>Diamond knife for trimming 90&amp;deg;</td><td>Diatome</td><td>DTB90</td><td></td></tr><tr><td>Jumbo diamond knife for sectioning</td><td>Diatome</td><td>DUJ3530</td><td></td></tr><tr><td>Silicon wafer (pieces)</td><td>Si-Mat</td><td>Custom Made</td><td>Doping: P/Bor, orientation: &amp;lt;100&gt;, thickness: 525 &amp;plusmn; 25 &amp;micro;m, resistivity: 1-30 &amp;Omega;-cm&lt;br/&gt; http://si-mat.com/silicon-wafers.html</td></tr><tr><td>ITO-coated coverslips</td><td>Balzers</td><td>Type Z</td><td>22 &amp;times; 22 &amp;times; 0.17 mm&lt;br/&gt; https://www.opticsbalzers.com/de/produkte/deckglas-fenster/corrslide.html</td></tr><tr><td>Aluminium carrier</td><td>Custom Made</td><td></td><td>76 &amp;times; 26 mm</td></tr><tr><td>Wafer forceps</td><td>Ideal-tek</td><td>34A.SA</td><td></td></tr><tr><td>Stubs forceps</td><td>Dumont</td><td>0103-2E1/2-PO-1</td><td>Dumoxel-H 2E 1/2</td></tr><tr><td>Diamond scriber</td><td>Plano</td><td>T5448</td><td></td></tr><tr><td>Eyelash/very soft cat&#039;s hair</td><td>Selfmade</td><td></td><td>Alternative: Plano</td></tr><tr><td>Brush</td><td>Selfmade</td><td></td><td></td></tr><tr><td>Pattex contact adhesive</td><td>Pattex</td><td>PCL3C</td><td>Kraftkleber Classic (the yellowish one)</td></tr><tr><td>Fixogum</td><td>Marabu</td><td>290110001</td><td>for fixing substrate to carrier</td></tr><tr><td>Adhesive tape</td><td>3M</td><td>851</td><td>for fixing substrate to carrier</td></tr><tr><td>Isopropanol</td><td>Bernd Kraft</td><td>07029.4000</td><td></td></tr><tr><td>Xylene</td><td>Carl Roth</td><td>4436</td><td>thinner for glue mixture</td></tr><tr><td>Rotihistol</td><td>Carl Roth</td><td>6640</td><td>alternative, limonene based thinner</td></tr><tr><td>&lt;strong&gt;Name&lt;/strong&gt;</td><td>&lt;strong&gt;Company&lt;/strong&gt;</td><td>&lt;strong&gt;Catalog Number&lt;/strong&gt;</td><td>&lt;strong&gt;Comments&lt;/strong&gt;</td></tr><tr><td>&lt;strong&gt;Software&lt;/strong&gt;</td><td></td><td></td><td></td></tr><tr><td>Image processing</td><td>Open source</td><td>Fiji (http://fiji.sc/#download)</td><td></td></tr><tr><td>Image acquisition</td><td>Zeiss</td><td>Atlas 5 AT&lt;br/&gt; (module for Zeiss SEM)</td><td>Alternative for automated image acquisition: WaferMapper: https://software.rc.fas.harvard.edu/lichtman/LGN/WaferMapper.html</td></tr><tr><td>&lt;strong&gt;Name&lt;/strong&gt;</td><td>&lt;strong&gt;Company&lt;/strong&gt;</td><td>&lt;strong&gt;Catalog Number&lt;/strong&gt;</td><td>&lt;strong&gt;Comments&lt;/strong&gt;</td></tr><tr><td>&lt;strong&gt;Staining&lt;/strong&gt;</td><td></td><td></td><td></td></tr><tr><td>Propidiumiodide</td><td>Sigma-Aldrich</td><td>P4170</td><td>Stock solution: 1.5 mM in 0.1 % sodium azide</td></tr><tr><td>Uranylacetate</td><td>Science Services</td><td>E22400</td><td></td></tr><tr><td>Lead(II) Nitrate</td><td>Merck</td><td>107398</td><td></td></tr><tr><td>Tri Sodium Citrate Dihydrate</td><td>Merck</td><td>106448</td><td></td></tr><tr><td>NaOH pellets</td><td>Merck</td><td>106469</td><td></td></tr><tr><td>1M NaOH solution</td><td>Bernd Kraft</td><td>01030.3000</td><td></td></tr><tr><td>Glass petri dish</td><td>Duran</td><td>23 755 56</td><td></td></tr><tr><td>&lt;strong&gt;Name&lt;/strong&gt;</td><td>&lt;strong&gt;Company&lt;/strong&gt;</td><td>&lt;strong&gt;Catalog Number&lt;/strong&gt;</td><td>&lt;strong&gt;Comments&lt;/strong&gt;</td></tr><tr><td>&lt;strong&gt;Mounting&lt;/strong&gt;</td><td></td><td></td><td></td></tr><tr><td>Stubs</td><td>Plano</td><td>G301F</td><td></td></tr><tr><td>Carbon pads</td><td>Plano</td><td>G3347</td><td></td></tr><tr><td>Copper tape</td><td>Plano</td><td>G3397</td><td>double-sided adhesive, conductive</td></tr><tr><td>Silver paint</td><td>Plano</td><td>G3692</td><td>Acheson Elektrodag 1415M</td></tr><tr><td>&lt;strong&gt;Name&lt;/strong&gt;</td><td>&lt;strong&gt;Company&lt;/strong&gt;</td><td>&lt;strong&gt;Catalog Number&lt;/strong&gt;</td><td>&lt;strong&gt;Comments&lt;/strong&gt;</td></tr><tr><td>&lt;strong&gt;Solutions/mixtures&lt;/strong&gt;</td><td></td><td></td><td></td></tr><tr><td>Adhesive mixture for coating blocks</td><td></td><td></td><td>Pattex contact adhesive /xylene as thinner, ratio 1:3.&lt;br/&gt; (Alternative for xylene: Rotihistol)</td></tr><tr><td>Reynolds lead citrate</td><td></td><td></td><td>50 mL:&lt;br/&gt; Dissolve 1.33 g of lead(II) nitrate in 10 mL of dH&lt;sub&gt;2&lt;/sub&gt;O.&lt;br/&gt; Dissolve 1.76 g of tri-sodium citrate dihydrate in 10 ml dH&lt;sub&gt;2&lt;/sub&gt;O.&lt;br/&gt; Mix both and add 1 M sodium hydroxide until the solution is clear.&lt;br/&gt; Fill up with dH&lt;sub&gt;2&lt;/sub&gt;O to 50 mL.</td></tr><tr><td>Propidium iodide staining solution</td><td></td><td></td><td>Prepare 1:1500 dilution from stock in dH&lt;sub&gt;2&lt;/sub&gt;O.&lt;br/&gt; Vortex for adequate mixing.</td></tr><tr><td>Aqueous uranyl acetate</td><td></td><td></td><td>Dissolve 3 % uranyl acetate in dH&lt;sub&gt;2&lt;/sub&gt;O (mix thoroughly).</td></tr></tbody></table>

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.

The workflow described here (Figure 1) starts with a sample embedded in a resin block. During the sample preparation, some heavy metal should be introduced into the tissue, but it is not necessary to use protocols optimized for rather strong metallization. Figure 1A shows a plant root (cress) block-stained conventionally with 1% OsO4 and 1% uranyl acetate, while the Arabidopsis root in Figure 1B is only weakly metalized using 0.5% uranyl acetate. The latter sample type is best suited for correlative approaches as some heavy metals tend to quench fluorescence. With a dedicated substrate holder (Figure 2), arrays of several hundred sections can be produced (Figure 1C). After fluorescent labeling, such arrays are imaged in a standard wide-field FLM (Figure 1D), then stained with heavy metal solutions and imaged in a SEM at different resolutions (Figure 1E-G).
Important tools for the reproducible generation of arrays, particularly when placing several ribbons from the microtome&#39;s knife boat onto a substrate, are the substrate holder (Figure 2A, custom-designed in the authors&#39; laboratory) and a Jumbo diamond knife with a boat large enough to accommodate microscope slides (Figure 2B). A flat meniscus, allowing good observation of the ribbons, is necessary and can be achieved by plasma cleaning of the substrate: A small droplet of distilled water should not form a lens-like structure on the substrate as in Figure 2C (untreated substrate), but a thin film (Figure 2D, plasma activated substrate). Under these conditions, ribbons attached to the dry part of an ITO-coated coverslip are easily visualized (Figure 2E) and can be observed and controlled during the lift-out of the substrate from the water.
As an example, arrays stained with propidium iodide to label the plant cell walls were imaged with a standard wide field FLM (Figure 3A). Since the sections are only 100 nm thick, even over-staining as shown here introduces little blurring. After registration, the two cells completely enclosed in the reconstructed volume were selected from the image stack (Figure 3B) for high-resolution imaging in 3D (see also Supplemental Movie S1). Following additional staining with uranyl acetate and lead citrate, the arrays were imaged in the SEM. Figure 3C shows an overview, recorded with 60 nm image pixels; the dark square in the center of the image indicates the position where the autofocus functions were executed, and the additional dose led to slight contamination. Appropriate ROIs in those serial sections (slices 51 to 248 of 435 slices in total) containing the two target cells selected in the FLM stack were then recorded with a 5 nm image pixel size (Figure 3D; see also Supplemental Movie S2).
Automated hierarchical imaging of the arrays in the SEM described here was done with the software/hardware platform solution ZEISS Atlas 5. First, an overview of the whole array was created using the SE detector, with very large (1,000 nm) image pixels and very low dwell time (Figure 4A). An ROI outlining only the tissue was placed on the first section and propagated to all other sections of the array. This section set was then recorded with 60 nm image pixels using a longer dwell time (Figure 4B). Finally, a site set, containing the two target cells plus one &#34;layer&#34; of surrounding cells to account for stage inaccuracy, was set up with the following parameters: ESB (Energy Selective Backscatter) detector, 5 nm image pixels, very long (40 &#181;s) dwell time (Figure 4C). Zooming in to such an image shows subcellular detail (Figure 4D) such as vacuoles (V), mitochondria (M), nucleus (N), and endoplasmic reticulum (arrows). See also the Supplemental Movie S3 for zooming in from an overview of the whole array to the subcellular detail of one target cell.
The array shown here (200 sections) plus an additional one of 250 sections took about 8 h to produce, one night to stain for LM, and one day to record (manually) at the FLM. Post-staining takes about 1-2 h in total, depending on the number of individual arrays. For SEM recording, a few hours are required to set up the Atlas run, and automated recording was 3&#8211;4 h for the intermediate resolution (60 nm pixel size) section set (200 sections, 450 x 200 &#181;m2) and about 5 days for the high-resolution (5 nm pixel size) ROI containing the two target cells (200 sections, 55 x 30 &#181;m2). Note that due to the low metal content of the sample shown here, a very slow scanning speed had to be used to reach a good signal-to-noise detection, which implied (for the currently available detector) a dwell time of 40 &#181;s for the high-resolution ROI.
There are several steps in the whole workflow prone to pitfalls: Ideally ribbons should be more or less straight and placed in the right order (Figure 5A). However, bent (Figure 5B), curved (Figure 5D), or even broken ribbons are often produced. This can result due to incorrect trimming (leading and trailing edges not exactly parallel), or non-uniformly applied adhesive, but also from an asymmetric or unevenly infiltrated sample. Particularly troublesome are samples containing both soft and very hard components. The latter components may be difficult to infiltrate such as the cell wall of the plant roots shown here (Figure 5C). In that case, folds (arrowheads) can easily be caused by variable compression and relaxation during sectioning. For automated imaging in the SEM, curved ribbons are not a great problem, since the ROIs can be rotated to accommodate the curvature of the ribbon.
Another critical step in the protocol is staining: Inadequate washing can lead to residues on the section (Figure 5E, F), and in the worst case, cover the most interesting area (circle on one of the two target cells in Figure 5F). Also, dust (Figure 5D, strongly light scattering particles) introduced into the knife boat, e.g., with a dirty substrate carrier, can cause serious problems: In the FLM, dust can be highly fluorescent (cf. some slices in Supplemental Movie S1) to such an extent that some registration algorithms do not function. The &#34;align&#34; function in TrakEM17 however, can handle such stacks as demonstrated in Supplemental Movie S1.
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/57059/57059fig1.jpg" /> 
Figure 1: Workflow for the correlative hierarchical imaging. Starting from a sample embedded in a resin block (A, strongly metalized sample), the sample is first trimmed (B, weakly metalized sample) and then arrays consisting of several ribbons of serial sections (C), here placed on an ITO-coated coverslip, are produced using an ultramicrotome. After staining with a fluorescent dye, stacks of images are recorded in a wide-field FLM (D). After further staining rounds with heavy metal salts, stacks are imaged in a SEM (E-G) at different resolutions (image pixel sizes). <a href="https://cloudflare-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/57059/57059fig1large.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/57059/57059fig2.jpg" /> 
Figure 2: Tools for the preparation of arrays. Substrate holder assembled from micromanipulators with seven axes of movement attached to a standard ultramicrotome (A): the screws, highlighted with circles, are for vertical (1) and horizontal (2) movement and for tilting (3) of the substrate carrier. The jumbo diamond knife with an oversize boat to accommodate large substrates (arrow), here with a piece of plasma-activated silicon wafer mounted onto a slide-sized aluminum carrier (B). 20 &#181;L drops of distilled water placed onto an untreated silicon wafer substrate (C) or on a plasma-activated substrate (D). Four ribbons floating in the knife boat, attached to an ITO-coated coverslip by their lower ends. <a href="https://cloudflare-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/57059/57059fig2large.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/57059/57059fig3.jpg" /> 
Figure 3: Correlation of the LM data with SEM data. Overviews (A, C) and target cells (B, D) recorded with FLM (A, B) and SEM (C, D). (B) is a software zoom, and the original data were recorded with a 40X objective lens on a 1,388 x 1,040 pixel camera chip, while (C) is recorded with 60 nm image pixel size, and (D) with 5 nm image pixel size, illustrating the true increase in resolution in the SEM. <a href="https://cloudflare-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/57059/57059fig3large.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/57059/57059fig4.jpg" /> 
Figure 4: Hierarchical imaging in the SEM using ZEISS Atlas 5 AT. Overview of an array recorded with 1,000 nm image pixels using the SE detector (A). Section sets with the ROI placed on tissue in every section and recorded with 60 nm image pixels (B). Site sets with a serial ROI placed on target cells and recorded with 5 nm image pixels (C). When zooming into such high-resolution images (D), intracellular membrane compartments such as vacuoles (V), nucleus (N), mitochondria (M), and endoplasmic reticulum (arrows) become visible. <a href="https://cloudflare-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/57059/57059fig4large.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/57059/57059fig5.jpg" /> 
Figure 5: Typical problems. 1. Arising from the sectioning process: Ribbons placed on ITO-coated coverslips are ideally straight (A), but irregular compression during sectioning may cause bent (B) or curved ribbons (D), or even folds (C). 2. Caused by handling substrate and ribbons in water, e.g., during sectioning and staining: Light scattering particles on substrate (D), rim of droplets on section (circle in E), or dirt smeared out over tissue due to inadequate washing after staining (F). <a href="https://cloudflare-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/57059/57059fig5large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
Supplemental Movie S1: FLM image stack. 435 images aligned in Fiji16 using TrakEM17 and saved as movie file (.avi). <a href="https://cloudflare-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/57059/WackerJoVE_movieS1.avi">Please click here to download this file.</a>
Supplemental Movie S2: SEM image stack. 210 images aligned in Fiji16 using TrakEM17. The original stack (300 images) of this data set was 15 GB. To downsize the stack from 3.3 GB (after alignment and cropping to only the two target cells), it was scaled in x and y by a factor of 0.2 using Fiji and then saved as .avi movie. <a href="https://cloudflare-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/57059/WackerJoVE_movieS2.avi">Please click here to download this file.</a>
Supplemental Movie S3: Zooming with different resolution levels in the SEM. Movie created in and exported from the Atlas 5 software in .mp4 format. <a href="https://cloudflare-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/57059/WackerJoVE_movieS3.mp4">Please click here to download this file.</a>

Watch this Scientific Journal Video about Multimodal Hierarchical Imaging of Serial Sections for Finding Specific Cellular Targets within Large Volumes at JoVE.com

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

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 ...

multimodal-hierarchical-imaging-serial-sections-for-finding-specific

Nanyang Technological University

Research

JoVE Journal

Developmental Biology

10.4K Views.  Universität Heidelberg. Dit protocol is gericht op specifieke cellen in weefsels voor beeldvorming op de nanoschaal resolutie met behulp van een Scannende Elektronen Microscoop (SEM). Groot aantal seriële secties uit biologisch materiaal hars-embedded zijn eerste beeld in een lichte Microscoop te identificeren van het doel, waarna in een hiërarchische wijze in de SEM.

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

Multimodal Imaging of Stem Cell Implantation in the Central Nervous System of Mice

During the past decade, stem cell transplantation has gained increasing interest as primary or secondary therapeutic modality for a variety of diseases, both in preclinical and clinical studies. However, to date results regarding functional outcome and/or tissue regeneration following stem cell transplantation are quite diverse. Generally, a clinical benefit is observed without profound understanding of the underlying mechanism(s)1. Therefore, multiple efforts have led to the development of different molecular imaging modalities to monitor stem cell grafting with the ultimate aim to accurately evaluate survival, fate and physiology of grafted stem cells and/or their micro-environment. Changes observed in one or more parameters determined by molecular imaging might be related to the observed clinical effect. In this context, our studies focus on the combined use of bioluminescence imaging (BLI), magnetic resonance imaging (MRI) and histological analysis to evaluate stem cell grafting.
BLI is commonly used to non-invasively perform cell tracking and monitor cell survival in time following transplantation2-7, based on a biochemical reaction where cells expressing the Luciferase-reporter gene are able to emit light following interaction with its substrate (e.g. D-luciferin)8, 9. MRI on the other hand is a non-invasive technique which is clinically applicable10 and can be used to precisely locate cellular grafts with very high resolution11-15, although its sensitivity highly depends on the contrast generated after cell labeling with an MRI contrast agent. Finally, post-mortem histological analysis is the method of choice to validate research results obtained with non-invasive techniques with highest resolution and sensitivity. Moreover end-point histological analysis allows us to perform detailed phenotypic analysis of grafted cells and/or the surrounding tissue, based on the use of fluorescent reporter proteins and/or direct cell labeling with specific antibodies.
In summary, we here visually demonstrate the complementarities of BLI, MRI and histology to unravel different stem cell- and/or environment-associated characteristics following stem cell grafting in the CNS of mice. As an example, bone marrow-derived stromal cells, genetically engineered to express the enhanced Green Fluorescent Protein (eGFP) and firefly Luciferase (fLuc), and labeled with blue fluorescent micron-sized iron oxide particles (MPIOs), will be grafted in the CNS of immune-competent mice and outcome will be monitored by BLI, MRI and histology (Figure 1).

This article describes an optimized sequence of events for multimodal imaging of cellular grafts in rodent brain using: (i) in vivo bioluminescence and magnetic resonance imaging, and (ii) post mortem histological analysis. Combining these imaging modalities on a single animal allows cellular graft evaluation with high resolution, sensitivity and specificity.

During the past decade, stem cell transplantation has gained increasing interest as primary or secondary therapeutic modality for a variety of diseases, ...

Neuroscience

Quantitative Multispectral Analysis Following Fluorescent Tissue Transplant for Visualization of Cell Origins, Types, and Interactions

With the desire to understand the contributions of multiple cellular elements to the development of a complex tissue; such as the numerous cell types that participate in regenerating tissue, tumor formation, or vasculogenesis, we devised a multi-colored cellular transplant model of tumor development in which cell populations originate from different fluorescently colored reporter gene mice and are transplanted, engrafted or injected in and around a developing tumor. These colored cells are then recruited and incorporated into the tumor stroma. In order to quantitatively assess bone marrow derived tumor stromal cells, we transplanted GFP expressing transgenic whole bone marrow into lethally irradiated RFP expressing mice as approved by IACUC. 0ovarian tumors that were orthotopically injected into the transplanted mice were excised 6-8 weeks post engraftment and analyzed for bone marrow marker of origin (GFP) as well as antibody markers to detect tumor associated stroma using multispectral imaging techniques. We then adapted a methodology we call MIMicc- Multispectral Interrogation of Multiplexed cellular compositions, using multispectral unmixing of fluoroprobes to quantitatively assess which labeled cell came from which starting populations (based on original reporter gene labels), and as our ability to unmix 4, 5, 6 or more spectra per slide increases, we&#39;ve added additional immunohistochemistry associated with cell lineages or differentiation to increase precision. Utilizing software to detect co-localized multiplexed-fluorescent signals, tumor stromal populations can be traced, enumerated and characterized based on marker staining.1

Complex tissue masses, from organs to tumors, are composed of various cellular elements. We elucidated the contribution of cellular phenotypes within a tissue utilizing multi-labeled fluorescent transgenic mice in combination with multiparameter immunofluorescent staining followed by spectral unmixing to decipher cell origin as well as cell characteristics based on protein expression.

With the desire to understand the contributions of multiple cellular elements to the development of a complex tissue; such as the numerous cell types that ...

Medicine

Three-Dimensional Imaging of Tumor-Bearing Tissue Using the Iterative Bleaching Extends Multiplexity Approach

The spatial heterogeneity of the tumor microenvironment (TME) is a critical determinant of therapeutic response, particularly for immune-oncology agents, where success depends on the distribution of specific immune cell subpopulations. Over the past decade, multiple sophisticated technologies have been introduced to achieve detailed resolution of the TME using two-dimensional sections from either formalin-fixed, paraffin-embedded (FFPE) or fixed-frozen tissues. While these thin sections are easier to procure and analyze, they lack the three-dimensional architecture needed to reliably and comprehensively characterize a tumor. To address this limitation, a tissue mounting and imaging technique was developed to enable the three-dimensional analysis of tumor lesions in their native in vivo state. This protocol outlines the procurement of human tumor tissue, the mounting of samples on custom-printed platforms, and staining procedures for post-fixation samples. The multiplexed immunofluorescence technique, IBEX (Iterative Bleaching Extends Multiplexity), was adapted to characterize the three-dimensional TME with up to 15 markers for tumor, immune, and stromal cells using commercially available antibodies. Imaging depths of up to 100 &#181;m were achieved using an inverted white-light laser confocal microscope with a custom-printed imaging adaptor and commercial glass-bottom dishes to ensure optimal tissue orientation. This protocol highlights the potential of the IBEX method to expand multiplexed immunofluorescence studies, providing a more comprehensive understanding of TME composition.

This article describes a protocol for multiplex immunofluorescence optimized to characterize the three-dimensional architecture of peritoneal metastases.

The spatial heterogeneity of the tumor microenvironment (TME) is a critical determinant of therapeutic response, particularly for immune-oncology agents, ...

Cancer Research

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

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

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

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

Visualization, Quantification, and Mapping of Immune Cell Populations in the Tumor Microenvironment

The immune landscape of the tumor microenvironment (TME) is a determining factor in cancer progression and response to therapy. Specifically, the density and the location of immune cells in the TME have important diagnostic and prognostic values. Multiomic profiling of the TME has exponentially increased our understanding of the numerous cellular and molecular networks regulating tumor initiation and progression. However, these techniques do not provide information about the spatial organization of cells or cell-cell interactions. Affordable, accessible, and easy to execute multiplexing techniques that allow spatial resolution of immune cells in tissue sections are needed to complement single cell-based high-throughput technologies. Here, we describe a strategy that integrates serial imaging, sequential labeling, and image alignment to generate virtual multiparameter slides of whole tissue sections. Virtual slides are subsequently analyzed in an automated fashion using user-defined protocols that enable identification, quantification, and mapping of cell populations of interest. The image analysis is done, in this case using the analysis modules Tissuealign, Author, and HISTOmap. We present an example where we applied this strategy successfully to one clinical specimen, maximizing the information that can be obtained from limited tissue samples and providing an unbiased view of the TME in the entire tissue section.

Here, we describe a simple and accessible strategy for visualizing, quantifying, and mapping immune cells in formalin-fixed paraffin-embedded tumor tissue sections. This methodology combines existing imaging and digital analysis techniques with the purpose of expanding the multiplexing capability and multiparameter analysis of imaging assays.

The immune landscape of the tumor microenvironment (TME) is a determining factor in cancer progression and response to therapy. Specifically, the density ...

Immunology and Infection

A Rapid Method for Multispectral Fluorescence Imaging of Frozen Tissue Sections

Multispectral fluorescence imaging on formalin-fixed paraffin-embedded (FFPE) tissues enables the detection of multiple markers in a single tissue sample that can provide information about antigen coexpression and spatial distribution of the markers. However, a lack of suitable antibodies for formalin-fixed tissues may restrict the nature of markers that can be detected. In addition, the staining method is time-consuming. Here we describe a rapid method to perform multispectral fluorescence imaging on frozen tissues. The method includes the fluorophore combinations used, detailed steps for the staining of mouse and human frozen tissues, and the scanning, acquisition, and analysis procedures. For staining analysis, a commercially available semiautomated multispectral fluorescence imaging system is used. Through this method, up to six different markers were stained and detected in a single frozen tissue section. The machine learning analysis software can phenotype cells that can be used for quantitative analysis. The method described here for frozen tissues is useful for the detection of markers that cannot be detected in FFPE tissues or for which antibodies are not available for FFPE tissues.

We describe a rapid staining method to perform multispectral imaging on frozen tissues.

Multispectral fluorescence imaging on formalin-fixed paraffin-embedded (FFPE) tissues enables the detection of multiple markers in a single tissue sample ...

In vivo Clonal Tracking of Hematopoietic Stem and Progenitor Cells Marked by Five Fluorescent Proteins using Confocal and Multiphoton Microscopy

We developed and validated a fluorescent marking methodology for clonal tracking of hematopoietic stem and progenitor cells (HSPCs) with high spatial and temporal resolution to study in vivo hematopoiesis using the murine bone marrow transplant experimental model. Genetic combinatorial marking using lentiviral vectors encoding fluorescent proteins (FPs) enabled cell fate mapping through advanced microscopy imaging. Vectors encoding five different FPs: Cerulean, EGFP, Venus, tdTomato, and mCherry were used to concurrently transduce HSPCs, creating a diverse palette of color marked cells. Imaging using confocal/two-photon hybrid microscopy enables simultaneous high resolution assessment of uniquely marked cells and their progeny in conjunction with structural components of the tissues. Volumetric analyses over large areas reveal that spectrally coded HSPC-derived cells can be detected non-invasively in various intact tissues, including the bone marrow (BM), for extensive periods of time following transplantation. Live studies combining video-rate multiphoton and confocal time-lapse imaging in 4D demonstrate the possibility of dynamic cellular and clonal tracking in a quantitative manner.

In vivo Clonal Tracking of Hematopoietic Stem and Progenitor Cells Marked by Five Fluorescent Proteins using Confocal and Multiphoton Microscopy

Combinatorial 5 fluorescent proteins marking of hematopoietic stem and progenitor cells allows in vivo clonal tracking via confocal and two-photon microscopy, providing insights into bone marrow hematopoietic architecture during regeneration. This method allows non-invasive fate mapping of spectrally-coded HSPCs-derived cells in intact tissues for extensive periods of time following transplantation.

We developed and validated a fluorescent marking methodology for clonal tracking of hematopoietic stem and progenitor cells (HSPCs) with high spatial and ...

Biology

Quantitation of Protein Expression and Co-localization Using Multiplexed Immuno-histochemical Staining and Multispectral Imaging

Immunohistochemistry is a commonly used clinical and research lab detection technique for investigating protein expression and localization within tissues. Many semi-quantitative systems have been developed for scoring expression using immunohistochemistry, but inherent subjectivity limits reproducibility and accuracy of results. Furthermore, the investigation of spatially overlapping biomarkers such as nuclear transcription factors is difficult with current immunohistochemistry techniques. We have developed and optimized a system for simultaneous investigation of multiple proteins using high throughput methods of multiplexed immunohistochemistry and multispectral imaging. Multiplexed immunohistochemistry is performed by sequential application of primary antibodies with secondary antibodies conjugated to horseradish peroxidase or alkaline phosphatase. Different chromogens are used to detect each protein of interest. Stained slides are loaded into an automated slide scanner and a protocol is created for automated image acquisition. A spectral library is created by staining a set of slides with a single chromogen on each. A subset of representative stained images are imported into multispectral imaging software and an algorithm for distinguishing tissue type is created by defining tissue compartments on images. Subcellular compartments are segmented by using hematoxylin counterstain and adjusting the intrinsic algorithm. Thresholding is applied to determine positivity and protein co-localization. The final algorithm is then applied to the entire set of tissues. Resulting data allows the user to evaluate protein expression based on tissue type (ex. epithelia vs. stroma) and subcellular compartment (nucleus vs. cytoplasm vs. plasma membrane). Co-localization analysis allows for investigation of double-positive, double-negative, and single-positive cell types. Combining multispectral imaging with multiplexed immunohistochemistry and automated image acquisition is an objective, high-throughput method for investigation of biomarkers within tissues.

Immunohistochemistry is a powerful lab technique for evaluating protein localization and expression within tissues. Current semi-automated methods for quantitation introduce subjectivity and often create irreproducible results. Herein, we describe methods for multiplexed immunohistochemistry and objective quantitation of protein expression and co-localization using multispectral imaging.

Immunohistochemistry is a commonly used clinical and research lab detection technique for investigating protein expression and localization within ...

High-Resolution 3D Reconstruction of Human Brain with SHORT and LSFM

Optical Clearing and Labeling for Light-sheet Fluorescence Microscopy in Large-scale Human Brain Imaging

Despite the numerous clearing techniques that emerged in the last decade, processing postmortem human brains remains a challenging task due to its dimensions and complexity, which make imaging with micrometer resolution particularly difficult. This paper presents a protocol to perform the reconstruction of volumetric portions of the human brain by simultaneously processing tens of sections with the SHORT (SWITCH - H2O2 - Antigen Retrieval - 2,2'-thiodiethanol [TDE]) tissue transformation protocol, which enables clearing, labeling, and sequential imaging of the samples with light-sheet fluorescence microscopy (LSFM). SHORT provides rapid tissue clearing and homogeneous multi-labeling of thick slices with several neuronal markers, enabling the identification of different neuronal subpopulations in both white and grey matter. After clearing, the slices are imaged via LSFM with micrometer resolution and in multiple channels simultaneously for a rapid 3D reconstruction. By combining SHORT with LSFM analysis within a routinely high-throughput protocol, it is possible to obtain the 3D cytoarchitecture reconstruction of large volumetric areas at high resolution in a short time, thus enabling comprehensive structural characterization of the human brain.

Author Spotlight: Advancing 3D Cytoarchitecture Analysis - Rapid Volumetric Reconstruction of the Human Brain 

The present protocol provides a step-by-step procedure for rapid and simultaneous optical clearing, muti-round labeling, and 3D volumetric reconstruction of tens of postmortem human brain sections by combining the (SWITCH - H2O2 - Antigen Retrieval - 2,2'-thiodiethanol [TDE]) SHORT tissue transformation technique with light-sheet fluorescence microscopy imaging in a routinely high-throughput protocol.

Despite the numerous clearing techniques that emerged in the last decade, processing postmortem human brains remains a challenging task due to its ...

Single-Cell Resolution Three-Dimensional Imaging of Intact Organoids

Organoid technology, in vitro 3D culturing of miniature tissue, has opened a new experimental window for cellular processes that govern organ development and function as well as disease. Fluorescence microscopy has played a major role in characterizing their cellular composition in detail and demonstrating their similarity to the tissue they originate from. In this article, we present a comprehensive protocol for high-resolution 3D imaging of whole organoids upon immunofluorescent labeling. This method is widely applicable for imaging of organoids differing in origin, size and shape. Thus far we have applied the method to airway, colon, kidney, and liver organoids derived from healthy human tissue, as well as human breast tumor organoids and mouse mammary gland organoids. We use an optical clearing agent, FUnGI, which enables the acquisition of whole 3D organoids with the opportunity for single-cell quantification of markers. This three-day protocol from organoid harvesting to image analysis is optimized for 3D imaging using confocal microscopy.

The entire 3D structure and cellular content of organoids, as well as their phenotypic resemblance to the original tissue can be captured using the single-cell resolution 3D imaging protocol described here. This protocol can be applied to a wide range of organoids varying in origin, size and shape.

Organoid technology, in vitro 3D culturing of miniature tissue, has opened a new experimental window for cellular processes that govern organ development ...

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

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